NUCLEAR TRANSFER TECHNIQUES FOR VISUALIZATION OF CHROMOSOMES IN POLAR BODIES AND BLASTOMERES
Verlinsky Y., PhD, Kuliev A., MD, PhD, Ahshina M.., PhD
Reproductive Genetics Institute, 836 W.Wellington, Chicago, Illinois, USA
The introduction of the fluorescence in situ hybridization (FISH) technique allows chromosomes to be enumerated on interphase cell nuclei for testing of polar bodies and blastomeres. However, the number of chromosomes studied by FISH is limited to the number of chromosome-specific probes available. Even with the currently available methods for re-hybridization of interphase nuclei for the second and the third time, complete karyotyping will not be realistic in the near future. On the other hand, current developments in nuclear transfer techniques make it possible to convert the interphase nucleus of a single blastomere and second polar body (PB2) into metaphase, to perform preimplantation genetic diagnosis (PGD) by full karyotyping.
In contrast to PB2 and blastomeres, the first polar body (PB1) consists of metaphase chromosomes. According to our observation, immediately after extrusion, PB1 chromosomes are uncountable if analyzed. After 2—3h of in vitro culture, however, and during the following 2—3h, the individual PB1 chromosomes become recognizable and countable, with degeneration occurring 6—7h after extrusion. This makes PB1 particularly useful for spectral karyotyping (SKY), or application of whole chromosome-specific fluorescence probes or other chromosome segment-specific probes for testing of chromosomal translocations. This method for PGD of translocations of maternal origin has been applied in more than two dozen clinical cycles. Although the method resulted in a significant reduction of spontaneous abortions in the PGD cycles and led to the birth of more than a dozen unaffected babies, it is sensitive to malsegregation and/or recombination between chromatids. Therefore, visualization of PB2 chromosomes, in combination with PB1 FISH analysis, will ideally be required for accurate PGD of chromosomal abnormalities. However, it is known that the PB2 forms a nucleus and never transforms into metaphase. To visualize PB2 chromosomes, Modlinsky and McLaren transplanted the mouse PB2 into a fertilized egg, which in some cases transformed the PB2 nucleus into a haploid group of mitotic chromosomes, but the success rate was very low, and even when the chromosomes were visualized they were unsuitable for karyotyping.
We have developed various techniques for converting the PB2 into the metaphase stage. One of these approaches involved electrofusion of the mouse PB2 with intact and/or enucleated mouse zygotes, and resulted in PB2 nucleus transformation into the metaphase plate in 34% of cases. The same results were obtained by electrofusion of the PB2 with a foreign one-cell mouse embryo, with the proportion of metaphase plates reaching 65% when the recipient one-cell stage mouse embryo was enucleated. The other approach involved the treatment of the one-cell stage mouse embryos with okadaic acid (a specific inhibitor of phosphates 1 and 2A), leading to visualization of PB2 chromosomes in up to 80% of cases. The visualized PB2 chromosomes were unichromatid G1 premature condensed chromosomes of good quality, suitable for differential staining. However, in contrast to the mouse data, okadaic acid treatment of human PB2 led to further condensation of already pycnotic PB2 nuclei, so we developed a special technique to convert PB2 into metaphase chromosomes, which is described below.
Conversion of second polar body into metaphase chromosomes
The technique for removal of PB2 was described in a previous chapter, with the only modification here that the biopsy is performed in a medium without sucrose. The PB2 is introduced into oocyte cytoplasts, which are usually obtained by enucleation of metaphase II oocytes, which remained unfertilized after intracytoplasmic sperm injection (ICSI), or of those metaphase II oocytes that were matured for 24—48h in vitro from immature oocytes. Prior to enucleation, these ooplast recipients were incubated for 10-15 min in medium with lmg/ml cytochalasin D, 0.3mg/ml nocodazole and 0.5mg/ml Hoechst 33342. As shown in the procedure steps for oocyte enucleation, PB1 is removed together with the nucleus of the oocytes. Because the oocytes with failure of fertilization are used, special attention is paid to remove both the meiotic metaphase II spindle and the sperm chromosomes. The enucleated oocytes are then washed and transferred into culture medium for at least 1 h to recover, before PB2 injection, which is essentially the same as in ICSI. The pipette for intracytoplasmic PB2 injection is prepared in the same way as for biopsy tools. The resulting pipette has a tip broken perpendicularly, without any irregularities and with an inner diameter of 7-10 mm. In contrast to the biopsy tools, these pipettes are not flame-polished. The same microforge is used to bend the tool to the desired angle. Prior to use, the micropipettes are treated with non-ionic detergent (NP10).
PB2 is aspirated into an injection pipette, with care taken that the PB2 plasma membrane is broken, and the pipette is brought into the perivitelline space of the recipient cytoplast through the partial zona dissection (PZD) slit made during oocyte enucleation, and moved into the center of the cytoplast. Cytoplasm is aspirated into the pipette until the plasma membrane is broken, and the PB2 nucleus is expelled into the cytoplast. The reconstructed haploid embryos are cultured in standard medium for at least 1 h.
Oocyte activation is performed by electrofusion, with the use of custom-made electrostimulation apparatus in a fusion chamber consisting of two platinum wire electrodes glued to the bottom of a glass dish with a gap of 0.33mm. Electrofusion medium consists of 0.3 mol mannitol, 0.lmmol MgSO4, 0.05mmol CaCI2 and 0.5% polyvinylpyrrolidone, dissolved in HPLC-grade water. The pH of the medium is adjusted to 7.4 by titration with 0.1 mol/N NaOH.
