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Molecular and Cellular Endocrinology

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Controls of meiotic signaling by membrane or nuclear progestin receptor inzebrafish follicle-enclosed oocytes

Richard N. Hanna1, Yong Zhu ∗

Department of Biology, East Carolina University, Howell Science Complex, 1000 E. 5th Street, Greenville, NC 27858, USA

a r t i c l e i n f o

Article history:Received 10 September 2010Received in revised form 24 January 2011Accepted 3 February 2011

Keywords:Membrane progestin receptormPRPaqrNuclear progestin receptornPRPgrProgestinOocyteMaturation

a b s t r a c t

Both membrane progestin receptors (mPRs) and the nuclear progestin receptor (nPR or Pgr) decodethe non-genomic progestin signaling (NGPS) in vertebrates. However, the receptor for decipheringextracellular NGPS and initiating meiosis resumption in vertebrate oocytes is still contested hotly. Westudied the roles of nPR and mPRs by determining their localization, changes of expression, and activationof NGPS during final oocyte maturation (FOM) in zebrafish. The nPR transcript and protein were expressedabundantly in follicular cells that were surrounding stage IV oocytes, but nPR transcript appeared absentwithin stage IV oocytes. The most significant daily changes of nPR transcript were observed in stage IV fol-licular cells, with the highest level observed just prior to ovulation. In contrast, the expressions of mPR�and mPR� transcripts and proteins were abundant and increased significantly in late stage denudedoocytes prior to oocyte maturation, consistent with the purported role of mPRs in interpreting NGPS.Moreover, over-expression of mPR� in follicle-enclosed oocytes significantly increased the activity ofMAPK, the production of cyclin B protein, and the number of oocytes that underwent FOM without exoge-nous progestin, while over-expression of mPR� or nPR alone had no such effect. Intriguingly, significantacceleration of FOM was observed when follicle-enclosed oocytes were incubated with the maturationinducing steroid, 4-pregnen-17, 20�-diol-3-one (DHP) following over-expression of nPR or mPR�.Interestingly, this acceleration in oocyte maturation was observed approximately 1 h later in oocytesover-expressing nPR compared to those over-expressing mPR�. Importantly, the acceleration of matura-tion in the nPR injected group was blocked by treatment with the transcription inhibitor actinomycin D,implying a requirement of the genomic signaling pathway, while the same treatment did not affect theaccelerated rate of maturation in mPR˛ injected oocytes. Taken together, these results imply that nPRand mPR� are unlikely receptors for inducing FOM, while mPR� is the long-sought-after nongenomicprogestin receptor that deciphers extracellular NGPS to initiate meiosis resumption in follicle-enclosedzebrafish oocytes.

© 2011 Elsevier Ireland Ltd. All rights reserved.

1. Introduction

The nongenomic actions of steroids have been generally rec-ognized, but the receptor(s) initiating these nongenomic actionshas been disputed hotly (Filardo et al., 2008; Prossnitz et al., 2008;Stormshak and Bishop, 2008; Thomas, 2008; Zhu et al., 2008).Resumption of cell cycle in meiosis arrested oocytes resulting

Abbreviations: DHP, 17�,20�-dihydroxy-progesterone; FOM, final oocyte mat-uration; GVBD, germinal vesicle breakdown; MIS, maturation-inducing steroid;mPR, membrane progestin receptor; MPF, meiosis/mitosis promoting factor; NGPS,nongenomic progestin signaling; Ngpr, nongenomic progestin receptor; nPR,nuclear progestin receptor; Paqr, progestin and AdipoQ receptor.

∗ Corresponding author. Tel.: +1 252 328 2712; fax: +1 252 328 4178.E-mail address: zhuy@ecu.edu (Y. Zhu).

1 Current address: La Jolla Institute for Allergy and Immunology, 9420 AthenaCircle, La Jolla, CA 92037, USA.

in final oocyte maturation (FOM) in fish and amphibian speciesis one of the best studied nongenomic actions of progestins.A non-genomic progestin receptor (Ngpr) located at the oocyteplasma membrane is required for transmitting non-genomic pro-gestin signaling (NGPS) into the oocytes. The Ngpr deciphersextracellular NGPS into a rapid cascade of changes in intracellularsecond messengers including down-regulation of cAMP, activa-tion of the mitogen-activated protein kinase (MAPK) pathway, andthe de novo synthesis of cyclin B protein leading to the resump-tion of meiosis (Kondo et al., 1997; Nagahama and Yamashita,2008; Thomas et al., 2002; Yamashita et al., 2000; Yoshikuni andNagahama, 1994; Zhu et al., 2008). Both the nuclear progestinreceptor (nPR or Pgr) and a pair of membrane progestin receptors(mPR� and mPR�) have been suggested to be the long-sought-afterNgpr decoding extracellular signaling of progestins and resulting inFOM (Bayaa et al., 2000; Josefsberg-Ben-Yehoshua et al., 2007; Tianet al., 2000; Tokumoto et al., 2006; Zhu et al., 2003a,b). The mPRs

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have seven transmembrane domains, localize at the oocyte mem-brane, couple to G proteins, bind specifically to progestins, and gen-erate rapid intracellular signaling, such as activation of MAPK andinhibition of cAMP (Hanna et al., 2006; Josefsberg-Ben-Yehoshuaet al., 2007; Thomas et al., 2007; Tokumoto et al., 2006; Zhu et al.,2003a,b). Intriguingly, the nPR has also been suggested to mediateNGPS during oocyte maturation in Xenopus (Bayaa et al., 2000; Tianet al., 2000). Injection of nPR mRNA into Xenopus follicle-enclosedoocytes accelerated progesterone-induced activation of the MAPKpathway and cell cycle proteins leading to oocyte maturation. Cur-rently it is not known whether these Ngpr candidates are workingtogether complementary, or whether they can independently acti-vate NGPS leading to resumption of meiosis. It is also uncertainto what extent each Ngpr candidate can activate the NGPS duringmeiosis, and if one receptor is “preferred” or “essential” for the pro-cess. Additional comparison studies are required for clarifying theroles of mPR and nPR during nongenomic signaling of progestinsand progestin-induced oocyte maturation (Zhu et al., 2008).

We have previously identified and characterized both mPRs(mPR� and mPR�) and nPR, which are expressed abundantly inzebrafish ovaries (Hanna et al., 2006, 2010; Hanna and Zhu, 2009).Both mPRs and nPR have a high binding affinity for progestinsand were able to activate either intracellular or nuclear signaling,respectively (Hanna et al., 2006, 2010). Since different developmen-tal stages of oocytes can be collected daily, zebrafish have becomean excellent model for studying oocyte maturation (Selman et al.,1993). Oocyte maturation in zebrafish occurs prior to the onsetof light, while ovulation and spawning occurs 1 h following theonset of light. Based on oocyte size and development, ooctyes canbe classified into five stages: stages I and II are previtellogenic(<340 �m), stage III are early vitellogenic (340–650 �m), stage IVare late vitellogenic and maturational competent (>650 �m), andstage V are mature eggs (Selman et al., 1993). The maturation ofstage IV oocytes can be induced ex vivo by exogenous hormones ifthe oocytes have not experienced sufficient stimuli of maturationsteroids in vivo (Selman et al., 1993, 1994).

