Hum. Reprod. Advance Access originally published online on May 16, 2006
Human Reproduction 2006 21(9):2319-2328; doi:10.1093/humrep/del157
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Comparative genomic hybridization analysis of human oocytes and polar bodies
1 Department of Obstetrics and Gynaecology, UCL Centre for Preimplantation Genetic Diagnosis, University College London, London, UK 2 Department of Obstetrics and Gynecology, Yale University School of Medicine, New Haven, CT, USA 3 The London Fertility Centre 4 The Assisted Conception Unit, University College London Hospitals, Eastman Dental Hospital, London and 5 Department of Molecular and Cellular Pathology, Ninewells Hospital and Medical School, University of Dundee, Dundee, Scotland, UK
6 To whom correspondence should be addressed at: Department of Obstetrics and Gynaecology, UCL Centre for Preimplantation Genetic Diagnosis, 86–96 Chenies Mews, University College London, London, WC1E 6HX, UK. E-mail: efragouli{at}hotmail.com
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Abstract |
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BACKGROUND: Classical cytogenetic methods and fluorescent in situ hybridization (FISH) have been employed for the analysis of chromosomal abnormalities in human oocytes. However, these methods are limited by the need to spread the sample on a microscope slide, a process that risks artefactual chromosome loss. Comparative genomic hybridization (CGH) is a DNA-based method that enables the investigation of the entire chromosome complement. We optimized and evaluated a CGH protocol for the chromosomal analysis of first polar bodies (PBs) and oocytes. The protocol was then employed to obtain a detailed picture of meiosis I errors in human oogenesis. METHODS: 107 MII oocyte–PB complexes were examined using whole genome amplification (WGA) and CGH. RESULTS: Data was obtained for 100 complexes, donated from 46 patients of average age 32.5 (range 18–42). 22 complexes from 15 patients were abnormal, giving an aneuploidy rate of 22%. CONCLUSIONS: The results presented in this study more than double the quantity of CGH data from female gametes currently available. Abnormalities caused by whole chromosome non-disjunction, unbalanced chromatid predivision and chromosome breakage were reliably identified using the CGH protocol. Analysis of the data revealed a preferential participation of chromosome X and the smaller autosomes in aneuploidy and provided further evidence for the existence of age-independent factors in female aneuploidy.
Key words: aneuploidy/chromosome abnormalities/comparative genomic hybridization/oocyte/polar body
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Introduction |
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Data suggest that less than half of all human conceptions result in a live birth, a flaw that becomes more pronounced with increasing maternal age. The effects are apparent in IVF, where many embryos cease development before they can be transferred to the mother. Of those embryos that do survive to transfer, only about 31%, are thought to result in a live birth (Society for Assisted Reproductive Technology and the American Society for Reproductive Medicine, 2004
). There are many factors that negatively influence embryo survival, but one of the most important is the high incidence of chromosomal abnormalities. Some 0.3% of live borns are chromosomally abnormal, with trisomy of chromosome 21 or trisomy for a sex chromosome being the most common forms (Hassold and Jacobs, 1984). However, the incidence of aneuploidy at conception and during the first few days post-fertilization is believed to be two orders of magnitude higher. The most comprehensive cytogenetic studies of human preimplantation embryos have revealed that more than half of all embryos contain aneuploid cells (Voullaire et al., 2000; Wells and Delhanty, 2000). Most of the chromosomal errors present during the preimplantation stage are not compatible with development to term.
It has been demonstrated that most trisomies and/or monosomies detected in prenatal samples and spontaneous abortions are due to errors arising during the first maternal meiotic division (Hassold et al., 1996). It appears that maternal meiosis is more error-prone compared with its male counterpart, perhaps as a consequence of the prolonged arrest that begins at the dictyotene stage during fetal life and only ends upon ovulation, which may occur several decades later. Karyotyping studies of human oocytes remaining unfertilized after regular IVF have revealed two main mechanisms of non-disjunction, both leading to aneuploidy of maternal origin: the first involves the segregation of homologous chromosomes to the same pole during meiosis I, leading to the generation of disomic and nullisomic daughter cells (Zenzes and Casper, 1992). The second was proposed by Angell (1991; Angell et al., 1994), whose observations indicated that chromosome imbalance may be caused by predivision of the chromosome centromere before anaphase I, with the two sister chromatids running the risk of random segregation to either pole.
