Differential effects of repeated ovarian stimulation on cytoplasmic and spindle organization in metaphase II mouse oocytes matured in vivo and in vitro

Jonathan Van Blerkom1,2,3 and Patrick Davis1,2

1 Department of Molecular, Cellular and Developmental Biology, University of Colorado, Boulder, CO 80309 and 2 Colorado Reproductive Endocrinology, Rose Medical Center, Denver, CO 80220, USA


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Acknowledgements
 References
 
The effects of four rounds of ovarian stimulation spaced 1–6 weeks apart on the normality of metaphase II (MII) spindle formation, chromosomal alignment and cytoplasmic organization were examined in intact ovulated mouse oocytes and at MII for oocytes obtained at the germinal vesicle stage from the same ovaries and matured in vitro. The terminal deoxynucleotidyl transferase-mediated dUDP nick-end labelling assay was used to identify DNA strand breaks in chromosomes, and histological studies of ovaries between and at each round of ovarian stimulation were performed. The results demonstrate a progressive and significant increase in the frequency of spindle defects with each round of ovarian stimulation, including those spaced weeks apart. Oocytes with spindle defects were also characterized by the occurrence of detached chromosomes and cytoplasmic asters. In contrast, in-vitro matured oocytes derived from the same ovaries were normal. No evidence of DNA strand breaks with repeated rounds of ovarian stimulation was detected in ovulated or in-vitro matured oocytes. The development and persistence of nodules of hypertrophied granulosa in regions where follicular growth occurs suggest that a progressively increasing proportion of oocytes in the ovulatory pathway may experience an intrafollicular milieu that has negative consequences for competence. The results are discussed with respect to ovarian and oocyte biological ageing and possible adverse implications for human oocyte competence with repeated hyperstimulation.

Key words: meiotic chromosomes/meiotic spindle/oocyte competence/ovarian stimulation


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Acknowledgements
 References
 
Ovarian hyperstimulation and ovulation induction permits the growth and development of supernumerary follicles and the ability to time the initiation of pre-ovulatory oocyte maturation. The success of clinical IVF has been largely attributed to the generation of such follicles and the ability to retrieve metaphase II (MII) oocytes at high frequency. In mammals such as mice, rats and hamsters, reduced fertility and pre- and post-implantation mortality have been indicated as consequences of a single round of ovarian stimulation using standard doses of gonadotrophins (Vanderhyden et al., 1986Go; Fossum et al., 1989Go; de Boer et al., 1991Go; Vogel and Spielmann, 1992Go; Ertzeid et al., 1993Go; McKiernan and Bavister, 1998Go). At higher doses, increased frequencies of oocyte aneuploidy (Mailhes et al., 1995Go), embryo mortality, fetal growth retardation and congenital abnormalities have been reported (Edgar et al., 1987Go; Sakai and Endo, 1987Go; Ma et al., 1997Go). Reduced viability with ovarian stimulation is often attributed to adverse `maternal' factors such as inadequate uterine synchrony or receptivity (Elmazar et al., 1989Go; Fossum et al., 1989Go; de Boer et al., 1991Go). The studies of Vogel and Spielmann (1992) indicate that ovarian stimulation may also be associated with chromosomal defects in the oocyte whose lethality is expressed during the preimplantation stages (Vogel and Spielmann, 1992Go).

In clinical IVF, it is common for women to have undergone several cycles of ovarian stimulation before pregnancy is achieved, or, if these attempts fail, to use oocytes obtained from women of known fertility. Outcome data from some studies indicate no significant change in pregnancy rates after as many as eight cycles of ovarian stimulation and IVF (Padilla and Garcia, 1989Go; Yovel et al., 1994Go; Dor et al., 1996Go). However, maternal age-related depletion of the ovarian reserve (Scott et al., 1995Go; Lass et al., 1997Go) that normally begins in women after about age 30–33 years (Navot et al., 1991Go; Hull et al., 1996Go; Lim and Tsakok, 1997Go; Stolwijk et al., 1997Go) is a significant factor in the quality and competence of oocytes and preimplantation embryos in both normal and stimulated cycles (Navot et al., 1991Go). For example, several recent studies have demonstrated that a diminished ovarian reserve that is age-related (Freeman et al., 2000Go), or occurs prematurely in women with early age menopause (Kline et al., 2000Go), is associated with increased frequencies of trisomic pregnancies. In this respect, physiological changes in the ovary have been suggested to adversely influence oocyte competence (Freeman et al., 2000Go). In contrast, a rapid decline in fecundity after the second IVF cycle in women of different ages has been observed (Copperman et al., 1995Go) and these authors suggested that regardless of age, this number of cycles may represent an important threshold related to the probability of pregnancy occurring on subsequent attempts. The extent to which oocyte competence declines after repeated cycles of ovarian stimulation in some women undergoing infertility treatment, or is associated with adverse changes in ovarian function that are stimulation-related, is unknown.

