1 Departments of Obstetrics and Gynecology and 2 Molecular and Human Genetics, Baylor College of Medicine, Houston, Texas, USA
3 To whom correspondence should be addressed at: Baylor College of Medicine, Department of Obstetrics and Gynecology, 6550 Fannin Street, Suite 885, Houston, TX 77030, USA. Email: bischoff{at}bcm.tmc.edu
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Abstract |
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Key words: aneuploidy/chromosomal nuclear localization/preimplantation genetic diagnosis
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Introduction |
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The nuclear organization of every human chromosome in diploid lymphoblasts and primary fibroblasts was studied by Boyle et al. (2001). As we would have predicted, this group found the most gene-rich chromosomes (HSA-1, 16, 17, 19 and 22) to be localized within the centre of the nucleus; gene-poor chromosomes (HSA-2, 3, 4, 7, 8, 11, 13 and 18) were concentrated towards the nuclear periphery. Chromosomes 5, 6, 10, 14, 15, 20 and 21 were not found to have a significant bias either to the peripheral or to central locations. Boyle et al. (2001)
found no correlation between chromosome size and position within the nucleus, although gene density clearly contributed to the nuclear positioning of individual chromosomes. Thus, disruption of the nuclear organization of chromosomes should result in altered interaction between chromatin and the nuclear membrane with subsequent deregulation of gene expression. Alternatively, both nuclear location and gene expression could reflect perturbation of the same underlying event.
To distinguish between these two possibilities, we studied blastomeres derived from preimplantation embryos of subjects undergoing preimplantation genetic diagnosis (PGD). Using fluorescence in situ hybridization (FISH) chromosome-specific probes, we hypothesized that the nuclear organization of individual chromosomes may reflect or play a role in cell division and embryonic development. Thus, we predicted altered nuclear position in aneuploid and morphologically abnormal but not in morphologically normal (euploid)embryos.
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Materials and methods |
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FISHPGD probes
Direct-labelled chromosome-specific probes (Vysis, Inc.) were used in analysis of all blastomeres. Our FISH strategy involved two sequential hybridizations. In the first hybridization, probes specific to chromosomes 13 (13q14 locus-specific, LSI 13; SpectrumGreen-labelled), 18 (18p11.1-q11.1 alpha satellite centromere, CEP18; 1:1 equal mixture of SpectrumOrange:SpectrumAqua-labelled), 21 (21q22.13-q22.2 locus-specific, LSI 12; SpectrumOrange-labelled), X (Xp11.1-q11.1 alpha satellite centromere, CEPX; 1:1 equal mixture SpectrumOrange:SpectrumGreen-labelled) and Y (Yq12 satellite III sequence-specific, CEPY; SpectrumAqua-labelled) were used simultaneously to identify blastomeres/embryos with the correct number of FISH signals corresponding to each chromosome. Blastomeres having two signals for each of the autosomes (13, 18 and 21) and either two signals for the X chromosome alone (XX) or one signal for both the X and Y chromosomes (XY) were classified as euploid (normal) and, hence, subjected to a second hybridization using probes for chromosomes 16 (16q11.2 satellite II centromere, CEP16; SpectrumOrange labelled), 22 (22q11.2 locus-specific, LSI 22; SpectrumGreen-labelled) and chromosome 18 again. The second hybridization for chromosome 18 was perfomed as a positive hybridization control (alpha satellite centromere, CEP18; SpectrumAqua-labelled).
For the two reciprocal translocation cases (nos. 11 and 12), only blastomeres (n=4 total) identified as balanced/normal for the rearrangement were subsequently assessed for chromosomes 13, 18, 21, X and Y. Of the 13 blastomeres/embryos examined in the 21:21 robertsonian translocation case (no. 10), only one was found to be balanced/normal; all 13 were assessed for chromosomes 13, 18, 21, X and Y.
FISH: denaturation and hybridization
Following our previously described protocol (McKenzie et al., 2003), probes were mixed, denatured for 5 min at 70°C, applied to coverslips and then mounted onto blastomeres. Slides with probe mixtures were simultaneously denatured for 5 min using a 76°C hot plate and then transferred to a humidity chamber for 69 h. Slides were post-washed in 0.4 x sodium salt citrate at 72°C and counterstained with DAPI (4',6-diamidino-2-phenylindole II; Vysis, Inc.).
To re-probe a blastomere nucleus, slides were placed in 0.4 xstandard saline citrate for 5 min at 70°C to remove probes from first hybridization. Slides were then dehydrated in serially increasing concentrations of ethanol (70, 90 and 100%), and air-dried for the second round of hybridization. Probe mixture was denatured separately at 70°C for 5 min and then sealed onto slides followed by simultaneous denaturation on a 76°C hot plate for 5 min. Following a 68 h hybridization, slides were post-washed as described above.
Microscope analysis
FISH signals were visualized using a Zeiss Axioskop microscope equipped with multi-bandpass filters that allow simultaneous visualization of different colors. A digital imaging system (Applied Imaging, USA) with a cooled charged couple device (gray-scale) was used for image capture and data collection.
Localization of hybridization signals
Eight concentric maps with four quadrants each were created using computer graphics. Shapes and sizes of these maps reflect the various nuclear morphologies expected following the biopsy and fixation procedure. Given that all blastomere FISH experiments were image-captured, each grid map was copied onto transparency sheets that were then placed directly on the computer monitor. For each blastomere nucleus, a concentric map that best matched the nuclear border was chosen and used to map each hybridization signal. Each of the four quadrants was designated to be equally distant from each other, representing the most central area (Q1) and radiating out toward to the nuclear periphery (Q4) (Figure 1). If a signal was equally present on the grid line between two quadrants, a value of one-half signal was assigned to both quadrants. Location of each hybridization signal was recorded for both euploid (normal) and aneuploid cells.
