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Advancing maternal age predisposes to mitochondrial damage and loss during maturation of equine oocytes in vitro
Theriogenology 81 (2014) 959–965
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Advancing maternal age predisposes to mitochondrial damage and loss during maturation of equine oocytes in vitro B.P.B. Rambags 1, D.C.J. van Boxtel, T. Tharasanit 2, J.A. Lenstra, B. Colenbrander, T.A.E. Stout* Department of Equine Sciences, Faculty of Veterinary Medicine, Utrecht University, Yalelaan 112, 3584 CM Utrecht, The Netherlands
a r t i c l e i n f o
a b s t r a c t
Article history: Received 16 August 2013 Received in revised form 8 January 2014 Accepted 16 January 2014
In many mammalian species, reproductive success decreases with maternal age. One proposed contributor to this age-related decrease in fertility is a reduction in the quantity or functionality of mitochondria in oocytes. This study examined whether maternal age or (in vitro maturation). IVM affect the quantity of mitochondria in equine oocytes. Oocytes were collected from the ovaries of slaughtered mares categorized as young (<12 years) or aged (12 years) and either denuded and prepared for analysis immediately (not-IVM) or matured in vitro for 30 hours before preparation (IVM). The mean oocyte mitochondrial DNA copy number was estimated by quantitative polymerase chain reaction and found to be significantly lower in oocytes from aged mares and that had been subjected to IVM than in any other group. Transmission electron microscopy demonstrated that mitochondria in aged mare oocytes subjected to IVM experienced significantly more swelling and loss of cristae than in other groups. We conclude that maternal aging is associated with a heightened susceptibility to mitochondrial damage and loss in equine oocytes, which manifests during IVM. This predisposition to mitochondrial degeneration probably contributes to reduced fertility in aged mares. Ó 2014 Elsevier Inc. All rights reserved.
Keywords: Horse Maternal age Mitochondrion Oocyte In vitro maturation
1. Introduction In many mammalian species, there is a threshold maternal age beyond which reproductive success decreases markedly [1]. In women, fertility declines after 35 years [2], largely as a result of an increase in the incidence of spontaneous abortion [3]. The decline culminates in the cessation of fertility in most women at around 41 years, even though the menopause does not onset until approximately 51 years [4]. Similarly, the likelihood of a live birth per embryo * Corresponding author. Tel.: þ31 30 2531350; fax: þ31 30 2537970. E-mail address: [email protected] (T.A.E. Stout). 1 Present address: uQinisa, Houten, The Netherlands. 2 Present address: Department of Obstetrics, Faculty of Veterinary Science, Chulalongkorn University, Bangkok, Thailand. 0093-691X/$ – see front matter Ó 2014 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.theriogenology.2014.01.020
transferred in human IVF programs decreases from 24% at maternal ages below 30 years to 8% at 42 years and 4% at 45 years or more [5]. The underlying causes of reproductive senescence have been proposed to include a decrease in the size of the resting follicle pool, a decrease in oocyte quality, and impaired endometrial receptivity. The size of the resting follicle pool certainly affects fertility in women, especially just before the menopause [6,7]. However, a reduction in oocyte number does not explain why many women are cyclic but unable to conceive in the last 10 years before the menopause [4]. Similarly, because the negative effects of advanced maternal age on IVF success can largely be overcome using an oocyte donated by a younger woman (35 years), impaired endometrial receptivity seems to be a relatively minor contributor to the premenopausal decline in fertility [5,8]. Instead, the primary contributor to reduced
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fertility is thought to be reduced oocyte quality; previous studies have reported a relatively low expression of spindle assembly checkpoint mRNA [9] and high incidences of spindle aberrations [10] and chromosomal abnormalities [11,12] in oocytes from aged women. Similarly, oviductal transfer of oocytes from aged mares (20–26 years) produced significantly fewer embryonic vesicles (31%) in inseminated young mare recipients than transfer of oocytes from young (6–10 years) donors (92%) [13]. One postulated underlying cause of the age-dependent decrease in oocyte quality is a decline in mitochondrial function [14,15]. Mitochondria play several important metabolic roles in eukaryotic cells, including energy generation by oxidative phosphorylation, steroid production, b-oxidation, and calcium homeostasis. However, mitochondria are also implicated in processes associated with cell deterioration, such as the production of potentially harmful reactive oxygen species (ROS) [16,17]. Because mtDNA is located close to the site of ROS generation, it may be particularly at risk for accumulating oxidative damage. Moreover, mtDNA is more sensitive to mutagens than nuclear DNA because it lacks introns, protective histones [18], and DNA repair mechanisms. Indeed, the mutation rate of mtDNA is more than 10 times higher than that of nuclear DNA [18], and point mutations, deletions, and duplications have been reported to accumulate in mtDNA over time, particularly in slowly or nondividing, postmitotic tissues with high energy demands, such as brain and muscle [14,19–22]. Mammalian oocytes are also nondividing, postmitotic cells; however, their energy demands are modest because they arrest after entering meiosis and remain in a resting phase until reactivation during final follicle development before ovulation. Nevertheless, because the resumption of meiosis may occur many years later, oocytes may accumulate mitochondrial DNA damage with increasing maternal age (for review, see Eichenlaub-Ritter et al. [23]). However, studies that have examined a possible age-related decline in oocyte mtDNA quality have produced conflicting results [24–30]. Even so, it was recently reported that increasing maternal age is accompanied by a decrease in oocyte mtDNA quantity in mice [31]. In this respect, earlier studies may have been biased by using relatively small populations of oocytes obtained from aged women attending IVF clinics for fertility problems. The aim of the present study was to determine whether maternal age significantly affects either mitochondrial number, estimated via mtDNA copy number, or electron microscopic morphological quality of equine oocytes before and after maturation in vitro. The mare was an attractive animal model because the horse is a monovulatory species in which fertility decreases markedly with advancing maternal age [32], oocytes can be obtained from slaughtered animals, and the time interval to reproductive senescence is more comparable with women than that of laboratory species such as mice.
