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BOR - Papers in Press, published online ahead of print January 14, 2004.
Biol Reprod 2004, 10.1095/biolreprod.103.025684
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BIOLOGY OF REPRODUCTION 70, 1465–1474 (2004)
DOI: 10.1095/biolreprod.103.025684
© 2004 by the Society for the Study of Reproduction, Inc.

Gamete Biology

Estradiol and Its Membrane-Impermeable Conjugate (Estradiol-Bovine Serum Albumin) During In Vitro Maturation of Bovine Oocytes: Effects on Nuclear and Cytoplasmic Maturation, Cytoskeleton, and Embryo Quality

Anna R. Beker-van Woudenberg1, Helena T.A. van Tol, Bernard A.J. Roelen, Ben Colenbrander, and Mart M. Bevers2

Department of Farm Animal Health, Faculty of Veterinary Medicine, Utrecht University, 3584 CL Utrecht,The Netherlands


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
In various cell types, there is increasing evidence for nongenomic steroid effects, i.e., effects that are not mediated via the classical steroid receptors. However, little is known about the involvement of the nongenomic pathway of estradiol (E2) on mammalian oocyte in vitro maturation (IVM). The aim of this study was to investigate whether the effects of E2 on bovine oocyte IVM are mediated via a plasma membrane receptor (nongenomic). First, we investigated the expression of estradiol (classical) receptor alpha (ER{alpha}) and beta (ERß) mRNA in oocytes and cumulus cells (CC). We also studied the effects of different exposure times to E2 (before and after germinal vesicle breakdown, GVBD) on nuclear maturation. To study the possible involvement of the putative estradiol plasma membrane receptor on the IVM of oocytes, we used E2 conjugated with bovine serum albumin (E2-BSA), which cannot cross the plasma membranes. Our results demonstrate that oocytes expressed ERß mRNA, while CC expressed both ER{alpha} and ERß mRNA. Exposure to E2 during the first 8 h of culture (before GVBD) induced a block at the metaphase I stage (MI). However, the presence of E2 after GVBD induced an increase of oocytes with nuclear aberrations. Meiotic spindle organization was severely affected by E2 during IVM and multipolar spindle was the most frequently observed aberration. Exposure of oocytes to E2-BSA did not affect nuclear maturation, blastocyst formation rate, nor embryo quality. Our results suggest that the detrimental effects of E2 on in vitro nuclear maturation of bovine oocyte are not exerted via a plasma membrane receptor.

apoptosis, embryo, estradiol, estradiol receptor, meiosis


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
In vivo, bovine oocytes resume meiosis after the preovulatory LH peak, while resumption of meiosis occurs spontaneously when cumulus-oocyte complexes (COCs) are removed from their follicle and cultured in vitro under suitable conditions [1, 2]. During meiotic maturation, the oocyte undergoes germinal vesicle breakdown (GVBD), which involves a gradual chromatin condensation, disappearance of the nucleolus, and disintegration of the nuclear membrane [3]. In bovine oocytes matured in vitro, GVBD occurs around 6–8 h after the onset of the culture [4, 5]. Before GVBD, transcription and translation can take place, but after GVBD, transcription strongly declines [6]. It is suggested that, when the chromatin is condensed, the transcription system is practically inactive [7]. Therefore, it is likely that cellular changes that occur after GVBD are not dependent on transcription and are consequently considered as nongenomic effects.

It is well known that steroids, including estradiol (E2), exert their function by binding to intracellular receptors with subsequent direct activation of gene transcription and protein synthesis, a so-called classical genomic pathway (for review, see [8, 9]). The classical estrogen receptor (ER) exists as two subtypes: ER-alpha (ER{alpha}) and ER-beta (ERß), and these subtypes exhibit different tissue localization patterns and can have distinct biological functions. ER{alpha} mRNA is predominantly expressed in the uterus, mammary gland, testis, pituitary, liver, kidney, heart, and skeletal muscle; whereas ERß transcripts are significantly expressed in the ovary and prostate (for review, see [10]). Additionally, a third type of ER (gamma) has been described in teleost fish [11], although it has not been found in mammals.

In addition to the genomic effect, indication of nongenomic effects exerted by steroids has been described in a wide variety of cell types, such as pancreatic cells, neurons, sperm, endometrial cells, granulosa cells, and oocytes (for reviews, see [12–14]). Involvement of nongenomic steroid pathways in oocytes was first described in amphibians and fish, providing some of the most convincing examples known of rapid, nontranscriptional regulation by steroid hormones [15, 16]. In Xenopus, for instance, progesterone binding to a plasma membrane-receptor triggers oocyte maturation [17]. However, although these mechanisms are well described in lower vertebrates, little is known about the plasma membrane-mediated effects of E2 during in vitro maturation of bovine oocytes. Additionally, specific nongenomic responses seem to depend on type of steroid, cell, tissue, and species.