In this study, we attempt to determine whether mPR�, mPR�,or nPR could decipher the extracellular signaling of progestinsand initiate resumption of meiosis in maturational competentstage IV oocytes. We compare the expression of mPR�, mPR�,and nPR in oocytes and surrounding follicles, examine the effectsof over-expression of mPR�, mPR� or nPR on oocyte matura-tion and activation of intracellular signaling, and determine ifthe progestin response occurs through genomic or nongenomicpathways.

2. Materials and methods

2.1. Zebrafish maintenance, collection of oocytes, and follicular layer removal

Experimental protocols were approved by The Institutional Animal Care andUse Committee at East Carolina University. Zebrafish were purchased from a localpet store and maintained at 28.5 ◦C on a 14-h light cycle with lights on at 8:00.Unless indicated, follicle-enclosed oocytes were collected from well-fed, healthyand mature female fish in the evening time (20:00) when few oocytes maturespontaneously without exogenous stimuli. The responses of these oocytes to thestimulation of exogenous hormones were optimal around evening time. Ovarieswere removed immediately following an appropriate anesthetic overdose (MS-222:200 mg/L in buffered solution) and placed in 50% L-15 media (Sigma) containing15 mM HEPES (pH 7.2). Individual follicle-enclosed oocytes were teased away fromeach other by gently pipetting up and down several times using a Pasteur pipette.Following methods similar to Selman et al. (1994), maturationally competent stageIV follicle-enclosed oocytes (>650 �m) were collected following two washes withmedia. For removal and isolation of follicular cells, stage IV follicle-enclosed oocyteswere chilled on ice for 30 min in 50% L-15 medium, and then the follicular cellswere removed manually with fine forceps, and removal was confirmed as describedbelow.

In brief, individual follicle-enclosed oocytes (stage IV) were held down usinga fine forceps, and the surface of the oocyte was scratched gently multiple timesby another fine forceps under a dissecting microscope (model MZ6, Leica, Wetzlar,

Germany), until the fibrous amorphous follicular layer disengaged from the oocytes.Then, one forceps held down the edge of the disengaged follicular layer, and anotherforceps rolled the oocyte till the whole follicular layer was stripped off completelyfrom the oocyte (supplemental Fig. 1).

To verify the effectiveness of the follicular removal procedure, the surface offreshly prepared oocytes were scanned and imaged immediately by a scanningelectronic microscope (SEM, Quanta 200 Mark I variable pressure scanning elec-tronic microscope, FEI, Hillsboro, OR, USA), or fluorescent microscopes. In order toobtain the best quality images, samples were also fixed overnight by 10% bufferedformalin (Fisher Scientific, Pittsburgh, PA, USA), 4% glutaraldehyde (ElectronMicroscopy Science, Hatfield, PA, USA), or ethanol diluted in modified Cortland’smedium (pH 7.8, Wolf and Quimby, 1969). The samples were shaken continuouslyand gently in a solution on a slow-rotating shaker (Slow Speed Roto Mix, Ther-molyne, Dubuque, IA, USA). Then, samples were kept in 70% ethanol until analysesfollowing a procedure of partial dehydration by an increased concentrations ofethanol (30%, 50%, 70%, 15 min each). For SEM analyses, samples were furtherdehydrated by increase concentration of the ethanol (95% once, 100% 3 times,15 min each). These samples were subjected to critical point drying (model CPD030,Bal-Tec) following replacement of ethanol solution with liquid CO2 by washing sam-ples with liquid CO2 for 8–10 times for 5 min each time. Then, samples were coatedwith gold and palladium (Hummer 6.6, Anatech, Union City, CA, USA). High qualitydigital SEM images were obtained at various magnifications using backscatteredelectrons or secondary electrons, depending on sample treatments. For fluorescentmicroscope images, the fresh or fixed samples in 70% ethanol were stained withDAPI (100 �g/ml) or propidium iodide (PI) staining (1 mg/ml) in Cortland’s mediumfor 10 min. These stained samples were examined and imaged immediately usingan up-right fluorescence microscope (model: Microphot-FX, Nikon Inc., Melville,NY, USA) or an inverted-fluorescent-spin-disk-confocal-scanning microscope(model: IX2-DSU, Olympus America, Center Valley, PA, USA).

We found that samples stained with PI showed similar follicular staining con-sistently compared to those stained by DAPI. Therefore, we chose PI staining forthe selection of denuded oocytes because of our preference for a low permeabledye, with lessened potential damage of signaling molecules. In short, all denudedoocytes for experiments were selected individually under a fluorescent microscopefollowing a quick 10-min PI staining.

2.2. Quantitative real time PCR (qRT-PCR) measurement of receptors

The levels of mPR or nPR transcripts were determined by quantitative real-timePCR (qRT-PCR) using SYBR green dye (Stragtagene, La Jolla, CA, USA) and a CepheidSmart Cycler MX4000 (Cepheid, Sunnyvale, CA, USA). Total RNA was extracted fromfreshly collected ovarian samples by homogenization in TRIzol reagent (Invitrogen)using a sonicator (Sonic Dismembrator Model 100, Fisher) and was purified fol-lowing the manufacturer’s instructions. Then 1 �g of total RNA of each sample wasreverse transcribed into cDNAs in a 10 �l reaction using SuperScript II (Invitrogen).As control, 1 �g total RNA of each sample was processed in identical conditions withthe same reaction mixture without reverse transcriptase. The qRT-PCR reactionwas conducted for 40 cycles with an annealing temperature of 62 ◦C using the mPRand nPR specific primers (mPR˛ – Forward: 5′-CGCTCAAGTGCGAACTTTTT-3′

Reverse: 5′-CGTACTTGCCATAGCAGCAG-3′; mPRˇ – Forward: 5′-ACGTCAAGCCACAGTACACG-3′ Reverse: 5′-TCCTGATGCACTGGACGATA-3′; nPR– Forward 5′-ACAGACAGCATACACCGC-3′ Reverse: 5′-TCCACAGGTCAGAACTCC-3′).The PCR mixture (25 �l) consisted of a 1× Cepheid enhancer additive (1 mM Tris,pH 8.0; 0.1 mg/ml bovine serum albumin, non-acetylated; 0.75 M trahalose; 1%tween-20), 10 �l MasterMix (2.5×) (Eppendorf), 500 nM forward and reverseprimers, and 0.25× SYBR green dye. The transcript levels, expressed as fmol/�gtotal RNA, were determined using Ct values of samples and a standard curvegenerated from serial known concentrations of plasmids corresponding to mPR˛,mPRˇ or nPR. The efficiency of the PCR and authentic PCR products was confirmedby analyses of melting curve, gel electrophoresis, and sequencing.