Investigation of a very large number of spare meiosis II (MII) oocytes and/or their corresponding first polar bodies (PBs) has taken place with the use of various cytogenetic methods, such as chromosome banding (Veiga et al., 1987; Kamiguchi et al., 1993; Pellestor et al., 2003), fluorescent in situ hybridization (FISH) (Dailey et al., 1996; Mahmood et al., 2000; Cupisti et al., 2003; Pujol et al., 2003), spectral karyotyping (SKY) (Sandalinas et al., 2002) and multicolour FISH (M-FISH) (Clyde et al., 2003). Data from such studies led to clear confirmation of a direct relationship between advancing maternal age and non-disjunction of both whole chromosomes and single chromatids (Sandalinas et al., 2002; Pellestor et al., 2003). The more frequent participation of chromosome X and those equal to or smaller than chromosome 13 was shown (Mahmood et al., 2000; Cupisti et al., 2003), along with the identification of a third mechanism leading to aneuploidy of maternal origin, the latter involving the presence of a trisomic cell line in the gonads of some patients (germinal/gonadal mosaicism) (Mahmood et al., 2000; Cupisti et al., 2003; Pujol et al., 2003).
The reported incidence of chromosome abnormalities in human oocytes and PBs varies greatly between different research studies, for example, Pellestor et al. (2003) reported an aneuploidy rate of 11% in the largest R-banding investigation of MII oocytes. In contrast, that found by Kuliev et al. (2003) in their FISH analysis for diagnostic purposes of first and second PBs biopsied from 6733 MII oocytes from patients of advanced maternal age was 52.1%. This variation may be the consequence of various factors such as the average age of the examined patient group, patient history and the type of analysed material (MII unfertilized oocytes versus in vitro matured to MII or PBs alone). However, a major technical parameter contributing to this variability is the fact that all the above methods require the spreading of the MII oocyte and/or PB(s) on a microscope slide, a process that increases the risk of artefactual chromosome loss. In general, hyperhaploidy can be reliably demonstrated by techniques that require spreading of an oocyte and/or PB on a microscope slide. The estimation of hypohaploidy, however, is problematic for methods involving spreading of cells, because of the possibility of artefactual loss of chromosomal material.
We have described a technique capable of detecting imbalance of any chromosome in a single cell, which avoids fixation and spreading of the sample. The method is based on the amplification of the entire DNA content of the cell followed by comparative genomic hybridization (CGH) (Wells et al., 1999; Wells and Delhanty, 2000). CGH is related to FISH and employs a competitive hybridization of differentially labelled DNA samples (test, green; chromosomally normal reference, red) to normal metaphase chromosomes. The ratio of green : red fluorescence along the length of each chromosome indicates whether there has been any gain or loss of chromosomal material in the test sample. Initially this approach was successfully applied to the analysis of blastomeres derived from normally developing preimplantation embryos (Voullaire et al., 2000; Wells and Delhanty, 2000). Since that time, our group and others have sought to utilize this methodology for investigating oocytes and PBs, initially in a research context and ultimately for the purpose of PGD (Wells et al., 2002; Gutiérrez-Mateo et al., 2004a; Fragouli et al., 2006b).
We report our cumulative results obtained after the CGH analysis of 93 MII oocyte–PB complexes and 14 single first PBs. Our results confirm and extend those presented in other cytogenetic investigations of human female gametes, more than doubling the amount of CGH data existing for this type of material. Various mechanisms leading to aneuploidy of maternal origin, especially those affecting younger patients were identified. We were able to accurately assess the incidence of chromosome abnormalities in the investigated MII oocyte–PB complexes because of our larger sample size. Importantly, our aneuploidy rate of 22% is much lower compared with that of 48% reported by the smaller MII oocyte–PB CGH investigation carried out by Gutiérrez-Mateo et al. (2004a). The reliability of CGH to detect chromosome abnormalities in both MII oocytes and PBs is demonstrated, along with its ability to distinguish between whole chromosome and single-chromatid anomalies.
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Participating patients
Forty-five women with no known chromosome abnormalities and one female balanced carrier of a reciprocal translocation [46,XX,t(X; 4)(q26; p16.1)] consented to donate their unused oocytes for research. Their age range was between 18 and 42 years, with an average age of 32.5 years. These patients were undergoing routine infertility treatment at four different IVF Centres: the Assisted Conception Unit at Ninewells Hospital, Dundee, Scotland (Centre 1, 15 patients; Fragouli et al., 2006b), the Assisted Conception Unit at University College London Hospitals Trust, London (Centre 2, 14 patients), the Mayo Clinic, Rochester, MN, USA (Centre 3, 13 patients) and the Yale Fertility Centre, Yale University School of Medicine, New Haven, CT, USA (Centre 4, 4 patients). The participating patients were being treated because of various indications, which are as follows: male factor, 27/46; unexplained, 4/46; tubal/uterine factors, 5/46; polycystic/multicystic ovaries, 3/46; repeated IVF failure, 1/46; preimplantation genetic screening (PGS), 2/46; PGD for a reciprocal translocation [46,XX,t(X; 4)(q26; p16.1)], 1/46; frozen oocytes before chemotherapy, 1/46; advanced maternal age, 1/46 and oocyte donation, 1/46. Details of the maternal age ranges, average ages and patient indications for each of the four centres are summarized in Table I.