Many of the steroid-mediated pathways are suggested to be initiated at the plasma membrane and involve conventional second messenger cascades [13]. For instance, a rapid transient increase in the intracellular free-calcium concentration ([Ca2+]i) occurs in human oocytes [18] as well as in sperm [19] shortly after exposure to E2. However, the nature and characteristics of the mediating plasma membrane receptor remains controversial. In addition, direct steroid-membrane interactions without receptor involvement have also been described, e.g., an intercalation of the steroid into the membrane of target tissues, which might alter physicochemical membrane properties such as the fluidity and the microenvironment of membrane receptors [20].

A complete meiotic maturation includes the extrusion of the first polar body, which requires, apart from other events, proper meiotic spindle formation. These structural changes are associated with the reorganization of microtubules and microfilaments, which are the major cytoskeletal components in mammalian oocytes that provide the framework for chromosomal movement and cell division [21–23]. Defects of the microtubular system in the oocyte can induce loss or gain of chromosomes, leading to aneuploidy [24], resulting in abnormal embryonic development following fertilization. Recently, we reported that, in bovine, the presence of E2 during in vitro maturation (IVM) has a detrimental effect (both in denuded oocytes and COCs) on the nuclear maturation, including abnormal dispersion of chromosomes and on subsequent embryo development [25]. The adverse effects of E2 on nuclear maturation might be due to an improper spindle organization during IVM. Disruption of the meiotic spindle by synthetic estrogen analogues has been reported [26, 27]. One of the reasons for impairment of embryo development is an inadequate cytoplasmic maturation, resulting in an increase of apoptosis during postfertilization development. Although studies in endothelial cells [28], male germ cells [29], and cardiac myocytes [30] suggested that E2 inhibits apoptosis, little is known about the effect of the presence of E2 during IVM on apoptosis in subsequent bovine embryo development.

Estradiol is often used in maturation protocols, but the mechanisms of action of E2 on the oocyte IVM are poorly understood. The aim of this study was to examine whether the observed detrimental effects of the presence of E2 during IVM [25] is a consequence of binding to the classical receptor (genomic), which regulates transcription, or via a transcription-independent manner (nongenomic) by binding to a plasma membrane-receptor. For this, E2 conjugated with bovine serum albumin (E2-BSA), which cannot cross the plasma membranes of living cells [31], was used during IVM to assess whether E2 acts via a plasma membrane- bound receptor. In addition, we investigated the effect of exposure of oocytes to E2 either before or after GVBD, i.e., before or after transcription had ceased. We also studied the expression of ER{alpha} and ERß mRNA in germinal vesicle stage (GV) oocytes, by reverse transcriptase-polymerase chain reaction (RT-PCR). In addition, to elucidate the nuclear abnormalities and low embryo development (percentage of blastocyst formation) induced by E2, we investigated the effect of E2 during IVM on meiotic spindle organization and on DNA fragmentation/apoptosis following subsequent in vitro embryo development.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Collection and Culture of Oocytes

Bovine ovaries, obtained at a local slaughterhouse, were transported to the laboratory in a thermoflask within 2 h after slaughter. Antral follicles, between 2 and 8 mm in diameter, were aspirated and COCs were recovered from the follicular fluid aspirates after sedimentation for 15 min. Only intact COCs with a compact and multilayered cumulus investment were used in this study. In experiments requiring denuded oocytes, those COCs had their cumulus cells removed by vortexing. Subsequently, the oocytes were washed in Hepes buffered M199 with Earle salts and glutamine (catalog # 20011-011; Gibco BRL, Paisley, UK), then in M199 with Earle salts and glutamine (catalog # 31100-027; Gibco BRL) supplemented with 26.2 mM NaHCO3 (defined as M199) and randomly allocated in groups of 35 per well in four-well culture plates (Nunc A/S, Roskilde, Denmark). Culture of the oocytes was carried out in 500 µl M199 per well, at 39°C in a humidified atmosphere of 5% CO2 in air, for 22 h.

In Vitro Fertilization and Embryo Culture

Both in vitro fertilization (IVF) and in vitro embryo culture (IVC) took place at 39°C in a humidified atmosphere of 5% CO2 in air. All experiments were carried out using frozen semen from the same batch of the same bull. Spermatozoa were thawed in a 37°C water bath for 1 min and washed in a discontinuous Percoll gradient prepared by adding 1 ml of 90% (v/v) Percoll under 1 ml of 40% (v/v) Percoll in a 15-ml centrifuge tube (Greiner Bio-One, Frickenhausen, Germany). The semen samples were added on top of the Percoll gradient and centrifuged for 30 min at 27°C at 700 x g. The pellet was resuspended in 70 µl modified Tyrode medium. Groups of 35 oocytes were washed twice in M199 and transferred to a well of four-well culture plates containing 430 µl of fertilization medium (Fert-Talp), as described by Parrish et al. [32] and modified by Izadyar et al. [33]. Twenty microliters of sperm suspension (final concentration 0.5 x 106 spermatozoa/ml) plus 20 µl of heparin (final concentration 1.8 IU/ml; Sigma Chemical Co., St. Louis, MO) and 20 µl PHE (20 µM D-penicillamine, 10 µM hypotaurine, 1 µM epinephrine) were added. After 20 h of incubation, groups of 35 presumptive zygotes were randomly placed in a coculture system of 500 µl M199 supplemented with 10% (v/ v) fetal calf serum (FCS; Gibco BRL) on a monolayer of buffalo rat liver (BRL) cells in each well of four-well culture plates. At the fourth day of culture, the noncleaved embryos were removed. At the fourth and eight day of culture, embryos were transferred to fresh cocultures.