Here, we examined whether repeated ovarian stimulation affected the `quality' of mouse oocytes at the nuclear and cytoplasmic levels, and whether the same types of defects associated with incompetent or compromised human oocytes occurred in gonadotrophin-treated mice. Reduced competence for the human oocyte has been associated with spindle malformations and chromosomal malalignment (Battagalia et al., 1996; Battaglia and Miller, 1997Go; Van Blerkom et al., 1997Go). Nuclear and cytoplasmic organization in intact, ovulated MII oocytes from each of four ovarian stimulation cycles spaced 1–6 weeks apart were compared with fully grown germinal vesicle (GV) stage oocytes recovered from the same ovaries and matured in vitro. Whether ovarian stimulation was associated with loss of DNA strand integrity was assessed by terminal deoxynucleotidyl transferase-mediated dUDP nick-end labelling (TUNEL assay) of chromosomes in MII stage oocytes. Histological analysis of ovaries was undertaken between and after rounds of ovarian stimulation. The results demonstrate that repeated rounds of ovarian stimulation, including those spaced weeks apart, have persistent effects on ovarian organization and are associated with an apparent reduction in ovarian reserve and a progressive increase in the frequency of intact, TUNEL-negative MII oocytes with spindle and cytoplasmic defects. In contrast, fully grown germinal vesicle (GV) stage oocytes obtained from the same ovaries after ovulation and matured in vitro appeared normal at MII.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Acknowledgements
 References
 
Ovarian stimulation schedule, in-vitro maturation, immunofluorescent analysis and TUNEL labelling
Cycles of ovarian stimulation were initiated in 5–6 week old ICR mice according to the standard protocol of 5 IU of pregnant mare serum gonadotrophin (PMSG; Sigma Chemical Co., St Louis, MO, USA) administered s.c. followed in 48 h by 5 IU of human chorionic gonadotrophin (HCG; Novarel, Ferring Pharmaceuticals Inc., Tarrytown, NY, USA). Four cycles of ovarian stimulation were performed with intervals of 1, 4 and 6 weeks between stimulation. For some studies, a fifth round was included at intervals of 1 and 4 weeks. Ovulated oocytes were collected from the oviductal ampulla at 14 h post HCG on each round, and fully grown GV stage oocytes, retrieved from the same stimulated ovaries, were matured in vitro as previously described (Van Blerkom and Runner, 1984Go), and examined at 14 h after the initiation of culture. Cumulus masses were disassociated with hyaluronidase (Sigma) and adherent coronal cells removed by repeated passage of oocytes through a glass micropipette. Representative ovaries from each stimulation cycle were fixed in formaldehyde and prepared for histological analysis. Ovaries were also removed for light microscopic histological analysis between stimulations at each interval.

Methods for chromosomal staining (Van Blerkom and Runner, 1984Go), tubulin immunofluoresence (Van Blerkom et al., 1995Go) and TUNEL labelling (Van Blerkom and Davis, 1998Go) used reageants and procedures previously described. Stained oocytes were examined first under conventional epifluorescence microscopy with appropriate narrow-band filter combinations and subsequently by scanning laser confocal microscopy (SLCM; Van Blerkom et al., 1995). For TUNEL, oocytes were prepared as whole mounts with partial zona pellucida thinning in acidic Tyrode's solution (15 s) in order to retain the first polar body (Van Blerkom et al., 2001Go). The sensitivity of the assay was confirmed by restaining after mechanical removal of the zona remnant, which often resulted in loss of the polar body (Van Blerkom and Davis, 1998Go). TUNEL-negative oocytes were exposed to DNase or hydrogen peroxide and restained (Van Blerkom et al., 2001Go).

Statistical Analysis
Frequencies of spindle malformation with each round of ovarian stimulation at each time interval were analysed statistically by the {chi}2-test. P < 0.05 was considered significant.


    Results
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Acknowledgements
 References
 
Normality of MII spindle in ovulated oocytes
The normality of chromosomal alignment and the position and shape of the MII spindle were assessed by anti-ß-tubulin immunofluorescence microscopy in oocytes recovered from the oviducts of mice at 14 h post HCG. For each round of stimulation, at least 100 ovulated oocytes were examined (see Table IGo). Figure 1AGo shows a normal MII oocyte stained with anti-ß-tubulin with fluorescence associated with the spindle (MII) and first polar body (PB1). Figure 1BGo shows the normal equatorial alignment of chromosomes (arrow) and associated MII spindle microtubules after dual staining with DAPI and anti-tubulin. The absence of organized cytoplasmic microtubules is a characteristic feature of normal MII mouse oocytes. On rounds 2–4, ~30% of oocytes with malformed spindles (e.g. Figure 1C–EGo) also contained chromosomes present as clusters (c, Figure 1FGo) and as detached, individual elements within the cytoplasm (arrow, Figure 1GGo). Figure 1C and DGo are representative epifluorescent and SLCM anti-tubulin images, respectively, of abnormal spindles in MII oocytes collected from oviducts after the second (arrow, Figure 1CGo) and third rounds of ovarian stimulation (arrow, Figure 1DGo) spaced 4 weeks apart. Typically, malformed spindles were either elongated (arrow, Figure 1C, DGo) or showed an asymmetric organization with higher densities of microtubules located at one pole (e.g. Figure 1MGo; small arrow, Figure 1EGo, round 4 with 4 week interval). Disorganized aggregates of cytoplasmic microtubules (asterisk, 1D) and microtubular asters (inset, Figure 1DGo; asterisks, Figure 1EGo) were detected in ~5% of MII mouse oocytes with malformed spindles after the second round of ovarian stimulation, and in virtually all abnormal oocytes on rounds 3 and 4.


View this table:
[in this window]
[in a new window]
 
Table I. Frequency (n, %) of spindle defects in intact metaphase II (MII) mouse oocytes matured in vivo or in vitro after each of four rounds of ovarian stimulation spaced either 1 or 4 weeks apart
 


View larger version (137K):
[in this window]
[in a new window]
 
Figure 1. DAPI fluorescence (B, F, G, L), anti-ß-tubulin immunofluorescence (A, C, D, inset D, E, J and M), and TUNEL staining of DNA (I, K, N) in mature mouse oocytes ovulated after repeated ovarian stimulation. (A, B, D, inset D) Scanning laser confocal microscopy; (C, E, FL) conventional epifluorescence illumination. (H) Representative light microscopic image of the type of intact, ovulated MII oocyte examined in this study. The asterisks in D and E indicate regions of disorganized microtubular staining and cytoplasmic asters, respectively. The equatorial alignment of chromosomes is shown by an arrow in B. The MII spindle is noted by an arrow in CE. The asterisk in N and the arrow in K indicate chromosomal TUNEL fluorescence after DNase treatment. TUNEL positive fluorescence in the first polar body (arrow, H) is noted by an arrow in I. c = chromosomes;MII = metaphase II spindle. Original magnifications: A, x350;B, x1000; C, x330; D, x370; D inset x1100; E, x310; F, x290; G, x360; H, x320; I, x320; J, x325; K, x310; L, x160;M, x310; N, x310.

