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Print Posted on 08/08/2017 in Fertility Treatment Options

Invasive Method of Embryo Selection Process: Embryo Biopsy in Close Focus

Invasive Method of Embryo Selection Process: Embryo Biopsy in Close Focus

Abstract: Invasive Method of Embryo Selection Process: Embryo Biopsy in Close Focus [an Embryology Dimension]. The present article focuses on representing, analyzing, comparing and discussing the peculiarities of the new method, proposed to evaluate embryo viability, its capacity to develop to blastocyst stage, and to implant and develop normally – EMBRYO BIOPSY. It is extremely vital to emphasize that variability of technical expertise in embryo biopsy, blastomere fixation, fluorescence in situ hybridization analysis, along with suboptimal laboratory quality control and inappropriate patient selection may impact preimplantation genetic diagnosis outcomes. That is why at the present stage, perfecting the existing systems of identifying high–quality embryo during embryo selection process and developing new higher–precision ones are among priorities in the theory and practice of improving expert systems for special purposes of morphological evaluation, which is a considerable challenge for embryologists. The special emphasis is given to the representation of the the process of embryo biopsy in close focus: zona pellucida opening for blastomere removal can be done by mechanical or chemical means, or by laser. Opening the zona pellucida is required irrespective of the stage of development.

INTRODUCTION

Embryo biopsy or preimplantation genetic diagnosis (PGD) is performed in couples at risk of genetic disease, to avoid transfer of embryos which are affected by a monogenic disease or which carry chromosomal aberrations in accordance with exclusive criteria. The oocytes’ nuclear maturity, normal fertilization, embryo quality, and embryo development are different steps that may jeopardize the number of embryos available for biopsy. As in all in–vitro fertilization (IVF) cycles, supernumerary non–affected good–quality embryos may be available after preimplantation genetic diagnosis (PGD). These embryos can be transferred or cryopreserved. Although preimplantation genetic screening (PGS) is widely offered, there are contradictory reports on the clinical merit of this procedure. Any gain from embryo selection following aneuploidy screening must significantly outweigh the impact of the procedure. Variability of technical expertise in embryo biopsy, blastomere fixation, fluorescence in situ hybridization analysis, along with suboptimal laboratory quality control and inappropriate patient selection may impact preimplantation genetic screening (PGS) outcomes.

In order for preimplantation genetic screening (PGS) to benefit patients, any gain from embryo selection following aneuploidy screening must significantly outweigh the impact of the embryo biopsy procedure. Removal of one or more blastomeres from the developing embryo has the potential to reduce embryo viability [19]. In addition, the culture conditions in which an embryo is exposed to are known to affect developmental viability [7].

One of the most important and unsolved problems in in–vitro fertilization treatment is to decide which embryos are more suitable to implant and therefore should be transferred and which embryos should be excluded. The development of preimplantation genetic diagnosis (PGD) techniques [40] has given many couples with a high risk of transmitting genetic pathology the chance to have a baby without the disease [38]. Allowing biopsied blastomeres to multiply in vitro will increase the number of cells available for analysis and thus improve the results of the genetic study [34]; moreover, PGD might be offered to a greater number of patients, increasing the range of indications. Also important is the fact that creating a hole in the zona pellucida (assisted hatching) might improve the implantation rates as previously demonstrated [17].

One criticism of this technique is the need to micromanipulate all embryos. This point, however, is not very important nowadays as many IVF centres worldwide are performing ICSI, PGD or assisted hatching, methods that are based on micromanipulation techniques.

Molecular cytogenetic analysis of interphase nuclei by fluorescent in situ hybridization (FISH) has been the most frequently used technique for the analysis of chromosomal abnormalities in human embryos. Since the advent of embryo biopsy and the application of FISH procedures to interpret chromosome aneuploidy in single blastomeres, the methodology has been continuously evaluated in an endeavour to minimize the detrimental effect on the embryo and optimize the outcomes for patients [19]. In order to reduce the number of embryos transferred, it is vital to have very efficient selection criteria to identify those embryos that are more likely to implant [embryos with high implantation potential] and develop into pregnancy. Moreover, it is also important to select spare embryos with the ability to implant for cryopreservation and future transfer.

A great number of methods have been suggested to evaluate and choose embryos suitable for future transfer. Routinely, morphological and development assessment [23; 30; 67] has been used to eliminate arrested or degenerate embryos with minimal potential for implantation. In addition, other methods have been proposed to measure several metabolic parameters of the embryos, for example: pyruvate uptake [20], O2 consumption [55], secretion of platelet activation factor, and the production of interleukin–1α [66]. These methods however, have not led to increased implantation and pregnancy rates, especially when more than three similar “good” (whatever the criteria used) embryos are present.

That is why at the present stage, perfecting the existing systems of identifying high–quality embryo during embryo selection process and developing new higher–precision ones are among priorities in the theory and practice of improving expert systems for special purposes of morphological evaluation, which is a considerable challenge for embryologists. In the connection with the above, the key objective of morphological evaluation is to establish which embryo score variables are the most essential for high–quality embryo selection process for further single embryo transfer (SET) at the early cleavage stage.

All the above determines formulating a deeply actual embryologic mission of our time, consisting in modelling or in other words in creating a funclional model and implementing it in practice, which explicates the timeliness of our research. The timeliness of our theoretical work can also be explained by the fact of its focusing on blastomere development after embryo biopsy because the scientific elite hasn’t even started creating the most accurate, transparent and inclusive general embryologic theory, which can represent the answer for one essential question: which of the identical embryos of good morphology, good cleavage rate and with adequate metabolic milieu should be selected for transfer?

This objective presupposes realizing the following tasks: (1) to distinguish the essentiality of embryo biopsy; (2) to represent the essence of the significance of mosaicism for subsequent embryo development; (3) to characterize the basic morphological characteristics [scoring] of the top–quality embryo which should be taken into consideration when selecting embryo for further transfer [based on the correct scientific interpretation of the fluorescent in situ hybridization (FISH) analysis as the integrative constituent of the preimplantation genetic diagnosis]; (4) to reveal the peculiarities of the zona pellucida opening by mechanical or chemical means, or by laser; (5) to evaluate if the use of a laser to open the zona pellucida in cases of preimplantation genetic diagnosis has any influence on the results of the biopsy procedure, further embryo development in vitro or the outcome after embryo replacement, compared with chemical zona drilling; (6) to describe how the result of embryo biopsy can be represented; (7) to compare the results of embryo biopsy after zona drilling using acid Tyrode medium or a laser and (8) to describe the blastomere development after biopsy.