BRL-Cell Culture

Buffalo rat liver cells separated from the BRL cell line from the American Type Culture Collection (ATCC) were cultured routinely in a 1:1 mixture of Ham F12 medium (Gibco BRL) and Dulbecco modified Eagle medium (Gibco BRL) supplemented with 7.5% (v/v) FCS (Gibco BRL) and 0.1% (v/v) penicillin/streptomycin (Gibco BRL). These cells differ from those currently available from ATCC in that they exhibit contact inhibition of growth.

Assessment of Nuclear Maturation

After culture, the oocytes were fixed for 15 min in 2.5% (w/v) glutaraldehyde (Merck, Darmstadt, Germany) in phosphate-buffered saline (PBS), washed twice with PBS, stained with 0.285 µM 4,6-diamino-2- phenylindole (DAPI; Sigma Chemical Co.) in PBS and mounted on microscope slides [34]. Evaluation of the nuclear status was done by epifluorescence microscopy (BH2-RFCA; Olympus, Tokyo, Japan). The oocytes were classified into four categories: 1) germinal vesicle (GV), oocytes with diffuse or slightly condensed chromatin; 2) metaphase I (MI), oocytes with clumped or strongly condensed chromatin that formed an irregular network of individual bivalents or a metaphase plate but no polar body; 3) metaphase II (MII), oocytes with either a polar body or two chromatin spots; and 4) aberrations, oocytes with an abnormal chromosomal organization, e.g., when chromosomes were disperse (not aligned along the metaphase plate) or when chromatin was persistent in the central region at the time of the late anaphase/telophase.

Assessment of Embryo Development

At Day 4 of culture (Day 0 = day of fertilization), the total cleavage rate and the number of cleaved embryos consisting of eight or more cells were evaluated. The embryos were examined by morphology at Day 9 and the percentage of blastocysts and hatched blastocysts was expressed on the basis of the number of oocytes at the onset of the culture.

TUNEL Assay

Terminal deoxynucleotidyl transferase (TdT) mediated dUTP nick-end labeling (TUNEL) assay was used as a way of evaluating bovine blastocyst quality and performed using a commercial kit (In Situ Cell Death Detection Kit, Fluorescein; Roche, Mannheim, Germany) as described by Gavrieli et al. [35]. Day 9 embryos were collected and rinsed twice in PBS + 0.1% (w/v) polyvinyl alcohol (PBS-PVA), stained for 5 min with 0.004 mM ethidium homodimer-1 (EthD-1; Molecular Probes, Inc., Eugene, OR) in PBS, then rinsed in PBS-PVA and fixed in 4% (w/v) paraformaldehyde in PBS overnight. Embryos were rinsed twice in PBS-PVA, permeabilized for 5 min on ice in PBS + 0.1% (v/v) Triton X-100 (Sigma Chemical Co.) and 0.1% (w/v) sodium citrate. Following washing in PBS-PVA, embryos were incubated in prewarmed (37°C) microdrops (30-µl drop per five embryos) of the TUNEL solution, partially covered with mineral oil (Sigma Chemical Co.) for 1 h, at 37°C, under a humidified atmosphere in the dark. Subsequently, embryos were rinsed twice in PBS-BSA and stained for 10 min with 0.285 µM DAPI in PBS. Finally, embryos were rinsed twice in PBS-PVA, once in plain PBS, mounted on a microscope slide with antifade mounting medium (Vectashield; Vector Laboratories, Burlingame, CA) and sealed under the coverslip with nail polish. Slides were evaluated under an epifluorescent microscope using filters at 500/ 580/525 nm.

Cells showing DNA-fragmented nuclei (TUNEL-positive staining) were termed apoptotic and cells with only plasma membrane damaged (ethidium-positive staining) were considered as dead. The total number of cells per embryo was determined by counting DAPI-positive nuclei. Apoptotic and dead cell indexes were calculated as the percentage of apoptotic or dead cells relative to the total number of cells, respectively. Only embryos showing at least one apoptotic or dead cell were included in the indexes.

Microtubules/Microfilaments Staining

All chemicals were purchased from Sigma Chemical Co. unless specified otherwise.