 
The frequency of intact MII oocytes with spindle malformations detected at ovulation for each of four rounds of ovarian stimulation spaced 1 and 4 weeks apart is presented in Table IGo. The results indicate the following: (i) after round 1, the frequency of malformed spindles increased progressively with each round of ovarian stimulation, such that on round 4, over half of the normal-appearing MII oocytes (e.g. Figure 1HGo) were affected, and (ii) the frequency of spindle abnormalities with ovarian stimulations spaced 4–6 weeks apart was comparable to those spaced 1 week apart (results for 4 week interval shown in Table IGo). With rounds of ovarian stimulation spaced 1 week apart, the average number of intact MII stage oocytes recovered/mouse on rounds 1 (n = 8 mice), 2 (n = 10), 3 (n = 10) and 4 (n = 12) was 35, 35, 20 and 10 respectively. Very similar recovery rates were observed with ovarian stimulation spaced 4 or 6 weeks apart, with comparable numbers of mice used at each interval. For example, with a 4 week interval, the average number of intact oocytes recovered at ovulation/mouse on rounds 3 and 4 was 23 and 11 respectively. For some studies, a fifth round of ovarian stimulation was performed. However, these efforts were discontinued owing to too few intact MII oocytes recovered for meaningful analysis. To determine whether maternal age in the oldest animals we examined could contribute to the frequency of malformed spindles observed after ovarian stimulation, a single round of stimulation was performed on naturally cycling 20-28 week old mice (n = 16). The frequency of spindle abnormalities or detached chromosomes, or both, in 308 normal-appearing MII oocytes at 14 h after HCG administration was ~2% (7/308). This finding suggests the absence of a maternal age contribution to spindle anomalies at 28 weeks of life. The increased frequency of spindle malformation observed on rounds 2–4 was statistically significant at each of the indicated time intervals between stimulation [{chi}2(3) = 428, P < 0.001].

Normality of MII spindle formation after in-vitro maturation
The normality of spindle organization in MII oocytes that matured in vitro from the GV stage after being removed from stimulated ovaries on rounds 1–4 of ovarian stimulation was assessed as described above. For each round, at each interval, the frequency of maturation to MII after 14 h of culture was ~91% and a minimum of 50 oocytes was examined by immunofluorescence and DAPI staining at each round (see Table IGo). In contrast to their ovulated counterparts, virtually all in-vitro matured oocytes (Table IGo) showed normally formed spindles with normal chromosomal alignment and no detectable cytoplasmic asters (images comparable to Figure 1A, BGo).

TUNEL analysis of DNA integrity
The TUNEL assay was used to determine whether multiple rounds of ovarian stimulation may be associated with breaks in DNA strand integrity that cannot be detectable by conventional DNA fluorescence microscopy. Intact, ovulated MII oocytes from each of four rounds of ovarian stimulation, and in-vitro matured oocytes derived from the same ovaries were assayed by TUNEL. For each group and at each interval, comparable numbers of oocytes were examined (e.g. in-vivo matured, 1 week interval: round 1 = 149, round 2 = 86, round 3 = 82, round 4 = 59; in-vitro matured, 1 week interval: round 1 = 32, round 2 = 62, round 3 = 50, round 4 = 101). The only TUNEL fluorescence detected in MII oocytes (e.g. Figure 1HGo) that had matured either in vivo or in vitro was associated with the first polar body (arrow, Figure 1IGo, same oocyte as in 1HGo). The MII chromosomes in all TUNEL negative oocytes showed intense fluorescence after DNAase treatment or exposure to hydrogen peroxide (e.g. Figure 1N, KGo). DAPI labelling, tubulin immunofluorescence, and TUNEL staining were examined in 109 representative ovulated oocytes obtained on round 4 with 1 week intervals between stimulation. We considered this regimen to be the most likely to have pathological effects on oocytes, including loss of DNA strand integrity, and therefore oocytes (Figure 1H–NGo) with normal (n = 31) and abnormal spindles (n = 78) were analysed. None of the 109 selected oocytes showed chromosomal TUNEL staining, including 31 oocytes with detached chromosomes. Comparable results, albeit with fewer oocytes, were obtained on round 4 after 4 (n = 31) and 6 week intervals (n = 44). However, after DNase treatment, intense TUNEL fluorescence was observed in both spindle-associated and detached chromosomes in all MII oocytes. For example, chromosomes in the oocyte shown in Figures 1HGo were TUNEL negative (Figure 1IGo). The appearance of the MII spindle in this apparently normal oocyte as observed by immunofluorescence is shown in Figure 1JGo, and associated chromosomes identified by DAPI staining are indicated by an arrow in Figure 1LGo. After exposure to DNase and re-assayed by TUNEL, intense chromosomal fluorescence was observed (arrow, Figure 1KGo). Figure 1MGo shows a representative oocyte with an abnormal spindle derived from the same cohort as the normal oocyte presented in Figure 1HGo. The TUNEL- negative chromosomes are contained within the region denoted by an asterisk in Figure 1MGo. After DNase treatment, intense chromosomal TUNEL fluorescence was detected in this region (asterisk, Figure 1NGo). The detection of TUNEL-positive fluorescence in polar bodies in untreated oocytes, and for MII chromosomes after DNase treatment or hydrogen peroxide exposure, confirms the ability of this assay to detect DNA strand breaks under the conditions used.

