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Gamete Biology |
Institute of Reproductive and Developmental Biology,3 Imperial College London, Hammersmith Hospital, London W12 0NN, United Kingdom
Department of Mathematics,4 Imperial College London, London SW7 2AZ, United Kingdom
Department of Anatomy and Developmental Biology,5 University College London, London WC1E 6BT, United Kingdom
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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cumulus cells, follicle-stimulating hormone, gamete biology, meiosis, oocyte development
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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When oocytes and their associated cumulus cells are removed from the follicular environment, spontaneous maturation occurs [8, 9]. This approach is now commonly used for the in vitro maturation of oocytes in domestic species and is stimulating considerable interest as an approach for increasing the number of oocytes available for human-assisted reproduction [10]. As subsequent development of in vitro-matured oocytes is generally compromised [10–12], gonadotropins are routinely added to the culture medium to augment maturation [10, 13] and provide oocytes that are more developmentally competent [13–18]. However, with no clear consensus on the optimum concentration of FSH needed, there is considerable variation in the dose range used (10 IU eCG/ml or 10–75 mIU eCG/ml–1 human menopausal gonadotropin for human in vitro maturation [IVM]; 10–1000 ng/ml or 0.2 IU eCG/ml–1 FSH for mouse IVM) [10, 16, 19–21]. Even following maturation in the presence of gonadotropins, live birth rates remain low. The decreased developmental competence of oocytes matured in vitro highlights the importance of the maturation environment in the production of healthy oocytes.
In the antral follicle, FSH alters the environment of the maturing oocyte by stimulating granulosa cell division and differentiation [4], modulating steroidogenesis [15, 22, 23], granulosa cell metabolism [24, 25], and protein synthesis [7, 22] and inducing cumulus expansion [26]. At this time, the oocyte is undergoing critical cellular events that enable the egg to resume meiosis and complete cytoplasmic maturation, raising concerns about possible deleterious effects of elevated levels of FSH. Indeed, animal studies have shown that exogenous gonadotropins (superovulation) can have a detrimental effect on both preimplantation [22, 27– 30] and postimplantation development [27, 30–36]. Collectively, these studies suggest that exposure to high levels of FSH in vivo may have an adverse effect on follicular function and oocyte health and prompted us to examine the effects of varying doses of FSH in vitro on the maturation of mouse oocytes, using energy substrate uptake and production as a marker of the health of the cumulus-oocyte complex.
Our aim in this study was to ascertain whether increasing concentrations of FSH alter the nutritional environment of the maturing oocyte by changing uptake and production of the energy substrates pyruvate, glucose, and lactate by murine cumulus-oocyte complexes (COCs). Pyruvate, as an energy source, is obligatory for resumption of meiosis [37, 38] and is supplied by surrounding cumulus cells through the glycolysis of glucose [39, 40]. Levels of these energy substrates can have a profound effect on oocyte maturation [41], and adequate levels of pyruvate and glucose are important for progression to metaphase II [19, 42]. Cumulus-enclosed oocytes can be prevented from undergoing spontaneous maturation using inhibitory agents such as cyclic AMP and purines [5, 6]. FSH overcomes this inhibition, resulting in ligand-induced maturation [6]. The hypoxanthine-arrested cumulus-enclosed oocyte model has been used to study the role of FSH in modulating substrate metabolism and resumption of meiosis [6, 24, 25, 43] and has demonstrated, for example, that glucose is important for both maintenance of meiotic inhibition and ligand-induced maturation [44].
In this study, to model conventional in vitro-maturation protocols, we have taken the alternative approach of looking at the effect of a wide range of doses of FSH on the consumption and production of substrates by spontaneously maturing COCs. Furthermore, previous studies that have examined the effects of FSH on COC metabolism have used COCs from primed mice that have been administered FSH 48 h previously to increase the yield of antral follicles. In contrast, in this study, we have measured substrate production and uptake by individual COCs isolated from the ovaries of unstimulated, prepubertal mice [21], which had not been previously exposed to endogenous or exogenous FSH, and related the maturation status of the oocyte to the metabolism of naive cumulus cells.
This work differs from previous studies on the relationship between COC metabolism and the regulation of meiotic resumption. First, we have analyzed COCs from unprimed mice that have not been exposed to altered levels of FSH in vivo. Second, we have analyzed individual COCs, allowing us unequivocally to relate nutrient uptake and production to resumption and completion of meiosis. Third, we have examined the role of the PI3-kinase pathway in FSH-stimulated glucose uptake and metabolism. Finally, we have used a system to model IVM, where cumulus-enclosed oocytes are allowed to undergo spontaneous maturation in the presence of FSH. We have examined the response of spontaneously maturing COCs to a range of doses of FSH that encompass doses previously used.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Unstimulated 25- to 28-day-old F1 (C57BL/6 x CBA/Ca) female mice (Harlan Olac Ltd., Oxon, UK) were killed by cervical dislocation, the ovaries were removed and transferred to M2 medium [45] supplemented with 4 mg/ml bovine serum albumin (BSA, Fraction V; Sigma, Poole, UK). Large antral follicles were punctured with sterile acupuncture needles (AcuMedic, London, UK) to release COCs. Only COCs that consisted of an oocyte surrounded by at least two complete layers of cumulus cells were selected for further culture. Mice were housed and maintained in accordance with the Animals (Scientific Procedures) Act 1986 and associated codes of practice.