Staining of microtubules/microfilaments was performed as described by Tremoleda et al. [36]. Briefly, denuded oocytes were washed three times in PBS and permeabilized for 1 h at 39°C with medium M [37]. Prior to the microtubule staining, oocytes were washed three times in PBS + 0.1% (w/v) BSA (PBS-BSA). Microtubules were then labeled by incubating fixed oocytes for 90 min at 37°C in a 1:250 solution of a mouse monoclonal anti-{alpha}-tubulin antibody (catalog # T-5168; Sigma) in PBS- BSA + 0.01% (v/v) Triton-X (PBT). After this incubation, the oocytes were washed three times in PBT and then incubated at 37°C for 1 h in a blocking solution as described by Albertini et al. [38]. Next, the oocytes were exposed for 1 h at 37°C to a goat anti-mouse antibody conjugated to tetramethylrhodamine isothiocyanate (TRITC), diluted 1:250 in PBT. After washing twice in PBT and then twice more in PBS, the oocytes were incubated for 1 h at 37°C with Alexa Fluor 488 phalloidin (15 IU/ml; Molecular Probes, Inc.) to enable detection of the microfilaments. Finally, the oocytes were washed twice in PBS-BSA and, to visualize DNA, oocytes were incubated with monomeric cyanine nucleic acid stain (TO- PRO3, 1 mM in PBS; Molecular Probes, Inc.) for 15 min.

Spindles were classified as bipolar, having two intact poles equidistant from the metaphase plate, or multipolar, in which greater than two microtubule arrays were detected in a single oocyte [39].

Confocal Laser Scanning Microscopy

Oocytes were examined using a laser scanning confocal microscope (CLSM; Leica TCS MP; Leica, Heidelberg, Germany) mounted on an inverted microscope (Leica DM IRBE) equipped with 40 and 100x oil- immersion objectives. The CLSM was equipped with a krypto-argon ion laser for simultaneous excitation of Alexa Fluor 488 (microfilaments), TRITC (microtubules), and TO-PRO3 (DNA) using 488/568/650-nm excitation/barrier filter combinations.

Extraction of Total RNA and Reverse Transcription

Cumulus cells were separated from the oocytes using a narrow-bore Pasteur pipette. Denuded oocytes and cumulus cells were washed four times in PBS, collected in Eppendorf tubes, and stored at –80°C until RNA extraction. Ten denuded oocytes or cumulus cells from 10 COCs were collected per tube. Five independent replicates of each tissue sample were made.

Isolation of total RNA combined with on-column DNase digestion was performed using the RNeasy Mini Kit and the RNase-free DNase Set (Qiagen, Valencia, CA) according the manufacturer's instructions. Briefly, 300 µl lysis buffer was added to the frozen samples and vortexed. The lysates were then diluted (1:1) with 70% ethanol and applied to the mini column. After binding of the RNA to the column, DNA digestion was performed for 15 min at room temperature with RNase-free DNase (340 Kunitz units/ml). After washing of the membrane-bound RNA three times, the RNA was eluted with 30 µl RNase-free water.

Prior to the reverse transcription reaction, the RNA samples were incubated for 5 min at 70°C, vortexed, and chilled on ice. Reverse transcription was done in a total volume of 20 µl containing 10 µl of the sample RNA, 4 µl 5x reverse transcriptase buffer, 8 U RNAsin (Promega, Leiden, The Netherlands), 150 U Superscript II reverse transcriptase (Invitrogen, Breda, The Netherlands), 0.036 U random primers (Invitrogen), and final concentrations of 10 mM dithiothreitol and 0.5 mM of each dNTP (Promega). The mixtures were incubated for 1 h at 42°C, for 5 min at 95°C, and stored at –20°C. Minus RT blanks were prepared under the same conditions but without reverse transcriptase.

Amplification of ER{alpha} and ERß cDNA by PCR

PCR reactions were carried out in a total volume of 25 µl as described previously by Tremoleda et al. [40]. Amplification with ER-alpha-specific primers was performed in two stages; for the first round of amplification, the primers were ER-alpha L1 (5'-CATGATCAGGTCCACCTTC-3'; sense, GenBank identification number 334, position 9–28) and ER-alpha R1 (5'-GTGATCTTGTCCAGGACTCG-3'; antisense, position 322–341). To increase the recovery and specificity of the final product, a second round of heminested PCR was performed using primer ER-alpha L2 (5'- GCCTGGCTAGAGATCCTGA-3'; sense, position 36–55) and ER-alpha R1. Amplification with ER-beta-specific primers was performed in one round with primers ER-beta L1 (5'-ATCCATTGCCAGCCGTCAG-3'; sense, GenBank identification number 27806640, position 102–121) and ER-beta R1 (5'-TGTCGGCCAGCTTGGTGAG-3'; antisense, position 724–743).

The thermal cycling profile for the first round was initial denaturation and activation of the polymerase for 15 min at 94°C, followed by 40 cycles of 15 sec at 94°C, 30 sec at 55°C, and 45 sec at 72°C. Final extension was for 10 min at 72°C. For nesting, 1 µl of the first round product was transferred to 24 µl amplification mix and amplified for 30 cycles according to the same profile. A standard sequencing procedure (ABI PRISM 310 Genetic analyser; Applied Biosystems, Foster City, CA) was used to verify the analytical specificity of the PCR products.