Multiple ovarian stimulation and ovarian histology
Initial histological analysis involved a survey of selected sections taken at 200 µm intervals through ovaries on each round of ovarian stimulation. For specific follicles or regions, all sections taken at 20 µm intervals through these areas were examined. Representative sections of ovarian stimulation rounds 2–5 are presented in Figure 2A–DGo (4 week interval between stimulation) respectively. Very similar results were observed with intervals of 1 and 6 weeks between stimulation. For each series, both ovaries from four animals were examined. At the first cycle of ovarian stimulation, ovarian histology was comparable to the image shown for round 2 in Figure 2AGo. For rounds 1–4, histological characteristics of fully grown GV stage oocytes and associated granulosa cells in small antral follicles (round 2, Figure 2HGo; round 3, Figure 2IGo; round 4, Figure 2JGo) appeared normal. We assume that GV stage oocytes recovered for in-vitro maturation from such ovaries are derived from follicles of this type. A morphometric analysis of the distribution of follicles at different stages of development was beyond the scope of the present study. However, with respect to the occurrence of different follicle types at each round, the following characteristics were observed, regardless of whether ovarian stimulation was spaced 1, 4 or 6 weeks apart: (i) at round 2, the relative abundance of primary, secondary and tertiary (antral) follicles was similar to that observed at round 1 (arrows, Figure 2AGo), (ii) at round 3, antral follicles were less evident than previously, with primary-to-early antral follicles present in clusters between nodules of hypertrophied granulosa (asterisk, Figure 2BGo), (iii) at round 4, most of the follicles examined were classified as primary (arrowheads) or secondary (arrow, Figure 1CGo), and (iv) at round 5 (Figure 2DGo), the ovary consisted of an occasional primary or secondary follicle (Figure 2FGo). For rounds 4 and 5, some unruptured antral follicles were detected. However, virtually all contained either a fragmented oocyte with no evident corona or cumulus cell investment (Figure 2KGo), or appeared to be devoid of an oocyte or oocyte remnant, as determined by examination of consecutive sections (round 5, arrow, Figure 1DGo; asterisk, Figure 1EGo). For example, the fragmented oocyte shown in Figure 2KGo is presumed to be at the GV stage owing to a characteristic peri-nucleolar configuration of DNA detected by DAPI staining (Figure 2LGo) in the region indicated by an arrow in Figure 2KGo. Some unruptured follicles detected 14 h after HCG on rounds 3 and 4 contained GV stage oocytes closely opposed to the follicular wall and enclosed by an unexpanded granulosa cell layer (e.g. Figure 2GGo).



View larger version (186K):
[in this window]
[in a new window]
 
Figure 2. (A-D) Representative histological images of paraffin sections of mouse ovaries at ovarian stimulation rounds 2, 3, 4 and 5 respectively. (E-J) Higher magnification images of immature oocytes present after two to four rounds of ovarian stimulation.(K) Fragmented oocyte observed in an antral follicle; (L) DAPI-stained image of the region indicated by an arrow in K suggesting that this oocyte was at the GV stage. (F) One of the few remaining primary follicles observed on round 5 of ovarian stimulation, and unruptured follicles apparently devoid of an oocyte are indicated by an arrow and asterisk in D and E, respectively. Nodules of hypertrophied granulosa are indicated by an asterisk in A-D.A = antrum; GV = germinal vesicle stage oocyte. A, x27;B, x34; C, x32; D, x28; E, x92; F, x250, G, x170; H, x330;I, x310; J, x180; K, x350; L, x1600.

 
A hypertrophied and apparently luteinized granulosa was a common feature of each round, and after round 2, the ovary consisted of multiple, discrete nodules of virtually confluent granulosa, with follicles located between nodules or at the periphery (arrowheads, Figure 2B, CGo). The presence of nodules of hypertrophied granulosa persisted between rounds of ovarian stimulation, including those spaced 4–6 weeks apart. By light microscopy, both follicular thecal cells and regions of confluent granulosa cells appeared intact, nucleated, and with no apparent evidence of zones of cellular deterioration or necrosis within nodules.


    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Acknowledgements
 References
 
Ovarian stimulation is used to increase the numbers of oocytes ovulated and embryos produced in a variety of mammals, and is one of the foundations of current treatments of human infertility. As noted above, studies with animal systems have indicated that a single cycle of ovarian stimulation can have adverse effects on competence during early development, and may well have downstream effects on the normality of fetal growth and development. Whether developmental defects could result from chromosomal malsegregation during ovarian stimulation-induced meiotic maturation has been examined in the mouse system. Hansmann and El-Nahass (1979) compared oocytes obtained after spontaneous or hormonally induced ovulation and found no increase in the incidence of non-disjunction (<1%) after a single round of ovarian stimulation (Hansmann and El-Nahass, 1979Go). Golbus (1981) found no increase in oocyte aneuploidy after ovarian stimulation in young mice from inbred and random-bred strains (Golbus, 1981Go). In contrast, cytogenetic analysis of pronuclear stage mouse eggs after single cycle of ovarian stimulation by Vogel and Spielmann (1992) showed chromosomal aberrations largely confined to the female pronucleus, indicating developmental compromise prior to fertilization (Vogel and Spielmann, 1992Go). McKiernan and Bavister (1998) reported comparable rates of meiotic maturation and in-vitro development to the blastocyst stage for normally cycling and PMSG-stimulated (no HCG) hamsters, but a significant decline in implantation frequency after transfer to surrogates (McKiernan and Bavister, 1998Go). These investigators proposed that PMSG stimulation in hamster might generate lethal defects during meiotic maturation or early embryogenesis that are not manifest until the peri-implantation stages. Sengoku and Dukelow (1988) found comparable frequencies of aneuploidy for cleavage-stage hamster embryos produced in natural and PMSG-stimulated cycles, indicating that peri-implantation mortality may have an epigenetic origin (Sengoku and Dukelow, 1988Go).