2.3. Synthesizing progesterone receptor transcripts

Expression vectors with an enhanced green fluorescent protein (EGFP) taggedat the C-terminal end of mPR˛, mPRˇ or nPRr were generated by fusion of thecoding region of the receptors in front of EGFP in the pCS2 + EGFP vector (RZPD– German Science Centre for Genome Research). GFP fused mPR˛, mPRˇ, ornPR transcripts were produced using the SP6 mMESSAGE mMACHINE transcrip-tion kit (Ambion, Austin, TX), following linearization of the expression vectorsusing the Not I enzyme. For comparison, the mRNAs encoding mPR˛, mPRˇor nPR without EGFP were also generated from the pcDNA 3.1 vector (Invitro-gen) constructs described previously (Hanna et al., 2006, 2010; Hanna and Zhu,2009) using the T7 mMESSAGE mMACHINE transcription kit (Ambion, Austin,TX) following linearization using Pme I. Unless indicated below, the transcriptssynthesized in vitro used for microinjection in follicle-enclosed oocytes or trans-fection of mammalian cells were fused transcripts of progestin receptors andEGFP.

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2.4. Expression of receptors in a mammalian cell line

We also expressed progestin receptors of zebrafish in human embryonic kidneycells (HEK293) for comparison of the cellular locations of the expressed transgenicprogestin receptors in non-native cells. (i.e., mammalian cells) as well as nativecells (i.e., oocytes). Our aim was for clarification of experimental controversysurrounding the ability of over-expressed mPRs protein to be trafficked properlyto the cell surface. The expression and localization of mPR�, mPR� or nPR werevisualized in human embryonic kidney cells (HEK293; American Type CultureCollection) transiently transected with the EGFP fused constructs of mPRs ornPR using Lipofectamine 2000 reagent (Invitrogen, Carlsbad, CA) following themanufacturer’s instructions. Transfected cells were cultured for three days inDMEM/F-12 media without phenol red (Sigma, Saint Louis, MO) containing 10%FBS (Gibco, Carlsbad, CA) at 37 ◦C with 5% CO2. The expression patterns of GFPfusion proteins were analyzed under an inverted tandem spinning disk confocalmicroscope (Olympus, model: IX2-DSU).

2.5. Western blotting

Membrane or cytosolic proteins were isolated from freshly collected follicularlayer enclosed oocytes, denuded oocytes or follicular layers as described previously(Hanna et al., 2010). In brief, samples were sonicated with 10 short bursts on asonicator (Sonic Dismembrator, Fisher Scientific) in PBS and centrifuged at 500 × gfor 5 min to remove any remaining nuclear material. The resulting supernatant wascentrifuged at 20,000 × g for 20 min to obtain the plasma membrane fraction andtwice at 150,000 × g for 1 h to produce a cytosolic fraction. The plasma membranewas further purified by centrifuging the pellet three times with a sucrose pad (1.2 Msucrose) at 6,500 × g for 45 min. The membrane or cytosolic fractions were mixedwith 2× SDS sample buffer (0.125 M Tris HCl pH 6.8, 4% SDS, 20% glycerol, 10% 2-mercaptoethanol) at a 1:1 ratio, and were boiled and cooled on ice for 5 min. Proteinsof each sample (40 �g, determined by the RC DC protein assay, Bio-Rad, Hercules,CA) were separated on a 7% SDS PAGE gel and were transferred to a nitrocellulosemembrane. The membrane was incubated for 1 h in a blocking solution containing5% nonfat powdered milk in TBST (50 mM Tris, 100 mM NaCl, 0.1% Tween 20, pH7.4). The membrane was incubated with a primary antibody [GFP, phopho-p44/42MAPK and p44/42 MAPK: 1:1000 dilution, Cell Signaling cat#2555, 910S, and 9102respectively; cyclin B: B112 (Kondo et al., 1997), 1:2500 dilution; nPR, 1:2500 dilu-tion (Hanna and Zhu, 2009); mPR�: 1:1000; and mPR�: 1:2000 dilution (Hanna andZhu, 2009) in the blocking solution overnight. The following day, the membraneswere washed five times for 5 min each with TBST, incubated for 1 h with horseradishperoxidase conjugated secondary antibody (1:5000 dilutions, goat anti-mouse anti-body for cyclin B detection or goat anti-rabbit antibody for other primary antibodies),and finally washed 5 times for 5 min each with TBST. The membranes then weredeveloped using Super Signal West Extended Dura Substrate (Pierce, Rockford, IL)and visualized using a Fluor Chem 8900 imaging station (Alpha Innotech, San Lean-dro, CA). Protein size was determined by comparing to a biotinylated protein ladder(Cell Signaling).

2.6. Maturation assays and microinjections

Microinjection was similar to the injection of goldfish oocytes describedpreviously (Hirai et al., 1992). In short, stage IV follicle-enclosed oocytes (>650 �m)were collected from females at approximately 20:00 and aligned in wells of 2%agarose submerged with 50% L-15 media, and injected with approximately 50 pgmRNA of mPR˛, mPRˇ or nPR in 1 nl volume with 0.05% phenol dye. Each treatmentgroup contained approximately 60 injected follicle-enclosed oocytes in a 5 cm Petridish containing 5 ml of fresh 50% L-15 media. For the experiments of transcriptionalinhibition or antiserum blocking, microinjected oocytes were pre-incubated witha transcriptional inhibitor (3 �g/ml of actinomycin D), progesterone receptorantiserum (1:100), or control serum/media for 2 h prior to the addition of variousconcentrations of the maturation steroid (4-pregnen-17, 20�-diol-3-one: i.e., DHP).Damaged follicle-enclosed oocytes were removed after 2 h of culture at 28 ◦C.Thereafter, the numbers of follicle-enclosed oocytes that underwent final oocytematuration were determined every 30 min for clearing and disappearance of thegerminal vesicle (GVBD) under a dissecting microscope as an indicator of FOMfollowing various treatments. For Western analysis, 10 oocytes were removed andlysed immediately in SDS buffer at various time points.

2.7. Statistical analyses

Data was analyzed by a one-way analysis of variance (ANOVA) followed by aNewman–Keuls multiple comparison test using the GraphPad Prism Program (SanDiego, CA). P value <0.05 was considered statistically significant.

3. Results

We first attempted to determine the expression levels of mPR�,mPR�, and nPR in late stage maturational competent follicle-enclosed oocytes (stage IV) at three critical time points during the

daily cycle of oocyte maturation. The 6:00 sampling time was 2 hbefore the onset of the lights and prior to oocyte maturation. Atthe 6:00 time point, rates of oocytes maturation were variable, anda high percentage of oocytes were undergoing maturation spon-taneously, without exogenous stimuli likely due to the exposureof high levels of endogenous hormones produced in the ovary. The13:00 time point was after spawning, and stage IV oocytes collectedat this time normally had poor responses to stimulation of exoge-nous hormones to undergo maturation. The 21:00 time point wasoptimal for ex vivo maturation experiments since oocytes begin torespond best to the stimulation of exogenous hormones with fewoocytes maturing spontaneously. Protein levels of mPR� and nPRwere highest in follicular cell enclosed oocytes in the early morn-ing (6:00) prior to oocyte maturation and decreased considerablyin the afternoon (13:00) following spawning (Fig. 1A). The expres-sion of mPR� protein was highest at night time (21:00) and alsodecreased following spawning (13:00; Fig. 1A).