using standard parameters for equine dental eruption and wear [33,34]. Animals for which the age could not be estimated reliably were excluded from the study. The ovaries from young (<12 years) and aged mares (12 years) were transported separately to the laboratory in thermos flasks at 30 C. On arrival at the laboratory within 4 hours of slaughter, the ovaries were washed with tap water (30 C), freed of extraneous tissue, and maintained at 30 C in 0.9% (wt/vol) saline supplemented with 0.1% (vol/vol) penicillin and streptomycin (Gibco BRL, Life Technologies, Paisley, UK). Cumulus oocyte complexes (COCs) were then recovered by aspirating, flushing, and scraping the contents of 5to 30-mm follicles [35]. For each mare age group, recovered oocytes were divided randomly into 2 groups. In the first group (not-IVM), 127 oocytes from young mares and 106 from aged mares were immediately denuded of their cumulus cells by vortexing for 4 minutes in calcium- and magnesium-free Earles’ balanced salt solution containing 0.25% (vol/vol) trypsin–EDTA (Gibco BRL, Life Technologies). After vortexing, oocytes were examined under an inverted stereomicroscope to confirm the absence of cumulus cells; denuded oocytes were stored for further analysis, and incompletely denuded oocytes were discarded. In the second group (in vitro matured: IVM), 114 COCs from young mares and 154 from aged mares were matured in vitro in maturation medium consisting of M199 tissue culture medium supplemented with 10% FCS, 0.01 units/mL porcine FSH, and 0.01 units/mL equine LH (SigmaAldrich Chemicals BV) for 30 hours at 38.7 C in a humidified atmosphere of 5% CO2 in air [35], before denuding and storage. Next, IVM oocytes were examined with an inverted stereomicroscope to confirm that cumulus removal was complete and determine whether a first polar body was visible. IVM oocytes were then divided into those that had reached the MII stage, i.e., had a visible first polar body (MII) and those that had not (not MII). The oocytes were then washed three times in phosphate buffered saline (PBS) containing 0.1% (wt/vol) polyvinyl alcohol (Sigma-Aldrich Chemicals BV) and three times in (Tris-EDTA) TE, consisting of 10 mM Tris (MP Biomedicals Inc., Eschwege, Germany) and 0.1 mM EDTA (BDH Ltd., Poole, UK) in double-distilled water, before being placed individually in eppendorf tubes in 5 mL of TE and stored at 20 C until analysis.
2. Materials and methods
Two series of reference samples were produced to ensure that values obtained in different quantitative polymerase chain reaction (QPCR) plates were comparable. First, a DNA sequence of 428 bp, spanning the fragment used for QPCR, was amplified (Table 1, #1) and purified using the Qiaquick Purification Kit (Qiagen, Venlo, The
2.1. Collection and culture of cumulus oocyte complexes Ovaries were recovered from 268 horse (Equus caballus) mares immediately after slaughter. Age was estimated
2.2. DNA extraction Oocytes were lysed by adding 5 mL of an alkaline lysis buffer (200 mM KOH and 50 mM dithiothreitol), incubating them at 65 C for 10 minutes and vortexing. After addition of 5 mL of neutralization buffer (0.9 M Tris–HCl, 0.3 M KCl, and 0.2 M HCl), the lysates were further diluted to a total volume of 150 mL and stored at 20 C. 2.3. Preparation of reference samples and internal controls
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Netherlands). PCR copy number was determined by measuring absorbance at 260 nm. Although after QPCR, serial dilutions gave a linear standard curve, results for the lower concentrations became less reliable after storage at 20 C. Therefore, a stock of lysed oocytes enriched with purified genomic DNA isolated from peripheral equine lymphocytes was prepared. Ten-fold serial dilutions were amplified by QPCR and calibrated using freshly prepared dilutions of the PCR product described above and analyzed on the same microtiter plate. This reference series was reproducible after a period of storage at 20 C and was aliquoted, stored at 20 C, and subsequently used on all microtitre plates. Internal assay controls were prepared by pooling a number of lysed denuded equine oocytes and storing aliquots at 20 C either undiluted (IC high) or after 10-fold dilution (IC low). 2.4. Determining oocyte mtDNA copy number by QPCR QPCR was performed in 96-well plates using a real-time PCR detection system (MyIQ Single-Color Real-Time PCR Detection System; Bio-Rad Laboratories, Veenendaal, The Netherlands). To minimize plate-dependent effects, 96-well plates included triplicate samples of the 2 negative controls (water), the 2 internal controls (IC high and IC low), the 6 reference samples, and 11 not-IVM oocytes and 11 IVM oocytes. The location of the not-IVM (n ¼ 233) and IVM samples (n ¼ 268) was alternated in successive plates. PCR primers (Table 1, #2) were designed for the quantification of oocyte mtDNA content. The PCR reaction mixture contained 1 mL of 1:150 diluted oocyte lysate, 2.5 mL GeneAmp 10 PCR Gold Buffer (Applied Biosystems, Nieuwerkerk a/s IJssel, The Netherlands), 2 mM MgCl2 (Applied Biosystems), 10 pM fluorescein (Bio-Rad Laboratories), 5 mL of a 1:20,000 dilution of SyberGreen (Bio Wittaker, Inc., Walkersville, MD, USA), 0.2 mM of each dNTP (Promega Benelux BV, Leiden, The Netherlands), 0.4 mM of each primer (Isogen Bioscience BV, Maarsen, The Netherlands), and 0.625 IU Amplitaq Gold (Applied Biosystems) made up to a total volume of 25 mL with double distilled water. After denaturation by heating to 95 C for 5 minutes, 40 PCR cycles consisting of incubation at 95 C for 20 seconds, 67.7 C for 30 seconds, and 72 C for 30 seconds were followed by a further 5 minutes at 72 C. The PCR product was checked by melting curve analysis, gel electrophoresis, and amplicon sequencing. To determine mtDNA copy number in each oocyte, sample threshold cycles were plotted against DNA concentration. The internal control samples (IC low and IC high) were used to monitor the intra- and interassay coefficients of variation, which were 13.1% and 26.7%, respectively, similar to in previous reports [36,37].