In Vitro Maturation of Oocytes
Minimum essential medium (MEM) Eagle (M2279; Sigma) containing 5.55 mM glucose (but no lactate) and supplemented with 0.23 mM sodium pyruvate (Sigma), 3 mg/ml BSA (Sigma), 50 U/ml penicillin, 50 µg/ml streptomycin, and 0.125 µg/ml amphotericin (catalog number 15240–039; GibcoBRL, Paisley, UK) was the basic culture medium used for in vitro maturation (IVM) and measurement of substrate uptake and production. This was supplemented with recombinant FSH (Puregon; Organon, Cambridge, UK) at concentrations of 0, 2, 20, 200, and 2000 ng/ml. One nanogram of Puregon contains 0.01 IU FSH. COCs were taken through four washes of MEM before being cultured individually in 5-µl drops of preequilibrated medium laid down using a positive displacement pipette (SMI digital adjust micropipettor, accuracy ± 1%; Alpha Laboratories, Eastleigh, Hants, UK) and overlaid with silicone oil (Dow Corning 200/ 50cs; BDH, VWR International, Lutterworth, UK) in an atmosphere of 5% CO2 in air at 37°C. The length of incubation was precisely noted and ranged from 15 to 16 h. Similar 5-µl drops of medium alone were incubated alongside the COC-containing drops and served as controls.
Scoring of Nuclear Stage of Maturation
At the end of the culture period, COCs were removed from the drop and the oocyte denuded of cumulus cells by repeated gentle pipetting. Oocytes were scored for stage of nuclear maturation and classified as being at the germinal vesicle (GV) stage if the nuclear membrane remained intact and was clearly visible by light microscopy, metaphase I if there was no visible nuclear membrane, or metaphase II following extrusion of the first polar body (Fig. 1, A—C).
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The oocytes were held in similar FSH-containing medium for up to 2 h before preparation for chromosomal analysis, to confirm the stage of nuclear maturation. Oocytes were spread and fixed as described by Tarkowski [46]. Briefly, oocytes were treated with hypotonic solution (1% sodium citrate) for 6–7 min and fixed in 5:1:4 methanol:acetic acid:water for 1 min or until the zona pellucida had disappeared. Oocytes were individually transferred to glass microscope slides and 13 µl of 3:1 methanol:acetic acid fixative was gently dropped onto the oocyte. As the fixative dried, further drops were added; this process was repeated 2–3 times. The oocyte was constantly observed throughout the fixation process using a Leica MZ12 dissecting microscope (Leica Microsystems, Milton Keynes, UK). Slides were dehydrated through an ethanol series (70%, 90%, and 100%; 5 min in each), air dried, and mounted in antifade Vectashield (Vector Laboratories, Peterborough, UK) containing 1.5 µg/ml DAPI (4,6-diamidino-2-phenylindole). Slides were analyzed using a Axioskop fluorescence microscope (Zeiss, Welwyn Garden City, UK) and images captured with a digital camera (Axiophot 2; Zeiss).
Measurement of Pyruvate, Lactate, and Glucose Uptake or Production
Individual 3-µl aliquots of culture medium from the experimental and control incubation drops were diluted with 597 µl of a 5 µM lactate solution (lactate standard; Sigma). All samples were stored at –70°C until analysis. The depletion of pyruvate or glucose and the production of lactate were measured by analyzing the difference between nutrient concentrations in the control and incubation drops as described previously [47].
Briefly, the assay is based on the conversion of substrate to product with the concurrent oxidation of NADH (pyruvate assay), reduction of NAD+ (lactate assay), or reduction of NADP+ (glucose assay) [48]. NADH and NADPH are highly fluorescent; hence, the amount of fluorescence emitted is proportional to the amount of NADH or NADPH present, which in turn is proportional to the amount of substrate in the sample. The fluorescence of the reactions was quantified using a COBAS Bio centrifugal autoanalyzer (Roche Products, Welwyn Garden City, UK) fitted with an FIA option, running assay programs as detailed previously [49]. Reactions were carried out at 25°C. Light of a particular excitation wavelength (340 nm) passes through the reaction mix in multiple cuvettes almost simultaneously, resulting in emission of secondary light, which is detected by a photomultiplier, with signals measured by a photometer. All chemicals were obtained from Boehringer Mannheim (Lewes, UK) unless otherwise stated.