Although findings from animal systems have been suggested to be clinically relevant in understanding potential reproductive toxicity of ovarian stimulation and ovulation induction (Tucker, 1996Go), comparatively few studies have examined whether the frequency of incompetent oocytes and embryos increases with repeated ovarian stimulation. It was reported (Lubbadeh et al., 1980Go) that with four cycles of ovarian stimulation in the dairy cow using either PMSG/HCG or FSH/LH, lengthening the time between stimulation by several weeks produced a consistent ovulatory response (number of ovulated oocytes). A similar finding was described for the mouse (Kanayama and Osada, 2000Go): there was a significant increase in morphologically abnormal oocytes after the third of five rounds of ovarian stimulation spaced 5 days apart, with ~33 and 50% of the recovered oocytes classified as `cleaved' after rounds 4 and 5 respectively. However, when rounds 4 and 5 were spaced 10 and 20 days apart, the frequency of cleaved oocytes was ~23 and 15% respectively, suggesting that `prolongation' of the interval between stimulations reduces the yield of abnormal oocytes. Whether cleaved oocytes obtained from the oviduct resulted from spontaneous parthenogenetic activation or fragmentation, or both, is unknown. However, for both cow and mouse, the occurrence of nuclear and cytoplasmic defects in ovulated oocytes was not determined.

Here, we examined cytoplasmic and spindle organization in morphologically normal MII oocytes ovulated after each of four rounds of ovarian stimulation. The results demonstrate that spindle malformation affects a significant and increasing proportion of intact oocytes ovulated between rounds 2 and 4, with no apparent reduction in incidence with intervals of up to 6 weeks between stimulation. Although MII oocytes with malformed spindles and scattered chromosomes were scored as abnormal, we did not determine whether any of DNA fluorescent structures were actually chromatids resulting from premature centromeric separation, a condition documented as a cause of aneuploidy in human oocytes (Angell et al., 1993Go). For the mouse, it was reported (Mailhes et al., 1998Go) that post-ovulatory ageing of mouse oocytes from stimulated ovaries was associated with a significant increase in the occurrence of premature centromeric separation, with frequencies of 2.8, 13.1 and 23.5% observed at 15, 20 and 25 h after ovulation induction respectively. The occurrence of cytoplasmic chromatids at 14 h in ovulated or in-vitro matured oocytes obtained after multiple rounds of ovarian stimulation is currently under investigation.

After round 2, cytoplasmic asters were detected in virtually all ovulated oocytes with spindle abnormalities. The occurrence of cytoplasmic asters, malformed spindles and detached chromosomes is very similar to the situation described for post-ovulatory mouse oocytes in vivo (Eichenlaub-Ritter et al., 1986Go), and for CBA/Ca strain mouse oocytes, which show an accelerated and rapid `ageing' in vitro that is characterized by the same defects (Eichenlaub-Ritter et al., 1988Go). An age-related increase in chromosomal malalignment in stimulated mouse oocytes has been observed (Saito et al., 1993Go); the authors suggested that this defect may be a underlying cause of reduced fertility. However, no increase in the frequency of malformed spindles or chromosomal malalignment in CBA/Ca oocytes ovulated by mice approaching the end of their reproductive life (>=9 months for the strain used) has been found (Eichenlaub-Ritter et al., 1988Go), indicating the absence of an evident ovarian age effect on competence. Here, analysis of ovulated oocytes in normally cycling mice given ovarian stimulation for the first time at 28 weeks showed no increase in the frequency of spindle or cytoplasmic abnormalities when compared with younger mice. This finding is consistent with the conclusion of Eichenlaub-Ritter et al. (1988) that maternal age alone in the mouse may not contribute significantly to spindle or chromosomal defects detected in oocytes that mature in vivo. However, our findings with young mice suggest the possibility that biological `ageing' of the oocyte cytoplasm during meiotic maturation may be accelerated in an increasing proportion of oocytes produced by repeated ovarian stimulation. The occurrence of these oocytes indicates an aetiology that seems to be unrelated to maternal age, but rather, one that may be influenced by conditions unique to the follicles in which the affected oocytes matured. For the human, however, it has been found that ~80% of MII oocytes aspirated from antral follicles of naturally cycling women between 40 and 45 years contained abnormally organized spindles with malaligned or detached chromosomes (Bataggalia et al., 1996). Unpublished studies of these oocytes also demonstrated the occurrence of cytoplasmic asters, structures that were not observed in the oocytes of younger women (20–25 years) which, in addition, rarely contained malformed spindles (D.Battagalia, personal communication).

It has been proposed (Battagalia and Miller, 1997) that an increased prevalence of age-related `intrinsic defects' (oocyte-specific) or adverse `extrinsic factors' (follicle-specific) may result from abnormal or asynchronous nuclear and cytoplasmic maturation, the consequence of which for women of advanced reproductive age may be high frequencies of oocyte aneuploidy. This finding raises the intriguing possibility that for the mouse, biological ageing of the oocyte cytoplasm is accelerated by ovarian stimulation, with the proportion of affected oocytes increasing with each round of stimulation, while for the human, a similar phenomenon may be age-related.