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2.5. Assessment of mitochondrial morphology Oocyte mitochondrial morphology was examined by transmission electron microscopy. Directly after collection or IVM, COCs were fixed for 35 to 65 hours in Karnovsky fixative (2% paraformaldehyde, 2.5% glutaraldehyde, .08 M Na-cacodylate buffer [pH 7.4], 0.25 mM CaCl2, and 0.5 mM MgCl2). After fixation, the COCs were washed for 10 minutes in 0.1 M Na-cacodylate buffer (pH 7.4) and immersed for 2 periods of 1 hour in 2% osmium tetroxide in the same buffer. Subsequently, the samples were stained for 1 hour with 2% aqueous uranyl, before being dehydrated by passage through a series of acetone solutions, and embedded in Durcupan ACM (Fluka, Buchs, Switzerland). At intervals of 50 mm through each block, exploratory semi-thin sections (1 mm) were cut using a glass knife on a Reichert Ultracut S microtome (Leica Microsystems B.V., Rijswijk, The Netherlands). These sections were stained with toluidine blue and screened for COC presence by light microscopy. If the COC was detected, further semi-thin sections were cut at 10 mm intervals until the maximum COC diameter was reached. Subsequently, ultrathin sections (50 nm) were cut with a diamond knife and mounted on single-slot formvar–carbon-coated copper grids. After staining with lead citrate for 2 minutes, the sections were examined and electron micrographs were taken at random locations within the equatorial plane of the oocyte using an electron microscope (Philips CM 10; Philips, Eindhoven, The Netherlands) at 80 kV and magnifications of approximately 3000 for overviews and 30,000 for detailed morphological analysis. Negatives were scanned with a Linoscan 1450 scanner (Heidelberger Druckmaschinen AG, Heidelberg, Germany) at 600 dpi resolution. The micrographs were subsequently used to examine and describe mitochondrial morphology, whereas mitochondrial size was measured by point hit analysis [38]. 2.6. Statistical analysis Statistical analysis was performed using SPSS 12.0.1 for Windows (SPSS Inc., Chicago, IL, USA). The influence of maternal age, oocyte maturation stage, and their interaction on the (natural logarithm of the) mtDNA copy number was examined by analysis of variance (ANOVA). A post hoc Bonferroni test was used to determine the source (age by maturation stage group) of any differences in mtDNA copy number. Differences in mitochondrial diameter between groups were also analyzed using ANOVA and a post hoc Bonferroni test. Normality of distribution for mitochondrial diameters and for the natural logarithms of mtDNA copy number was confirmed using quantile–quantile plots; in a normally distributed data set, the plot tends to a straight
Table 1 Primer pairs used for quantification of mtDNA in equine oocytes and for the production of a reference PCR product. ID
Forward primer (50 -30 )
Positiona
Reverse primer (50 -30 )
Positiona
Ta
Purpose
#1 #2
AAGAAAACCCCACAAAACTA CATGATGAAACTTCGGCTCCCT
14051 14273
GTGAATGAAGAGGCAGATAAAA TGAGTGACGGATGAGAAGGCAG
14478 14390
55 C 67.7 C
Reference PCR product mtDNA quantification
Abbreviations: PCR, polymerase chain reaction; Ta, annealing temperature. a bp position in equine mtDNA (GenBank entry: NC_001640).
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line. Differences between groups were considered statistically significant if P is less than 0.05. 3. Results 3.1. Oocyte mtDNA quantities In total, 501 oocytes were analyzed for mtDNA quantity (Fig. 1). The mean mtDNA copy number per oocyte was 1.13 106, but there was considerable variation between oocytes (range: 4.68 103 to 3.82 106; SEM ¼ 3.53 104). No significant differences in mtDNA copy number were found between oocytes recovered from young mares, irrespective of whether they had been processed immediately after recovery (not-IVM: 1.15 106) or matured in vitro and either reached MII (IVM/MII: 1.38 106) or not (IVM/notMII: 1.46 106). Oocytes recovered from aged mares and not subjected to IVM contained similar mtDNA copy numbers (1.41 106) as oocytes from young mares. By contrast, mtDNA copy number in oocytes recovered from aged mares and incubated in vitro (IVM/non-MII: 0.65 106; IVM/MII: 0.78 106) were lower than in all other groups (P < 0.001). 3.2. Oocyte mitochondrial morphology Oocyte mitochondrial morphology was examined in 20 oocytes (5 per group for young vs. aged combined with notIVM vs. IVM). Mitochondria in oocytes from young mares and those from aged mares’ oocytes that had not been subjected to IVM were comparable in size and morphology (Fig. 2A–C) with intact mitochondrial membranes, clearly appreciable transverse cristae, and a mitochondrial matrix of moderate electron density. By contrast, mitochondria in IVM oocytes from aged mares appeared swollen (Fig. 2D). The apparent enlargement was confirmed by hit point analysis, which revealed a larger mean surface area than for mitochondria in not-IVM oocytes from both young and aged mares (Table 2). In addition, the internal architecture
Fig. 1. Mean (SEM) mtDNA copy number in oocytes recovered from young (<12 years) and aged (12 years) mares as determined by quantitative polymerase chain reaction. Within age classes, the oocytes are divided into three groups: (i) those that were not matured in vitro, (ii) those that were matured in vitro but did not reach the M-II stage, and (iii) those that were matured in vitro and did reach M-II. The number of oocytes per group is indicated on the column. Groups with different superscript letters had significantly different mtDNA copy numbers (P < 0.001).
of most of the mitochondria was altered. Of the 37 mitochondria examined in the five aged mare IVM oocytes, 21 exhibited severe disruption with no visible cristae remaining and a relatively low mitochondrial matrix density (Fig. 2D), 11 were moderately affected and retained some cristae, whereas only 5 mitochondria retained a grossly normal morphology. Overall, in 3 of the 5 oocytes from aged mares, all the mitochondria examined were severely or moderately disrupted, whereas the remaining two oocytes contained either normal or only moderately disrupted mitochondria. 4. Discussion We used QPCR and transmission electron microscopy to demonstrate that maternal age predisposes to mitochondrial degeneration in equine oocytes. In fact, oocyte mitochondrial degeneration manifested only after IVM of the oocytes from aged mares and was characterized by swelling of mitochondria and a reduction in mtDNA copy number. These results are reminiscent of previous reports that oocytes from aged mice contain fewer mitochondria with structural alterations [31] and that, after oocyte maturation, the mitochondria are more sensitive to experimentally induced damage than those in MII oocytes from younger animals [39]. The increased susceptibility to oocyte mitochondrial degeneration with increasing maternal age may include a role for ROS produced by the mitochondria during oxidative phosphorylation (for review, see Eichenlaub-Ritter et al. [23]). ROS can damage various cellular components including proteins, lipid membranes, and DNA, and the damage can accumulate during an individual’s lifetime [40]. On the basis of the current findings and previous studies in aged women [41] and mice [39], it appears that mitochondria in oocytes in the resting (nonrecruited) pool may accumulate ROS-induced damage during a female’s lifetime. This damage may be subtle or “subclinical” and only become apparent when the metabolic demands of the oocyte increase with the onset of follicle growth and oocyte maturation [42–45]. The damaged mitochondria may not be able to meet the increased demands for energy, leading the oocyte to enter a vicious circle of spiraling ROS production and mitochondrial damage that culminates in acute severe oxidative stress characterized by swelling, rupture, and loss of the mitochondria [46–48]. Alternatively or additionally, it has been proposed that, as a result of aging, oocytes suffer a decrease in the content of critical anti-apoptotic proteins (such as members of the Bcl-2 family) by degradation or inactivation, rendering the oocytes more susceptible to apoptosis and turning the calcium oscillations triggered by sperm penetration and required for fertilization into apoptosis-triggering signals [49]. In the present study, mtDNA copy number did not differ between not-IVM and IVM oocytes within the young mare group. A similar finding was reported for human oocytes [50], and it suggests that mitochondrial replication in the female germ cell line [51] is completed before the oocyte embarks on its final maturation. Furthermore, because ATP content does not differ between in vitro matured mouse
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Fig. 2. Transmission electron micrographs of mitochondria in equine oocytes. Mitochondria in young mare oocytes that were not (A) or were (B) matured in vitro. The mitochondria have intact membranes and transverse cristae. (C) A mitochondrion in an aged mare’s oocyte that had not been matured in vitro. Its appearance is comparable to that of mitochondria in the oocytes of young mares. (D) A mitochondrion in an aged mare’s oocyte that had been matured in vitro. The mitochondrion is grossly swollen, has lost its transverse cristae, and has a low mitochondrial matrix density (scale bar ¼ 200 nm).