Pyruvate assay Sixty microliters of diluted medium was automatically sampled and added to 200 µl of 50 mM phosphate buffer (30 mM K2HPO4 [BDH] and 20 mM NaH2PO4 [BDH]) containing 10 µM NADH. Ten microliters of lactate dehydrogenase (LDH) (300 IU/mg; 1500 IU/ml) diluted 1:80 with distilled water was added before fluorometric measurement.
Lactate assay Diluted medium (40 µl) was automatically sampled and added to 200 µl buffer containing 0.54 M glycine (Sigma), 0.2 M hydrazine sulphate (Sigma), 2.7 mM EDTA (BDH), 0.54 M NaOH (BDH), and 0.6 mM NAD. Twenty microliters of LDH, diluted 1:10 in distilled water, was added before fluorometric measurement.
Glucose assay Diluted medium (70 µl) was automatically sampled and added to 200 µl buffer containing 82 mM Tris (Sigma), 16.4 mM MgCl2 (BDH), 0.03 mM NADP, and 0.068 mM ATP. Hexokinase/glucose-6-phosphate dehydrogenase (40 µl), diluted 1:80 with distilled water, was added before fluorometric measurement.
Inhibition of PI3-Kinase Pathway with LY294002
The PI3-kinase inhibitor LY294002 (Sigma) was initially solubilized in 100% methanol (BDH, Analar grade) to make a stock of 16.27 mM before storage at –80°C. Mouse cumulus-oocyte complexes were collected and classified as described above. COCs were divided evenly between five treatment groups (Table 1).
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Preincubation COCs were initially preincubated for 30 min in the absence of FSH in medium with and without LY294002 (Table 1). COCs were washed through four drops of MEM supplemented with 0, 10, or 25 µM LY294002, before being individually cultured in 5-µl drops of the same medium overlaid with silicone oil (Dow Corning 200/50cS; BDH) in an atmosphere of 5% CO2 in air at 37°C. Control conditions included medium alone and medium with methanol carrier (Table 1). The concentration of methanol did not exceed 0.33%. At the end of the 30-min preincubation, COCs were moved to fresh drops of medium for overnight incubation with FSH (Puregon; Organon), continuing in the same concentration of LY294002 (see below).
Overnight incubation The culture medium for overnight incubation was supplemented with 2 ng/ml FSH (Puregon; Organon) and 10 or 25 µm LY294002. In addition, three control media contained 1) medium only, 2) medium and methanol carrier alone, and 3) 2 ng/ml FSH and methanol carrier (Table 1). COCs were washed through four washes of the final culture medium before being individually cultured in 5-µl drops of medium overlaid with silicone oil in an atmosphere of 5% CO2 in air at 37°C for 15–16 h. At the end of the culture period, the oocytes were removed from their surrounding cumulus cells and scored for stage of nuclear maturation. Spent culture medium was analyzed for glucose uptake and lactate production as described above.
Immunohistochemical Detection of GLUT4
Mouse COCs, collected as described above, were immediately individually processed for immunohistochemistry. All PBS (Oxoid Ltd., Basingstoke, Hants, UK) was filtered with a 0.22-µm filter (Millipore, Billerica, MA). COCs were fixed with 4% paraformaldehyde (Sigma) (1 h, room temperature) before washing once with PBS. COCs were then individually placed in 2% liquid agarose gel (low gelling temperature, <30°C) (Sigma) and left overnight at room temperature to set. A small section of the gel containing the COC was excised with a scalpel (
5 mm x 5 mm), serially dehydrated through alcohol (Analar; BDH; 70%, 1 h; 90%, 1 h; 100%, 1.5 h) and placed in xylene (Histoclear; National Diagnostics, Hessle, UK) for 1 h. Specimens were embedded in paraffin wax (Surgipath Europe Ltd., Peterborough, UK), serially sectioned (5 µm), and retrieved on poly-L-lysine-coated microscope slides (BDH). Sections were deparaffinized in two changes of xylene (10 min each) and then serially hydrated through alcohol (Analar; BDH; 10 min each 100%, twice; 90%, twice; 70%, twice; and finally, 5 min in distilled water, twice). Mouse skeletal muscle tissue (processed as described above) was used as a positive control.