In order to determine the major production sites of progestinreceptors, we separated oocytes from the surrounding follicu-lar cells, and confirmed the effectiveness and efficiency of ourseparation procedure. We did not observe any follicular cells indenuded oocytes using a scanning electronic microscope (SEM,Fig. 2, supplemental Fig. 2) or two different fluorescent micro-scopes (Fig. 2). In denuded oocytes, chorionic pores could be seenclearly in all areas of the oocytes surface by SEM imaging (Fig. 2H,supplemental Fig. 2D). In contrast, a fibrous amorphous layer and alayer of follicular nucleus were observed clearly in follicle-enclosedoocytes (Fig. 2A and B, supplemental Fig. 2A and C). In agreementwith the results obtained from SEM, nuclear staining of follicularcells was observed clearly in follicle-enclosed oocytes stained withDAPI (Fig. 2C, supplemental 3A and 4A) or PI (Fig. 2E, supplementalFig. 3B and 4C). In contrast, denuded oocytes showed no nuclearstaining around the oocyte surfaces (Fig. 2I and K; supplemental Fig.4E and G). In some cases, PI showed better staining than DAPI (Fig. 2,supplemental Fig. 4). Similar results were obtained when stainedoocytes were observed under an upright fluorescent microscope(supplemental Fig. 4), or when the same procedure was appliedto earlier stage III oocytes (supplemental Fig. 2). Freshly preparedsamples without fixation had similar results as those fixed samplesin all SEM and fluorescent microscope analyses (data not shown).Therefore, we could conclude with certainty that our denudedoocytes were free from any contamination of follicular cells, andthat a low permeable dye (PI) was suitable for staining and for theselection of live follicular cells in this case.

Next, we attempted to determine the major production sites ofthe progestin receptor in denuded oocytes and detached follicularcells at 6:00 just prior to maturation. The protein levels of mPR� andmPR� were noticeably higher in the denuded oocytes comparedto those in follicular cells (Fig. 1A). Concurrently, the transcriptsof the mPRs, particularly mPR˛, were significantly higher in thedenuded oocytes prior to oocyte maturation in the morning (6:00)and decreased following spawning (Fig. 1B). In contrast, the expres-sion of nPR transcript was restricted entirely to the follicular layerssurrounding the stage IV oocytes and was significantly higher instage IV follicular cells at 6:00, prior to ovulation (Fig. 1A and B).

To clarify the localization of the progestin receptor expressedin non-native mammalian cells (Fig. 1), and as a comparison tothose in native oocytes (Fig. 3, supplemental Fig. 5), we examinedthe over-expression of GFP tagged mPRs and nPR in transientlytransfected human embryonic kidney cells (HEK 293). The mPR�expressed clearly on the cell membrane, though some of thereceptor appeared to be trapped within intracellular membranecompartments (Fig. 1C). The mPR� expression was rather variable,localizing to membrane, intracellular, or cytoplasmic areas withincells. In contrast, the nPR expression was found exclusively in thenuclei and cytoplasm of transfected cells (Fig. 1C).

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Fig. 1. Expressions of membrane progestin receptor � (mPR�), mPR� and nuclear progestin receptor (nPR or Pgr) in zebrafish oocytes and transfected human embryonickidney 293 cell line (HEK293). (A) Representative Western analyses of mPR�, mPR� and nPR proteins in follicle-enclosed stage IV oocytes (>650 �m) sampled at 13:00(after spawning), 21:00 (evening), or 6:00 h (before spawning). The same analysis was also repeated in denuded oocytes and detached follicular cell layers collected at 6:00(right panel in A). (B) Changes of mPR˛, mPRˇ, and nPR transcripts in detached follicular layers and denuded stage IV oocytes (n = 8) analyzed by real-time quantitative PCR(qRT-PCR). Different letters above the error bars indicate that those groups are significantly different from each other at P < 0.05. UD: under detection limit. (C) Localizationof zebrafish mPR�-EGFP, mPR�-EGFP, and nPR-EGFP proteins in transfected HEK293 cells. White arrows indicate plasma membrane localization; black arrows indicateintracellular compartment localization; and asterisks indicate cytoplasmic/nuclear localization. Each experiment was repeated at least three times.

Then, we attempted to determine whether up-regulating pro-gestin receptors could affect nongenomic signaling of progestinand the outcome of final oocyte maturation in stage IV follicle-enclosed oocytes by expressing in vitro synthesized transcripts ofEGFP tagged mPR˛, mPRˇ, or nPR. Clearly, these exogenous tran-scripts of mPR˛, mPRˇ, and nPR could be translated successfully instage IV oocytes. The proteins of mPR� and mPR� were localizedat the oocyte membrane by probing the plasma membrane frac-tion of oocytes using a specific GFP antibody (Fig. 3) or fluorescentimages (Fig. 3A, supplemental Fig. 5A and B). In contrast, nPR-GFPfusion or control GFP protein was found only in the cytosolic frac-tion of the oocytes (Fig. 3B, supplemental Fig. 5C and D). In addition,microinjection did not significantly affect the capacity of oocytes toundergo oocyte maturation in response to DHP stimulation com-pared to un-injected oocytes (supplemental Fig. 6).

Interestingly, over-expression of mPR� increased the num-ber of follicle-enclosed oocytes that underwent maturation in theabsence of exogenous progestin, while no significant differencewas observed in nPR or mPRˇ injected groups compared to thecontrol groups (Fig. 4A). Furthermore, over-expression of mPR�increased MAPK activation (p-ERK: phosphorylated MAPK) andcyclin B production in follicle-enclosed oocytes concomitantly withthe increase in maturation. In contrast, no significant change ofthese signaling molecules was observed in mPRˇ or nPR injectedgroup compared to control injected oocytes (Fig. 4B).

Over-expressions of mPR� or nPR in follicle-enclosed oocytessignificantly acceleratedDHP induced oocyte maturation comparedto mPR� and control injected oocytes (Fig. 5A and B). A significant

acceleration of maturation in oocytes over-expressing mPR� or nPRbecame apparent at 1 h and 2 h, respectively, following DHP stim-ulation (Fig. 5A and B). Interestingly, actinomycin D blocked nPRinduced acceleration, while the mPR� initiated acceleration wasunaffected by treatment with actinomycin D (Fig. 5C and D).