oocytes at the germinal vesicle breakdown, MI, and MII stages [52], it appears that neither an increase in mitochondrial number nor activity is required for final oocyte maturation. The significance of the predisposition to mitochondrial damage and loss in the oocytes of aged mares relates to the likely consequences for oocyte developmental competence, i.e., the ability to yield a viable embryo/pregnancy. For example, low mtDNA copy numbers have been associated with reduced oocyte fertilizability in women [36] and sows [53], whereas severe mitochondrial disfunction has been shown to impair normal germinal vesicle breakdown, Table 2 Relative size (mean SEM) of mitochondria in oocytes from aged (12 years) or young (<12 years) horse mares that were either processed immediately or after IVM. Maternal age
Oocyte maturation stage
Mitochondrial sizea
<12 <12 12 12
Not-IVM IVM Not-IVM IVM
40.6 61.3 53.1 99.7
x,y
y y y y
0.7x (n 3.4y (n 1.9x (n 5.9y (n
¼ ¼ ¼ ¼
25) 37) 36) 37)
Indicates values that do not differ significantly (P < 0.05). Abbreviation: COC, cumulus oocyte complex. a Relative mitochondrial size was determined by hit point analysis (n ¼ 5 COCs/group; total number of mitochondria measured per group are indicated in parentheses).
meiotic spindle formation, and chromosome disjunction [54,55] and to increase the incidence of maternally inherited chromosomal abnormalities [23,56]. Low energy production [52] and mitochondrial damage [39,56] in preovulatory oocytes have also been associated with poor embryo development and increased pre-blastulation embryonic death. Suggestively, the incidence of meiotic spindle aberrations [10,56,57,58] and chromosomal abnormalities in both oocytes [11,12] and blastocysts [59] are higher in aged than younger females. Critically, an embryo’s mitochondria are almost exclusively maternally inherited (i.e., oocyte derived) [60–63], and postfertilization, embryonic mitochondrial replication does not appear to be initiated until relatively late, the early gastrula stage in mice [64–67] and the expanded blastocyst stage in pigs [68]. Larsson et al. [66] illustrated this concept by disrupting Tfam, a gene critical for mitochondrial replication, in mice; although embryos homozygous for the disrupted gene reached gastrulation, they died soon afterward. Given the late onset of mitochondrial replication, early embryonic cell divisions must involve partitioning of a finite number of oocyte-derived mitochondria over an increasing number of blastomeres. Furthermore, the energy requirements of early embryonic development must be met by this increasingly subdivided pool of mitochondria. The decrease in mitochondrial quantity detected in
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aged mare oocytes after IVM in the present study could result in insufficient mitochondria per embryonic cell to support normal development and, thereby, predispose to embryonic death. Aged mares do, indeed, suffer a higher incidence of early embryonic death than younger animals [32]. Reduction of the number of mitochondria per cell may also explain aberrant chromosomal segregation during mitosis [23,55], where aneuploid embryonic cells have been described for many mammalian species, including man [69,70] and the horse [71]. In conclusion, the present study demonstrates that maternal aging predisposes to mitochondrial damage and loss during IVM of equine oocytes. It is postulated that when an aged oocyte is reactivated, damage to the mitochondrial membranes triggers a ROS-associated cascade leading to swelling and loss of mitochondrion. A predisposition to mitochondrial damage may contribute to the age-dependent decrease in fertility in mares. Acknowledgments The authors would like to thank A. Klarenbeek (Department of Equine Sciences) for helping collect ovaries and estimate mare ages, L. van Weeren and A. Ultee (Department of Biochemistry and Cell Biology) for their assistance with transmission electron microscopy, Professor W. Stoorvogel (Department of Biochemistry and Cell Biology) and Dr. G. Posthuma (University Medical Center) for help with the point hit analysis, and J. van den Broek for assistance with statistical analysis. References [1] Heffner LJ. Advanced maternal agedhow old is too old? N Engl J Med 2004;351:1927–9. [2] Menken J, Trussell J, Larsen U. Age and infertility. Science 1986;233: 1389–94. [3] Nybo Andersen AM, Wohlfahrt J, Christens P, Olsen J, Melbye M. Maternal age and fetal loss: population based register linkage study. BMJ 2000;320:1708–12. [4] te Velde ER, Pearson PL. The variability of female reproductive ageing. Hum Reprod Update 2002;8:141–54. [5] Templeton A, Morris JK, Parslow W. Factors that affect outcome of in-vitro fertilisation treatment. Lancet 1996;348:1402–6. [6] Faddy MJ, Gosden RG. A mathematical model of follicle dynamics in the human ovary. Hum Reprod 1995;10:770–5. [7] Faddy MJ, Gosden RG, Gougeon A, Richardson SJ, Nelson JF. Accelerated disappearance of ovarian follicles in mid-life: implications for forecasting menopause. Hum Reprod 1992;7:1342–6. [8] Sauer MV. Infertility and early pregnancy loss is largely due to oocyte aging, not uterine senescence, as demonstrated by oocyte donation. Ann N Y Acad Sci 1997;828:166–74. [9] Steuerwald N, Cohen J, Herrera RJ, Sandalinas M, Brenner CA. Association between spindle assembly checkpoint expression and maternal age in human oocytes. Mol Hum Reprod 2001;7:49–55. [10] Battaglia DE, Goodwin P, Klein NA, Soules MR. Influence of maternal age on meiotic spindle assembly in oocytes from naturally cycling women. Hum Reprod 1996;1:2217–22. [11] Pellestor F, Andreo B, Arnal F, Humeau C, Demaille J. Maternal aging and chromosomal abnormalities: new data drawn from in vitro unfertilized human oocytes. Hum Genet 2003;112:195–203. [12] Pellestor F, Anahory T, Hamamah S. Effect of maternal age on the frequency of cytogenetic abnormalities in human oocytes. Cytogenet Genome Res 2005;111:206–12. [13] Carnevale EM, Ginther OJ. Defective oocytes as a cause of subfertility in old mares. Biol Reprod Monogr Ser 1995;1:209–14. [14] Nagley P, Wei YH. Ageing and mammalian mitochondrial genetics. Trends Genet 1998;14:513–7.