Estradiol Preparation

A 734 µM estradiol (Sigma Chemical Co.) stock solution was prepared in ethanol (Merck) and stored at –20°C. Because membrane filters bind steroids, E2 was added aseptically from the stock solution after sterilization of the culture medium. On the day of the experiment, the stock solution was added to a Petri dish, the ethanol was evaporated, and only then M199 was added to the Petri dish. The dish was kept stirred for at least 5 h at 4°C. Then the medium was transferred to a four-well plate (500 µl per well) and put in the incubator (39°C in a humidified atmosphere of 5% CO2 in air) for at least 2 h before use. The final concentration of E2 was 3.67 µM and was determined by validated solid-phase 125I RIA method (TKE: Diagnostic Products Corporation, Los Angeles, CA) as described previously [41]. Control media underwent the same treatment without E2.

Estradiol-BSA Conjugate Preparation

A 1 µM stock solution of estradiol-BSA conjugate (E2-BSA; Sigma Chemical Co.) was prepared in M199 and stored at –20°C. On the day of the experiment, the stock solution was diluted in M199 to a final concentration of 0.1 µM of E2-BSA (equivalent to 3.67 µM of free E2 because one molecule BSA is conjugated with 35 molecules E2). Because the final concentration of BSA in the maturation medium supplemented with E2- BSA was 0.1 µM, the control maturation medium was supplemented with BSA to obtain an equivalent concentration.

Experiment 1: Different Time Exposure of Estradiol During Oocyte Maturation

Transcription is practically absent in oocytes after GVBD [6]. Therefore, to examine whether the observed impairment on nuclear maturation in the presence of E2 during IVM [25] is a consequence of binding to the classical receptor (genomic), which regulates transcription, or is via a transcription-independent manner (nongenomic), denuded oocytes were cultured in the presence of 3.67 µM estradiol at different times of exposure: a) during the whole culture period of 22 h (E2: 0–22 h), b) from 0 to 8 h of the culture (E2: 0–8 h, i.e., before GVBD), or c) from 8 to 22 h of the culture (E2: 8–22 h, i.e., after GVBD). After 8 h of culture, the oocytes were washed twice in M199 and then transferred to a new four-well dish containing the subsequent medium (according to the treatment). Media were prepared as described above. Cultures without E2 (during 22 h) served as controls. The percentage of oocytes at GV, MI, and MII and the percentage of nuclear aberrations were assessed at the end of the culture. Three independent experiments were performed.

Experiment 2: Effect of Estradiol on Meiotic Spindle

Denuded oocytes were cultured in the presence of 3.67 µM estradiol in the conditions described. Cultures without E2 served as controls. At the end of the culture, the oocytes were stained for microtubule, microfilament, and DNA and analyzed by confocal laser microscopy as described previously. Three independent experiments were performed.

Experiment 3: Effect of Estradiol-BSA Conjugateon Nuclear Maturation of Bovine Oocytes

Estradiol conjugated with BSA cannot cross the plasma membrane and therefore cannot exert an effect via binding to the classical intracellular ER. However, it can bind to a receptor on the plasma membrane, if present, with specificity and affinity similar to the free steroid form [42]. To examine how E2 could affect oocyte maturation, denuded oocytes were matured in the presence of a) 0.1 µM BSA, b) 0.1 µM E2-BSA conjugate, or c) 3.67 µM estradiol + 0.1 µM BSA under the conditions described above. Cultures without (free or conjugated) estradiol and BSA served as controls. The percentage of oocytes at GV, MI, and MII and the percentage of nuclear aberrations were assessed at the end of the culture. Five independent experiments were performed.

Experiment 4: Estradiol-BSA Conjugate During IVMand Its Effect on Subsequent Embryo Developmentand Embryo Quality

To determine whether exposure of COCs to E2 during IVM has an effect on subsequent embryo development and embryo quality and if those effects are exerted via a plasma membrane receptor, COCs were matured in M199 in the presence of a) 0.1 µM BSA, b) 0.1 µM E2-BSA conjugate, or c) 3.67 µM estradiol under the conditions described above. Prior to in vitro fertilization, the COCs were denuded and only oocytes showing an extruded polar body (PB) were fertilized. Subsequently, the presumptive zygotes were cultured for 9 days. Cultures in only M199 served as controls. The percentage of cleaved zygotes, embryos of eight or more cells, blastocysts, and hatched blastocysts were assessed. In addition, TUNEL- staining was performed on the blastocysts at the end of the culture. Five independent experiments were performed.

Statistical Analysis

The number of oocytes per stage (GV, MI, MII, and aberrations), of cleaved and embryos of eight or more cells at Day 4 and of blastocysts at Day 9 were analyzed by the chi-square test (SPSS for windows version 10.0.05; SPSS Inc., Chicago, IL). Cell numbers of embryos were analyzed by ANOVA (SPSS). Apoptotic and dead cell indexes were analyzed by logistic regression with overdispersion (Statistical software: R version 1.6.2; [www.r-project.org]). Differences were considered significant when P < 0.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Expression of mRNA for ER{alpha} and ERß on Bovine Oocyte and Cumulus Cells

Amplification of cDNA from cumulus cells primed with specific primers for ER{alpha} resulted in all samples in a specific DNA product after two rounds of amplification, while amplification of cDNA from oocytes yielded no specific product (Fig. 1). PCR analysis also demonstrated the presence of a specific band with expected size for ERß after one round of amplification in all five replicates generated from, respectively, oocytes and cumulus cells. Amplification of RT blanks yielded no specific products (Fig. 1). Therefore, our results demonstrate that bovine cumulus cells express both ER{alpha} and ERß mRNA, but bovine oocytes express only ERß mRNA. This different pattern of mRNA expression for ER might have implications on the mechanisms regulated by E2 because ER{alpha} and ERß can have distinct biological functions [10].