Slides were examined under an Eclipse TE300 inverted light microscope (Nikon UK, Kingston-upon-Thames, UK) and those containing sections through the COC or muscle were identified. All incubations were carried out in a humidified chamber at ambient temperature (unless otherwise stated). Positive control sections of skeletal muscle were incubated for 15 min with 10 µg/ml pepsin (Sigma) to unmask antigen before processing in parallel with COC-containing slides. Slides were incubated for 15 min in 0.3% hydrogen peroxide (Merck) in water to quench endogenous peroxidase activity, washed with PBS (5 min), incubated with goat-blocking serum (Rabbit Unitect Immunohistochemistry System; Oncogene Research Products, VWR International, Lutterworth, UK) (20 min), washed again with PBS, and then incubated overnight at 4°C with primary rabbit polyclonal antibody to GLUT 4 (Santa Cruz: Sc 7938; Insight Biotechnology Limited, Wembley, UK). Antibodies were used at a dilution of 10 µg/ml (1:20 dilution, as determined by dilution curve) in 1% w/v PBS-BSA (Sigma). Some sections of skeletal muscle and COCs were incubated with rabbit IgG (10 µg/ml; Rabbit Unitect Immunohistochemistry System) in 1% w/v PBS-BSA (Sigma). Sections were rinsed three times in PBS (5 min each) before addition of biotinylated goat anti-rabbit IgG secondary antibody (Rabbit Unitect Immunohistochemistry System), diluted according to the manufacturer's instructions, for 30 min, then washed three times in PBS followed by application of peroxidase-conjugated avidin-biotin complex (Rabbit Unitect Immunohistochemistry System) for 30 min. Sections were washed in five changes of PBS, rinsed in 1% (v/ v) Triton-X 100/PBS (Sigma) (30 sec) and visualized with 3,3'-diaminobenzidine tetrahydrochloride (DAB; 60–90 sec; Zymed Laboratories Inc, San Francisco, CA). Sections were then counterstained with Harris Hematoxylin (BDH) (COCs, 20 sec; skeletal muscle, 2 min). Sections were finally dehydrated through a 70%, 90%, 100% ethanol series as described before and mounted in DPX (BDH).
Sections were examined using an Eclipse E600 light microscope (Nikon UK). Images were captured at the same light intensity using a DXM1200 digital camera (Nikon UK) and Lucia digital image analysis software (Nikon UK).
Confocal Analysis of Whole-Mount COCs
Mouse COCs were collected as described above and immediately processed for immunohistochemistry. COCs were washed in PBS, briefly (1 min) fixed in 100% methanol (BDH, Analar grade) and washed again in PBS. Fixed COCs were then incubated for 30 min in 0.1 M lysine (Sigma) in PBS containing 0.1% Triton-X-100 (Sigma). After three washes in PBS, COCs were transferred into a 10-µl drop of 10% swine-blocking serum (DAKO, Glostrup, Denmark) made up in PBS at room temperature for 30 min. COCs were rinsed in PBS before incubation in 10-µl drops of primary antibody (4 µg/ml; GLUT 4 Santa Cruz: Sc 7938; Insight Biotechnology Ltd.) diluted 1:50 in 1.5% v/v PBS-swine serum (DAKO) overnight at 4°C. After brief washes in PBS, COCs were transferred to 10-µl PBS drops containing a 1:50 dilution of FITC-labeled swine-anti-rabbit secondary antibody (DAKO) and incubated at room temperature for 1 h. Exposure to light was minimized during and following secondary antibody incubation. COCs were washed in PBS before being individually mounted in Vectashield containing 1.5 µg/ml DAPI (Vector Laboratories, Peterborough, UK) on a slide (BDH) within a ring of clear nail varnish (three coats thick). A coverslip was applied and sealed with nail varnish. Slides were examined by laser scanning confocal microscopy using a Leica SP2 laser scanning confocal microscope (Leica Microsystems). The DAPI was stimulated with the 351- and 363-nm laser lines and then the FITC was separately stimulated with 488-nm laser lines to eliminate any bleed through of the emission spectra. Images were stored digitally and reproduced using Adobe Photoshop (Adobe Systems, San Jose, CA).
Statistical Analysis
Numbers of oocytes cultured in all concentrations of FSH that reached metaphase II were compared using 
2 analysis for multiple groups. If the difference was significant (P < 0.05), differences between individual pairs of groups were compared using 2 analysis, and the P value was then corrected using the Bonferroni correction (multiplying the P value by the number of comparisons initially made, i.e., n(n – 1)/2, where n is the number of groups). These analyses were performed using GraphPad InStat version 3.0a for Macintosh (GraphPad Software, San Diego, CA; http://www.graphpad.com).