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Advancing maternal age predisposes to mitochondrial damage and loss during maturation of equine oocytes in vitro B.P.B. Rambags 1, D.C.J. van Boxtel, T. Tharasanit 2, J.A. Lenstra, B. Colenbrander, T.A.E. Stout* Department of Equine Sciences, Faculty of Veterinary Medicine, Utrecht University, Yalelaan 112, 3584 CM Utrecht, The Netherlands
a r t i c l e i n f o
a b s t r a c t
Article history: Received 16 August 2013 Received in revised form 8 January 2014 Accepted 16 January 2014
In many mammalian species, reproductive success decreases with maternal age. One proposed contributor to this age-related decrease in fertility is a reduction in the quantity or functionality of mitochondria in oocytes. This study examined whether maternal age or (in vitro maturation). IVM affect the quantity of mitochondria in equine oocytes. Oocytes were collected from the ovaries of slaughtered mares categorized as young (<12 years) or aged (12 years) and either denuded and prepared for analysis immediately (not-IVM) or matured in vitro for 30 hours before preparation (IVM). The mean oocyte mitochondrial DNA copy number was estimated by quantitative polymerase chain reaction and found to be significantly lower in oocytes from aged mares and that had been subjected to IVM than in any other group. Transmission electron microscopy demonstrated that mitochondria in aged mare oocytes subjected to IVM experienced significantly more swelling and loss of cristae than in other groups. We conclude that maternal aging is associated with a heightened susceptibility to mitochondrial damage and loss in equine oocytes, which manifests during IVM. This predisposition to mitochondrial degeneration probably contributes to reduced fertility in aged mares. Ó 2014 Elsevier Inc. All rights reserved.
Keywords: Horse Maternal age Mitochondrion Oocyte In vitro maturation
1. Introduction In many mammalian species, there is a threshold maternal age beyond which reproductive success decreases markedly [1]. In women, fertility declines after 35 years [2], largely as a result of an increase in the incidence of spontaneous abortion [3]. The decline culminates in the cessation of fertility in most women at around 41 years, even though the menopause does not onset until approximately 51 years [4]. Similarly, the likelihood of a live birth per embryo * Corresponding author. Tel.: þ31 30 2531350; fax: þ31 30 2537970. E-mail address: [email protected] (T.A.E. Stout). 1 Present address: uQinisa, Houten, The Netherlands. 2 Present address: Department of Obstetrics, Faculty of Veterinary Science, Chulalongkorn University, Bangkok, Thailand. 0093-691X/$ – see front matter Ó 2014 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.theriogenology.2014.01.020
transferred in human IVF programs decreases from 24% at maternal ages below 30 years to 8% at 42 years and 4% at 45 years or more [5]. The underlying causes of reproductive senescence have been proposed to include a decrease in the size of the resting follicle pool, a decrease in oocyte quality, and impaired endometrial receptivity. The size of the resting follicle pool certainly affects fertility in women, especially just before the menopause [6,7]. However, a reduction in oocyte number does not explain why many women are cyclic but unable to conceive in the last 10 years before the menopause [4]. Similarly, because the negative effects of advanced maternal age on IVF success can largely be overcome using an oocyte donated by a younger woman (35 years), impaired endometrial receptivity seems to be a relatively minor contributor to the premenopausal decline in fertility [5,8]. Instead, the primary contributor to reduced
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fertility is thought to be reduced oocyte quality; previous studies have reported a relatively low expression of spindle assembly checkpoint mRNA [9] and high incidences of spindle aberrations [10] and chromosomal abnormalities [11,12] in oocytes from aged women. Similarly, oviductal transfer of oocytes from aged mares (20–26 years) produced significantly fewer embryonic vesicles (31%) in inseminated young mare recipients than transfer of oocytes from young (6–10 years) donors (92%) [13]. One postulated underlying cause of the age-dependent decrease in oocyte quality is a decline in mitochondrial function [14,15]. Mitochondria play several important metabolic roles in eukaryotic cells, including energy generation by oxidative phosphorylation, steroid production, b-oxidation, and calcium homeostasis. However, mitochondria are also implicated in processes associated with cell deterioration, such as the production of potentially harmful reactive oxygen species (ROS) [16,17]. Because mtDNA is located close to the site of ROS generation, it may be particularly at risk for accumulating oxidative damage. Moreover, mtDNA is more sensitive to mutagens than nuclear DNA because it lacks introns, protective histones [18], and DNA repair mechanisms. Indeed, the mutation rate of mtDNA is more than 10 times higher than that of nuclear DNA [18], and point mutations, deletions, and duplications have been reported to accumulate in mtDNA over time, particularly in slowly or nondividing, postmitotic tissues with high energy demands, such as brain and muscle [14,19–22]. Mammalian oocytes are also nondividing, postmitotic cells; however, their energy demands are modest because they arrest after entering meiosis and remain in a resting phase until reactivation during final follicle development before ovulation. Nevertheless, because the resumption of meiosis may occur many years later, oocytes may accumulate mitochondrial DNA damage with increasing maternal age (for review, see Eichenlaub-Ritter et al. [23]). However, studies that have examined a possible age-related decline in oocyte mtDNA quality have produced conflicting results [24–30]. Even so, it was recently reported that increasing maternal age is accompanied by a decrease in oocyte mtDNA quantity in mice [31]. In this respect, earlier studies may have been biased by using relatively small populations of oocytes obtained from aged women attending IVF clinics for fertility problems. The aim of the present study was to determine whether maternal age significantly affects either mitochondrial number, estimated via mtDNA copy number, or electron microscopic morphological quality of equine oocytes before and after maturation in vitro. The mare was an attractive animal model because the horse is a monovulatory species in which fertility decreases markedly with advancing maternal age [32], oocytes can be obtained from slaughtered animals, and the time interval to reproductive senescence is more comparable with women than that of laboratory species such as mice.