(1)           WHY EMBRYO BYOPSY SOMETIMES IS SO ESSENTIAL? AND WHAT IS MOSAIC EMBRYO?

Post–zygotic chromosome segregation errors are very common in embryos after in vitro fertilization, resulting in mosaic embryos. However, the significance of mosaicism for the developmental potential of early embryos is unknown. Therefore, it is considered essential to assess chromosomal constitution and development of embryos from compaction to the peri–implantation stage [58].

Most of the current knowledge concerning the chromosomal constitution of preimplantation embryos comes from the analysis of cleavage–stage embryos by preimplantation genetic screening (PGS) performed 3 days after fertilization, when embryos are usually composed of 6–10 cells (blastomeres). Molecular cytogenetic analysis of interphase nuclei by fluorescent in situ hybridization (FISH) has been the most frequently used technique for the analysis of chromosomal abnormalities in embryos [58]. Data obtained by such studies have indicated that more than 50% of cleavage–stage embryos generated by in vitro fertilization (IVF) contain chromosomally abnormal cells [25]. These abnormalities may arise from an error during meiosis, resulting in a uniform abnormality present in all cells, or from segregation errors occurring during the first mitotic divisions. The latter event results in chromosomal mosaicism, defined as the coexistence of karyotypically distinct cell lineages derived from a single zygote [58].

Mosaic embryos can be composed of a mixture of chromosomally normal and abnormal cells or of abnormal cells with different abnormalities. Mosaicism has been reported to affect up to 91% of preimplantation embryos if all cells are investigated [3; 10; 16; 57; 72; 78; 81]. Studies using comparative genomic hybridization (CGH) and array comparative genomic hybridization (CGH) in preimplantation embryos, allowing the screening of all chromosomes, have confirmed the high prevalence of chromosomal mosaicism at this early stage of development and also demonstrated the high incidence of structural abnormalities [72; 78; 81].

These recent findings have changed the understanding of the cytogenetic processes occurring at the cleavage stage. However, the implications of chromosomal mosaicism for further embryonic development and implantation remain unclear. So far, the majority of the studies investigating the chromosomal constitution of blastocysts have suggested no definite selection against most of the chromosomal abnormalities observed at the cleavage stage [3; 8; 10; 22; 31; 32; 54; 62; 63]. Although there are reports of an increase in the proportion of blastocysts showing chromosomal mosaicism, compared with early cleavage–stage embryos [10; 63], the proportion of aneuploid cells within an embryo seems to decline towards the blastocyst stage [10; 62].

In spite of the high frequency observed in preimplantation embryos, a low percentage (0.3%) of aneuploidy is found at term birth [43]. Although up to 91% of preimplantation embryos are mosaic [72], the incidence of mosaicism in spontaneous miscarriage specimens is significantly lower (<10%). First trimester diagnoses in chorionic villi from viable pregnancies show an even lower incidence of mosaicism (1–2%) [52]. It therefore seems that the majority of mosaic embryos disappear prior to the period of first trimester, either due to a selection against mosaic embryos, or “normalization”, due to selection against abnormal cells within the embryo [52]. However, a ‘black box’ remains concerning events surrounding implantation [53].

At present, the scientific knowledge of the fate of the mosaic embryo is limited to the blastocyst stage at Day 6 post–fertilization. Clearly, if the significance of mosaicism for subsequent development is to be understood, events subsequent to this stage need to be elucidated. Therefore, the main aim of embryology’s experts was to assess the prevalence of mosaic embryos and how the chromosomal constitution of embryonic cells evolves from compaction to the peri–implantation period at Day 8 post–fertilization, using an in vitro implantation model.

FISH (fluorescence in–situ hybridization) analysis of embryos in order to assess the frequency of chromosomal mosaicism can be performed at three different stages of development (morula, preimplantation blastocyst and peri–implantation blastocyst). Perfect quality, frozen–thawed morula–stage embryos can be either fixed with all cells analysed or biopsied and allowed to develop in vitro. Reanalysis should be performed either at Day 5, in case of developmental arrest, or at Day 8. This approach allows the scientists to evaluate the chromosomal constitution of embryos during different stages of development and to determine how chromosomal constitution may influence the developmental capacity of embryos.

(2)           WHAT IS FLUORESCENCE IN–SITU HYBRIDIZATION (FISH) ANALYSIS? 

(2.1)       Pre–fluorescence in–situ hybridization (FISH) analysis’ procedures

Margarida Avo Santos, Gijs Teklenburg, Nick S. Macklon, Diane Van Opstal, G. Heleen Schuring–Blom, Pieter–Jaap Krijtenburg, Johanna de Vreeden–Elbertse, Bart C. Fauser, Esther B. Baart in their scientific study “The fate of the mosaic embryo: chromosomal constitution and development of Day 4, 5 and 8 human embryos” proposed perfect strategy for the most accurate performance of preimplantation genetic diagnosis (PGD). The recommended criteria for this procedure were described by the scientists as following: Prior to biopsy, embryos should be washed twice in calcium/magnesium–free medium (G–PGD medium) and then incubated in the same medium for 5 minutes at 37°C, allowing decompaction. The biopsy should be performed on the heated stage of a Nikon IX–70 microscope, equipped with micromanipulation tools. An infrared diode laser system with appropriate software should be used for dissection of the zona pellucida prior to biopsy. The retrieved blastomeres should be dissolved in lysis buffer and the nuclei were fixed on poly–L–lysine coated slides using methanol: acetic acid (3:1), as described before [2; 28]. Non–biopsied Day 4 and biopsied embryos that either were arrested at Day 5 or developed until Day 8 should be also dissolved using lysis buffer to remove the zona pellucida and the cytoplasm. Nuclei should be washed by gentle agitation of the lysis solution until clear and good spreading of nuclei was evident to minimize overlapping. Finally, a drop of methanol: acetic acid (3:1) should be added for fixation. Fixed nuclei from biopsied blastomeres and whole embryos should be viewed using a phase–contrast microscope and their location marked with a diamond pen. Preparations should be air–dried and stored at −20°C for up to 6 months prior to fluorescence in–situ hybridization (FISH) analysis [58].