Follicle-enclosed oocytes over-expressing mPR� underwentoocyte maturation significantly faster at lower concentrations ofDHP compared to those of control injected oocytes without exoge-nous stimulus (Fig. 5E, supplemental Table 1). Expression of mPR�alone increased significantly the numbers of oocytes that under-went final oocyte maturation compared to the control injection(Fig. 5E, supplemental Table 1). Interestingly, incubation of mPR�antibodies decelerated DHP induced oocyte maturation (Fig. 5F,supplemental Fig. 7 and its Table 2), while incubation of oocyteswith multiple mPR� or nPR antibodies did not affect the rate ofoocyte maturation (supplemental Fig. 7). Activation of the MAPKpathway and production of cyclin B appeared to be greater and ear-lier in follicle-enclosed oocytes over-expressing mPR�-EGFP thanin control oocytes (Fig. 6A). An apparent earlier increase in p-MAPKwas also observed in oocytes over-expressing nPR, but there wasno observed difference in cyclin B production compared to the con-trol. Similar results were obtained in stage IV oocytes injected withthe mRNAs of mPR�, mPRˇ, or nPR without the GFP tag (results notshown).

Interestingly, the transcript levels of mPR˛ and nPR in stage IVfollicle-enclosed oocytes were 10–100 times higher than those ofmPRˇ (Fig. 6B). A greater than two fold increase in the amount ofmPR� and mPR� transcripts and an approximate 50% reduction

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Fig. 2. Representative surface images of follicle-enclosed stage IV oocytes (A–F)compared to denuded stage IV oocytes (G–L), photographed using a scanningelectronic microscope (SEM) or a confocal scanning microscope. (A), (B), (G) and(H) are recorded under the SEM; (B) and (H) show the magnified surface of the samefollicle-enclosed oocyte in picture A or the denuded oocyte in picture G. Images of(C–F) and (I–L) were photographed under a confocal microscope. (C), (E), (I) and (K)are projected confocal images of follicle-enclosed oocytes (C and E) compared todenuded oocytes (I and K) stained with 4’,6-diamidino-2-phenylindol (DAPI, C andI) or propidium iodide (PI, E and K). (D), (F), (J) and (L) were taken using differen-tial interference contrast (DIC), corresponding to the same oocytes in the picturesdirectly above (C), (E), (I) and (K), respectively.

in nPR transcript were observed in un-injected follicle-enclosedoocytes at 90 min following the incubation with DHP (Fig. 6).

4. Discussion

This study demonstrates that mPR� is the nongenomic pro-gestin receptor (Ngpr) that interprets extracellular signaling ofprogestins and initiates meiosis resumption in zebrafish. Combinedwith results from previous studies, supporting evidence includeslocalization of mPR� at the oocyte membrane, up-regulation ofthe receptor prior to oocyte maturation, specific MIS binding, rapidactivation of secondary messengers through a Gi coupled pathway,blocking of oocyte maturation by mPR� specific antibodies, accel-eration of oocyte maturation by over-expression of mPR�, and thelack of effect of the transcription inhibitor actinomycin D on mPR�induced oocyte maturation (Hanna et al., 2006; Hanna and Zhu,2009; Thomas et al., 2007; Tokumoto et al., 2006; Tubbs et al., 2010;Zhu et al., 2003a,b).

We have summarized the current understanding of mPR� andnPR initiated nongenomic signaling leading to resumption of cellcycle in Fig. 7. To our knowledge, we have shown for the firsttime that up-regulating mPR� alone initiates meiosis resumptionwithout exogenous progestins, due to increased phosphorylation ofMAPK (ERK1/ERK2) and the production of cyclin B proteins. Trans-lation of cyclin B protein from pre-stored mRNA is required forassembling the cdc2/cyclin B complex (maturation-promoting fac-tor) and the meiosis resumption in fish oocytes (Hirai et al., 1992;Kishimoto, 2003; Kondo et al., 1997; Nagahama, 1997). Therefore,our results support the role of zebrafish mPR� as an Ngpr thatinitiates meiosis resumption. One possible interpretation for themPR� accelerated maturation is that there is a “signal transduc-tion repressor” that can be dissociated by an increased amount ofthe receptor. The increased amount of the receptor is sufficient inactivating signaling pathways by dislodging a signal transductionrepressor such as the inhibitory subunit of protein kinase K. Thishypothesis is supported in part by recent reports on human mPRsstudies in the yeast model. Yeast does not produce endogenousprogestin nor human mPRs. Lyon and his co-workers found mPRconstitutively transduced signals without ligand when expressedat high levels, but did require an agonist when expressed at lowlevels (Kupchak et al., 2009; Smith et al., 2008; Villa et al., 2009).Alternatively, mPR� may constitutively produce very low quanti-ties of a second messenger that are too low to activate downstreameffectors under normal conditions. When mPR� is up-regulated oractivated, the net production of the messenger increases above thethreshold necessary to initiate downstream signaling moleculesthat initiate oocyte meiosis. It is also possible that residual amountof steroids such as DHP produced by follicles endogenously dur-ing the incubation period is able to bind over-expressed mPR� toactivate downstream signaling, and therefore induces final oocytematuration. Further studies are required to test these hypotheses.

Increased maturational competence by luteinizing hormonein oocytes might partly be due to up-regulation of the matu-ration steroid receptor (MIS) in surrounding follicular cells (Ge,2005; Pang and Ge, 2002; Patino and Thomas, 1990). Indeed, up-regulation of mPR� protein levels in oocytes by gonadotropinsoccurs concomitantly with the increased sensitivity of oocytes toMIS in several fish species including seatrout, goldfish, Atlanticcroaker, and zebrafish (Tan et al., 2009; Tokumoto et al., 2006;Tubbs et al., 2010; Zhu et al., 2003b). Current results on theincreased number of oocytes undergoing FOM when mPR� is over-expressed are consistent with our previous findings (Zhu et al.,2003a; Hanna et al., 2006). Furthermore, the mPR� initiated activa-tion of meiotic signaling appears to be specific, as mPR� antibodiesblocked this activation in follicle-enclosed oocytes. However, theseresults should be interpreted cautiously, as it is unclear whether

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Fig. 3. (A) Expression of mPR�-GFP fusion proteins in a representative oocyte microinjected with the transgenic transcript. Follicular cells were removed for better imagingunder a fluorescent microscope; (B) Western analyses of expressions of transgenic proteins for membrane progestin receptor � (mPR�), mPR�, or nuclear progestin receptor(nPR) in protein extractions of purified plasma membrane (for mPRs) or cytosolic fraction (for nPR) from stage IV zebrafish oocytes. These oocytes were microinjected ex vivowith fused transcripts generated by fusing mPR�, mPR�, or nPR in front of enhanced green fluorescent protein. Samples were collected following 5-h incubation in a culturemedium. Representative results are shown as positive reactions (bands) to an anti-GFP antibody Con: control oocytes injected with a GFP carrier plasmid.

these antibodies are acting directly on the oocytes through thefollicular layer, or whether there is a likely blockage in steroidalresponses at the follicular level where the mPR� is also abundantlypresent.