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The ovarian stimulation protocol was similar in all four centres and consisted of pituitary down-regulation using GnRH agonist Synarel or GnRH analogue Suprefact. Follicular development stimulation was achieved by administering daily injections of 150–450 IU of gonadotrophin. Follicular growth was observed by transvaginal ultrasonography, and 10 000 IU of hCG was administered between 12 and 14 days. Collection of oocytes took place 36 h later by ultrasound transvaginal aspiration.
Ethical approval
Donation of oocytes from all the patients occurred only after their informed consent. This research work was carried out under license from the Human Fertilization and Embryology Authority and was also approved by the Research Ethics Committees of University College London Hospital and Tayside Trusts and the Institutional Review Boards of the two USA IVF Centres.
Oocytes and corresponding PBs
One hundred and seven MII oocytes and/or corresponding first and possibly second PBs (189 single cells) were investigated during this study. Of these, 36 MII oocytes remained unfertilized after sperm exposure either via IVF (15 cells) or via ICSI (21 cells), two of which consisted of both first and second PBs, 16 MII oocytes were unexposed to sperm and 55 were either at the germinal vesicle (GV, 26 cells) or meiosis I (MI, 29 cells) stage at the time of retrieval were left to mature in vitro (over 24–48 h), and were again unexposed to sperm. An additional two immature oocytes (one GV and one MI) were also examined. Table I summarizes the different types of investigated oocytes and the IVF Centres that donated them.
Oocyte and PB separation and processing
Zona pellucida removal and oocyte and corresponding PB separation were achieved using either Acid Tyrode’s or Pronase (Sigma, Gillingham, UK). The single cells were processed as outlined in Fragouli et al. (2006b). PB biopsy took place for the separation of MII oocytes and their corresponding first PBs from Patient A (UCL). The procedure is described in detail in Fragouli et al. (2006). Cell lysis took place by incubating samples in 2 µl proteinase K (PK) (125 µg/mL) and 1 µl sodium dodecyl sulphate (SDS) (17 mM) (Sigma), at 37°C for 1 h, followed by an incubation at 95°C for 15 min to inactivate the PK enzyme.
Whole genome amplification
Method 1—degenerate oligonucleotide primed-PCR
The degenerate oligonucleotide primed-PCR (DOP-PCR) was employed for the whole genome amplification (WGA) of most (104 of the 107) of the MII oocytes and their PBs. The protocol used was as described by Wells et al. (2002), with modifications as described in Fragouli et al. (2006b).
Amplifications were carried out initially in a Hybaid Omnigene and later in a 9700 PE thermocycler (Applied Biosystems, Warrington, UK).
Method 2—multiple displacement amplification
The multiple displacement amplification (MDA) was employed for the WGA of three MII oocyte–PB pairs (O-1/O-1PB, E-1/E-1PB and E-5/E-5PB) from Centre 2. Amplification was achieved using bacteriophage 
29 DNA polymerase, exonuclease-resistant phosphorothioate-modified random hexamer oligonucleotide primers and reaction buffer, according to the manufacturer’s instructions (REPLI-G kit, Qiagen, Crawley, UK). The total volume was 50 µl, whereas the reactions took place by incubating all samples at 30°C for 6 h, followed by a 3-min incubation at 65°C to inactivate the 29 DNA polymerase. The 9700 PE thermocycler was employed during the MDA amplifications.
At the end of either the DOP-PCR or the MDA, 45 µl of amplified product underwent the remainder of the CGH procedure as described below, and 5 µl was kept for agarose gel analysis. Strict precautions to avoid contamination were taken during oocyte and PB processing, as described in Wells and Sherlock (1998). Two microlitres of the final drop of phosphate-buffered saline/polyvinyl alcohol (PBS/PVA) into which each of the oocytes and PBs was washed before transferring into microfuge tubes along with the rest of the DOP-PCR or MDA reagents was used as negative controls for each experiment. WGA and CGH were employed for these negative controls, and absence of contaminating DNA was seen as absence of fluorescent signals on metaphase chromosomes, during analysis.
The reference DNA against which the amplified oocyte and PB DNAs were hybridized came from clumps of 3–5 buccal cells from 46,XX individuals. These were processed in exactly the same way as the oocytes and PBs.
Fluorescent labelling of amplified products
Amplified oocyte, PB and buccal cell DNAs were fluorescently labelled using the Nick Translation kit (Abbott, Maidenhead, UK), according to the manufacturer’s instructions. The test oocyte or PB DNA was labelled using Spectrum Green-dUTP (Abbott), whereas the reference buccal cell DNA was labelled using Spectrum Red-dUTP (Abbott). Both red and green DNAs were ethanol co-precipitated with 30 µg of human Cot-1 DNA (GIBCO/BRL, Paisley, UK). Pellets were air-dried and resuspended in 6 µl of hybridization buffer [50% formamide, 2x saline sodium citrate (SSC), 10% dextran sulphate, pH 7].
