Pregnancies following blastocyst stage transfer in PGD cycles at risk for ß-thalassaemic haemoglobinopathies

G.A. Palmer1,3, J. Traeger-Synodinos2, S. Davies1, M. Tzetis2, C. Vrettou2, M. Mastrominas1 and E. Kanavakis2

1 Embryogenesis Fertility Clinic, Kifissias Avenue, Athens 15125 and 2 Medical Genetics, Athens University, St. Sophia's Children's Hospital, Athens 11527, Greece


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Discussion
 References
 
BACKGROUND: Preimplantation genetic diagnosis (PGD) usually involves blastomere biopsy 3 days post-insemination (p.i.), followed by genetic analysis and transfer of unaffected embryos later on day 3 or 4. We evaluate a strategy involving embryo biopsy on day 3 p.i., genetic analysis on day 4 and, following culture in blastocyst sequential media, transfer of unaffected embryos on day 5 p.i. METHODS: PGD cycles were initiated in 15 couples at risk of transmitting ß-thalassaemia major. Oocyte retrieval and ICSI were performed according to standard protocols. Embryo culture used blastocyst sequential media. Embryos were biopsied on day 3 p.i. using acid Tyrode's for zona drilling, and the single blastomeres were genotyped by a protocol involving nested polymerase chain reaction and denaturing gradient gel electrophoresis analysis. RESULTS: Forty of 109 (37%) embryos biopsied on day 3 p.i. developed to blastocysts by day 5 p.i., with at least one blastocyst available for transfer in 12 cycles (80%). Genotype analysis characterized 51/109 (47%) embryos unaffected for ß-thalassaemia major, of which 28 were blastocysts. Transfer of 37 day 5 p.i. embryos (blastocysts and non blastocysts) initiated eight clinical pregnancies. Implantation rate per embryo transferred was 12/37 (32%). CONCLUSIONS: Embryo biopsy on day 3, followed by delayed transfer until day 5 p.i. offers a novel and effective strategy to overcome the time limit encountered when performing PGD, without compromising embryo implantation.

Key words: ß-thalassaemia/blastocyst stage transfer/Ca2+Mg2+ free medium/preimplantation genetic diagnosis


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Discussion
 References
 
In Greece, ~10% of the population are carriers for ß-thalassaemia and related haemoglobinopathies, and our experience indicates that many couples who have an unsuccessful reproductive history may prefer a diagnostic procedure which avoids the termination of an affected pregnancy identified by prenatal diagnosis. Preimplantation genetic diagnosis (PGD) offers such a possibility through the selection of unaffected IVF embryos for transfer (Handyside et al, 1992Go). PGD usually involves blastomere biopsy from cleavage stage embryos, and for monogenic disorders, subsequent diagnosis applies protocols based on the polymerase chain reaction (PCR). Several approaches for the genetic diagnosis of ß-thalassaemia in single cells have been described (Monk et al., 1990; Varawalla et al., 1991Go; El-Hashamite et al., 1996Go; Ray et al., 1996Go; Kuliev et al, 1999Go) but clinical application has been limited. We established a genotyping method based on denaturing gradient gel electrophoresis (DGGE), which provides a reliable method for detection of the embryonic genotype (Kanavakis et al., 1999Go; Vrettou et al., 1999Go). Embryo biopsy and transfer is usually performed on the day 3 p.i., although pregnancies after day 4 embryo transfer have been reported (Grifo et al., 1997Go). The merits of blastocyst stage over cleavage stage embryo transfer have been extensively debated (Tsirigotis, 1988Go; Edwards and Beard, 1999Go) and recent reports indicate improved pregnancy rates and high implantation potential following further embryo culture (Gardner et al., 1998Go). We present the results from 15 PGD cycles (15 couples) in which we biopsied cleavage embryos on day 3 p.i. and then used blastocyst sequential media to facilitate development of blastocysts for blastocyst stage transfer. Eight pregnancies resulted, providing support for the view that this strategy has the advantage of extending the time allowed for embryo biopsy and genetic analysis for clinical PGD cycles without compromising pregnancy outcome.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Discussion
 References
 