using standard parameters for equine dental eruption and wear [33,34]. Animals for which the age could not be estimated reliably were excluded from the study. The ovaries from young (<12 years) and aged mares (12 years) were transported separately to the laboratory in thermos flasks at 30 C. On arrival at the laboratory within 4 hours of slaughter, the ovaries were washed with tap water (30 C), freed of extraneous tissue, and maintained at 30 C in 0.9% (wt/vol) saline supplemented with 0.1% (vol/vol) penicillin and streptomycin (Gibco BRL, Life Technologies, Paisley, UK). Cumulus oocyte complexes (COCs) were then recovered by aspirating, flushing, and scraping the contents of 5to 30-mm follicles [35]. For each mare age group, recovered oocytes were divided randomly into 2 groups. In the first group (not-IVM), 127 oocytes from young mares and 106 from aged mares were immediately denuded of their cumulus cells by vortexing for 4 minutes in calcium- and magnesium-free Earles’ balanced salt solution containing 0.25% (vol/vol) trypsin–EDTA (Gibco BRL, Life Technologies). After vortexing, oocytes were examined under an inverted stereomicroscope to confirm the absence of cumulus cells; denuded oocytes were stored for further analysis, and incompletely denuded oocytes were discarded. In the second group (in vitro matured: IVM), 114 COCs from young mares and 154 from aged mares were matured in vitro in maturation medium consisting of M199 tissue culture medium supplemented with 10% FCS, 0.01 units/mL porcine FSH, and 0.01 units/mL equine LH (SigmaAldrich Chemicals BV) for 30 hours at 38.7 C in a humidified atmosphere of 5% CO2 in air [35], before denuding and storage. Next, IVM oocytes were examined with an inverted stereomicroscope to confirm that cumulus removal was complete and determine whether a first polar body was visible. IVM oocytes were then divided into those that had reached the MII stage, i.e., had a visible first polar body (MII) and those that had not (not MII). The oocytes were then washed three times in phosphate buffered saline (PBS) containing 0.1% (wt/vol) polyvinyl alcohol (Sigma-Aldrich Chemicals BV) and three times in (Tris-EDTA) TE, consisting of 10 mM Tris (MP Biomedicals Inc., Eschwege, Germany) and 0.1 mM EDTA (BDH Ltd., Poole, UK) in double-distilled water, before being placed individually in eppendorf tubes in 5 mL of TE and stored at 20 C until analysis.
2. Materials and methods
Two series of reference samples were produced to ensure that values obtained in different quantitative polymerase chain reaction (QPCR) plates were comparable. First, a DNA sequence of 428 bp, spanning the fragment used for QPCR, was amplified (Table 1, #1) and purified using the Qiaquick Purification Kit (Qiagen, Venlo, The
2.1. Collection and culture of cumulus oocyte complexes Ovaries were recovered from 268 horse (Equus caballus) mares immediately after slaughter. Age was estimated
2.2. DNA extraction Oocytes were lysed by adding 5 mL of an alkaline lysis buffer (200 mM KOH and 50 mM dithiothreitol), incubating them at 65 C for 10 minutes and vortexing. After addition of 5 mL of neutralization buffer (0.9 M Tris–HCl, 0.3 M KCl, and 0.2 M HCl), the lysates were further diluted to a total volume of 150 mL and stored at 20 C. 2.3. Preparation of reference samples and internal controls
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Netherlands). PCR copy number was determined by measuring absorbance at 260 nm. Although after QPCR, serial dilutions gave a linear standard curve, results for the lower concentrations became less reliable after storage at 20 C. Therefore, a stock of lysed oocytes enriched with purified genomic DNA isolated from peripheral equine lymphocytes was prepared. Ten-fold serial dilutions were amplified by QPCR and calibrated using freshly prepared dilutions of the PCR product described above and analyzed on the same microtiter plate. This reference series was reproducible after a period of storage at 20 C and was aliquoted, stored at 20 C, and subsequently used on all microtitre plates. Internal assay controls were prepared by pooling a number of lysed denuded equine oocytes and storing aliquots at 20 C either undiluted (IC high) or after 10-fold dilution (IC low). 2.4. Determining oocyte mtDNA copy number by QPCR QPCR was performed in 96-well plates using a real-time PCR detection system (MyIQ Single-Color Real-Time PCR Detection System; Bio-Rad Laboratories, Veenendaal, The Netherlands). To minimize plate-dependent effects, 96-well plates included triplicate samples of the 2 negative controls (water), the 2 internal controls (IC high and IC low), the 6 reference samples, and 11 not-IVM oocytes and 11 IVM oocytes. The location of the not-IVM (n ¼ 233) and IVM samples (n ¼ 268) was alternated in successive plates. PCR primers (Table 1, #2) were designed for the quantification of oocyte mtDNA content. The PCR reaction mixture contained 1 mL of 1:150 diluted oocyte lysate, 2.5 mL GeneAmp 10 PCR Gold Buffer (Applied Biosystems, Nieuwerkerk a/s IJssel, The Netherlands), 2 mM MgCl2 (Applied Biosystems), 10 pM fluorescein (Bio-Rad Laboratories), 5 mL of a 1:20,000 dilution of SyberGreen (Bio Wittaker, Inc., Walkersville, MD, USA), 0.2 mM of each dNTP (Promega Benelux BV, Leiden, The Netherlands), 0.4 mM of each primer (Isogen Bioscience BV, Maarsen, The Netherlands), and 0.625 IU Amplitaq Gold (Applied Biosystems) made up to a total volume of 25 mL with double distilled water. After denaturation by heating to 95 C for 5 minutes, 40 PCR cycles consisting of incubation at 95 C for 20 seconds, 67.7 C for 30 seconds, and 72 C for 30 seconds were followed by a further 5 minutes at 72 C. The PCR product was checked by melting curve analysis, gel electrophoresis, and amplicon sequencing. To determine mtDNA copy number in each oocyte, sample threshold cycles were plotted against DNA concentration. The internal control samples (IC low and IC high) were used to monitor the intra- and interassay coefficients of variation, which were 13.1% and 26.7%, respectively, similar to in previous reports [36,37].