Comparative genomic hybridization
The remainder of the CGH procedure took place as described in Fragouli et al. (2006a) with certain modifications. Hence, both slides and probes were denatured in a waterbath set between 73 and 75°C, and hybridization took place for 42–72 h. Post-hybridization washes took place as described by Wells et al. (2002), followed by dehydration of the slides through a 70, 90 and 100% ethanol series. The chromosomes were counterstained with antifade medium, containing diamidophenylindole (DAPI). The combination of DOP-PCR and CGH, applied to single cells, has been previously validated by our group (Wells et al., 1999) as well as others (Voullaire et al., 1999).
Microscopy, image analysis and interpretation
Metaphase spreads were observed using an Olympus BX 40 fluorescent microscope with a cooled charge-coupled device (CCD) system and filters for the fluorochromes used. Ten metaphases were captured on average per hybridization. Analysis and interpretation of the captured images was feasible by using Vysis Quips CGH software (Vysis/Abbott, UK) that converted fluorescent intensities into a red–green ratio for each chromosome. The study was not carried out blind. Initially, the scoring of chromosomal losses and gains was determined according to strict criteria (i.e. pre-set red : green ratio thresholds from the CGH software used for the analysis and interpretation). However, in a couple of cases, suboptimal hybridization led to some uncertainties in interpretation. In these rare cases, consideration of the corresponding oocyte/PB was very useful in confirming that apparent losses/gains observed were indeed genuine.
Equal sequence copy number between the test and the reference DNAs was seen as no fluctuation of the ratio profile from 1:1. Test sample underrepresentation was seen as fluctuation of the ratio profile in favour of the red colouration (below 0.80), whereas test sample overrepresentation was seen as fluctuation of the ratio profile towards the green colouration (above 1.20). Such fluctuations were respectively scored as losses or gains in the test sample compared to the reference sample. Distinction between loss of whole chromosomes and single chromatids was possible. This was achieved initially by comparing the fluorescence intensity of the green fluorochrome (sample DNA) on the deficient chromosome with that on the euchromatic region of the Y chromosome. The Y chromosome is absent from both the test and the reference DNAs; hence, the slight fluorescence observed on this chromosome could be attributed to background fluorescence unrelated to hybridization of unique sequences from the test and reference DNA samples. This served as an indicator of the amount of fluorescence expected on a chromosome that had been entirely lost. When some green fluorescence was visible and the chromosome in question was relatively bright compared with the Y chromosome, but fainter when compared with the rest of the chromosomes, then the loss would be attributed to a single chromatid. The difference in fluorescence intensity, which allowed chromatid loss to be distinguished from whole chromosome loss, was unambiguous and easily scored by eye. However, to remove any subjectivity from the scoring procedure, we also assessed the average red : green fluorescence ratio of pixels along the axis of each chromosome. Loss of an entire chromosome (confirmed by analysis of the corresponding PB or oocyte) resulted in average red : green ratios of >2.8:1. Chromatid loss produced red : green ratios ranging from 1.3:1 to 2.0:1.0. This contrasted with balanced chromosomes that displayed ratios of 1.15:1 to 1:0.87. Gain of a single chromatid would only be distinguished from whole chromosome gain in the cases where the corresponding cell was characterized as having lost this chromatid. Examples of the green fluorescence intensity observed during the loss of a single chromatid that scored when a whole chromosome is missing and that seen for a balanced chromosome compared with the green fluorescence of the Y chromosome can be seen in Figure 1, in Fragouli et al. (2006b). Heterochromatic, centromeric and telomeric regions were excluded from analysis, as they tend to show an artefactual deviation of the ratio profile.
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Results |
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Ninety-three MII oocyte–PB complexes and 14 single first PBs donated from 46 women were investigated during this study. Of these women, 45 were considered to be of normal 46,XX karyotype, whereas one was a balanced carrier of a reciprocal translocation [46,XX,t(X; 4)(q26; p16.1)]. These women were undergoing routine assisted reproductive technology (ART) procedures combined with either PGS or PGD in some cases, in four different IVF Centres, for various indications, as detailed in the Materials and methods section, and Table I. Their age range was between 18 and 42 years, and the average maternal age was 32.5 years.
Most (71) of the examined MII oocyte–PB complexes were unexposed to sperm, 16 of which were at MII stage upon retrieval and 55 being scored as immature (GV or MI) during oocyte collection and being cultured to mature in vitro. An additional 36 MII oocytes that were considered to be unfertilized after sperm exposure either via IVF or via ICSI were also analysed using CGH.