Patient history and treatment.
The partners in all 15 cycles were heterozygous for ß-thalassaemia, {delta}ß thalassaemia or sickle cell anaemia (HbS) as confirmed by haematological analysis and DNA characterization (Kanavakis et al, 1997Go) (Table IGo). They received counselling and subsequently initiated IVF/PGD treatment. All couples had a history of infertility and entered the IVF programme expressing a wish to avoid an affected pregnancy (Table IIGo). Patients received HMG stimulation (Metrodin HP; Serono, Bari, Italy). The GnRH agonist (Suprafact; Hoechst, Germany) was administered in the long (down regulation) protocol. Follicular development was monitored by ultrasonograghy and estradiol concentrations. HCG (Profasi; Serono) was administered (10 000 IU), according to our IVF protocol, when there were three or more follicles measuring >=17 mm in diameter and estradiol concentration was >3500 pmol/l. Luteal phase support was with 600 mg progesterone (Utrogestan; Bestins Iscovesco, France) administered vaginally from the day of oocyte retrieval until the pregnancy test.


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Table I. Parental ß-globin mutations
 

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Table II. PGD for ß-thalassaemia; outcomes of day 5 transfer
 
Oocyte retrieval, insemination and embryo culture
Oocyte retrieval was performed transvaginally, ~36 h after HCG administration, using ultrasound guidance. Cumulus oocyte complexes were identified in the laboratory and placed in 1 ml pre-equilibrated IVF-50 Medium (Scandinavian IVF Sciences AB, Gothenburg, Sweden). ICSI was performed to eliminate contamination by sperm when performing subsequent embryo biopsy. The ICSI protocol was performed as previously described (Van Steirteghem et al., 1993Go). Culture of embryos was in incubators at 37°C and in a humidified atmosphere of 5% CO2 under mineral oil [Sigma-Aldrich (OM) Ltd, Greece]. Oocytes exhibiting two pronuclei (PN) 16 h post-insemination were considered to be normally fertilized and were placed in pre-equilibrated G1.2 medium (Scandinavian IVF Sciences) in 4-well Nunclon dishes (Nalge Nunc International, Roskilde, Denmark). After embryo biopsy the embryo was rinsed and placed in G2.2 medium (Scandinavian IVF Sciences) until embryo transfer.

Embryo biopsy
Approximately 68 h following insemination the quality of the embryos was evaluated by observation using a binocular microscope (Nikon). Embryos were then scored (Dawson et al., 1995Go). Only embryos of good morphology were used (grade 1 or 2). Biopsies were performed on day 3 p.i. when the majority of embryos had reached 6–8 cells. Biopsies were undertaken using a Nikon diaphot inverted microscope equipped with warming stage and Narishige micromanipulater. All micropipettes were manufactured by Cook IVF. The aspiration pipette had an internal diameter of 40 µm, the zona drilling pipette had an internal diameter of 5 µm and the holding pipette an external diameter of 110 µm with a 20 µm internal diameter at the tip. All embryos were pre-incubated in Ca2+/Mg2+ free medium (EB10; Scandinavian IVF Sciences) for 10–15 min prior to embryo biopsy. Embryos were placed individually in 50 µl drops of HEPES buffered Earle's Balanced Salt solution (EBSS; Gibco BRL, Life Technologies, USA) under mineral oil. While the embryo was immobilized by suction to the holding pipette, a hole was created in the zona pellucida by acidic Tyrode's solution (ZD10; Scandinavian IVF Sciences). The drilling pipette was removed and replaced by an embryo biopsy pipette and a single blastomere was aspirated from the embryo by gentle suction into the surrounding medium. After this procedure the biopsied embryos were placed in separate numbered wells of Nunclon dishes containing G2.2 media.