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2.5. Assessment of mitochondrial morphology Oocyte mitochondrial morphology was examined by transmission electron microscopy. Directly after collection or IVM, COCs were fixed for 35 to 65 hours in Karnovsky fixative (2% paraformaldehyde, 2.5% glutaraldehyde, .08 M Na-cacodylate buffer [pH 7.4], 0.25 mM CaCl2, and 0.5 mM MgCl2). After fixation, the COCs were washed for 10 minutes in 0.1 M Na-cacodylate buffer (pH 7.4) and immersed for 2 periods of 1 hour in 2% osmium tetroxide in the same buffer. Subsequently, the samples were stained for 1 hour with 2% aqueous uranyl, before being dehydrated by passage through a series of acetone solutions, and embedded in Durcupan ACM (Fluka, Buchs, Switzerland). At intervals of 50 mm through each block, exploratory semi-thin sections (1 mm) were cut using a glass knife on a Reichert Ultracut S microtome (Leica Microsystems B.V., Rijswijk, The Netherlands). These sections were stained with toluidine blue and screened for COC presence by light microscopy. If the COC was detected, further semi-thin sections were cut at 10 mm intervals until the maximum COC diameter was reached. Subsequently, ultrathin sections (50 nm) were cut with a diamond knife and mounted on single-slot formvar–carbon-coated copper grids. After staining with lead citrate for 2 minutes, the sections were examined and electron micrographs were taken at random locations within the equatorial plane of the oocyte using an electron microscope (Philips CM 10; Philips, Eindhoven, The Netherlands) at 80 kV and magnifications of approximately 3000 for overviews and 30,000 for detailed morphological analysis. Negatives were scanned with a Linoscan 1450 scanner (Heidelberger Druckmaschinen AG, Heidelberg, Germany) at 600 dpi resolution. The micrographs were subsequently used to examine and describe mitochondrial morphology, whereas mitochondrial size was measured by point hit analysis [38]. 2.6. Statistical analysis Statistical analysis was performed using SPSS 12.0.1 for Windows (SPSS Inc., Chicago, IL, USA). The influence of maternal age, oocyte maturation stage, and their interaction on the (natural logarithm of the) mtDNA copy number was examined by analysis of variance (ANOVA). A post hoc Bonferroni test was used to determine the source (age by maturation stage group) of any differences in mtDNA copy number. Differences in mitochondrial diameter between groups were also analyzed using ANOVA and a post hoc Bonferroni test. Normality of distribution for mitochondrial diameters and for the natural logarithms of mtDNA copy number was confirmed using quantile–quantile plots; in a normally distributed data set, the plot tends to a straight
Table 1 Primer pairs used for quantification of mtDNA in equine oocytes and for the production of a reference PCR product. ID
Forward primer (50 -30 )
Positiona
Reverse primer (50 -30 )
Positiona
Ta
Purpose
#1 #2
AAGAAAACCCCACAAAACTA CATGATGAAACTTCGGCTCCCT
14051 14273
GTGAATGAAGAGGCAGATAAAA TGAGTGACGGATGAGAAGGCAG
14478 14390
55 C 67.7 C
Reference PCR product mtDNA quantification
Abbreviations: PCR, polymerase chain reaction; Ta, annealing temperature. a bp position in equine mtDNA (GenBank entry: NC_001640).
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line. Differences between groups were considered statistically significant if P is less than 0.05. 3. Results 3.1. Oocyte mtDNA quantities In total, 501 oocytes were analyzed for mtDNA quantity (Fig. 1). The mean mtDNA copy number per oocyte was 1.13 106, but there was considerable variation between oocytes (range: 4.68 103 to 3.82 106; SEM ¼ 3.53 104). No significant differences in mtDNA copy number were found between oocytes recovered from young mares, irrespective of whether they had been processed immediately after recovery (not-IVM: 1.15 106) or matured in vitro and either reached MII (IVM/MII: 1.38 106) or not (IVM/notMII: 1.46 106). Oocytes recovered from aged mares and not subjected to IVM contained similar mtDNA copy numbers (1.41 106) as oocytes from young mares. By contrast, mtDNA copy number in oocytes recovered from aged mares and incubated in vitro (IVM/non-MII: 0.65 106; IVM/MII: 0.78 106) were lower than in all other groups (P < 0.001). 3.2. Oocyte mitochondrial morphology Oocyte mitochondrial morphology was examined in 20 oocytes (5 per group for young vs. aged combined with notIVM vs. IVM). Mitochondria in oocytes from young mares and those from aged mares’ oocytes that had not been subjected to IVM were comparable in size and morphology (Fig. 2A–C) with intact mitochondrial membranes, clearly appreciable transverse cristae, and a mitochondrial matrix of moderate electron density. By contrast, mitochondria in IVM oocytes from aged mares appeared swollen (Fig. 2D). The apparent enlargement was confirmed by hit point analysis, which revealed a larger mean surface area than for mitochondria in not-IVM oocytes from both young and aged mares (Table 2). In addition, the internal architecture
Fig. 1. Mean (SEM) mtDNA copy number in oocytes recovered from young (<12 years) and aged (12 years) mares as determined by quantitative polymerase chain reaction. Within age classes, the oocytes are divided into three groups: (i) those that were not matured in vitro, (ii) those that were matured in vitro but did not reach the M-II stage, and (iii) those that were matured in vitro and did reach M-II. The number of oocytes per group is indicated on the column. Groups with different superscript letters had significantly different mtDNA copy numbers (P < 0.001).
of most of the mitochondria was altered. Of the 37 mitochondria examined in the five aged mare IVM oocytes, 21 exhibited severe disruption with no visible cristae remaining and a relatively low mitochondrial matrix density (Fig. 2D), 11 were moderately affected and retained some cristae, whereas only 5 mitochondria retained a grossly normal morphology. Overall, in 3 of the 5 oocytes from aged mares, all the mitochondria examined were severely or moderately disrupted, whereas the remaining two oocytes contained either normal or only moderately disrupted mitochondria. 4. Discussion We used QPCR and transmission electron microscopy to demonstrate that maternal age predisposes to mitochondrial degeneration in equine oocytes. In fact, oocyte mitochondrial degeneration manifested only after IVM of the oocytes from aged mares and was characterized by swelling of mitochondria and a reduction in mtDNA copy number. These results are reminiscent of previous reports that oocytes from aged mice contain fewer mitochondria with structural alterations [31] and that, after oocyte maturation, the mitochondria are more sensitive to experimentally induced damage than those in MII oocytes from younger animals [39]. The increased susceptibility to oocyte mitochondrial degeneration with increasing maternal age may include a role for ROS produced by the mitochondria during oxidative phosphorylation (for review, see Eichenlaub-Ritter et al. [23]). ROS can damage various cellular components including proteins, lipid membranes, and DNA, and the damage can accumulate during an individual’s lifetime [40]. On the basis of the current findings and previous studies in aged women [41] and mice [39], it appears that mitochondria in oocytes in the resting (nonrecruited) pool may accumulate ROS-induced damage during a female’s lifetime. This damage may be subtle or “subclinical” and only become apparent when the metabolic demands of the oocyte increase with the onset of follicle growth and oocyte maturation [42–45]. The damaged mitochondria may not be able to meet the increased demands for energy, leading the oocyte to enter a vicious circle of spiraling ROS production and mitochondrial damage that culminates in acute severe oxidative stress characterized by swelling, rupture, and loss of the mitochondria [46–48]. Alternatively or additionally, it has been proposed that, as a result of aging, oocytes suffer a decrease in the content of critical anti-apoptotic proteins (such as members of the Bcl-2 family) by degradation or inactivation, rendering the oocytes more susceptible to apoptosis and turning the calcium oscillations triggered by sperm penetration and required for fertilization into apoptosis-triggering signals [49]. In the present study, mtDNA copy number did not differ between not-IVM and IVM oocytes within the young mare group. A similar finding was reported for human oocytes [50], and it suggests that mitochondrial replication in the female germ cell line [51] is completed before the oocyte embarks on its final maturation. Furthermore, because ATP content does not differ between in vitro matured mouse
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Fig. 2. Transmission electron micrographs of mitochondria in equine oocytes. Mitochondria in young mare oocytes that were not (A) or were (B) matured in vitro. The mitochondria have intact membranes and transverse cristae. (C) A mitochondrion in an aged mare’s oocyte that had not been matured in vitro. Its appearance is comparable to that of mitochondria in the oocytes of young mares. (D) A mitochondrion in an aged mare’s oocyte that had been matured in vitro. The mitochondrion is grossly swollen, has lost its transverse cristae, and has a low mitochondrial matrix density (scale bar ¼ 200 nm).