To determine whether defects observed at high frequency after repeated rounds of ovarian stimulation were oocyte-related or associated with maturation in vivo, we compared nuclear and cytoplasmic characteristics of MII oocytes derived from the same ovaries after as many as four rounds of ovarian stimulation, and after ovulation (14 h post HCG) or maturation from the GV stage in vitro (14 h). At each round, the frequency of maturation in vitro was similar, and spindle organization, chromosomal alignment, and cytoplasmic organization for virtually all in-vitro matured oocytes were indistinguishable from MII oocytes ovulated in natural cycles (Van Blerkom and Runner, 1984Go). The studies of Golbus indicate that in-vitro maturation of mouse oocytes is not associated with a significant increase in aneuploidy (Golbus, 1981Go). In comparison to values obtained from MII oocytes produced in natural cycles, or after one round of ovarian stimulation (1.1%), a slight increase in the frequency of aneuploidy (2.4%, hyperhaploidy only) after maturation in vitro of mouse oocytes obtained at the GV stage from young animals (6-8 weeks) was observed. Although the absence of evident spindle defects and detached chromosomes in the in-vitro matured oocytes we examined is suggestive of normalcy, whether multiple rounds of ovarian stimulation influence ploidy in these oocytes remains to be determined.

The TUNEL assay has been used to assess the extent of DNA fragmentation in mouse oocytes from stimulated ovaries with respect to maternal age (Fujino et al., 1996Go). The results indicated extensive chromosomal fragmentation in ~25% of oocytes ovulated by mice of comparatively advanced reproductive age (40–48 weeks). These investigators proposed that positive TUNEL staining was indicative of DNA strand breaks resulting from apoptotic-induced lesions in the chromosomes, and suggested this cell death process as a possible aetiology of chromosomally based reproductive failure in older women. Here, we asked whether multiple rounds of ovarian stimulation could be associated with loss of DNA strand integrity that may not be detectable by standard fluorescent-probe analysis which, in the present study, identified chromosomes that appeared to be intact. None of the oocytes that had maturated in vivo or in vitro showed DNA TUNEL staining; in particular, oocytes with abnormal spindle organization, as determined by anti-tubulin immunostaining, or detached chromosomes identified by DAPI staining, were TUNEL negative. The specificity of the assay was demonstrated by TUNEL-positive fluorescence in first polar body of TUNEL-negative oocytes, and for the same oocytes, TUNEL-positive fluorescence after DNase treatment or hydrogen peroxide exposure (Van Blerkom and Davis, 1998Go; Van Blerkom et al., 2001Go). This finding indicates that breaks in DNA strand integrity detectable by the TUNEL assay are not associated with repeated rounds of ovarian stimulation. Similar studies using the single-cell alkaline gel electrophoresis (`comet') assay to detect DNA fragmentation are in progress (Van Blerkom et al., 2001Go).

The apparent normality of in-vitro matured oocytes derived from the same ovaries where more than half of the ovulated oocytes were abnormal suggests that follicle-specific conditions during pre-ovulatory maturation may be a proximate cause of the defects identified in this study. For the human, clinical findings indicate that each fully grown pre-ovulatory follicle has unique properties, and that some properties are related to the competence of the corresponding oocyte. For example, follicle-specific differences in perifollicular blood flow indices have been correlated with chromosomal normality for the oocyte (Van Blerkom et al., 1997Go), embryo morphology during early cleavage (Huey et al., 1999Go), and outcome after transfer (Bahl et al., 1999Go; Coulam et al., 1999Go). Inadequate expansion of the perifollicular capillary bed, especially for women in the latter years of reproductive life, has been suggested to create conditions in the antral follicle that adversely effect oocyte physiology resulting in spindle malformation and aneuploidy (Gaulden, 1992Go).

If extrinsic factors differentially influence the normality of meiotic maturation in vivo, the mouse may offer a clinically relevant system in which to ask why only some MII oocytes are affected. Our histological findings demonstrate a persistent and aberrant ovarian morphology characterized by nodules of hypertrophied granulosa which become more abundant with repeated cycles of ovarian stimulation, even with cycles spaced weeks apart. Whether proximity of pre-antral or early antral follicles to these regions may be an important factor in determining the normality of pre-ovulatory maturation during subsequent natural or stimulated cycles is currently under investigation. In this regard, it was found that multiple rounds of ovarian stimulation in the rat altered hormonal homeostasis (Szoltys et al., 1994Go), and if other thecal and granulosa cell activities are also perturbed, these changes may have a direct effect on oocyte competence.

For the human, successful outcomes after multiple cycles of ovarian stimulation and ovulation induction for intrauterine insemination or IVF, or both, can imply that oocyte competence is not adversely influenced by the number or regimens of ovarian stimulation experienced prior to conception. This assumption may be incorrect if, relative to the total number of oocytes retrieved and fertilized, the proportion of competent oocytes and embryos declines with repeated cycles of stimulation, especially when issues related to ovarian reserve and maternal age are taken into consideration. Although extrapolations from findings obtained in the mouse to the human should be made with caution, one possibility is that similar cytoplasmic and spindle defects detected in the mature oocytes of both species reflect a common response of the maturing oocyte to adverse intrafollicular conditions. Whether or how disorders in ovarian structure and physiology associated with repeated stimulation may influence oocyte competence remains to be determined. However, an understanding of the relationship between oocyte competence and ovarian responses to stimulation in the mouse may provide insights into the origin of oocyte defects and the biology of ooplasmic ageing that that could be of clinical relevance in the diagnosis and treatment of human infertility.