Genotype analysis
Following biopsy each single blastomere was placed directly into a 0.2 ml Eppendorf tube in 10 µl of double-distilled sterile water that was overlaid with mineral oil (all DNAse and RNAse free). A total of 5 µl of PCR-grade Proteinase K (Roche Molecular Biochemicals, Mannheim, Germany), diluted in double-distilled water was added to the 10 µl of water containing the selected cell, to a final concentration of 50 µg/ml. To lyse the blastomeres, the Proteinase K was activated by incubation at 37°C for 1 h followed by 65°C for 10 min, and inactivated by heating to 95°C for 10 min (Vrettou et al., 1997). A first round of amplification was carried out following the addition of 35 µl of PCR premix (50 µl final PCR reaction volume) containing 0.2 mmol/l of dNTP's, 1.5 mmol/l MgCl2, 1 µmol/l of each primer for Region A or Region B according to the parental genotypes (see Figure 1Go) and 4 IU of amplitaq gold in buffer provided by the manufacturer. PCR cycling times were 95°C for 8 min, 96°C for 2 min followed by 20 cycles of 96°C for 30 s, 60°C for 40 s, 72°C for 30 s and 20 cycles of 95°C for 30 s, 60°C for 20 s, 72°C for 30 s. Following the first PCR, nested PCR reactions were carried out to produce amplicons suitable for analysis using DGGE as follows: 1 µl aliquot of the first PCR reaction was re-amplified in a 50 µl volume containing a pre-mixed solution of buffer/dNTPs/MgCl2/Taq polymerase provided by the manufacturer (Supermix; Gibco BRL Life Technologies) and 0.4 µmol/l of each ß-gene PCR primer, which were selected according to the mutations under investigation in the PGD cycle (Figure 1Go). PCR cycling conditions included 95°C for 2 min followed by 30 cycles of 95°C for 30 s, 58°C for 20 s and 72°C for 30 s. Following the nested PCR, amplification products were checked by agarose gel electrophoresis and analysed by DGGE as previously described (Kanavakis et al., 1997Go) (Figure 2Go). Precautions against contamination were most stringent. Manipulation of cells and PCR set-up were carried out in separate UV-treated laminar flow hoods, with dedicated equipment. PCR set-up employed exclusive PCR pipettes and pipette tips with filter. Three negative controls (lysis mixture only) and two blanks were included through the first round PCR and two additional blanks were included in the nested PCR set-up in all sets of PGD analyses.



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Figure 1. PCR strategy for amplification of ß-globin gene sequences. Position of ß-thalassaemia mutations carried by the 15 couples are indicated by number codes above the gene (for mutation codes see legend to Table IGo). Mutation 5 (IVSII-745C>G) was outside the region covered by the PGD genotyping method, but linked polymorphisms in Fragment 3 were used for indirect genotyping in for this mutation and in those cases with {delta}ß- thalassaemia deletions. * = primers with `GC clamp' at the 5' end (5'-CGCCCGCCCCGCCCCCGTGCCC CCCGCGCCGCCCGCCCCGCCCCC-3').

 


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Figure 2. Image of DGGE gel stained with ethidium bromide and viewed with ultraviolet light. Both parents (couple number 11) had the IVSI-nt110(G>A) ß-thalassaemia mutation. Lane 8 is one of the heterozygous parents; lanes 1–7 are from single blastomere samples from seven separate embryos. For the IVSI-nt110(G>A) mutation the lower homoduplex band (Hm) represents the normal allele (N); the upper homoduplex band (Hm) represents the pathological allele (M); heteroduplexes bands are above these. Only embryos with indication of at least one normal homoduplex were deemed suitable for transfer i.e. lanes 2, 3, 4, 5 and 7.

 
Embryo transfer
Embryo development was categorized at transfer on day 5 p.i. (~120 h) as follows: poor (little or no development since biopsy); morula; cavitating (showing signs of a cavity); expanding blastocyst; expanded blastocyst; hatching blastocyst; hatched blastocyst (Table IIIGo). Embryos diagnosed as unaffected for thalassaemia major were transferred to the recipient using a Wallace catheter (Sims Portex Ltd., Hythe, UK).


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Table III. Development of embryos biopsied
 
Results

A total of 214 cumulus–oocyte complexes were retrieved from the 15 patients, of which 184 were at metaphase II (86%) and subjected to the ICSI procedure. A total of 119 (65%) showed two pronuclei on day 1 p.i. (Table IIGo). Embryo biopsy was performed on 109 of these embryos (Table IIGo) at the cleavage stage (4–8 cells), the remaining were deemed unsuitable for embryo biopsy due to embryo fragmentation and/or slow development (10/119; 8.4%). Embryos were biopsied 68–75 h p.i., however, only 63% (69/109) embryos had developed to the stages of 8-cells or more. The remaining embryos (40/109; 37%) were at the 4–7 cell stage even after delaying biopsy to the maximum permissible time. The number of embryos after embryo biopsy reaching the blastocyst stage was 40/109 (37%). The remaining were morula/early cavitation stages (55/109; 50%) or had undergone little or no development after the biopsy (14/109; 13%). From the embryos biopsied at 8-cells or more, 34/69 (49%) developed to blastocysts, but blastocyst development from embryos biopsied at slower developmental stages was only 6/40 (15%). Twenty-seven of all embryos biopsied (24.7%) had begun to hatch or had hatched at the time of embryo transfer ~120 h after insemination (Table IIIGo).