(2.2)       Fluorescent in situ hybridization analysis: practical performance

For the most accurate representation of the practical performance of fluorescent in situ hybridization analysis, we consider it is essential to focus closely on scientific investigation: two rounds of five colour FISH were applied to single blastomeres or embryos. In the first round, FISH was performed for chromosomes 1, 7, 15, X and Y and in the second round for chromosomes 13, 16, 18, 21 and 22. The DNA probes used in the first round were centromere probes for chromosomes 1 [21], 7 [79], 15 [15], X [82] and a Y chromosome heterochromatin probe [50]. These were labelled as described previously [4], but using a BioPrime DNA labelling kit. The second round of FISH was performed using a commercial ready–to–use probe mix containing centromere probes. Slides were prepared according to standard protocols and hybridized using the same protocol as for the embryonic cells. Chromosome localization of the probes was verified on metaphase spreads and FISH signals were counted in 100 interphase nuclei. In addition, the positions of 10 individual nuclei were recorded and images were obtained after each round to check for persisting signals from the first round [58].

Hybridization was performed as described previously [4]. Per slide, 0.2 µl (single blastomeres) or 0.4 µl (whole embryos) of hybridization mixture was applied. Slides were examined with a Zeiss Axio Observer epifluorescence–equipped inverted microscope, using appropriate filters. An “EMBRYO MAP” was drawn for whole embryos, marking the location and attributing a number to the individual nuclei. Images of representative nuclei were captured with AxioVision imaging system. After the second round, images of the same nuclei were recorded and compared with those from the first round to ensure these had not persisted. Overlapping nuclei were excluded from the fluorescent in situ hybridization (FISH) analysis [58].

(2.3)       How fluorescent in situ hybridization (FISH) signal analysis can be scientifically interpreted after its practical performance?

For both rounds, the scientists used the scoring criteria previously published [60]. On the basis of the analysis of two blastomeres per embryo, they classified Day 4 embryos as normal (both nuclei showing the normal amount of signals for the chromosomes investigated), mosaic (one normal nucleus and one abnormal or each nucleus showing a different abnormality) or abnormal (both nuclei carrying the same abnormality). Day 4 embryos in which only one blastomere was analysed were classified as chromosomally normal or abnormal. After analysis of all the cells from each embryo on Days 4, 5 or 8, the scientists used the following definitions on the basis of the results obtained. To distinguish between true aneuploidy and FISH artefact, an abnormal cell line was defined as at least 10% of the nuclei showing the same chromosome abnormality. This threshold is frequently used in cytogenetics, if control material is lacking. Applying this criterion resulted in embryos being classified as normal if at least 60% of nuclei showed a normal chromosome constitution, and more importantly, if <10% of the nuclei showed the same chromosome abnormality. Embryos were classified as aneuploid if >90% of nuclei showed the same abnormality and if <10% of the nuclei showed a normal or different abnormal chromosome constitution. Embryos were classified as mosaic, if composed of cells with either a normal or an abnormal chromosomal constitution, with between 10 and 90% of the cells showing the same chromosomal abnormality. Chaotic embryos were classified as such when almost all the cells showed different and complex chromosomal abnormalities. Embryos with >90% haploid, tetraploid or triploid nuclei were classed as such [58]. However, the scientists considered the occurrence of some tetraploid cells as a normal phenomenon of in vitro cultured embryos [10; 31] and counted them as normal cells [58].

After reanalysis on Day 5 and 8, confirmation rates were calculated to investigate the reliability of the Day 4 diagnosis [58]. The chromosomal constitution diagnosed at Day 4 was considered to be cytogenetically confirmed [58], when the chromosome constitution of the investigated blastomeres was reflected in at least 10% of the cells within the embryo analysed on Day 5 or 8 [3]. After the proportions of chromosomally normal cells per embryo were calculated, statistical analysis using Pearson correlation coefficient allowed testing for statistical significance, with P < 0.05 considered significant [58].

Analysis of non–biopsied Day 4 embryos

In order to assess the chromosomal constitution and incidence of mosaicism of compacting embryos, cryopreserved Day 4 embryos were thawed (n = 21) and fixed within 10 minutes after removal from liquid nitrogen. None of the Day 4 embryos successfully analysed by FISH (n = 18) consisted of only normal cells. Only one (6%) embryo was classified as normal, with 3/15 (20%) differently abnormal cells. Of the remaining thawed Day 4 embryos, 15 (83%) were diagnosed as mosaic and two (11%) were chaotic. The average percentage of chromosomally normal cells per embryo was 55% (range 0–83%) [58].

FISH analysis of blastomeres and biopsied embryos

In total, 91 blastomeres from 53 compacting embryos were analysed by FISH. From 18 embryos, one cell was available for analysis and for 33 embryos the analysis was based on two blastomeres. Embryos 38 and 50 showed partial compaction by the time of biopsy, and therefore 3–cell were inadvertently biopsied. From the group of embryos where a single blastomere was biopsied, 39% were normal. In the group of embryos with two or three blastomeres biopsied, 40% were normal, 54% were mosaic and 6% were abnormal. FISH analysis of two blastomeres is consistent with the analysis of all cells from non–biopsied morulas, revealing a high rate of mosaicism, with more than half (54%) of the embryos identified as mosaic [58].

The scientists found almost all embryos to be mosaic at the morula stage. This high rate of mosaicism is reflected in their results obtained after biopsy and FISH analysis of one or two cells from Day 4 embryos. They found that the incidence of mosaic embryos decreased over time, with a significant decrease between Day 4 and Day 8 blastocyst stage. Moreover, the scientists found a positive correlation between the total number of cells in the embryo and the proportion of chromosomally normal cells in developing Day 5 and Day 8 blastocysts, but not in Day 4 morulas and embryos arrested before cavitation. Finally, the scientists observed that FISH diagnosis on one or two blastomeres of Day 4 embryos was not predictive of subsequent developmental potential [58].

The high incidence of chromosomal abnormalities in cleavage–stage embryos has been brought to light by several studies employing PGS–FISH in recent years [25]. However, it is still unclear which mechanisms lead to such high aneuploidy rates. The inefficiency of the cell cycle checkpoints during the first cleavage divisions (before embryonic genome activation at around the 8–cell stage) [13; 69] has been suggested as a possible cause for improper chromosome segregation [52]. According to this hypothesis, activation of the embryonic genome and initiation of compaction could lead to the establishment of functional cell cycle checkpoints, resulting in prevention of new errors and the developmental arrest of chromosomally abnormal cells and/or the entire embryo. In the present study, the scientists investigated the frequency of chromosomal abnormalities after compaction, to ascertain whether the incidence of chromosomal abnormalities decreases after presumable activation of the embryonic genome [58].