The uptake and/or production of pyruvate, glucose, and lactate by COCs at all concentrations of FSH were found to be normally distributed (using the Kolmogorov-Smirnov test) for oocytes that reached metaphase I or metaphase II. However, the standard deviations (SDs) were not the same in all of the groups (tested using the Bartlett test). The presence of negative values, indicating substrate production rather than uptake, precluded log or reciprocal transformation of the data, so analysis of variance was not appropriate. The data were therefore analyzed using the regression command in STATA 8 (STATA Corporation, College Station, TX). Inspection of raw data showed that the relationships between the dose of FSH and substrate uptake and/or production were not linear, leading us to treat the response to increasing doses of FSH as categorical data. Robust variance estimates were used to allow for the significantly different SDs described earlier; these were calculated using the robust option in STATA 8. Initially, regression of substrate uptake and/or production against dose of FSH was carried out independently for each stage. A regression of substrate uptake and/or production was then carried out against both dose of FSH, stage of maturation reached, and interaction terms, which allowed us to examine whether the COCs where the oocyte reached metaphase II responded to FSH differently from the COCs where the oocyte only reached metaphase I. Finally, the dependence of glucose and lactate uptake on the dose of LY294002 was analyzed using ordinary linear regression (treating dose as a continuous variable). Again, the calculations were performed in STATA 8 with the regression command and the robust option.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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A total of 352 COCs (constituting 18 replicate experiments) from 61 mice were matured in vitro, with between 69 and 72 COCs being cultured in each concentration of FSH for 15–16 h overnight. At the end of the incubation, culture medium was retained for analysis of substrate concentration (see below), and oocytes were immediately denuded to determine their maturation status. In a preliminary study, COCs were immediately denuded following isolation from the ovary and all oocytes were confirmed to be at the GV stage.
Incubation with FSH significantly increased the percentage of oocytes that reached metaphase II by the end of the incubation (P = 0.012, 2 for multiple groups, GV/Met I vs. Met II; Fig. 2).
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In the hour following denuding, an additional 29 oocytes, which had not extruded their first polar body at the time of denuding, reached metaphase II. There was no significant difference in the length of time between denuding and spreading of oocytes matured at each dose of FSH (P = 0.99, ANOVA). By the time the oocytes were spread, a similar proportion of oocytes had reached metaphase II irrespective of the dose of FSH (73%, 79%, 80%, 76%, and 78% at 0, 2, 20, 200, and 2000 ng/ml FSH, respectively). The majority of oocytes that completed maturation after denuding had been cultured in the absence of or in low concentrations of FSH. The difference between the maturation rate observed at the time of denuding and when the oocytes were spread, on average 1 h later, suggests that high concentrations of FSH could result in faster maturation. However, a similar proportion of oocytes will ultimately complete maturation irrespective of the presence or dose of FSH, given sufficient time.
All oocytes scored as Met I by light microscopy were confirmed to be Met I by examination of the configuration of chromosomes following spreading (Fig. 1, D and E).
Uptake and Production of Pyruvate, Glucose, and Lactate
Metabolic studies were performed on 14 replicates (274 COCs). COCs where the oocyte completed nuclear maturation and reached metaphase II are termed Met II COCs (n = 194), while COCs where the oocyte failed to complete maturation and only reached metaphase I are termed Met I COCs (n = 71). Twelve oocytes failed to resume meiosis and remained at the GV stage.
FSH had a significant overall effect on the uptake or production of pyruvate (P = 0.0002), glucose (P = 0.0009), and lactate (P < 0.0001) by Met II COCs (Fig. 3). At all concentrations of FSH, glucose uptake and lactate production by Met II COCs were significantly higher compared with control (0 ng/ml; Fig. 3). However, within this range of FSH concentrations, there was a marked lack of any dose effect and the effect of FSH on glucose uptake (P = 0.6) or lactate production (P = 0.7) by Met II COCs was not dose dependent (Fig. 3). Pyruvate uptake and production was also significantly influenced by FSH (P = 0.0002); under control culture conditions, pyruvate was taken up by Met II COCs at a rate of 4.5 ± 2.1 pmols COC–1 h–1. However, in the presence of FSH, Met II COCs produced pyruvate (Fig. 3). At higher concentrations of FSH (200 and 2000 ng/ml), there were significant amounts of pyruvate produced compared with control (P = 0.00003 and P = 0.02, respectively) (Fig. 3).
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In Met I COCs, in marked contrast with Met II COCs, FSH had no effect on pyruvate uptake/production (P = 0.39), glucose uptake (P = 0.84), or lactate production (P = 0.39). Indeed, Met I COCs had a different metabolic profile from Met II COCs (Fig. 3). Overall, Met II COCs had higher pyruvate production, glucose uptake, and lactate production than Met I COCs at the same concentration of FSH (pyruvate, P < 0.0001; glucose, P = 0.009; lactate, P < 0.0001). Specifically, in the absence of and at 2 ng/ ml FSH, there was no significant difference between Met I COCs and Met II COCs in substrate uptake and production. However, at higher concentrations of FSH, Met II COCs produced significantly more lactate than Met I COCs at 20 (P = 0.004), 200, and 2000 (P < 0.0001) ng/ml FSH. Similarly, Met II COCs produced significantly more pyruvate than Met I COCs at 200 (P = 0.005) and 2000 (P < 0.0001) ng/ml FSH.