From a total of 109 embryos biopsied 51(46.7%) were deemed unaffected for ß-thalassaemia major (homozygous normal or heterozygous carrier) by the presence of a normal band on DGGE analysis. This number included 28 blastomeres. A total of 37 unaffected embryos was transferred (2.4 embryos transferred per patient). All patients had embryo transfer on day 5 p.i. and 12 (80%) patients had at least one unaffected blastocyst to transfer. A maximum of four embryos was transferred if available. The implantation rate per embryo transferred was 32% (12/37) (Table IIGo). Blastocyst-only transfers occurred in 7/15 cases (46.6%). Five became pregnant with an implantation rate per embryo transferred of 8/17 (47%). To maximize the chance of pregnancy a mix of blastocysts and slower developmental stages were transferred in the remaining eight cycles. In five of these patients (33.3%) the transfer cohort consisted of blastocysts and morulae/cavitating embryos. Three became pregnant (embryo implantation rate 4/14; 28%). No pregnancies were observed in the three patients with no blastocysts available for transfer (0/6). These patients had a mix of morulae and cleavage stage (possibly arrested) embryos.

The clinical pregnancy rate was 8/15 (53%). Three healthy children have been born, and three ongoing (one awaiting confirmation of genotype by amniocentesis). One pregnancy resulted in a missed abortion at 8 weeks and one pregnancy was terminated after chorionic villus sampling (CVS) at 12 weeks revealed the child was affected with ß-thalassaemia major.


    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Discussion
 References
 
One major problem of current PGD procedures is the limited time available for diagnosis while retaining embryo viability. Usually embryos are biopsied early on day 3 p.i. and transferred later the same day. Although the time needed for genetic analysis using DGGE is capable of giving a result within 24 h (day 4 p.i.) with reported pregnancies (Kanavakis et al., 1999Go) the decision was taken to biopsy throughout the day 3 p.i., to allow a maximum number of embryos to reach the 8-cell stage, with genotype analysis on the following day and transfer on day 5 p.i. This strategy allowed the embryo biopsy to be performed at a time more dependant on embryo development and, in addition, allowed genetic analysis to be arranged during more acceptable working hours.

While facilitating embryo biopsy, it has been reported that prolonged exposure to Ca2+/Mg2+ free medium is detrimental to embryo development (Ducibella et al, 1979), but we observed no adverse effect with 10–15 min exposure prior to embryo biopsy and report ongoing pregnancies and healthy births in accordance with previous observations (Kahraman et al., 2000Go).

A limitation to further embryo culture has always been the number and viability of blastocysts available for transfer. Recent studies have indicated that it is possible to obtain viable blastocysts in vitro without the need for feeder cells (Jones et al., 1998Go). Reports of high pregnancy rates and implantation potential have stirred debate over a potential role for blastocyst stage transfer in IVF (Tsirigotis, 1998Go; Edwards and Beard, 1999Go; Jones and Trounson, 1999Go). The emergence of commercially available medium (e.g. G1.2/G2.2; Scandinavian IVF Sciences and Blast-assist; Medicult) has led to an increasing number of clinics transferring embryos at the blastocyst stage, although there is a need for follow-up on children born, since blastocyst stage transfer has raised questions about the sex ratio (Ménézo et al., 1999Go) and fetal abnormalities in animal studies (Behboodi et al., 1995Go).

However, a recent randomized trial has shown no advantages over day 3 transfers (Coskun et al., 2000Go) but one obvious advantage of culturing to the day 5 p.i. is the increased time to execute PGD procedures. Biopsy of one or two blastomeres from an 8-cell embryo does not interfere with in-vitro development of embryos (Hardy, 1990) and therefore the outcome of IVF/PGD cycles should not be compromised by further embryo culture to day 5 p.i. A biopsy later on day 3 p.i. allows the operator to conduct the biopsy at an optimum time with regard to embryo development. Despite that, only 63.3% of embryos reached 8-cells or more at the time of biopsy in this study. In practice, PGD offers little choice in quality of embryos for biopsy, and in some cases embryos were also biopsied at the 4- and 5-cell stage, despite reports that this retards embryo development (Tarin, 1992). Embryo biopsy of a single blastomere was not detrimental to developmental potential; the blastocyst rate per biopsied embryo was 40/109 (36.6%) and only 14/109 (13%) were classified as poor and unsuitable for transfer based on morphology. Blastocyst development from the embryos biopsied at 8-cells or more was 34/69 (49%).