    Acknowledgements
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Acknowledgements
 References
 
This work was supported by a grant from the National Institutes of Health (HD-31907).


    Notes
 
3 To whom correspondence should be addressed at: Department of Molecular, Cellular and Developmental Biology University of Colorado, Boulder, CO 80309, USA. E-mail: vanblerk{at}spot.colorado.edu Back


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Acknowledgements
 References
 
Angell, R., Xian, J. and Keith, J. (1993) Chromosome anomalies in human oocytes in relation to age. Hum. Reprod., 8, 1047–1054.[Abstract]

Bahl, P., Pugh, N. Chui, D. et al. (1999) The use of transvaginal power Doppler ultrasonography to evaluate the relationship between perifollicular vascularity and outcome in in-vitro fertilization treatment cycles. Hum. Reprod., 14, 939–945.[Abstract/Free Full Text]

Battaglia, D. and Miller, M. (1997) The aging oocyte. Endocrinologist, 7, 1–5.[ISI]

Battaglia, D., Goodwin, P., Klein, N. and Soules, M. (1996) Influence of maternal age on meiotic spindle assembly in oocytes from naturally cycling women. Hum. Reprod., 11, 2217–2222.[Abstract]

Copperman, A., Selick, C., Grunfeld, L. et al. (1995) Cumulative number and morphological score of embryos resulting in success: realistic expectations from in vitro fertilization-embryo transfer. Fertil. Steril., 64, 88–92.[ISI][Medline]

Coulam, C., Goodman, C. and Rinehart, J. (1999) Colour Doppler indices of follicular blood flow as predictors of pregnancy after in-vitro fertilization and embryo transfer. Hum. Reprod., 14, 1979–1982.[Abstract/Free Full Text]

De Boer, P., van der Hoeven, F., Wolters, E. and Mattheij, J. (1991) Embryo loss, blastomere development and chromosomal constitution after human chorionic gonadotropin-induced ovulation in mice and rats with regular cycles. Gynecol. Obstet. Invest., 32, 200–205.[ISI][Medline]

Dor, J., Seidman, D. and Ben-Shlomo, I. (1996) Cumulative pregnancy rate following in-vitro fertilization: the significance of age and infertility aetiology. Hum. Reprod., 11, 425–428.[Abstract]

Edgar, D., Whallet, K. and Mills, J. (1987) Effects of high dose and multiple-dose gonadotropin stimulation on mouse oocyte quality assessed by preimplantation development following in vitro fertilization. J. In Vitro Fertil. Embryo Transfer, 4, 273–276.[ISI][Medline]

Eichenlaub-Ritter, U., Chandely, A. and Gosden, R. (1986) Alterations to the microtubular cytoskeleton and increased disorder of chromosome alignment in spontaneously ovulated mouse oocytes aged in vivo: an immunofluorescence study. Chromosoma, 94, 337–345.[ISI][Medline]

Eichenlaub-Ritter, U., Chandley, A. and Gosden, R. (1988) The CBA mouse as a model for age-related aneuploidy in man: studies of oocyte maturation, spindle formation and chromosome alignment during meiosis. Chromosoma, 96, 220–226.[ISI][Medline]

Elmazar, M., Vogel, R. and Spielmann, H. (1989) Maternal factors influencing development of embryos from mice superovulated with gonadotropins. Reprod. Toxicol., 3, 135–138.[ISI][Medline]

Ertzeid, G. and Storeng, R. (1992) Adverse effects of gonadotropin treatment in pre-and postimplantation development in mice. J. Reprod. Fertil., 96, 649–655.[Abstract]

Ertzeid, G., Storeng, R. and Lyberg, T. (1993) Treatment with gonadotropins impaired implantation and fetal development in mice. J. Assist. Reprod. Genet., 10, 286–291.[ISI][Medline]

Fossum, G., Davidson, A. and Paulson, R. (1989) Ovarian hyperstimulation inhibits embryo implantation in the mouse. J. In Vitro Fertil. Embryo Transfer, 6, 7–10.[ISI][Medline]

Freeman, S., Yang, Q., Allran, K. et al. (2000) Women with a reduced ovarian complement may have an increased risk for a child with Down Syndrome. Am. J. Hum. Genet., 66, 1680–1683.[ISI][Medline]

Fujino, Y., Ozaki, K., Yamamusi, S. et al. (1996) DNA fragmentation of oocytes in aged mice. Hum. Reprod., 11, 1480–1483.[Abstract/Free Full Text]

Gaulden, M. (1992) The enigma of Down syndrome and other trisomic conditions. Mutat. Res., 269, 69–88.

Golbus, M. (1981) The influence of strain, maternal age, and method of maturation on mouse oocyte aneuploidy. Cytogenet. Cell Genet., 31, 84–90.[ISI][Medline]

Hansmann, I. and El-Nahass, E. (1979) Incidence of nondisjunction in mouse oocytes. Cytogenet. Cell Genet., 24, 115–121.[ISI][Medline]

Huey, S., Abuhamad, A., Barroso, G. et al. (1999) Perifollicular blood flow Doppler indices, but not follicular pO2, pO2, or pH, predict oocyte developmental competence in in vitro fertilization. Fertil. Steril., 72, 707–712.[ISI][Medline]

Hull, M., Fleming, C., Hughes, A. and McDermott, A. (1996) The age-related decline in female fecundity: a quantitative controlled study of implantation capacity and survival of individual embryos after in vitro fertilization. Fertil. Steril., 65, 783–790.[ISI][Medline]

Kanayama, K. and Osada, H. (2000) The yield of abnormal unfertilized eggs observed after repeated gonadotropin-induced ovulation. J. Int. Med. Res., 28, 24–27.[ISI][Medline]