One hundred and eighty-nine single cells (oocyte or PB) had their genomes amplified using one of two WGA methods. DOP-PCR was employed for the initial amplification of 183 MII oocytes and/or PBs. Of these 70 MII oocytes, 87 first and 2 second PBs were successfully amplified (87%), yielding the expected 200–4,000 bp smears upon agarose gel analysis. Three MII oocyte–PB complexes were amplified by MDA, in an attempt to evaluate this method as an alternative WGA protocol. Successful amplification was accomplished for five of the six single cells (83.3%), with the amplified products resulting in smears with average fragment sizes of 10 000 bp.
CGH results were available from the oocyte or the PB or both for 88 MII oocyte–PB pairs. Additionally, CGH data were obtained from 12 single first PBs and 1 immature oocyte.
One hundred and fifteen single cells, those being 50 MII oocytes, 64 first and 1 second PBs, were characterized as haploid normal, 23,X. These single cells represented 78 MII oocyte–PB complexes, and paired results were obtained for 39 of these. As far as the developmental stage during oocyte retrieval was concerned, 22/78 were at the MII stage and were unfertilized, 13/78 were at the MII stage but were unexposed to sperm, whereas 43/78 were matured in vitro before CGH analysis.
Twenty-two MII oocyte–PB complexes were classified as abnormal, the latter corresponding to 12 oocytes, 17 first and 1 second PBs. In sixteen of these complexes, the oocyte was at the MII stage at retrieval (3 unexposed to sperm, 13 classified as unfertilized, 1 consisting of both the first and the second PBs), whereas for six cases, the oocyte was in vitro matured. The resulting aneuploidy rate was 22%. A summary of the results obtained after CGH analysis of the abnormal oocyte–PB complexes, including the maternal ages and the developmental stage during oocyte collection, is summarized in Table II.
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Forty-two anomalies were scored in total, with the fertilized oocyte at risk of trisomy in 14 cases versus the risk of monosomy in 13 cases. Three oocytes (B-4, B-5 and D-18) and five first PBs (A-3, B-5, D-18, N-6 and O-1) were characterized as extensively aneuploid with at least two chromosomes affected. Reciprocal losses and gains were seen for seven complexes (A-13, B-5, D-18, F-1, K-5, L-1 and M-2). CGH had failed for one of the two cells of the complex in five cases (A-3, E-5, I-1, L-2 and O-1), whereas the oocyte or PB was lost during processing for five (E-1, G-4, J-2, J-4 and N-6) of the abnormal pairs. The oocyte from complex C-2 and the PBs from complexes L-3 and L-4 were not available for investigation. Non-complementarity of CGH results for the oocyte–PB pair H-3 was attributed to germinal mosaicism.
Two main mechanisms leading to female aneuploidy were observed during this investigation. Thus, abnormalities attributed to both whole chromosome non-disjunction (12 complexes) and unbalanced predivision of single chromatids (nine complexes) were observed, either alone or in combination. It was not possible to distinguish between these two mechanisms in two oocytes and one PB with gain of chromosomal material (L-3, L-4 and C-2) because of non-availability of the corresponding cells. Among the abnormal MII oocyte–PB complexes that were considered to be unfertilized, we were able to identify one oocyte (K-5) that was initially aneuploid because of the presence of an extra chromatid 21, the latter being corrected by segregation of this extra chromatid to the second PB at the end of MII. We were also able to identify one patient who was germinal mosaic for trisomy 13, as mentioned above. It is unclear, however, whether this patient had extensive gonadal mosaicism or whether most of their germinal cells were normal.
Chromosomes of differing sizes were identified to be participating in aneuploidy events, including the larger 1, 2, 5 and 9, detected as abnormal in one MII oocyte–PB complex each. Chromosome X was most frequently involved in aneuploidy (10 complexes), followed by chromosomes 21 (four complexes), 20 (three complexes), 13 and 8 (two complexes each). Two different types of abnormalities involving chromosome 4 were identified in two MII oocyte–PB complexes. N-6 PB was scored as consisting of an extra chromosome 4, with its corresponding oocyte not being available for investigation. A structural abnormality in the form of a duplication of the chromosomal region 4q31.1-q35 was detected for the O-1PB. Interestingly, the breakpoint on chromosome 4 coincides with a previously defined chromosomal fragile site (Corso and Parry, 1999). Chromosomes belonging to groups A–C have been previously identified to be aneuploid in other oocyte–PB CGH investigations (Wells et al., 2002; Gutiérrez-Mateo et al., 2004a,b). The frequency of specific chromosome aneuploidy detected during this investigation is summarized in Table III. Figure 1 demonstrates the analyses of metaphases and the resulting interpretations for the D-oocyte–PB complex.