In contrast to previous reports (Bayaa et al., 2000; Josefsberg-Ben-Yehoshua et al., 2007; Tian et al., 2000), nPR appeared to playno major role in deciphering extracellular signaling of progestinand directing oocyte meiosis in zebrafish. In zebrafish, the level of

Fig. 4. Enhanced oocyte maturation and activities of meiotic signaling moleculeswithout aid of exogenous steroid in follicle-enclosed stage IV oocytes by over-expression of mPR� at 5 h following the microinjection. (A) Comparing percentagesof oocytes that underwent GVBD without exogenous DHP in oocytes microinjectedwith transcripts of mPR˛, mPRˇ, or nPR. * denotes significant difference comparingto the control groups (Con and UN, P < 0.05). Control (Con) oocytes were micro-injected with EGFP transcripts. UN: un-injected control. (B) Representative Westernblot analysis of activity of MAPK (p-ERK: phosphorylated MAPK) and production ofcyclin B protein. The total amount of MAPK (ERK) was determined as a loadingcontrol. The experiment was repeated three times.

nPR transcript was below the detection limit in late stage (stageIV) oocytes, although it increased dramatically prior to ovulationin the follicular layers surrounding late stage maturational com-petent oocytes. The major production site and striking changes ofnPR transcript in follicular cells are more conducive to nPR’s rolein ovulation than in oocyte maturation (Pinter and Thomas, 1995;Pinter and Thomas, 1997). A significant role of nPR in the ovulationand follicular layer function of rodents and humans is well recog-nized (Conneely et al., 2002; Svensson et al., 2000). However, theoocyte is a very unique cell that can import large proteins suchas vitellogenin (>100 kD) actively into the oocyte (Hyllner et al.,1994), and could possibly introduce nPR protein (69 kD) from thefollicle where it is present in high levels. Alternatively, nPR pro-tein could be synthesized in immature oocytes of earlier stagesas we have previously observed (Hanna et al., 2010). In addition,nPR has been suggested to play a role in the maturation of Xenopusoocytes (Bayaa et al., 2000; Josefsberg-Ben-Yehoshua et al., 2007;Tian et al., 2000); and the involvement of nongenomic progestinsignaling (NGPS) in cancer cell lines (Fu et al., 2008). Our currentexperimental results could not definitively exclude potential role(s)of nPR in NGPS in zebrafish oocytes. Our well designed experimentsincluding appropriate controls allow us to conclude that mPR� islargely responsible for extracellular NGPS during the FOM. How-ever, we should be cautious with this conclusion due to the constantexchanges of proteins and steroids between the follicular cells andoocytes. Further studies are required to address these questions.

The time delay of accelerated oocyte maturation induced byover-expression of nPR, compared to the effect induced by mPR�over-expression under stimulations of DHP, was likely due to theadditional time required for the nongenomic and/or transcriptionalactivities of nPR. Blocking of the nPR signaling pathway by thetranscription inhibitor actinomycin D in the current study sup-ports the requirement of transcriptional activity of nPR in inducingoocyte meiosis in zebrafish. Lack of cyclin B production in nPR over-expressing oocytes further supported this hypothesis, even thoughan earlier activation of MAPK was observed. Previous studies haveidentified that nongenomic signaling pathways of progestins are inpart due to activation of MAPK signaling by nPR in Xenopus oocytesand transfected breast cancer cells (Bayaa et al., 2000; Faivre et al.,2005; Shao et al., 2003; Tian et al., 2000). When Xenopus oocytesand cancer cells are compared with fish oocytes, one key differencefor the resumption of cell cycle and assembly of the MPF complexin fish oocytes is the requirement of the de novo synthesis of cyclin

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Fig. 5. Acceleration of oocyte maturation in follicle-enclosed stage IV oocytes by over-expression of progestin receptors. These oocytes were exposed to 100 nM exogenous17�,20�-dihydroxy-progesterone (DHP) unless indicated otherwise. (A and B) Percentage of oocytes underwent GVBD in oocytes microinjected with in vitro synthesizedtranscripts of mPR˛, mPRˇ, or nPR; (C and D): effects of the transcriptional inhibitor actinomycin D (AD) on DHP induced GVBD in oocytes microinjected with mPR˛ (C) or nPR(D); (E) dose-dependent acceleration of GVBD in oocytes microinjected with mPR˛ treated with 1–100 nM concentrations of DHP. * denotes significant difference in GVBDrates in oocytes injected with mPR˛ or nPR transcripts comparing to those control groups with the corresponding treatments (Con) (P < 0.05). See supplemental Table 1 foradditional details for (E). (F) Deceleration of GVBD in oocytes pre-incubated with mPR� antibodies. As controls, oocytes were pre-incubated with pre-serum or media (UN).* denotes significant decelerations in GVBD rates in oocytes pre-exposed with mPR� antiserum comparing to those control groups (untreated or treated with pre-serum)(P < 0.05). See supplemental Fig. 7 and Table 2 for additional details of (F). Over 60 oocytes were used for each treatment in each experiment. Each experiment was repeatedat least four times. The results show as average (mean ± SDE) of data from four representative experiments. Con: control oocytes were microinjected with EGFP transcript;UN: un-injected oocytes.

B, which cannot be primed by activation of MAPK alone (Kajiura-Kobayashi et al., 2000). Nevertheless, any potential nPR inducedNGPS does not appear to be sufficient to initiate oocyte meiosisand the resumption of cell cycle in the presence of actinomycin D,implying that an alternate genomic mechanism is required. In thisrespect, zebrafish could serve as an excellent model for future stud-ies of nongenomic and genomic signaling pathways of nPR, and itsinteractions with the nongenomic pathways of mPRs in control ofthe cell cycle.

Interestingly, nPR mRNA decreased with MIS stimulation. Thisdown-regulation may be attributed to the follicular layer expres-sion and the subsequent function of nPR in ovulation. Dramaticdown-regulation of nPR in granulosa cells undergoing ovulationand luteinization have previously been observed in mammals(Conneely et al., 2002). In contrast, an increase in mPR� and mPR�,protein and mRNA was observed at 60–90 min following MIS stimu-lation in oocytes concomitant with an all or none response of oocytematuration. This also demonstrates functional regulation of mPRsby the MIS ligand. On the other hand, the transcriptional blocker

actinomycin D did little to block MIS induced oocyte maturationin control injected oocytes, implying that the newly synthesizedmPR� transcripts were not required for resumption of meiosis instage IV oocytes.