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  		FIG. 1. Expression of bovine estrogen receptor alpha (ER{alpha}) and beta (ERß) mRNA in oocytes and cumulus cells as detected by reverse transcriptase-polymerase chain reaction (RT-PCR). Samples are indicated at the bottom and a 100-base pair DNA ladder is indicated as the marker for fragment size

Effect of Estradiol on Nuclear Maturation Dependson Timing of Exposure

When COCs were cultured in the presence of E2 or E2- BSA during the whole period of culture (22 h), there was a significant decrease of the percentage of oocytes that reached MII stage, compared with control oocytes (Table 1). The low percentage of MII oocytes resulted from a significantly higher percentage of MI oocytes and also from a significantly higher percentage of aberrations compared with controls. However, when E2 was present only for the first 8 h of culture (Table 1; group 3) the decrease of the percentage of MII oocytes resulted from a high percentage of MI oocytes without a significant increase in the percentage of oocytes with nuclear aberrations. In addition, E2 present only after the first 8 h of culture (Table 1; group 4) also showed a significant decrease in the percentage of MII oocytes compared with control oocytes, but it was mostly caused by an increased percentage of aberrations compared with control oocytes. These results suggest that, although we cannot rule out that, during the first 8 h of culture, E2 can affect the in vitro maturation via a genomic pathway, we concluded that E2 is affecting nuclear maturation also in a nongenomic manner because effects were observed after condensation of chromatin (GVBD), when the transcription system in oocytes is practically inactive.


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  		TABLE 1. Effect of different periods of exposure to 17ß-estradiol (3.67 µM) on in vitro nuclear maturation of bovine denuded oocytes

Effect of Estradiol on the Meiotic Spindle Organization

Control oocytes at the GV stage did not have a detectable microtubule network in the cytoplasm, and microfilaments were seen in the cortex where the vesicle was located (Fig. 2A). During prometaphase stage, microtubules were observed in association with each chromatin particle (Fig. 2B). MI spindles were barrel shaped and were often located at the periphery of the cell (not shown). During anaphase I and telophase I (Fig. 2, C–E), microtubules were found in the central part of the spindle and a microfilament furrow was observed between the two sets of chromatin (Fig. 2D, arrow), indicating a role of microfilaments in polar body extrusion. MII spindles were often located close to the polar body and slightly rotated compared with MI spindles, yet they maintained a typical barrel shape with chromosomes lined up on midplate (Fig. 2F). However, when oocytes were matured in the presence of E2, spindle morphology was impaired in 35% of the oocytes (n = 52). Multipolar spindles (Fig. 3, A–E and G) and the persistence of chromatin in the central region of late anaphase/telophase (Fig. 3F, arrows) were the principal forms of anomaly observed in E2-treated oocytes. Some oocytes showed multipolar spindles even when chromosomes were properly aligned on the metaphase plate (Fig. 3E). Occasionally, spindles were fragmented into several small pieces distributed throughout the cytoplasm (Fig. 3, H–I). These results demonstrate that the presence of E2 during IVM has adverse effects on the spindle organization of bovine oocytes.



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  		FIG. 2. Confocal laser scanning photomicrographs of normal organization of microtubules/microfilaments in bovine oocytes. Green, microfilaments; red, microtubules; blue, chromatin. A) At the germinal vesicle stage, an organized microtubule network was not detectable. B) at prometaphase I, microtubules were tightly associated with the condensed chromatin. C–E) at late anaphase/telophase I stage, microtubules were seen in the central part of the spindle. Note the microfilament furrow between the two separated set of chromosomes (arrow). PB, Polar body. F) at metaphase II stage, a barrel- shaped spindle with chromosomes aligned up on midplate. A–F) Original magnification x1000



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  		FIG. 3. Confocal laser scanning photomicrographs of the aberrant spindle configuration of bovine oocytes cultured in the presence of estradiol. Green, microfilaments; red, microtubules; blue, chromatin. A–E) Multipolar spindles in metaphase I stage oocytes. F) Late anaphase/telophase I with lagging chromatin between microtubule array (arrows). G) Multipolar spindle in metaphase II stage oocytes. PB, Polar body. H–I) Spindle fragmentation. A–E) Original magnification x800; (F–I) original magnification x400