oocytes at the germinal vesicle breakdown, MI, and MII stages [52], it appears that neither an increase in mitochondrial number nor activity is required for final oocyte maturation. The significance of the predisposition to mitochondrial damage and loss in the oocytes of aged mares relates to the likely consequences for oocyte developmental competence, i.e., the ability to yield a viable embryo/pregnancy. For example, low mtDNA copy numbers have been associated with reduced oocyte fertilizability in women [36] and sows [53], whereas severe mitochondrial disfunction has been shown to impair normal germinal vesicle breakdown, Table 2 Relative size (mean SEM) of mitochondria in oocytes from aged (12 years) or young (<12 years) horse mares that were either processed immediately or after IVM. Maternal age
Oocyte maturation stage
Mitochondrial sizea
<12 <12 12 12
Not-IVM IVM Not-IVM IVM
40.6 61.3 53.1 99.7
x,y
y y y y
0.7x (n 3.4y (n 1.9x (n 5.9y (n
¼ ¼ ¼ ¼
25) 37) 36) 37)
Indicates values that do not differ significantly (P < 0.05). Abbreviation: COC, cumulus oocyte complex. a Relative mitochondrial size was determined by hit point analysis (n ¼ 5 COCs/group; total number of mitochondria measured per group are indicated in parentheses).
meiotic spindle formation, and chromosome disjunction [54,55] and to increase the incidence of maternally inherited chromosomal abnormalities [23,56]. Low energy production [52] and mitochondrial damage [39,56] in preovulatory oocytes have also been associated with poor embryo development and increased pre-blastulation embryonic death. Suggestively, the incidence of meiotic spindle aberrations [10,56,57,58] and chromosomal abnormalities in both oocytes [11,12] and blastocysts [59] are higher in aged than younger females. Critically, an embryo’s mitochondria are almost exclusively maternally inherited (i.e., oocyte derived) [60–63], and postfertilization, embryonic mitochondrial replication does not appear to be initiated until relatively late, the early gastrula stage in mice [64–67] and the expanded blastocyst stage in pigs [68]. Larsson et al. [66] illustrated this concept by disrupting Tfam, a gene critical for mitochondrial replication, in mice; although embryos homozygous for the disrupted gene reached gastrulation, they died soon afterward. Given the late onset of mitochondrial replication, early embryonic cell divisions must involve partitioning of a finite number of oocyte-derived mitochondria over an increasing number of blastomeres. Furthermore, the energy requirements of early embryonic development must be met by this increasingly subdivided pool of mitochondria. The decrease in mitochondrial quantity detected in
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aged mare oocytes after IVM in the present study could result in insufficient mitochondria per embryonic cell to support normal development and, thereby, predispose to embryonic death. Aged mares do, indeed, suffer a higher incidence of early embryonic death than younger animals [32]. Reduction of the number of mitochondria per cell may also explain aberrant chromosomal segregation during mitosis [23,55], where aneuploid embryonic cells have been described for many mammalian species, including man [69,70] and the horse [71]. In conclusion, the present study demonstrates that maternal aging predisposes to mitochondrial damage and loss during IVM of equine oocytes. It is postulated that when an aged oocyte is reactivated, damage to the mitochondrial membranes triggers a ROS-associated cascade leading to swelling and loss of mitochondrion. A predisposition to mitochondrial damage may contribute to the age-dependent decrease in fertility in mares. Acknowledgments The authors would like to thank A. Klarenbeek (Department of Equine Sciences) for helping collect ovaries and estimate mare ages, L. van Weeren and A. Ultee (Department of Biochemistry and Cell Biology) for their assistance with transmission electron microscopy, Professor W. Stoorvogel (Department of Biochemistry and Cell Biology) and Dr. G. Posthuma (University Medical Center) for help with the point hit analysis, and J. van den Broek for assistance with statistical analysis. References [1] Heffner LJ. Advanced maternal agedhow old is too old? N Engl J Med 2004;351:1927–9. [2] Menken J, Trussell J, Larsen U. Age and infertility. Science 1986;233: 1389–94. [3] Nybo Andersen AM, Wohlfahrt J, Christens P, Olsen J, Melbye M. Maternal age and fetal loss: population based register linkage study. BMJ 2000;320:1708–12. [4] te Velde ER, Pearson PL. The variability of female reproductive ageing. Hum Reprod Update 2002;8:141–54. [5] Templeton A, Morris JK, Parslow W. Factors that affect outcome of in-vitro fertilisation treatment. Lancet 1996;348:1402–6. [6] Faddy MJ, Gosden RG. A mathematical model of follicle dynamics in the human ovary. Hum Reprod 1995;10:770–5. [7] Faddy MJ, Gosden RG, Gougeon A, Richardson SJ, Nelson JF. Accelerated disappearance of ovarian follicles in mid-life: implications for forecasting menopause. Hum Reprod 1992;7:1342–6. [8] Sauer MV. Infertility and early pregnancy loss is largely due to oocyte aging, not uterine senescence, as demonstrated by oocyte donation. Ann N Y Acad Sci 1997;828:166–74. [9] Steuerwald N, Cohen J, Herrera RJ, Sandalinas M, Brenner CA. Association between spindle assembly checkpoint expression and maternal age in human oocytes. Mol Hum Reprod 2001;7:49–55. [10] Battaglia DE, Goodwin P, Klein NA, Soules MR. Influence of maternal age on meiotic spindle assembly in oocytes from naturally cycling women. Hum Reprod 1996;1:2217–22. [11] Pellestor F, Andreo B, Arnal F, Humeau C, Demaille J. Maternal aging and chromosomal abnormalities: new data drawn from in vitro unfertilized human oocytes. Hum Genet 2003;112:195–203. [12] Pellestor F, Anahory T, Hamamah S. Effect of maternal age on the frequency of cytogenetic abnormalities in human oocytes. Cytogenet Genome Res 2005;111:206–12. [13] Carnevale EM, Ginther OJ. Defective oocytes as a cause of subfertility in old mares. Biol Reprod Monogr Ser 1995;1:209–14. [14] Nagley P, Wei YH. Ageing and mammalian mitochondrial genetics. Trends Genet 1998;14:513–7.
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