Finally, although the mPR� receptor was localized at the oocytemembrane and capable of regulating similar signaling moleculesin transfected cell lines as mPR� in a previous study (Hannaet al., 2006), the current study does not support the role of mPR�in directing progestin signaling and initiating meiosis resump-tion. There was no significant acceleration of the maturation ratewith mPR� over-expression. Additionally, the mPR� transcript wasexpressed in much lower levels (over 10 fold lower) than the mPR�in zebrafish oocytes. The mPR� may have a role in oocyte devel-opment since the expression levels peaked at night time priorto oocyte maturation. A similar role of mPR� in early germ celldevelopment has been implied in zebrafish testes, where mPR�expression is restricted to spermatogonia and early spermato-cytes, but not found in mature sperm or spermatids (Hanna andZhu, 2009). The mPR� may also inhibit the function and signal-

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Fig. 6. (A) Changes in the activities of MAPK (p-ERK) and production of cyclinB protein in stage IV follicle-enclosed oocytes microinjected with transcripts ofmembrane progestin receptor � (mPR˛) or nuclear progestin receptor (nPR) at thestart of incubation (0 time point), and at 30, 60 and 90 min in the presence of100 nM 17�,20�-dihydroxy-progesterone (DHP). The total amount of MAPK (ERK)was determined as a loading control. (B) The transcript abundance of mPR˛, mPRˇ,and nPR in un-injected ooctyes at the beginning of the incubation (0 time) and90 min after incubation with 100 nM DHP determined by quantitative real-time PCR(qRT-PCR). The data shows the means ± SEM (n = 6, *P < 0.05). Each experiment wasrepeated at least three times.

ing of mPR� since co-expression of mPR� and mPR� decreasedthe expression levels and signaling of either receptor comparedto mPR� expression alone (Hanna and Zhu, unpublished data).Interestingly, mPR� appears to be the major membrane progestinreceptor responsible for oocyte maturation in Xenopus, since mPR�appears to be absent in this species (Josefsberg-Ben-Yehoshua et al.,2007). Therefore, it is likely that mPR� in Xenopus plays a similarrole as mPR� in fish species. Further studies are required to definethe roles of mPR�, mPR� and nPR in oocyte maturation of other

Fig. 7. Generalized signaling pathways for membrane progestin receptor � (mPR�)and nuclear progestin receptor (nPR or Pgr) in a typical cell that expresses bothtypes of receptors. In response to the stimulus of extracellular signaling such as pro-gestin (17�, 20�-dihydroxy-progesterone, i.e., DHP), rapid nongenomic signalingis initiated by mPR� or nPR located at the cell membrane and cytoplasm, respec-tively. The binding of progestin to the mPR� initiates activation of an inhibitoryG-protein alpha subunit, which in turn down-regulates the activity of adenylylcyclase (Zhu et al., 2003a,b; Hanna et al., 2006; Thomas et al., 2007). This down-regulation sets off inhibition of cAMP production, which leads to de-activation ofprotein kinase A (PKA). The de-activation of PKA relieves the inhibitory effects ofPKA on MAPK and leads to the de novo synthesis of cyclin B protein from preexistingtranscript (see Nagahama and Yamashita, 2008 for detailed review). The activa-tion of MAPK also causes de-phosphorylation of pre-existing cell division controlprotein 2 (cdc2). Formation of active meiosis/mitosis promoting factor (MPF) fromnewly synthesized cyclin B protein and de-phosphorylated cdc2 results in re-entryof the cell cycle, including germinal vesicle breakdown (GVBD) and final oocytematuration (FOM) in fish and frogs. The nPR localized in the cytoplasm could alsoinitiate nongenomic responses under certain conditions, leading to up-regulationof MAPK. MAPK activation alone is sufficient in Xenopus oocytes and a breast can-cer cell line for initiating the resumption of the cell cycle due to pre-stored cyclinB protein. However, this MAPK activation alone is not sufficient for the forma-tion of MPF and initiation of meiosis in fish oocytes and likely the majority oftypes of animal cells due to the lack of de novo synthesis of cyclin B protein andother unidentified co-factors. For the cell cycle to continue, further nongenomicand/or genomic actions of progestin receptors, including production of novel fac-tors in the nucleus/cytoplasm, are required to facilitate the production of cyclin Bprotein Solid lines represent signaling pathways supported by experimental evi-dence. Dash lines represent possible signaling pathways that needs supportingevidence.

vertebrate species since discrepancies in their function are difficultto consolidate.

Further research is also needed to elicit the role of mPR�, to iden-tify additional factors activated by nPR, and to examine further theinteractions of mPRs and nPR in oocytes (Fig. 6). In humans andother mammals, no maturation inducers and receptors have beenidentified, though similar signaling through inhibition of cAMP andactivation of MAPK occurs during final oocyte maturation. Addi-tional research on signaling of mPRs and nPR in zebrafish shouldhelp identify analogous mechanisms directing oocyte maturationin mammals.

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Acknowledgements

This work was supported in part by the National Science Foun-dation Grant IBN-0315349 (Y.Z.) and East Carolina UniversityResearch and Creative Activity Grant (Y.Z.), East Carolina Univer-sity Thomas Harriot College of Arts and Sciences Research Award(Y.Z), and East Carolina University Division of Research & Gradu-ate Studies Research Development Awards (Y.Z.). We would like toacknowledge Dr. Thomas Fin, Dr. Tim Christensen, and Mr. MichaelReubens and Mr. Sean Daly for their technical assistance. Wewould also like to express our great appreciation to Dr. MasakaneYamashita at Hokkaido University, Japan, for sharing a goldfishcyclin B antibody, and Ms. Joyce Joines Newman for technical edit-ing.

Appendix A. Supplementary data

Supplementary data associated with this article can be found, inthe online version, at doi:10.1016/j.mce.2011.02.004.

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Supplemental Fig. 1: Showing the removal of follicular cells from an representative stage IV 1

oocyte at various steps of the de-follicular process, recorded digitally under a dissecting 2

microscope (Model: MZ76, Leica). A). The out-layer of the follicular layers of the oocyte was 3

disentangled slightly indicated by an arrow; Light reflections were adjusted differently from two 4

pictures showing below for a better viewing of the follicular layer. B) Half of the follicular layers 5

were pulled away from the same oocyte indicated by an arrow; C). a whole follicular layer was 6

pulled away completely from the same oocyte, which become a denuded oocyte. Arrows indicate 7

the follicular layer. 8

9

Supplemental Fig. 2: Representative surface images of a follicle-enclosed oocyte at an early 10

stage (stage III) (A & C) compared to a denuded oocyte at the same stage (B & D), photographed 11

under a scanning electronic microscope. Images C & D are the magnified surface of same 12

follicle-enclosed oocyte in picture A or the denuded oocyte in picture B at a high magnification. 13

The arrow in picture C indicates the most out-layer of the follicular layer. Neu: nucleus of 14

follicular cells. Arrows in picture C indicate likely deposited protein masses between the follicles 15

and the oocyte. * is the area that is magnified and inserted on top right corner of picture D 16

showing chronic pores of the chronic membrane. 17

18

Supplemental Fig. 3: Projected confocal images showing follicular nucleus in follicle-enclosed 19

oocytes stained by DAPI (A) or PI (B). * marks the relative position of 3 different oocytes in 20

each picture. 21

22

Supplemental Fig. 4: Comparison surface images of follicle-enclosed stage IV oocytes (A-D) 23

with denuded oocytes (E-H) stained with DAPI (pictures A & E) or with propidium iodide 24

nuclear staining (PI, pictures C & G), recorded under a regular-upright-fluorescent microscope 25