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The abnormal MII oocyte–PB complexes were generated from 15 patients whose average age was 31 years (age range: 18–40 years). A clear maternal age effect was not observed, mostly because of the unequal age distribution of the investigated population. Hence, 33 women below the age of 37 years donated 84 MII oocyte–PB complexes, and 19 were aneuploid (22.6%). The aneuploid cells were derived from 12 patients whose average age was 29 years (age range: 18–33 years). Conversely, only 13 women were either 37 years or older, donating 16 complexes; three of which were considered as abnormal (18.7%). The average age of these three patients was 38.6 years (age range: 37–40 years).
CGH results were obtained for 132 of the 164 single cells (80.5%), whose genome was initially successfully amplified using one of two WGA methods. No difference was seen as far as the fluorescence intensities between samples amplified by the DOP-PCR and with the MDA are concerned. CGH was slightly more efficient for the PBs rather than their corresponding oocytes (70/92 versus 62/72). This could be attributed to the nature of the PB, which is a much smaller cell compared with its oocyte counterpart; hence, the PB DNA is more easily accessible. Gutiérrez-Mateo et al. (2004a) also postulated that the high mitochondrial content of oocytes could result in fainter hybridization signals. Most importantly, however, CGH was identified to be a sensitive method, able to detect abnormalities attributed both to whole chromosome and single-chromatid non-disjunction and also structural abnormalities.
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Discussion |
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During this study, CGH was carried out to assess the full chromosome complement of 93 MII oocyte–PB complexes and 14 single first PBs. The results obtained effectively quadruple the amount of published oocyte CGH data and suggest an incidence of oocyte aneuploidy that is significantly lower than that reported using CGH previously (Gutiérrez-Mateo et al., 2004a
). Chromosome abnormalities were detected for 12 MII oocytes, 17 first and 1 second PBs. This represents data from 22 MII oocyte–PB complexes, as in some cases, a result was only obtained from the PB or the oocyte but not both. We were able to score losses and gains of whole chromosomes and losses of single chromatids with accuracy. Gains of single chromatids could be scored where both oocyte and PB were available. It has been suggested that the frequency of single-chromatid anomalies is almost equal to that of whole chromosome abnormalities (Clyde et al., 2001; Sandalinas et al., 2002; Cupisti et al., 2003; Pujol et al., 2003). This was also seen in this study, with whole chromosome non-disjunction being observed for 12 complexes and unbalanced predivision of single chromatids for nine.
Theoretically, it is expected that a single-chromatid abnormality will result in embryonic aneuploidy in 50% of cases, depending on whether the unattached chromatid enters the second PB after fertilization (Angell et al., 1993). This hypothesis was confirmed during our investigation. The ‘correction’ of a meiosis I error during MII was seen for oocyte K-5, as described in the Results section.
We observed an almost equal incidence of chromosome loss and gain in the abnormal oocytes (13 versus 14 cases, respectively). This is in accordance with the theoretical expectation that the risk of a monosomic conception would be equal to that of a trisomic one (te Velde and Pearson, 2002) but assumes that anaphase lag does not occur. One study of human preimplantation embryos in fact found the monosomies exceeded trisomies (Munné et al., 2003). Chromosomes from almost all groups were found to participate in aneuploidy events, with chromosome X and those smaller or equal in size to chromosome 13 being more frequently affected. This is in agreement with previous investigations, employing conventional karyotyping (e.g. Pellestor et al., 2003), FISH (e.g. Mahmood et al., 2000; Cupisti et al., 2003) and CGH (e.g. Gutiérrez-Mateo et al., 2004b; Fragouli et al., 2006b).
Chiasmata are crucial for the correct meiotic segregation of homologues (Tease et al., 2002). Because fewer chiasmata are formed in the case of the smaller bivalents, during the first prophase in female meiosis, they are at increased risk of segregation errors. Various studies have been carried out in an attempt to elucidate the patterns of female recombination that could potentially lead to chromosome malsegregation (Robinson et al., 1993; Sherman et al., 1994; Fisher et al., 1995; Hassold et al., 1995; Lamb et al., 1996; Nicolaidis and Petersen, 1998; Savage Brown et al., 2000). All these investigations came to the common conclusion that variations in the recombination patterns act as risk factors contributing to non-disjunction of chromosomes. Further confirmation came from a recent study, carried out by Lenzi et al. (2005). These authors demonstrated that up to one-third of human oocytes may fall below the optimal threshold of one or two crossovers per bivalent, increasing the non-disjunction risk.
The chromosome with the highest anomaly frequency in the investigated MII oocyte–PB complexes was X, with single-chromatid abnormalities predominating over those of whole chromosomes. This may be attributed to the absence of recombination affecting this bivalent in some cases. Thomas et al. (2001), in their investigation on the origin of trisomy X, observed that, in 56% of cases arising from errors taking place during meiosis I, there was no recombination between the two homologues. Additionally, it has been shown in the mouse that a univalent chromosome X can segregate intact to either pole at anaphase I or prematurely divide into its two sister chromatids, which in turn are also at risk of random segregation (Hunt et al., 1995).