Of the 274 COCs that were analyzed for substrate uptake or production, 71 contained an oocyte that had only reached metaphase 1 by the end of the culture. A proportion of these oocytes (15/71, 21%) completed maturation to metaphase II within 1 h 30 min of denuding, before being spread for chromosome analysis. Considering only COCs cultured in the presence of FSH (pooling data from 2 to 2000 ng/ml FSH), those that contained oocytes that matured after denuding produced twice as much lactate as COCs that contained oocytes that did not mature further (65.8 ± 12.7 pmol COC–1 h–1 compared with 30.1 ± 4.4 pmol COC–1 h–1; P = 0.02) (Fig. 4). Further linear regression analysis, taking account of both the stage of meiotic maturation and the dose, showed that the group that completed maturation after denuding had significantly different (P < 0.01) uptake/production of all three substrates compared with both those that had already matured by the time of denuding and those that never progressed beyond metaphase I.
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Expression of GLUT4 in Granulosa Cells
Immunohistochemistry of GLUT4, using either a biotinylated secondary antibody in paraffin sections of COCs (Fig. 1, F and G) or a fluorescently labeled secondary antibody in whole mount COCs (Fig. 1, H and I), demonstrated the presence of GLUT4 in the granulosa cells of COCs enclosing a GV-stage oocyte.
Effect of LY294002 on Nuclear Maturation
A total of 136 COCs (constituting seven replicate experiments) from 34 mice were matured in vitro, with between 26 and 28 COCs being cultured under each condition for 15–16 h overnight. At the end of the incubation, culture medium was retained for analysis of substrate concentration (see below), and oocytes were immediately denuded by gentle pipetting to determine their maturation status. The conditions under which the oocytes matured had a significant effect on the proportion of oocytes that reached metaphase II (P = 0.021, 2 for multiple groups, GV/Met I vs. Met II; Fig. 5). However, pairwise comparison showed that LY294002 had no significant effect on maturation (maturation in the presence of methanol carrier and 2 ng/ml FSH against the same media with 10 or 25 µM LY294002, P = 1 and 0.8, respectively, Bonferroni corrected).
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Effect of LY294002 on Uptake and Production of Glucose and Lactate
Metabolic studies were performed for all seven replicates (134 COCs: data for two Met 1 COCs not available). COCs where the oocyte completed nuclear maturation and reached metaphase II are termed Met II COCs (n = 90), while COCs where the oocyte failed to complete maturation and only reached metaphase I are termed Met I COCs (n = 38). Five oocytes failed to resume meiosis and remained at the GV stage and one oocyte had degenerated by the end of the culture period.
Glucose uptake by Met II COCs was similar under control conditions and in the presence of the methanol carrier (P = 0.9; two-tailed unpaired t-test) (Fig. 6) and was comparable with glucose uptake by the control group in the study described above (Fig. 3). Glucose uptake was again significantly increased above control values in the presence of 2 ng/ml FSH and the methanol control (P = 0.001, t-test). LY294002 significantly reduced FSH-stimulated glucose uptake in a dose-dependent manner (P = 0.02, linear regression) to the extent that glucose uptake in the presence of 25 µM LY294002 was similar to control levels, i.e., in the absence of FSH (P = 0.32, t-test) (Fig. 6).
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Lactate production by Met II COCs was also similar under control conditions and in the presence of the methanol carrier (P = 0.62, two-tailed unpaired t-test) (Fig. 6) and was comparable with lactate production by the control group in the study described earlier (Fig. 3). Lactate production was significantly increased above control values in the presence of 2 ng/ml FSH and the methanol carrier (P = 0.001, t-test). As seen for glucose uptake, LY294002 reduced lactate production in a dose-dependent manner, although this trend was not significant (P = 0.15, linear regression) (Fig. 6). Furthermore, lactate production remained significantly higher than controls in the presence of 25 µM LY294002 (P = 0.0001) (Fig. 6).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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In the absence of FSH, 57% of COCs in this study completed spontaneous maturation within 16 h, whereas a previous study showed higher spontaneous maturation rates (70–80%) in medium containing identical substrate concentrations [19]. In contrast with our results, the latter study [19] and others [16, 20], found that further supplementation of the medium with FSH did not increase maturation rates. The COCs from our study were, however, from unprimed immature mice, whereas all of the previous studies used mice that had been exposed to FSH in vivo before COC collection. Such priming, known to increase both antral follicle size and the developmental capacity of maturing oocytes [21], may be responsible for the higher control maturation rates and the absence of any further augmentation of maturation following exposure to FSH in vitro. It is possible that treating animals with FSH 48 h before retrieval of COCs primes the oocyte for maturation, perhaps by upregulating FSH-receptor expression in the cumulus cells [50], so that the COCs respond more rapidly to FSH in vitro.