Kline, J., Kinney, A., Levin, B. and Warburton, D. (2000) Trisomic pregnancy and earlier age menopause. Am. J. Hum. Genet., 67, 395–404.[Medline]

Lass, A. Silye, R., Abrams, D. et al. (1997) Follicular density in ovarian biopsy of infertile women: a novel method to assess ovarian reserve. Hum. Reprod., 12, 1028–1031.[ISI][Medline]

Lim, A. and Tsakok, M. (1997) Age-related decline in fertility: a link to degenerative oocytes? Fertil. Steril., 68, 265–271.[ISI][Medline]

Lubbadeh, W., Graves, C. and Spahr, (1980) Effect of repeated superovulation on ovulatory response of dairy cows. J. Anim. Sci., 50, 124–127.[ISI][Medline]

Ma, S., Kalousek, D., Yuen, B. and Moon, Y. (1997) Investigation of effects of pregnant mare serum gonadotropin (PMSG) on the chromosomal complement of CD-1 mouse embryos. J. Assist. Reprod. Genet., 14, 162–169.[ISI][Medline]

Mailhes, J., Marchetti, F. and Young, D. (1995) Synergism between gonadotropins and vinblastine relative to the frequencies of metaphase I, diploid and aneuploid mouse oocytes. Mutagenesis, 10, 185–188.[Abstract]

Mailhes, J., Young, D. and London, S. (1998) Postovulatory ageing of mouse oocytes in vivo and premature centromere separation and aneuploidy. Biol. Reprod., 58, 1206–1210.[Abstract]

McKiernan, S. and Bavister, B. (1998) Gonadotropin stimulation of donor females decreases post-implantation viability of cultured one-cell hamster embryos. Hum. Reprod., 13, 724–729.[Abstract]

Navot, D., Bergh, P., Williams, M. et al. (1991) Poor oocyte quality rather than implantation failure as a cause of age-related decline in female fertility. Lancet, 337, 1375–1377.[ISI][Medline]

Padilla, S. and Garcia, J. (1989) Effect of maternal age and number of in vitro fertilization procedures on pregnancy outcome. Fertil. Steril., 52, 270–273.[ISI][Medline]

Saito, H., Koike, K., Saito, T. et al. (1993) Aging changes the alignment of chromosomes after human chorionic gonadotropin stimulation may be a possible cause of decreased fertility in mice. Horm. Res., 39 (Suppl. 1), 28–31.[ISI][Medline]

Sakai, N. and Endo, A. (1987) Potential teratogenicity of gonadotropin treatment for ovulation in the mouse offspring. Teratology, 36, 229–233.[ISI][Medline]

Scott, R., Opsahl, M., Leonardi, M. et al. (1995) Life table analysis of pregnancy rates in a general infertility population relative to ovarian reserve and patient age. Hum. Reprod., 10, 1706–1710.[Abstract]

Sengoku, K. and Dukelow, R. (1988) Gonadotropin effects on chromosomal normality of hamster preimplantation embryos. Biol. Reprod., 38, 150–155.[Abstract]

Stolwijk, A., Zielhuis, G., Sauer, M. et al. (1997) The impact of a women's age on the success of standard and donor in vitro fertilization. Fertil. Steril., 67, 702–710.[ISI][Medline]

Szoltys, M., Galas, J., Jablonka, A. and Tabarowski, Z. (1994) Some morphological and hormonal aspects of ovulation and superovulation in the rat. J. Endocrinol., 141, 91–100.[Abstract]

Tucker, K. (1996) Reproductive toxicity of ovulation induction. Semin. Reprod. Endcrinol., 14, 345–353.

Van Blerkom, J. and Davis, P. (1998) DNA strand breaks and phosphatidylserine redistribution in newly ovulated and cultured mouse and human oocytes: occurrence and relationship to apoptosis. Hum. Reprod., 13, 1317–1324.[Abstract]

Van Blerkom, J. and Runner, M. (1984) Mitochondrial reorganization during resumption of arrested meiosis in the mouse oocyte. Am. J. Anat., 171, 335–355.[ISI][Medline]

Van Blerkom, J., Davis, J.P., Merriam, J. and Sinclair, J. (1995) Nuclear and cytoplasmic dynamics of sperm penetration, pronuclear formation, and microtubule organization during fertilization and early preimplantation development in the human. Hum. Reprod. Update, 1, 429–461.[Abstract]

Van Blerkom, J., Antczak, M. and Schrader, R. (1997) The developmental potential of the human oocyte is related to the dissolved oxygen content of follicular fluid: association with vascular endothelial growth factor levels and perifollicular blood flow characteristics. Hum. Reprod., 12, 1047–1055.[ISI][Medline]

Van Blerkom, J., Davis, P. and Alexander, S. (2001) A microscopic and biochemical study of fragmentation phenotypes in stage appropriate human embryo. Hum. Reprod., 16, 740–750.

Vanderhyden, B., Rouleau, A., Walton, E. and Armstrong, D. (1986) Increased mortality during early embryonic development after in-vitro fertilization of rat oocytes. J. Reprod. Fertil., 77, 401–409.[Abstract]

Vogel, R. and Spielmann, H. (1992) Genotoxic and embryotoxic effects of gonadotropin-hyperstimulated ovulation of murine oocytes and preimplantation embryos, and term fetuses. Reprod. Toxicol., 6, 329–333.[ISI][Medline]

Yovel, I., Geva, E., Lessing, J. et al. (1994) Analysis of the fourth to eighth in-vitro fertilization treatments after three previously failed attempts. Hum. Reprod., 9, 738–741.[Abstract]

Submitted on October 5, 2000; accepted on January 15, 2001.