FISH analysis of morula–stage embryos revealed that the great majority of Day 4 embryos (83%) are mosaic according to the scientific definition, and no embryo consisted of normal cells only. Thus, the compaction stage does not provide a developmental barrier for chromosomally abnormal or mosaic embryos. These results are concordant with data from a previous study by the scientific group, where FISH analysis of 15 chromosomes in cryopreserved compacted morulas (n = 12) revealed all embryos to be mosaic [4]. Cryopreservation has been suggested to possibly induce chromosomal abnormalities after thawing and subsequent culture [2; 46; 64]. Therefore, both in the present and in the previous study by Baart et al. (2007), embryos were fixed immediately after thawing to avoid changes in chromosome constitution. Furthermore, work done by Bielanska et al. [10; 11] using fresh embryos showed that more than half of morula–stage embryos are mosaic (58%), when screening for nine chromosomes.

Analysis of biopsied Day 5 and Day 8 embryos

The scientists aimed at finding indirect evidence supporting the model where apoptosis of chromosomally abnormal cells is initiated upon differentiation at the blastocyst stage, but not before this stage of embryonic development. Therefore, they tested for a correlation between the total cell number and the percentage of normal cells within an embryo at Day 4, Day 5 (arrested and blastocyst), and Day 8.

After embryo biopsy and FISH analysis, embryos were reanalysed at Day 5 if arrested at that time or at Day 8 and the diagnosis compared with the Day 4 analysis. From the 24 embryos arrested before cavitation that were fixed on Day 5, 20 were successfully analysed after two rounds of FISH, showing that 14 (70%) were mosaic, 4 (20%) normal and 2 (10%) chaotic. In 8% of the nuclei, some overlap was observed and these were excluded from the FISH analysis. From the 24 blastocysts put into co–culture, 8 had degenerated by Day 8 and could not be retrieved. The remaining 16 embryos were successfully fixed on that same day. However, four embryos were excluded from further analysis due to failed hybridization for the second round of FISH. In total, 2859 blastomere nuclei from 12 embryos were analysed for 10 chromosomes, diagnosed 7 Day eight blastocysts (58%) as normal and 5 (42%) as mosaic. No Day 8 embryo consisted of only normal cells, nor did the scientists find an embryo with uniform aneuploidy. In each embryo, a range of cells with different abnormalities was found, with each abnormality present in cell numbers not reaching the 10% threshold of scientific definition for a mosaic cell line. For instance, in embryo 43, several cells with either a monosomy 1, monosomy 15, monosomy 16, monosomy 18, trisomy 13, monosomy 13, trisomy 21, monosomy 21 or monosomy 22 were present (Supplementary data, Table S2). These nine different abnormalities cannot all have originated from errors arising during the first cleavage divisions. This may therefore indicate that new segregation errors have occurred in this embryo after Day 4. The average percentage of chromosomally normal cells per embryo was 66% (range 35–84%) [58].

When comparing the percentage of mosaic embryos at Day 4 and in Day 5 blastocysts, the scientists observed a significant decrease by Day 5 of development. This suggests that a proportion of mosaic Day 4 embryos do not reach the blastocyst stage. The scientists reported a decrease in the incidence of mosaic embryos over time, with the lowest incidence observed at Day 8. The decrease of mosaic cases over time may be partially caused by their definition of mosaicism, which requires at least 10% of the cells to carry the same chromosome abnormality. This criterion is currently the best available method to distinguish true aneuploidy from FISH artefacts, when control material is lacking. However, it may lead the experts to underestimate the proportion of mosaic embryos at Day 8 and overestimate at Day 4 and Day 5. An example was embryo 46, where only 63% of the cells were found to be normal, but none of the abnormalities reached the 10% threshold. It is currently not known if this embryo could be diagnosed as normal, since the minimal proportion of normal cells needed for further normal development is yet unknown [58].

The presence of chromosomally abnormal cells does not exclude blastocyst development [11; 22; 32; 63]. However, the scientific data also suggest that a significant proportion of mosaic embryos undergo developmental arrest before reaching the blastocyst stage. According to the model proposed by Evsikov and Verlinsky (1998), if the number of aneuploid cells at the morula stage reaches a certain threshold level, there is self–elimination (arrest) of the whole embryo. However, embryos with a number of aneuploid cells below the threshold level develop further and reach the blastocyst stage [58].

Why the reanalysis of the mosaic embryos must be performed on Day 8? 

What is vital to mention [on the basis of the scientific investigation to represent the most accurate, transparent and inclusive information concerning the false diagnosis of the mosaic embryos] is the summarization of the interpretation of the FISH analysis on Days 5 (embryos arrested before cavitation) and 8 (developing blastocysts) and comparison of the observations with the results from the biopsy on Day 4. The Day 4 diagnosis was not predictive of the potential of the embryo to develop until Day 8. Reanalysis of biopsied embryos that either arrested before cavitation or developed until Day 8 showed similar cytogenetic confirmation rates (40% for arrested embryos and 36% for Day 8 embryos). However, the proportion of embryos with a false positive diagnosis on Day 4 (i.e. embryos that were diagnosed as abnormal or mosaic on Day 4 but classified as normal after reanalysis) was higher on Day 8 embryos (4/7 = 57%) than on arrested embryos analysed at Day 5 (2/12 = 17%). Conversely, the incidence of a false negative diagnosis was higher on Day 5, where 6/8 (75%) embryos diagnosed as normal on Day 4 were found to be mosaic on Day 5, whereas 1/4 (25%) Day 8 embryos were falsely diagnosed as normal. This was largely due to the increased incidence of chromosomally normal embryos on Day 8, with the percentage of embryos diagnosed as mosaics falling from 70% on Day 5 to 42% on Day 8. Reanalysis of the FISH results of Day 5 good–quality blastocysts analysed in a study by Baart et al. (2007) showed a lower percentage of chromosomally mosaic Day 5 blastocysts (58%) than observed in the present study (70%), where embryos arrested before cavitation were analysed at Day 5. Overall, we identified seven cases (7/50 = 14%) where at least one of the abnormalities observed likely originated during meiosis [58].