Abnormalities affecting the larger chromosomes 1, 2, 4, 5, 8, 9 and 12 were also detected during our investigation. It seems that larger autosomes can malsegregate during meiosis, but this phenomenon is not as common as the one involving the non-disjunction of smaller autosomes. This is in agreement with Lynn et al. (2004) and with the findings of Roig et al. (2005) who examined the homologue-pairing process taking place during prophase I in 8603 human oocytes. From the data obtained, it was concluded that pairing errors affect all chromosome groups, but some chromosomes seem to be more prone to recombinational errors than others. Interestingly, in the current study, all eight aneuploidies affecting the ‘larger’ chromosomes (1–12) arose by a classical non-disjunction mechanism rather than by premature chromatid separation. This is in contrast to aneuploidy of the ‘smaller’ chromosomes (13–22 and X), which resulted from both mechanisms at a similar rate.
Twenty-seven percent of the abnormal oocyte complexes in this study had an abnormality affecting the A–C group of chromosomes (excluding the X chromosome abnormalities) versus 41% for the D–G group. This finding is of relevance to PGS of embryos, which is increasingly employed to assist in the identification and preferential transfer of viable embryos during IVF treatment. Virtually, all reported PGS methods employ FISH and screen between five and nine chromosomes. Chromosomes 1–12 are seldom assessed, and yet it is clear that a significant number of oocytes harbour presumably lethal defects involving these chromosomes.
One of the features of CGH that distinguishes it from FISH is the ability to detect imbalances affecting fragments of chromosomes as opposed to whole chromosomes only. A structural alteration of the long-arm subtelomeric region of chromosome 4 (4q31.1-q35) was identified in one first PB (O-1PB) with our CGH protocol. Alterations affecting chromosome fragments have been previously observed in blastomeres from embryos examined using CGH (Voullaire et al., 2000; Wells and Delhanty, 2000) and in two oocytes during the SKY investigation carried out by Sandalinas et al. (2002). However, this is the first CGH study to detect such an alteration in a PB or oocyte. Wells and Delhanty (2000) observed reciprocal losses and gains of parts of chromosomes 1, 2 and 7 in pairs of blastomeres from two different embryos. Breakpoints mapped to fragile chromosomal sites that are prone to breakage. The 4q31.1-q35 region observed as a gain in the O-1PB has also been characterized as a common fragile site (Corso and Parry, 1999). Such fragile sites could be induced by depletion of specific medium nutrients (Martin et al., 1990), a factor that should be considered during the development of oocyte maturation protocols.
The total aneuploidy rate for the current study was calculated to be 22%. This is comparable to the 20% aneuploidy rate suggested by Pellestor et al. (2005) in their data collection of various cytogenetic investigations of human oocytes and corresponding first PBs but much lower than the 48% reported by Gutiérrez-Mateo et al. (2004a) in their CGH investigation of 30 MII oocyte–PB pairs. The authors attributed this high aneuploidy rate to various reasons, one being the in vitro maturation (IVM) of their investigated group of cells. This was not observed for the in vitro matured oocytes in our data set. Hence, 32% (15 of 47) of the MII oocytes that were mature at the time of retrieval were aneuploid compared with an aneuploidy rate of 10.7% (6 of 56) in those that were immature at retrieval and left to mature in vitro. These data, although preliminary, provide reassurance that IVM of oocytes does not increase rates of chromosome abnormality.
It is noteworthy that most of the oocytes investigated during this study were derived from patients predicted to have good ovarian function (i.e. relatively young, no ovarian pathology, cause of infertility unrelated to ovulation or oocyte quality).
Various sources of information indicate that certain couples are particularly predisposed to the conception of aneuploid embryos. Women whose age is lower than 35 years (i.e. too young to be considered at high risk of aneuploid pregnancy) and who have had three or more conceptions with the same trisomy are good candidates for gonadal mosaicism as has been clearly demonstrated by several studies (Cozzi et al., 1999; Sachs et al., 1999; Somprasit et al., 2004). One such patient was identified in our group as well. CGH detected an extra chromosome 13 in the oocyte of Patient H, whereas the corresponding PB was characterized as being 23,X, suggesting gonadal/germinal mosaicism for trisomy 13.
Another class of patients with a predisposition to the generation of aneuploid oocytes are those displaying an apparent elevated meiotic error rate. In their report, Mahmood et al. (2000) describe the finding of three abnormal oocyte–PB pairs of the examined four, generated from a 26-year-old woman. Similarly in the study of MII oocyte–PB pairs by Gutiérrez-Mateo et al. (2004b), one patient aged 27 years had three of four PBs aneuploid for two different chromosomes. Six women from our investigated patient group, A, B, D, L, N and O, were considered to be candidates for a susceptibility to the generation of aneuploid oocytes. This observation was made based on the presence of either multiple abnormalities in one or more of their examined cells or the same type of abnormality in all of their cells. It is interesting to note that three (B, D and L) of the above six women were being treated for polycystic or multicystic ovaries. Seven of the 10 MII oocyte–PB complexes generated from these patients were highly abnormal. However, this is too small a sample size to draw any definite conclusions concerning an association of polycystic ovaries and aneuploidy.