(Microphot-FX, Nikon). Pictures B, D, F & H were taken under the dark field, corresponding to 26

those same oocytes directly above (A, C, E & G), respectively. 27

28

Supplemental Figure 5: Representative fluorescent images of zebrafish stage IV oocytes 29

microinjected with in vitro synthesized GFP fusion transcripts of progestin receptors, recorded at 30

5 hours after injection. Enhanced green fluorescence protein (EGFP) was fused at C-terminal end 31

of the receptors. Oocytes were microinjected with EGFP fused transcripts of membrane progestin 32

receptor α (mPRα, picture A); mPRβ (picture B); nuclear progestin receptor (nPR, picture C), or 33

EGFP (picture D). Arrows indicate cell surface expression of the mPRs in the ooctyes. nPR-34

EGFP and EGFP were expressed inside the oocytes as soluble proteins (C and D). Also see 35

figure 3A for surface expression of mPR. 36

37

Supplemental Fig. 6: (A) Representative microphotographs of un-injected or dye-injected 38

follicle-enclosed stage IV oocytes at 5 hours following the incubation with vehicle (-) or 100 nM 39

17α,20β-dihydroxy-progesterone (i.e., DHP with + sign). (B) Percentages of oocytes undergoing 40

GVBD in un-injected (UN) and dye-injected (INJ) follicle-enclosed stage IV oocytes at 5 hours 41

following the incubation with DHP or vehicle. The numbers in parentheses above the bars were 42

total numbers of oocytes examined in each group. * denotes significant difference in GVBD rates 43

in DHP exposed oocytes comparing to those without exogenous DHP (P<0.01). (C) Percentages 44

of oocytes undergoing GVBD examined in a time course in dye-injected (Con inj.) or un-45

injected (UN) follicle-enclosed stage IV oocytes incubated with exogenous 100 nM DHP (red 46

and pink lines) or vehicle (green line). 47

48

Supplemental Fig. 7: Decelerations of DHP induced GVBD in follicle-enclosed stage IV 49

oocytes by mPRα antibodies. These oocytes were pre-incubated in 50% L-15 culture medium 50

containing antibodies against mPRα, mPRβ or nPR (1:100 dilution) for 2 hours prior to the 51

addition of 100nM DHP. As controls, oocytes were pre-incubated with media containing pre-52

serum or no serum (Un: Untreated). Results show as averages of three representative 53

experiments (n=100 oocytes per treatment). Also see supplemental table 2 for additional details. 54

Sup

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Stage III Follicle-Enclosed Oocytes or Denuded Oocytes

Supplemental Fig.3. Hanna et al

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Fig.2 Hanna et al Supplemental Fig.5. Hanna et al

A

C

17,20β-DHP

B

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Supplemental Fig.7 Hanna et al

Supplemental Fig.7 Hanna et al

Note: The numbers inside table show percentages of oocytes undergoing GVBD at

examined time following addition of various concentrations of DHP. The numbers in

red show significant difference in GVBD rates in those mPRα injected oocytes treated

with DHP comparing to those control Injected oocytes (Con) treated with 100 nM DHP

(5th row). The numbers in blue (4th row) show significant difference in GVBD in those

mPRα injected oocytes with no exogenous DHP comparing those control injected

oocytes without exogenous DHP (last row). Control oocytes were either injected with

GFP transcript from a carrier vector or un-injected . Comparing to GFP injected

control groups, similar GVBD rates were observed in un-injected groups (data not

shown). See figure 5E and figure 5 legend for more details.

Supplemental Table 1. DHP dose responses in mPRα injected oocytes

Time (min) 0 30 60 90 120 150 180 210

mPRα+100nM DHP 0 9.63+/-0.76 25.9+/-2.87 59.7+/-7.97 78.3+/-9.66 88.8+/-5.59 94.4+/-3.34 94.4+/-12.7

mPRα+10nM DHP 0 8.3+/-2.42 18.1+/-7.37 45.4+/-11.5 65.7+/-13.5 80.3+/-6.97 90.0+/-3.59 95.6+/-10.0

mPRα+1nM DHP 0 8.52+/-1.19 12.5+/-5.3 28.7+/-10.0 45.5+/-13.0 65.0+/-4.25 77.7+/-4.09 88.8+/-8.49

mPRα 0 0 0 4.25+/-5.44 9.32+/-5.82 15.6+/-7.92 25.3+/-6.99 30.3+/-8.25

Con+100nM DHP 0 2.25+/-0.93 2.44+/-1.00 28.8+/-6.93 49.6+/-8.43 68.4+/-8.25 84.7+/-3.5 93.8+/-9.84

Control 0 0 0 1.66+/-0.48 1.72+/-0.49 1.72+/-0.94 3.44+/-1.81 9.67+/-5.44

Supplemental Table 2. Inhibition of DHP Induced GVBD in

Stage IV Follicle-Enclosed Oocytes by mPRα Antibodies. Time (min) 0 30 60 90 120 150 180

Untreated 0 0 0 33.3+/-6.93 58.7+/-8.43 73.3+/-8.25 84.6+/-3.58

Pre-Serum 0 0 0 30.3+/-5.44 55.7+/-5.83 70.6+/-1.92 80.0+/-5.99

mPRβ1 0 0 1.84+/-1.25 10.5+/-10.0 47.0+/-13.0 63.1+/-4.25 75.0+/-4.09

mPRβ2 0 0 0 19.2+/-11.5 50.2+/-13.5 66.6+/-6.97 80.8+/-3.59

mPRβ3 0 0 0 22.7+/-9.97 60.8+/-9.66 75.0+/-5.59 81.2+/-8.34

nPR1 0 0 2.76+/-1.01 13.6+/-9.60 62.5+/-10.5 73.8+/-2.36 84.6+/-6.36

nPR2 0 0 2.66+/-1.35 13.3+/-11.5 60.2+/-9.84 71.4+/-3.33 84.6+/-3.12

mPRα1 0 0 0 15.7+/-10.0 40.5+/-12.7 73.3+/-7.37 78.5+/-5.42

mPRα2 0 0 0 0 18.8+/-8.55 42.8+/-5.96 80.2+/-4.59

mPRα3 0 0 0 6.2+/-5.97 15.3+/-7.66 23.0+/-5.59 40.1+/-8.34

mPRα4 0 0 0 7.3+/-4.60 25.6+/-10.5 50.6+/-3.36 75.8+/-3.36

Note: Numbers inside the table show percentages of oocytes undergoing GVBD. Numbers in

red show significant difference in GVBD rates in those oocytes with pre-exposure of mPRα

antibodies comparing to those oocytes with pre-exposure of different antibodies or untreated

control group (P<0.05). See supplemental figure 7 and figure 7 legend for more details.