It is essential that for each reanalysed embryo, the proportion of chromosomally normal cells should be determined and correlated to the total number of cells. The scientists found a significant positive correlation (P = 0.034) between the total number of cells and the percentage of normal cells of Day 8 embryos [58].

The scientists found a significant positive correlation between the total number of cells and the percentage of chromosomally normal cells per embryo in Day 5 blastocysts and Day 8 peri–implanted embryos, but not in Day 4 morulas or embryos arrested before cavitation. The difference in the results for the two groups analysed at Day 5 (arrested embryos that failed to initiate cavitation and blastocysts) support the model. Thus, their scientific data provide indirect evidence that cavitation may be critical for the onset of a negative selection against abnormal
cells [31] and/or for the establishment of a growth advantage of the normal over the abnormal cells [22; 81].

Surprisingly, analysis of Day 8 embryos showed the persistence of high numbers of cells with different chromosomal abnormalities until this stage of development, although not falling within the range of scientific definition for mosaicism. The identification of numerous different segregation errors at Day 8 indicates that new abnormalities can arise after cavitation. This may explain the reported poor predictive value of FISH diagnosis on one or two blastomeres of Day 4 embryos. Out of the 50 embryos analysed, the scientists identified seven embryos (14%) where at least one of the abnormalities was likely to have been caused by a meiotic error [58].

Tetraploidy has been described as a normal phenomenon in embryonic trophoblast cells [29]. Furthermore, the occurrence of some tetraploid cells has been considered as a normal phenomenon of in vitro cultured
embryos [10; 31]. Therefore, we included tetraploid and near tetraploid cells into the group of chromosomally normal cells. However, the scientific definition may lead to an underestimation of the incidence of chromosomal abnormalities, as tetraploid cells can arise after aberrant cell division [14]. We have at the moment no method of distinguishing between these possibilities.

In conclusion, the scientific data suggest that a proportion of mosaic embryos undergo developmental arrest between compaction and cavitation. If the embryo continues to develop, reduced proliferation or cell death of aneuploid cells may be responsible for the increased proportion of chromosomally normal cells throughout development of human embryos. Although the biological implications of chromosomal mosaicism has not been well explored yet, emerging evidence illustrate that we may currently underestimate the impact on embryonic development and disease in later life [45].

(2)           THE PROCESS OF EMBRYO BIOPSY IN CLOSE FOCUS: PRACTICAL PERFORMANCE 

Most of scientific current knowledge concerning the chromosomal constitution of preimplantation embryos comes from the analysis of cleavage–stage embryos by preimplantation genetic screening (PGS) performed 3 days after fertilization, when embryos are usually composed of 6–10 cells (blastomeres).

Preimplantation genetic diagnosis (PGD) can be performed to avoid replacement of embryos affected by a monogenic disease or carrying chromosomal aberrations, in other words, it can be performed to exclude affected embryos through establishing strict exclusion criteria [71]. The diagnosis can be performed at different stages of embryo development. Zona pellucida opening for blastomere removal can be done by mechanical or chemical means, or by laser. Opening the zona pellucida (ZP) is required irrespective of the stage of development.

Before the introduction of preimplantation genetic diagnosis (PGD), different micromanipulation procedures had already been used to create an opening in the zona pellucida (ZP). These procedures were used to increase fertilization rates in cases of poor semen quality [33; 80] or to assist the hatching process [18; 49; 70]. The opening in the zona pellucida (ZP) was usually made either mechanically or chemically. The same methods of zona drilling have also been used with preimplantation genetic diagnosis (PGD) [39; 40; 76; 77].

In the last decade, the use of lasers as an alternative method of zona drilling has been evaluated and introduced into the IVF laboratory. The first lasers worked in the UV range and were contact–mode lasers [1; 51; 61]. These systems have several disadvantages, such as possible DNA damage due to absorption of ultraviolet (UV) light or the need for a complex set–up that delivers the laser energy directly to the target rather than being absorbed by aqueous solutions [68]. Relatively larger wavelengths are not absorbed by DNA or water, so that they can be used in a non–contact mode [68]. Lasers working in the infrared (IR) range are highly suitable in this respect. More recently, an application for assisted hatching has been published [6; 36; 56; 59], and the performance of laser zona drilling (LZD) was compared with other drilling procedures [5; 44]. So far, however, data on the use of this type of laser for zona drilling with a view to embryo biopsy for PGD are limited [12; 75].

(2.1)       Recommended pre–biopsy preparation procedures [embryo selection in accordance with exclusive criteria and preparation of the microtools for performing embryo biopsy]

Before performing embryo biopsy, the number of blastomeres should be counted and recorded for each individual embryo. In other cases, the term compaction should be used. Biopsy should not be performed in case of the following reasons: (I) very poor quality of the embryos (>50% of the embryos are filled with anucleated fragments) or (II) if none of the embryos reached the third mitotic division on day 3. Only embryos with <50% of their volume filled with anucleated fragments are considered suitable to undergo the biopsy procedure [48]. For performance of embryo biopsy, it is highly recommended to prepare the microtools. It is interestingly6 to note that according to recommendation, home–made microtools with the following characteristics should be prepared from washed and sterilized capillaries. The holding pipette should have an outer diameter of 100 µm and an opening of 25–30 µm. The biopsy pipette should have an inner diameter of 35–40 µm. The opening should be fire polished. The drilling pipette should have a more tapered shape, with an outer diameter of 10–12 µm and an opening of ∼5 µm. The microtools should be sterilized the day before the biopsy procedure [48].

(2.2)       Representation of the embryo biopsy procedure using acid Tyrode medium and laser for opening the zona pellucida (ZP)

For the most accurate, transparent and inclusive representation of the embryo biopsy procedure using acid Tyrode medium, we consider that the special emphasis should be given to the scientific study, where both methods are perfectly described through practical performance and its annotations into the corresponding protocols.

Dishes with 25 µl droplets of Ca2+–Mg2+–free medium covered with mineral oil should be prepared and incubated overnight in an environment of 6% CO2, 5% O2 and 89% N2. Just before the biopsy procedure, embryos selected for biopsy should be incubated in this medium for between 5 and 10 minutes, depending on the degree of compaction, unless there are obvious signs of total absence of compaction. The biopsy procedure should be performed in 50 µl droplets of HEPES–buffered Earle’s medium supplemented with 0.5% (w/v) human serum albumin (HSA) covered with mineral oil. In cases of zona drilling using medium (ATD), one extra droplet with acid Tyrode solution need to be present [48].