Here we showed that FSH at doses of 20 ng/ml or higher significantly increased the proportion of oocytes reaching metaphase II after 15–16 h culture. However, when the oocytes were scored an hour later, a similar proportion of oocytes had completed maturation under all conditions, irrespective of the dose of FSH. This strongly suggests that FSH, at doses of 20 ng/ml or higher, accelerates maturation.
The mechanism by which FSH accelerates maturation is unclear. Gonadotropins are known to stimulate glucose consumption by cumulus cells, leading to pyruvate production [51, 52]. In the mouse, an FSH-stimulated increase in glycolysis, resulting in increased pyruvate production, was thought to be responsible for the increase in spontaneous meiotic maturation seen when glucose was the only available energy source [19]. An additional mechanism by which FSH action and glucose metabolism might interact to regulate meiosis unfolded from an elegant series of studies carried out by Downs and coworkers [24, 43, 53]. Glucose is required for FSH-induced maturation [44]. FSH increases hexokinase activity in the cumulus oophorus [43], facilitating glucose uptake and its conversion to glucose-6-phosphate. Furthermore, it is the pentose phosphate pathway, and not glycolysis, that is the metabolic route that mediates ligand-induced resumption of meiosis [24, 43]. The pentose phosphate pathway provides precursors for the de novo purine pathway, which in turn is involved in meiotic induction [53]. Alternatively, as has recently been shown, ATP induces Ca2+ release from intracellular stores in cumulus cells, which is then transmitted via gap junctions to the oocyte [54]. While FSH itself had no effect on Ca2+ in COCs, it may be that, in our study, the FSH-induced increased glucose uptake resulted in increased glycolytically produced ATP, leading to Ca2+ release and a stimulation of maturation.
In the current study, FSH stimulated glucose consumption and lactate and pyruvate production (and hence glycolysis) by spontaneously maturing individually cultured COCs in a manner similar to group cultured complexes undergoing ligand (FSH)-induced maturation [25, 43]. The significantly higher production of pyruvate at 200 and 2000 ng/ml FSH, with no concomitant increase in glucose uptake (Fig. 3), suggests that glucose could be being diverted to the glycolytic pathway. While the exact fate of the glucose consumed by COCs could not be ascertained, there was a significant correlation (P < 0.0001) between glucose uptake and both pyruvate and lactate production by individual COCs, suggesting that the glucose consumed in this instance was being metabolized, via glycolysis, to pyruvate, which is then converted to lactate, as has been observed previously [25].
While the presence of FSH had a significant effect on the rate of glucose consumption or lactate production, the dose of FSH was immaterial, suggesting that there is a threshold level of less than 2 ng/ml for FSH-induced glucose uptake. Glucose enters cells via facilitated glucose transporters (GLUT transporters), and we have detected the insulin-sensitive transporter GLUT4 in cumulus cells by immunohistochemistry. The lack of dose effect of FSH on glucose uptake is most likely to be caused by saturation of these transporters at the lowest dose of FSH (2 ng/ml). In contrast, pyruvate metabolism did appear to be influenced by the dose of FSH, with a significant increase in pyruvate production seen when COCs were exposed to 200 and 2000 ng/ml FSH. Thus, high levels of FSH appear to alter the nutritional environment of the oocyte by making more pyruvate available.
Culture of individual COCs allows direct and unequivocal comparison between complexes containing oocytes at the same stage of nuclear maturation. Glucose uptake and lactate production are highest when COCs are both exposed to FSH and contain a metaphase II oocyte at the end of culture. Cumulus cells take up most of the glucose; oocytes alone have low or undetectable glycolytic activity [24, 52, 55], so the differences in glucose uptake with maturational stage of the enclosed oocyte are unlikely to be due to changing metabolism of the maturing oocyte itself. Recently, a relationship between pyruvate utilization by COCs and the nuclear stage of the oocyte has been reported, with shifts in uptake and production of pyruvate being observed in group-cultured COCs at specific time points throughout oocyte maturation [25, 56]. Here we have shown that, in the presence of FSH, COCs containing immature oocytes consume pyruvate, while those where the oocyte completed maturation produce pyruvate, consume more glucose, and produce more lactate. The increased glycolysis seen in COCs where the oocyte has completed maturation may well result from the changing energy requirements of the oocyte as it undergoes extensive chromatin and cytoplasmic remodeling during nuclear maturation [57].