Estradiol-BSA Conjugate Does Not Affect Nuclear Maturation

Although (free) E2 + BSA (Table 2; group 4) induced a significant decrease of the percentage of MII oocytes and a significant increase of the percentage of GV, MI, and aberrations compared with oocytes exposed to E2 in the conjugate form (Table 2; group 3), there was no significant difference in the percentage of GV, MII, and aberrations when bovine oocytes were cultured for 22 h in M199 supplemented with E2-BSA conjugate compared with oocytes cultured in the presence of BSA. Even when a 10-fold higher concentration of E2-BSA (1 µM) was used, there was no significant difference in the percentage of aberrations, GV, MI, or MII compared with oocytes cultured in the presence of 10-fold concentrated BSA (control) (4.5%, 4.5%, 12.0%, 79.0% and 3.1%, 3.6%, 14.7%, 78.6%, respectively). These results indicate that the observed effects of E2 on in vitro nuclear maturation are not exerted via a putative membrane receptor because no significant effect was observed when E2 was conjugated with BSA, which cannot cross the plasma membrane.


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  		TABLE 2. Effect of estradiol (free or BSA conjugated) during IVM on nuclear maturation of bovine denuded oocytes

Estradiol-BSA Conjugate, During IVM, Does Not Affect Subsequent Embryo Development

When COCs were cultured in the presence of E2 or E2- BSA for 22 h, denuded and only oocytes with an extruded polar body were fertilized in vitro, the presence of E2 (free or conjugated) did not affect the subsequent embryo development in terms of cleavage rate, eight or more cells, and (hatched) blastocyst at D9 (Fig. 4) because there were no significant differences between groups for all of the parameters studied. This suggests that E2, either free or conjugated, does not affect oocyte cytoplasmic maturation.



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  		FIG. 4. Effect of estradiol (free or conjugated) during IVM on subsequent bovine embryo development of oocytes. (IVF was performed only with oocytes showing extruded polar body; mean ± SEM)

Apoptotic and Dead Cell Indexes Were Not Increasedby the Presence of Estradiol

At least one cell with DNA-fragmented nucleus and/or plasma membrane damage was detected in 94% of all the Day 9 embryos studied (n = 141), and it was similar for all groups. There was no significant difference between different groups in the percentage of embryos showing DNA fragmentation (TUNEL positive) or membrane damage (dead cells). The number of cells per embryo was also not affected by any treatment. In addition, the average apoptotic cell and dead cell indexes were not different between groups (Table 3). These results indicate that exposure to E2 during IVM does not affect the embryo quality in terms of apoptotic/dead cells when only MII oocytes with an extruded polar body were fertilized.


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  		TABLE 3. Effect of estradiol (free or BSA conjugated) during IVM on the incidence of apoptosis and dead cells of Day 9 blastocyst originated from IVF of selected oocytes (only MII oocytes with an extruded polar body)


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The present study demonstrates that the presence of estradiol in the culture media, either partially or during the whole period of IVM, had a negative effect on the in vitro nuclear maturation of bovine denuded oocytes. The pattern of this negative effect was dependent on the timing of exposure of the oocytes to E2. Exposure of bovine oocytes to E2 during the first 8 h of IVM (before GVBD) induced a block at MI stage for a high proportion of the oocytes without increasing the percentage of nuclear aberrations. Because transcription still can occur during the first 8 h of culture [6], a genomic effect of E2 during this period cannot be excluded. On the other hand, addition of E2 to the culture medium after the first 8 h of culture (after GVBD) causes a significant increase of the percentage of oocytes showing nuclear aberrations. The aberrations varied from multipolar spindles to metaphase plates showing outliner chromosomes, indicating damage to the meiotic spindle apparatus. This result, associated with the assumption that transcriptional nuclear activities are absent from 8 h after resumption of meiosis (after GVBD) [6], indicates that the effect of E2 on the metaphase spindle is mediated nongenomically, possibly by a direct binding of E2 to the microtubules and not via a plasma membrane because E2-BSA conjugate failed to induce those aberrations. Various studies with 2-methoxyestradiol (2MOE2), an endogenous metabolite of estradiol, demonstrated that 2MOE2 inhibits tubulin polymerization by direct binding at the colchicine site [43, 44], suggesting that interactions of estrogens and/or estrogen metabolites with tubulin and microtubule assembly may play important epigenetic roles. In addition, Aizu-Yokota et al. [45] demonstrated that the microtubule-disruptive effect of E2 is not associated with newly synthesized proteins and mRNA because cycloheximide and actinomycin D had no preventive action on the effect of E2. Furthermore, microtubule disruption can be induced by estradiol in estrogen receptor-negative human breast cancer cell lines, clearly indicating that estradiol can induce microtubule disruption independent of its binding to estrogen receptors [46].