The microtools should be fixed and aligned on an inverted microscope equipped with Hoffman Modulation Contrast optics and a heated stage. A double tool holder carrying both the drilling and the aspiration pipette should be used in cases with acid Tyrode medium (ATD). Needles should be always replaced in cases of cell lysis. The embryo should be visualized and fixed on the holding pipette after aspiration of the acid into the drilling pipette. Drilling should be performed by releasing acid onto the the zona pellucida (ZP) between two blastomeres. Strong aspiration should be applied upon rupture of the the zona pellucida (ZP) [48].

In cases with laser zona drilling (LZD), embryos should be fixed on the holding pipette in a similar way. Two, or exceptionally three, pulses of 5–8 ms (1.48 µm) should be applied after correct positioning of the zona pellucida (ZP) using the target generator. Again, the opening should be made between two blastomeres [48].

After zona drilling by either acid Tyrode medium or laser, blastomeres should be aspirated gently, removed from the embryo and released into the medium. Biopsied embryos should be transferred to 25 µl droplets of G2.2 medium and cultured until the time of diagnosis and possible replacement. Two blastomeres should be aspirated from embryos containing at least seven cells. Exceptionally, a third blastomere should be aspirated in cases where lysis of one of the two other cells occurred. Only one blastomere should be aspirated from embryos with five or six blastomeres in some cases of fluorescence in‐situ hybridization (FISH) analysis [48].

The result of embryo biopsy can be represented in two different ways. The first is according to the definition of the ESHRE consortium on Preimplantation genetic diagnosis [42], the definition of successful biopsy is: “the removal of a cell without lysis such that the cell could be used for analysis”. The second way presents the ratio of intact blastomeres to the total number of aspirated blastomeres [48].

The diagnosis can be performed by polymerase chain reaction (PCR) in order to avoid replacement of embryos affected by a monogenic disease, or by FISH (translocation carriers or sexing) to avoid replacement of embryos with an abnormal chromosomal constitution or affected by an X–linked disease [71]. After diagnosis, the best quality embryos considered normal for the tested chromosomes or mutation, which have developed further between the biopsy and time of the diagnosis, should be replaced if selection is possible. Embryo replacement is performed on day 3 or day 4 after oocyte retrieval depending on the time necessary to obtain the results of the diagnosis [48].

The numbers of biopsied embryos, aspirated blastomeres and intact blastomeres, as well as the causes of lysis, should be summarized. Lysis of aspirated blastomeres after zona drilling do not occur when the laser is used, while ∼60% of the lysed cells after acid Tyrode medium (ATD) are a consequence of the drilling procedure only [48].

The actual time needed to create a hole in the zona pellucida (ZP) by blowing acid onto the zona or by activating the laser two or three times is in the same range for both procedures and usually takes between 10 and 20 s. In practice, one embryologist can perform all aspects of the biopsy procedure. The overall time required to perform the embryo biopsy procedure is in favour of laser zona drilling (LZD) because (I) changing of needles in cases of cell lysis occurred less frequently; (II) the alignment of two needles instead of three is easier and faster; and (III) the extra manipulations necessary to (re)fill the drilling needle with acid and the necessary precautions to avoid unnecessary blowing of acid are absent [48]. Using a laser for zona drilling means that embryo biopsy with laser zona drilling (LZD) takes less time. This advantage of laser zona drilling (LZD) has been confirmed by others [5; 44]. Nevertheless, differences in procedures [47] or differences in experience between experts may also influence outcome or the time needed.

(2.3)       Comparison of the results of embryo biopsy after zona drilling using acid Tyrode medium or a laser [in accordance with the article: “Comparison of the results of human embryo biopsy and outcome of PGD after zona drilling using acid Tyrode medium or a laser”, written by authors Joris H., De Vos A., Janssens R., Devroey P., Liebaers I., Van Steirteghem A. as a result of their scientific investigation]

The developmental stage and the quality of the biopsied embryos early on day 3 are similar in both groups (after zona drilling using acid Tyrode medium and after zona drilling using laser) [48].

A reduction in the number of blastomeres as a consequence of the biopsy procedure can be clearly observed, because of the higher damage rate after acid Tyrode medium (ATD) and the resulting removal of a third blastomere in embryos, therefore after the biopsy procedure, usually many embryos would be at 5–cell stage or at 6–cell stage on Day 3. There is no impact of the biopsy procedure on embryo quality. The patterns of development between the moment of biopsy and the evening of day 3 are similar in both groups after zona drilling using acid Tyrode medium and after zona drilling using laser), except for the degree of compaction [48].

After laser zona drilling (LZD), more embryos compact earlier than after acid Tyrode medium (ATD). Similar patterns in overall development can be observed. After acid Tyrode medium (ATD), most of the biopsied embryos have more than eight cells or are at the morula stage. This rate is very similar after laser zona drilling (LZD). The difference in initiation of compaction can be observed on the evening of day 3 and would be even more pronounced on day 4 (the prevalence would be given to biopsied embryos after laser zona drilling (LZD) [48]. What is essential to mention is that a tendency towards lower compaction rates after acid Tyrode medium (ATD) can be observed. When using sequential culture media, compaction may occur already on day 3 [74]. Zona drilling with acid Tyrode thus seems to influence embryo development in vitro more than laser zona drilling (LZD) does [48].

No single accurate, transparent and inclusive definition of success rates of embryo biopsy exists. It can be defined per embryo [42] or per individually aspirated blastomere. laser zona drilling (LZD) results in more intact cells than acid Tyrode medium (ATD). This difference is the result of the impact of acid Tyrode during the drilling procedure. Damage to blastomeres during aspiration into or expulsion from the biopsy pipette is similar in both groups (after zona drilling using acid Tyrode medium and after zona drilling using laser) [48]. Easier blastomere lysis after acid Tyrode medium (ATD) may be related to the influence of the acid solution [26]. Apart from technical failures related to the diagnostics procedure, lysis of blastomeres during embryo biopsy may be the cause of conflicting results when diagnosis is attempted on lysed cells [65].