One main concern of delaying embryo transfer is the possibility of there being no blastocyst available for transfer. In a prospective randomized study only 5% of patients did not have an embryo transfer and implantation rates of 50% were achieved (Gardner et al., 1998Go). As expected, our total figures show lower implantation rates (12/37; 32%), but our cohort included biopsied embryos from transfers with a mix of embryo development stages. However, in the seven patients who only had blastocysts transferred, an implantation rate of 8/17 (47%) was achieved. The three patients (20%) whose embryos did not achieve the blastocyst stage opted for embryo transfer despite poor prospects of pregnancy. Owing to the limited number of embryos available for transfer following PGD (after embryo biopsy of only morphologically `good' embryos and suitable genotype after genetic analysis) it has been recommended that cycles with less than nine oocyte–cumulus complexes should be cancelled (Liebaers et al, 1998Go). In PGD cases where high numbers of oocytes are retrieved it may be advantageous to culture biopsied embryos to the blastocyst stage despite fears of having no embryos available for transfer. It is interesting to note that the patients with no blastocysts for transfer had amongst the lowest number of oocytes collected (Table IIGo). The limitations of PGD are great but in Greece, where almost 1 in 10 of the population are carriers of ß-thalassaemia or Hbs, we prefer to offer PGD for couples at risk for transmitting ß-thalassaemia major as an additional therapy for patients requiring IVF. ß-thalassaemia is a monogenic recessive genetic disorder and therefore only 25% of embryos are likely to be affected. The proportion of embryos for transfer (homozygous normal or heterozygous carrier) are greater than that of a dominant genetic disorder which would otherwise have greatly diminished the embryos available. The genotyping method based on DGGE used in this study not only facilitates detection of the wide spectrum of mutations underlying ß-thalassaemia major and sickle cell thalassaemia but also addresses many of the inherent potential problems associated with PCR-based genotyping of single cells, which include total PCR failure, allelic drop-out (ADO) and contamination. DGGE is advantageous since it facilitates simultaneous analysis of more than one mutation in a single PCR fragment, detects the presence of normal as well as pathological alleles and in addition monitors the occurrence of allelic dropout through the expectation that heterozygous samples have more than one electrophoretic band on DGGE analysis (Figure 2Go). In practice only embryos with a normal electrophoretic DGGE band are considered to be unaffected and suitable for transfer, preventing misdiagnosis even if ADO has occurred. The nested PCR/DGGE protocol proved reliable, with a PCR success rate (and thus potential diagnosis) in 86% of blastomeres analysed (unpublished observations). It is more difficult to evaluate the accuracy of the genotyping results—the embryos that are not transferred are not reanalysed—but a single case of misdiagnosis has been detected amongst seven pregnancies so far evaluated with PND. On the basis that all embryos deemed unaffected have a normal DGGE band (Kanavakis et al, 1999Go; Vrettou et al, 1999Go), we conclude that the misdiagnosis was probably not due to inaccuracy of the genotyping method but rather to chance contamination of a tube(s) by extraneous genetic material introduced during processing. There are two potential sources of contamination when performing PGD using PCR based protocols: PCR carry-over contamination and operator contamination. There was no wider contamination of the system, as indicated by the PCR-negative controls and blanks at all stages in all cycles. Therefore we conclude that the misdiagnosis was due to operator contamination (during biopsy or PCR set-up), which is more likely to lead to the transfer of an affected embryo, although it is likely to be a rare event (Lewis et al., 2001Go).

Overall our experience indicates that culturing to the blastocyst stage does not compromise the outcome of PGD cycles and provides a solution to the time constraints of a busy IVF and molecular genetics unit. The increased time available after embryo biopsy allows procedures to be carried out during more acceptable working hours and could potentially allow for more extensive genetic analysis, such as cell recycling in PCR (Thornhill et al., 1994; He et al., 1999Go) or re-hybridization of the same cell in fluorescent in-situ hybridization (Gianaroli et al., 1999Go).


    Notes
 
3 To whom correspondence should be addressed at: Centre for Reproductive Medicine, Alpha Lab, Anastasiou 8, Athens 115 24, Greece. E-mail: gpalme{at}otenet.gr Back

Note added in proof

By the time of publication all embryos of the three ongoing pregnancies had been confirmed as unaffected by prenatal diagnosis. The triplet pregnancy was selectively reduced to twins at the request of the parents and all three ongoing pregnancies went to term, resulting in the birth of five more healthy babies.


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 Top
 Abstract
 Introduction
 Materials and methods
 Discussion
 References
 
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Submitted on June 7, 2001; accepted on October 8, 2001.