Exposure of oocytes to E2-BSA conjugate during IVM affected neither the nuclear maturation of bovine oocytes (in terms of percentage of mature oocytes), the embryo development (in terms of the percentage of blastocysts), nor the embryo quality (in terms of cell number and apoptotic index). Our results are in agreement with Tesarik and Mendoza [18], who, using human oocytes, also did not find significant differences in percentage of MII oocytes when E2-BSA conjugate was present during IVM compared with the control. In contrast with our results, however, an increase of cleavage rates was described after IVM in the presence of E2-BSA conjugate. Differences in results might be explained by the fact that, in our study, oocytes were used from 2- to 8-mm follicles, which did not include large (preovulatory) follicles, while Tesarik and Mendoza [18] used GV oocytes originating from ovum-pick-up of gonadotropin-stimulated women undergoing micromanipulated- assisted fertilization. In addition, differences between the species might be relevant for the observed differences.

Here we demonstrated the expression of both ER{alpha} and ERß mRNA in cumulus cells. In addition, the present study is, to our knowledge, the first report about the expression of ERß mRNA in bovine oocytes. Although ovaries express both ER{alpha} and ERß mRNA, different expression patterns can occur within the heterogeneous cell types composing the tissue (see review [10]). In rats, ERß is preferentially localized in granulosa cells, whereas ER{alpha} is detectable in the surrounding thecal cells [47]. In bovine, however, Schams and Berisha [48] demonstrated the expression of ER{alpha} and ERß mRNAs in both theca and granulosa cells. Because, in our present study, all maturation experiments were done using denuded oocytes, it is likely that all genomic effects of E2 on those oocytes were mediated via ERß. However, in vivo, the importance of ER{alpha} mRNA expression in cumulus cells should not be underestimated. We previously demonstrated that the adverse effects of E2 on nuclear maturation of bovine in vitro-cultured oocytes were more pronounced in denuded oocytes compared with COCs, suggesting that cumulus cells could have a protective role [25]. In addition, ER{alpha} and ERß form both heterodimers and homodimers [49], and these forms may interact differentially with response elements on genes, suggesting that ER{alpha} and ERß may play different roles in gene regulation [50]. Moreover, indication for certain distinct biological functions of the ER subtypes is presented by different phenotypes of ER{alpha} and ERß knockout mice [10].

In a previous study, nuclear maturation was impaired when COCs and denuded oocytes were cultured in the presence of E2, although E2 did not affect the cytoplasmic maturation in terms of blastocyst formation [25]. In the present study, we extended this information by demonstrating that also embryo quality, in terms of apoptotic and dead cell indexes, was not affected by the presence of E2 during IVM when only oocytes showing extruded polar body were fertilized. The apoptotic cell index was relatively low for all treatment groups (range of 2–6%), which is in agreement with other studies [51, 52]. However VanSoom et al. [53] reported a higher average of apoptotic cell index (16.5%) for Day 9 blastocysts. The differences in the level of apoptosis might be due to the different in vitro culture conditions. In our study, despite the low number of apoptotic cells per embryo, almost all blastocysts showed at least one cell with DNA-fragmentation and/or plasma membrane damage. Our results are in agreement with other studies, which also showed some cell death in almost all in vitro- produced embryos [51, 53, 54]. Not only in vitro but also in vivo embryos can show signs of apoptosis. Hardy [55] reported over 80% of mouse blastocysts freshly flushed from the uterus on Day 4 or 5 had one or more fragmented nuclei. Although apoptosis may result from suboptimal culture conditions or may be involved in the elimination of abnormal cells (see review [56]), the presence of cells in vivo with classic features of apoptosis [57] indicates a role for apoptosis in normal development.

In summary, the present study demonstrates that the pattern of the negative effect of E2 on in vitro nuclear maturation of bovine oocytes depends on the timing of exposure of the oocytes to E2. When E2 was present in the first 8 h of culture, a higher percentage of oocytes arrested at the MI stage was the main feature observed. While, when oocytes were cultured in the presence of E2 only after the first 8 h of culture, the main effect observed was the increase of the percentage of nuclear aberrations. We also showed that E2 severely affects the meiotic spindle organization by increasing the percentage of multipolar spindles, possibly in a nongenomic manner. However, we did not find evidence for a membrane-bound receptor because E2 conjugated with BSA affected neither the oocyte nuclear maturation, the subsequent embryo development, nor embryo quality. In addition, we demonstrated that the presence of E2 during IVM neither affects cytoplasmic maturation in terms of blastocysts formation nor embryo quality in terms of apoptosis. Our results suggest that the negative effect of E2 on in vitro nuclear maturation of bovine oocytes is not exerted via a plasma membrane receptor.


   ACKNOWLEDGMENTS
 
We thank Dr. Peter Hendriksen and Dr. Fariboz Izadyar for critical reading of the manuscript. We are also grateful to Dr. Hans Vernooij for statistical support, Elly Zeinstra for her excellent technical assistance and Eric Schoevers for his assistance with the confocal microscopy.


   FOOTNOTES
 
1 Correspondence: Anna R. Beker-van Woudenberg, Department of Farm Animal Health, Faculty of Veterinary Medicine, Utrecht University, Yalelaan 7, 3584 CL Utrecht, The Netherlands. FAX: 31 30 2534811;a.beker{at}vet.uu.nl Back

2 Deceased Back

Received: 18 November 2003.

First decision: 4 December 2003.

Accepted: 6 January 2004.


   REFERENCES
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 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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