It is essential to mention that safety aspects of laser zona drilling (LZD) are currently being discussed, and, in particular, thermal effects should be minimized. Being vital for outcoming results, these safety aspects should be always implemented in laser zona drilling (LZD) procedure. Therefore, short pulse durations (≤5 ms) and an appropriate distance from adjacent blastomeres (>8 µm) are recommended [27] to avoid damage. It is highly recommented that from the start of the laser zona drilling (LZD) procedure, similar safety measurements should be applied: experts should use short pulse times (5–8 ms) in an area between cells. In most cases, two pulses are enough to create a hole that is large enough for blastomere removal [48]. Even if membrane damage without lysis may occur to the cells close to the site of drilling, those cells should be removed and manipulated immediately for diagnosis [48]. Although the use of acid Tyrode is considered safe, a temporary but significant cytoplasmic acidification in oocytes following zona drilling has been demonstrated by Depypere and Leybaert (1994). Similar to laser zona drilling (LZD), acid Tyrode medium (ATD) may damage blastomeres close to the drilling site [73].

Consequently, it is possible to make the conclusions, that (1) preimplantation genetic diagnosis (PGD) is a complex procedure in which the embryo biopsy procedure is just one step in a cascade of events leading to the replacement of embryos healthy for the tested disease or chromosomal aberration. It is therefore very difficult to determine the impact of just the embryo biopsy procedure; (2) it was scientifically confirmed that the use of the 1.48 µm IR laser system results in an easier drilling procedure, but more important is the observation that fewer blastomeres are damaged; (3) laser zona drilling (LZD) does not influence further development in vitro and gives rise to similar pregnancy and implantation rates as after acid Tyrode medium (ATD); (4) Further follow–up is necessary to prove the safety of this laser zona drilling procedure.

(3)           BLASTOMERE DEVELOPMENT AFTER EMBRYO BIOPSY

Scientists Selmo Geber and Marcos Sampaio proposed a new method to evaluate embryo viability, its capacity to develop to blastocyst stage, and to implant and develop normally. They analysed the in–vitro development of isolated blastomeres, biopsied on Day 3 after fertilization, and compared it to the development of the original embryo within a Day 3 co–culture, in order to find a relationship that could show the embryo’s potential future development and so increase implantation rates [35]. This is the first known invasive method proposed to select embryos for transfer and presents a very close relationship with embryo development.

According to scientific recommendations, this invasive method proposed to select embryos for transfer should be performed at cleavage stage. Normally, fertilized embryos which had reached the 6– to 10–cell stages on Day 3 irrespective of grade are transferred into drops of HEPES–buffered medium in a Petri dish under mineral oil. Biopsy should be performed using a pair of micromanipulators in conjunction with an inverted microscope. Each embryo should be immobilized by suction with a flame–polished holding pipette held in one micromanipulator. The second micromanipulator with a double holder controlled a drilling pipette (internal diameter 10 μm) containing acid Tyrode’s solution (pH 2.2) and a sampling pipette (internal diameter 30 μm) containing buffered medium. The drilling pipette should be placed in close contact with the zona pellucida and a hole made with a controlled stream of acid Tyrode’s solution. Immediately the zona pellucida is penetrated this pipette should be removed, and the sampling pipette should be pushed through the hole. One or two cells judged to be at the equivalent of the 8–cell stage should be then removed by gentle suction. In all cases, an interphase nucleus must be observed in the isolated blastomeres [35].

To present the most accurate information concerning blastomere development after embryo biopsy, we would like to exemplify it by the experimental investigation, which was done by Selmo Geber and Marcos Sampaio. The biopsied embryos and blastomeres were co–cultured in 20 μl drops of Earle’s medium supplemented with 10% heat–inactivated maternal serum under mineral oil in a Petri dish at 37°C under a gas phase of 5% CO2 in air. The embryos and blastomeres were assessed daily for morphological development until day 6. Cavity formation was considered when fluid began to accumulate either intracellularly or in intercellular cavities between 2 or more cells. Cell number at the blastocyst stage was counted by Giemsa staining and the nuclei of biopsied blastomeres were labelled either by Giemsa or by vital labelling (Hoechst 33342) as described by Geber et al. (1995b). Cell numbers were estimated on the assumption that they were equivalent to the number of nuclei counted. Results: a total of 66 normally fertilized embryos which reached the 6– to 10–cells stage at day 3 were biopsied. Forty–four embryos had one cell biopsied and 22 had two cells biopsied [35].

Post–biopsy embryo development

A total of 33 (50%) biopsied embryos reached the blastocyst stages on Day 5 or Day 6. Twenty–one blastocysts developed from the 44 embryos which had one cell removed (47.7%) and 12 developed from 22 embryos which had two cells removed (54.5%). The remainder arrested at earlier stages. Sixteen out of these 33 blastocysts (48.5%) hatched out from the zona pellucida on day 5 (n = 3) or day 6 (n = 13) [35].

Blastomere development

Approximately 64% of the isolated blastomeres divided over 3 days in culture, i.e. 42 out of the 66. Among the rest (24 out of 66), isolated blastomeres failed to divide over the same period. In 33 cases, out of 66 biopsies (50%) the isolated blastomere cavitated within the 3 days of culture. Cavitation and division were observed in 30 out of the 66 biopsied blastomeres (45.5%). Of the blastomeres taken from embryos that developed to the blastocyst stage, 88% divided and 79% cavitated. In the group of blastomeres biopsied from embryos that subsequently arrested, 39% divided and only 21% cavitated. Nine per cent of blastomeres from the first group and 55% from the second group neither divided nor cavitated. In the first group 76% of the blastomeres divided and cavitated simultaneously, and in the second group 15% [35].

The scientists found that the proportion of blastomeres, biopsied from embryos that developed to blastocyst stage, that cavitated was significantly higher than that biopsied from embryos that arrested (P < 0.001). When they considered blastomere division, we also found a statistically significant difference between those biopsied from embryos that reached the blastocyst stage, and from arrested embryos (P < 0.001). A statistically significant difference was also found when the scientists compared the blastomeres that either cavitated and divided, between the groups of blastomeres originated from embryos that reached blastocyst stage and from embryos that arrested (P < 0.001). When analysing the blastomeres that neither divided nor cavitated the scientists also found a statistically significant difference (P < 0.001). More blastomeres divided cavitated and divided/cavitated when biopsied from embryos that reached blastocyst stage than those biopsied from embryos that arrested. Moreover, more blastomeres neither divided nor cavitated when biopsied from embryos that arrested [35].

One blastomere cavitated only (3%) and four divided only (12%), after 3 days in culture, in the group of blastomeres biopsied from embryos that reached blastocyst stage. In the other group, two blastomeres cavitated only (6%) and eight divided only (24%) [35].