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Results |
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Localization in blastomeres with structurally normal versus unbalanced chromosomes
For the 21q:21q robertsonian translocation, FISH for chromosomes 13, 18, 21, X and Y was performed on 13 blastomeres. Among 12 blastomeres classified as unbalanced, we observed no significant difference in localization of any chromosome, compared with either chromosomally normal/balanced blastomeres derived from non-homologous robertsonian and reciprocal translocation cases (n=5) or from euploid cases (n=30) (2=5.8 to 2.7, P
1.0, df = 2).
Correlation of chromosome localization to embryo morphology
Two of the 17 couples had no normal embryos and were therefore excluded from analysis. For each of the remaining 15, there was no difference (P=0.818) in localization of chromosomes when comparing morphologically normal embryos (grades 4 or 5) to morphologically abnormal embryos (grades 1, 2 or 3) (Figure 4).
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Discussion |
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The localization of chromosomes did not differ when comparing morphologically normal versus abnormal embryos. This is consistent with studies showing that morphological appearance does not necessarily predict a healthy embryo, up to 40% of embryos with normal morphology being aneuploid (Munné et al., 1993, 1994
).
Our results are based on extrapolation of data from the two-dimensional (2D) blastomere preparations. The pivotal assumption is that the relative organization of the nucleus is preserved. Indeed, Croft et al. (1999) compared location of hybridization signals for HSA-18 and -19 in human fibroblasts between confocal laser scanning microscopy and traditional 2D preparations: three-dimensionally (3D) preserved cells fixed with paraformaldehyde and not subjected to hypotonic swelling proved consistent with the orientations of 2D specimens. Similarly, data collected from hypotonically treated flattened nuclei in Drosophila were consistent with data collected from 3D preserved nuclei (Csink and Henikoff, 1998
).
Localization of signals to the periphery of the nucleus in presumed chromosomal aneuploidy suggests either that cells are undergoing programmed cell death (apoptosis) or that key functions (i.e. transcription of certain regulatory genes) maintaining nuclear organization may be altered. During apoptosis, chromosomal fragmentation and relocalization to membrane periphery (in preparation for packaging into apoptotic bodies) occurs prior to characteristic membrane changes. Thus, chromosomes localizing to the periphery may presage embryonic death. Conversely, genes regulating nuclear organization may be altered. As a result, inefficient or failed organizational maintenance in these nuclei can give rise to abnormal growth and/or errors in cell division leading to embryonic loss. For example, coordination of mitosis is mediated by cell cycle checkpoints under genetic control. The spindle assembly checkpoint in particular is crucial for ensuring fidelity of chromosome segregation. Several well-established cell cycle genes, such as centromeric protein Nuf2, are conserved in yeast, nematodes and humans and are critical for proper chromosome segregation (Nabetani et al., 2001). Overexpression or down-regulation of human Cdc14a phosphatase directly causes aberrant chromosome positioning in daughter cells and can lead to genomic instability (Mailand et al., 2002
).
Proper orientation of the embryonic axis may be pivotal for early embryo cleavage and may affect chromosome segregation. The embryonic axis appears to be dependent on proper polarization of both chromatin and nuclear precursor bodies (Van Blerkom et al., 1995; Edwards and Beard, 1997
; Payne et al., 1997
). Edwards and Beard (1997)
have postulated that pronuclei rotate within the ooplasm to orientate their axis towards the second polar body in order to achieve proper orientation for subsequent cleavage. Disruption of this orientation may manifest in uneven cellular division or chromosomal segregation, leading to aneuploidy in the resultant embryo (Gianaroli et al., 2003
).
A caveat in our studies is that results are based on a single probe for each chromosome. Given that the observed localization of each chromosome may vary based on the chromosomal region to which the probes hybridize, further evaluation using probes that span the length of each chromosome will be necessary to assess more precisely nuclear location of each chromosome.
A further pitfall is that we evaluated blastomeres from patients having infertility. Ideally, a control group of reproductively normal patients would be studied. However, these normal individuals would not be expected to undergo PGD, thus, such embryos would be difficult to collect. Alternatively, analysis of blastomeres from subjects undergoing PGD for single gene disorders (presumed to be reproductively normal) may prove to be more suitable as controls. Case no. 16 (X-linked chorioderemia) is such a suitable control; however, only 16 blastomeres were available for assessment. A correlative limitation is that data were generated by pooling all infertility diagnoses together. A better design would be stratification by indication. Although the limited number of patients in this study precluded such analysis, preliminary evaluation showed no significant difference between the types of indications (data not shown).
That we found perturbations in nuclear localization in embryos from women undergoing PGD for a variety of indications (Table I) supports the biological plausibility of peripheral localization indicating cellular disturbance. Of special interest is our finding of this in women with both prior trisomy and repeated IVF failure. This bears on the phenomenon of recurrent aneuploidy, which appears responsible for repetitive clinical pregnancy losses in otherwise normal women. Recurrent aneuploidy is known to extend to preimplantation embryos. In nine couples with repeated abortions, Pellicer et al. (1999) found that 58.5% of the PGD embryos were aneuploid compared to 16.7% of PGD embryos from controls with normal reproductive histories. Our data also suggest that mechanisms regulating nuclear organization are altered in infertile couples. Further investigation is warranted to elucidate the underlying mechanism associated with altered localization of chromosomes in preimplantation embryos.
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Acknowledgements |
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References |
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Submitted on April 20, 2004; accepted on June 3, 2004.