Inclusion of data on MII oocytes and/or PBs from patients with a predisposition to aneuploidy will influence the estimation of chromosome abnormality rates in smaller studies. This is not considered in the CGH study of Gutiérrez-Mateo et al. (2004a), rather the authors argue that their relatively high aneuploidy rate resulted from the fact that all 23 chromosomes were investigated. We also examined the entire chromosome complement of MII oocytes and their corresponding PBs, and suggest that the lower aneuploidy rate obtained is a consequence of the much larger number of samples tested. Results from a large sample size are less likely to be skewed by the presence of unusual patients with high aneuploidy rates.
The average age of the patients whose MII oocyte–PB complexes were characterized as aneuploid was 31 years (range: 18–40 years). We were unable to detect an advancing maternal age effect most likely because of the unequal age distribution of our patient group. Hence, most (33) of the investigated women were below the age of 37 years. Another explanation for an absence of a maternal age effect is the action of age-independent mechanisms leading to maternal aneuploidy. There are some data suggesting that such mechanisms operate in certain infertile patients (e.g. recurrent miscarriage patients, Munné et al., 2005). Furthermore, the study of Lenzi et al. (2005) demonstrated that genome-wide variation in recombination is an individual trait, something which implies a genetic basis. Combination of our data with that obtained from the two studies carried out by Gutiérrez-Mateo et al. (2004a,b) who used a similar protocol indicates that CGH is ready for a wider clinical application. The value of PGS with the use of FISH to score up to nine chromosomes (X, Y, 13, 15, 16, 17, 18, 21 and 22) has recently been demonstrated in two different studies. It was shown in both that such an approach is of benefit for IVF patients of advanced maternal age (over 35 years), with or without repeated miscarriages, as it leads to a reduction in spontaneous abortion rates and an increase in pregnancy rates (Munné et al., 2005; Platteau et al., 2005). These benefits were not seen for the younger women. A CGH-based approach could expand the range of patients benefiting from PGS by permitting detection of the entire chromosomal set, including chromosomes that do not show a strong association of age and aneuploidy. The most important drawback at this stage, however, is the labour-intensive nature of the existing protocols and the hybridization time required to yield results. Both in our study and that of Gutiérrez-Mateo et al. (2004a), there was no difference in the fluorescence intensities obtained after 42 and 72 h of incubation. Considering all the above, a PGS approach involving the screening of first PBs using CGH to predict the chromosome constitution of the corresponding oocytes should be considered for couples who have undergone at least one unsuccessful cycle of PGS–FISH. A CGH-based approach may be particularly valuable in cases where the prior PGS–FISH cycle has revealed an excess of meiotic versus post-zygotic errors. Such an approach would involve biopsy of first PBs before fertilization on day 0, with the CGH protocol lasting for approximately 60 h (6 h for the amplification and 42–48 h for the hybridization). Biopsy and FISH analysis of blastomeres should also be feasible on day 3, with both CGH and FISH results obtained on day 4 and embryo transfer either late on day 4 or day 5.
To conclude, we report data from the largest CGH study of MII oocytes and PBs to date. The information obtained during our investigation is of both clinical and scientific value. The reliability of CGH for the detection of whole chromosome and single-chromatid gains and losses and also de novo structural alterations because of chromosome breakage is clearly demonstrated. We were able to accurately assess the aneuploidy incidence in our investigated group of MII oocyte–PB complexes. The relatively large size of our sample set provides some protection from bias introduced by atypical patients with unusually high aneuploidy rates. The two main mechanisms of maternal aneuploidy, those being the malsegregation of entire chromosomes or single chromatids, were both observed, along with one case of gonadal mosaicism. The preferential involvement of the smaller autosomes was also seen. Our findings confirm and extend the data from other cytogenetic investigations of human female gametes. The information obtained from our study could be crucial for the further treatment and counselling of such couples. Moreover, we postulate that age-independent mechanisms could be acting in groups of younger IVF patients and may be influenced by their infertile etiology. Our data thus raise questions about the regulatory mechanisms of female meiosis, whose elucidation should provide a clearer picture of the complex issue of female infertility.
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The authors thank all the patients who agreed to donate oocytes towards this research project, and the consultant obstetricians and embryologists of the four participating IVF Centres for their support and help. Elpida Fragouli is a recipient of the 2003 Florence and William Blair-Bell Memorial Fellowship from WellBeing of Women under grant no. PG 603.
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Submitted on February 14, 2006; resubmitted on March 25, 2006; accepted on April 11, 2006.
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