The potential future development of the Day 3 biopsied embryos was calculated using the odds ratio. Embryos whose biopsied blastomeres showed cavitation after 3 days in culture were 13.8 times more likely to develop to the blastocyst stage. Embryos whose biopsied blastomeres presented cell division were 11.1 times more likely to develop to the blastocyst stage. Embryos whose biopsied blastomeres showed either cavitation or division were 17.5 times more likely to develop to blastocyst stage. Finally, embryos whose biopsied blastomeres did not cavitate or divide were 12.0 times more likely to arrest [35].

The data were also analysed with logistic regression for multivariate analysis. The results demonstrated that cell division and cavitation are significant factors for predicting the probability of embryos to develop to the blastocyst stage. Using this model, the scientists were able to estimate the probability of an embryo to develop to blastocyst stage. Embryos that had biopsied blastomeres showing cell division and cavitation had an 81.6% probability of developing to the blastocyst stage. If cavitation was the only observed phenomenon, the probability was 50%; if cell division was the only phenomenon, 37.5%; and if none of them occurred, 11.9% [35].

The experimental scientific investigation, which was done by Selmo Geber and Marcos Sampaio is the first published study that shows an invasive method to predict embryo development and to select the most suitable embryos to implant after IVF–embryo transfer. In this study, the scientists show a very close relationship between embryo development and its biopsied blastomere. This new technique can be used to select the embryos for transfer in the cases where many good morphology embryos are available and also for preimplantation genetic diagnosis (PGD). Moreover, the scientists established the possibility to evaluate whether it is worth keeping the remaining embryos frozen. In conclusion, it is worth being emphasized that that embryo biopsy for selection of the embryos for transfer can improve implantation rates after fresh or freezing–thawing good prognosis cycles [35].

CONCLUSION

The development of preimplantation genetic diagnosis (PGD) techniques [40] has given many couples with a high risk of transmitting genetic pathology the chance to have a baby without the disease [38]. Allowing biopsied blastomeres to multiply in vitro will increase the number of cells available for analysis and thus improve the results of the genetic study [34]; moreover, PGD might be offered to a greater number of patients, increasing the range of indications. Also important is the fact that creating a hole in the zona pellucida (assisted hatching) might improve the implantation rates as previously demonstrated [17]. One criticism of this technique is the need to micromanipulate all embryos. This point, however, is not very important nowadays as many IVF centres worldwide are performing ICSI, PGD or assisted hatching, methods that are based on micromanipulation techniques. It is well known that several factors might influence the results of IVF–embryo transfer and pregnancy and implantation rates. The two most important targets for research are the endometrium and the quality of the embryos suitable for transfer. For the latter, several selection criteria have been proposed [20; 23; 30; 55; 66; 67] to determine which embryos are more suitable to implant and develop. Traditionally morphological criteria have been mostly used, which are based on the number of blastomeres (cleavage rate), size and shape of the blastomeres and the amount of anuclear fragments. It has already been demonstrated that the majority of pregnancies result from the transfer of good morphology embryos with the expected number of blastomeres; however, only a few of those embryos implant and develop successfully.

Further retrospective analysis of the scientific studies concerning the issues, described in the objective of the present article, has shown that:

(1) preimplantation genetic diagnosis (PGD) is a complex procedure in which the embryo biopsy procedure is just one step in a cascade of events leading to the replacement of embryos healthy for the tested disease or chromosomal aberration. It is therefore very difficult to determine the impact of just the embryo biopsy procedure.

(2) post–zygotic chromosome segregation errors are very common in embryos after in vitro fertilization, resulting in mosaic embryos. However, the significance of mosaicism for the developmental potential of early embryos is unknown. Therefore, it is considered essential to assess chromosomal constitution and development of embryos from compaction to the peri–implantation stage and most of the current knowledge concerning the chromosomal constitution of preimplantation embryos comes from the analysis of cleavage–stage embryos by preimplantation genetic screening (PGS) performed 3 days after fertilization, when embryos are usually composed of 6–10 cells (blastomeres). Molecular cytogenetic analysis of interphase nuclei by fluorescent in situ hybridization (FISH) has been the most frequently used technique for the analysis of chromosomal abnormalities in embryos [58]. Data obtained by such studies have indicated that more than 50% of cleavage–stage embryos generated by in vitro fertilization (IVF) contain chromosomally abnormal cells [25]. These abnormalities may arise from an error during meiosis, resulting in a uniform abnormality present in all cells, or from segregation errors occurring during the first mitotic divisions. The latter event results in chromosomal mosaicism, defined as the coexistence of karyotypically distinct cell lineages derived from a single zygote [58]. Mosaic embryos can be composed of a mixture of chromosomally normal and abnormal cells or of abnormal cells with different abnormalities. Mosaicism has been reported to affect up to 91% of preimplantation embryos if all cells are investigated [3; 10; 16; 57; 72; 78; 81]. Studies using comparative genomic hybridization (CGH) and array comparative genomic hybridization (CGH) in preimplantation embryos, allowing the screening of all chromosomes, have confirmed the high prevalence of chromosomal mosaicism at this early stage of development and also demonstrated the high incidence of structural abnormalities [72; 78; 81].

(3) FISH (fluorescence in–situ hybridization) analysis of embryos in order to assess the frequency of chromosomal mosaicism can be performed at three different stages of development (morula, preimplantation blastocyst and peri–implantation blastocyst). Perfect quality, frozen–thawed morula–stage embryos can be either fixed with all cells analysed or biopsied and allowed to develop in vitro. Reanalysis should be performed either at Day 5, in case of developmental arrest, or at Day 8. This approach allows the scientists to evaluate the chromosomal constitution of embryos during different stages of development and to determine how chromosomal constitution may influence the developmental capacity of embryos. The definition of mosaicism requires at least 10% of the cells to carry the same chromosome abnormality.

(4) it was scientifically confirmed that the use of the 1.48 µm IR laser system results in an easier drilling procedure, but more important is the observation that fewer blastomeres are damaged; laser zona drilling (LZD) does not influence further development in vitro and gives rise to similar pregnancy and implantation rates as after acid Tyrode medium (ATD); Further follow–up is necessary to prove the safety of this laser zona drilling procedure.

(5) it was proved that there is a very close relationship between embryo development and its biopsied blastomere.

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