It is well established that a dialogue between the oocyte and surrounding somatic cells is necessary for successful follicle development [58–61], and the different metabolic patterns seen raise the possibility that a similar dialogue is taking place here, with either the cumulus cells directing maturation or the oocyte directing cumulus cell metabolism. The latter theory is supported by studies showing that glucose oxidation by the cumulus cells of bovine COCs was significantly decreased following removal of the oocyte [52]. The increase in glucose uptake by complexes that have completed nuclear maturation in the presence of FSH suggests that COCs containing a mature oocyte are more able to respond to FSH. Conversely, we must also consider the possibility that a reduced response to FSH by complexes containing an oocyte that fails to mature may simply be a reflection of COC health, with the COC having been isolated from an atretic follicle. However, a proportion of oocytes went on to complete nuclear maturation after denuding, and these oocytes resided in complexes that had a different metabolic profile from those that contained an oocyte that failed to mature (Fig. 4). These findings strengthen the hypothesis that substrate metabolism of the cumulus-oocyte complex is in some way related to meiotic maturation of the oocyte.
A major hormonal regulator of glucose uptake is insulin, which controls glucose uptake by modulating the partitioning of the insulin-sensitive facilitative glucose transporter GLUT4 within the cell [62]. Insulin stimulates the translocation of GLUT4 from intracellular stores to the membrane, hence increasing glucose uptake, and it is well established that this stimulation is mediated by the PI3-kinase pathway [63–65]. The observation that FSH can act in a similar manner in stimulating glucose uptake led us to examine whether FSH action was also mediated by the PI3-kinase pathway.
Using immunohistochemistry, we have demonstrated that GLUT4 is present in murine granulosa cells. Previous studies that have examined GLUT expression have produced conflicting results: GLUT4 expression has been detected by Northern blotting in sheep granulosa cells [66], but not in immature mouse [67] or rat [68] granulosa cells, using in situ hybridization or Western analysis, respectively. These variable results may be due to differences in methodology. Recently, we have demonstrated the presence of GLUT4 mRNA in human granulosa-lutein cells using reverse transcription-polymerase chain reaction, with sequence verification of the product (S. Rice, personal communication).
Incubation of FSH-stimulated mouse COCs in the PI3-kinase inhibitor LY294002 significantly decreased glucose uptake in a dose-dependent manner (Fig. 6), suggesting that FSH regulates glucose uptake via the PI3-kinase pathway. This is a likely scenario: it is now becoming clear that FSH can activate a variety of signaling cascades, including the PI3-kinase pathway. Signalling via the PI3-kinase pathway has been clearly demonstrated in rat granulosa cells by increased phosphorylation of protein kinase B (PKB/Akt) by FSH [69, 70]. Furthermore, FSH-stimulated cellular responses can by inhibited by PI3-kinase inhibitors [70, 71]. Our observations of GLUT4 protein in the granulosa cells (Fig. 1), coupled with the inhibition of FSH-stimulated glucose uptake by LY294002 in the intact COC (Fig. 6), suggests that FSH could activate the PI3-kinase pathway, resulting in translocation of GLUT4 to the granulosa cell membranes with a concomitant increase in glucose uptake.
Increased glycolytic activity by feline and bovine oocytes following IVM and before fertilization has been associated with improved preimplantation development [72, 73]. The ability of somatic cells to respond to FSH by increasing their glucose uptake and its subsequent metabolism may be an important factor not only in nuclear maturation, but may also significantly affect cytoplasmic maturation and subsequent preimplantation development, an important aspect of which must be the activation of metabolic pathways for the production of energy and the accumulation of the precursors of biosynthesis.
Here we have shown that FSH stimulates glucose uptake and pyruvate and lactate production in spontaneously maturing oocytes. The lack of a dose effect of FSH on glucose uptake suggests that only low physiological doses of FSH are required during IVM. FSH increases glucose uptake by activating the PI3-kinase pathway, and the presence of GLUT4 in the granulosa cells suggests a likely candidate for mediating this effect. High doses of FSH change the environment of the maturing oocyte by increasing pyruvate production by cumulus cells. Increased glucose uptake may be involved in the higher levels of maturation seen in high FSH concentrations, although whether this is regulated by increased availability of pyruvate, increased activity of the pentose phosphate pathway, or higher ATP levels remains unclear. Further studies are required to precisely evaluate the nature of the dialogue between the maturing oocyte and the surrounding cumulus cells in terms of changing FSH responsiveness and nutrient metabolism and to identify whether the cumulus cells or the oocyte directs this dialogue.
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2 Correspondence: Kate Hardy, Institute of Reproductive and Developmental Biology, Imperial College London, Hammersmith Hospital, Du Cane Road, London W12 0NN, UK. FAX: 44 0 207 594 2111; k.hardy{at}imperial.ac.uk
Received: 23 September 2003.
First decision: 2 October 2003.
Accepted: 3 March 2004.
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