The reconstructed embryos are checked every 30min starting at 24 h after activation, and fixed 45min after the disappearance of PB2 pronuclei. Alternatively, premature chromosome condensation of the resulting PB2 pronuclei in the reconstructed embryos is induced by 1-h exposure to okadaic acid (5—l0 mg/ml). To prevent embryo cytoplasm fragmentation, okadaic acid is diluted to the working concentration with phosphate-buffered saline containing 3mg/ml bovine serum albumin and 0.5mg/ml cytochalasin D. Hypotonic treatment of the embryos is avoided, to prevent overspreading of chromosomes. Because there is also no need to improve the spreading of chromosomes during fixation, the embryos transferred from the fixative to the slide are simply left to dry out.
The whole procedure is usually completed within 2 days of PB2 removal, and so can be realized well before the embryo transfer. The procedure is started by PB2 removal a few hours after oocyte exposure to sperm or ICSI, followed by its injection into enucleated oocytes, and electrostimulation. The following day the embryos without pronuclei are fixed or exposed to okadaic acid prior to fixation, should the PB2 pronucleus still be present. The method for karyotyping of the PB2 has currently been applied for the PGD of translocations.
Conversion of single blastomeres into metaphase chromosomes
A similar principle is used for visualization of individual blastomeres. Initially individual blastomere nuclei were transformed into metaphase chromosomes through blastomere fusion with enucleated human zygotes. However, the efficiency of this method was not high enough to be applicable for PGD. To improve the efficiency of the method, the individual blastomeres are currently fused with intact or enucleated mouse zygotes at the pronuclear stage, known to be at the S phase of the cell cycle.
Frozen mouse zygotes, from Charles River Laboratories (Wilmington, MA), may be used as recipient cytoplast to induce conversion of the blastomere nucleus into metaphase. Although initially mouse zygotes were enucleated, there is no need for this step for most of the cases, as mouse and human chromosomes may be clearly distinguished.
Although intact blastomeres may be inserted microsurgically into the perivitelline space, this has appeared to be traumatic, and is currently replaced by blastomere—zygote agglutination with phytohemagglutinin (Irvine Scientific, Santa Ana, CA). Before the procedure, the thawed mouse zygotes are freed of zonae pellucidae with acidic Tyrode's solution and pipetted through the flame-polished Pasteur pipettes with an internal diameter of 80 mm to separate the PB2. Using the flame-polished Pasteur pipettes with internal diameter of 100mm, blastomere-zygote pairs are brought together and agglutinated in 300mg/ml of phytohemagglutinin in protein-free human tubal fluid buffered with 20 mmol /l of HEPES in a four-well plastic dish (Nunc).
For electrofusion, blastomere-zygote pairs are oriented between electrodes by hand, with the final orientation achieved with alternating current (500kHz; 0.2 kV/cm for 2 /s). Cell fusion is induced with a single direct current pulse (1kV/cm for 500 ms), and the results are assessed in 20min.
When human blastomeres are fused with intact mouse zygotes, the heterokaryons entering mitosis are identified under the dissecting microscope. Because of the transparency of mouse cytoplasm, the disappearance of pronuclei and the formation of the joint metaphase plate are clearly visible. The heterokaryons with a persisting pronucleus are exposed for 1 h to 5 mmî1 of okadaic acid in phosphate-buffered saline containing 3 mg / ml of bovine serum albumin and 0.5 mg / ml of cytochalasin D. After 10-15 min of incubation in a hypotonic solution (0.1% sodium citrate and 0.6% bovine serum albumin) the resulting mitotic heterokaryons are fixed in a cold 3:1 solution of methanol and acetic acid in a four-well plastic dish. When the cytoplasm clears, heterokaryons are transferred onto slides and air-dried. Chromosome plates are assessed by phase contrast and then used for standard chromosome analysis. For FISH analysis the slides are pretreated with formaldehyde and pepsin (Visis Inc., Downers Grove, IL).
Although the overall success rate of the procedure is as high as 94%, its efficiency can be improved to almost 100% with experience. Similar results were obtained by Willadsen and colleagues, using bovine ooplasts for fusion with human blastomeres. Our data showed that some of the failures were simply due to the absence of the nucleus in biopsied blastomeres, or because the heterokaryons were fixed after they had already cleaved. It is also useful to perform blastomere biopsy no earlier than day 3 or day 4, to avoid the biopsy of two- and four-cell embryos, leading to accelerated heterokaryon cleavage. However, the success rate did not depend on whether mouse zygotes were enucleated before fusion with blas-tomeres. This allows simplification of the procedure by using intact mouse zygotes.
The procedure is quite simple and includes the following components. Mouse zygotes are thawed and freed of zonae pellucidae and PB2 1-2 h before electrofusion with human blastomeres. Four hours after fusion, heterokaryons are monitored for signs of the disappearance of pronuclei, and fixed at mitosis following hypotonic treatment. To avoid monitoring and a possible missing of mitosis, the heterokaryons may be cultured in the presence of microtubuli inhibitors, vinblastine or podophyllo-toxin. All the embryos left in the culture by the 9th hour after fusion are fixed following 1 h of pretreat-ment with okadaic acid.
This method has been applied for the PGD of paternally derived reciprocal translocations and for confirmation of the PGD of chromosomal abnormalities performed by PB1 and PB2 FISH analysis.