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Standard preimplantation genetic diagnosis (PGD) cannot be applied for de-novo mutations (DNM), because neither origin nor relevant haplotypes are available for testing in single cells. PGD strategies were developed for 80 families with 38 genetic disorders, determined by 33 dominant, three recessive and two X-linked DNM. All three recessive mutations were of paternal origin, while of 93 dominant mutations, 40 were paternal, 46 maternal and seven detected in affected children. The development of specific PGD strategy for each couple involved DNA analysis of the parents and affected children prior to PGD, including a mutation verification, polymorphic marker evaluation, whole and single sperm testing to establish the normal and mutant haplotypes and PGD by polar body analysis and/or embryo biopsy. Overall, 151 PGD cycles were performed for 80 families, for which a specific PGD design has been established. The application of these protocols resulted in pre-selection and transfer of 219 (1.72 per cycle) DNM-free embryos in 127 (84.1%) PGD cycles, yielding 63 (49.6%) unaffected pregnancies and birth of 59 (46.5%) healthy children, confirmed to be free of DNM. The data show feasibility of PGD for DNM, which may routinely be performed with accuracy of over 99%, using the established PGD strategy.
Preimplantation genetic diagnosis (PGD) is currently applicable when the mutation origin or relevant haplotypes are available for tracing its inheritance in oocytes and embryos. This makes difficult to offer PGD when parent(s) or affected children have a newly identified genetic condition, determined by de-novo mutation (DNM). We developed the PGD strategies for such families, and applied them to 80 families with 38 different genetic disorders, including 33 dominant, three recessive and two X-linked DNM. This involved DNA analysis of the parents and affected children prior to PGD, which included a mutation verification, polymorphic marker evaluation, whole and single sperm testing to establish the normal and mutant haplotypes, and PGD by polar body analysis and/or embryo biopsy. The technique was applied to 151 PGD cycles, resulting in pre-selection and transfer of 219 (1.72 per cycle) DNM-free embryos in 127 (84%) PGD cycles, yielding 63 (49.6%) unaffected pregnancies and birth of 59 (46.5%) healthy children, confirmed to be free of DNM tested. The method may routinely be performed with accuracy of over 99%.
Preimplantation genetic diagnosis (PGD) is an established procedure for couples at risk of producing offspring with inherited disorders, which has been applied for more than 200 different genetic conditions, resulting in the birth of thousands of unaffected children by the present time (
The Preimplantation Genetic Diagnosis International Society (PGDIS). 2008 Guidelines for good practice in PGD: program requirements and laboratory quality assurance. Reprod BioMed Online 16, 134–147.
The Preimplantation Genetic Diagnosis International Society (PGDIS). 2010 10th International Congress on Preimplantation Genetic Diagnosis. Reprod BioMed Online 20, S1–S42.
). With an increasing number of different genetic disorders for which PGD is being applied each year, it may presently be applicable for any inherited disorder for which sequence information or relevant haplotypes are available for the detection by direct mutation analysis or haplotyping in oocytes or embryos (
The Preimplantation Genetic Diagnosis International Society (PGDIS). 2009 Ninth International Congress on Preimplantation Genetics: PGD and Stem Cells. Reprod BioMed Online 18, S1–S32.
The range of PGD indications is also gradually expanding to include the risk for common diseases with genetic predisposition and non-genetic conditions, such as human leukocyte antigen (HLA) typing with the purpose of stem cell therapy for existing affected siblings in a family (
Rechitsky, S., Kuliev, A., 2010. Novel indications for preimplantation genetic diagnosis. 10th International Congress on Preimplantation Genetic Disgnosis, Montpellier, France, 5–8 May 2010. Reprod Biomed Online 20, Suppl. S1–S2.
PGD has also become applicable to population groups objecting to any type of embryo biopsy, through performing pre-embryonic diagnosis by polar body (PB) analysis (
). This became feasible with the improvement in DNA analysis, which can now be completed in less than 9 h after PB2 removal, to ensure that the oocytes are still at the pronuclei stage. So the oocytes predicted to have a mutant gene are frozen prior to the embryo formation, while the embryos originating from the mutation-free oocytes are allowed to develop further and are replaced after reaching the blastocyst stage.
According to the current guidelines, performing PGD requires knowledge of sequence information for Mendelian diseases, but may also be performed when the exact mutation is unknown, through the application of linkage analysis (
The Preimplantation Genetic Diagnosis International Society (PGDIS). 2009 Ninth International Congress on Preimplantation Genetics: PGD and Stem Cells. Reprod BioMed Online 18, S1–S32.
). With expanding use of single-nucleotide polymorphisms, linkage analysis may allow PGD for any genetic disease, irrespective of the availability of specific sequence information (
). This approach is more universal making it possible to track the inheritance of the mutation without actual testing for the gene itself.
The ongoing developments also allow the genetic analysis of single biopsied cells by DNA microarray technology, which provides the possibility for a simultaneous testing for a causative gene, multiple linked markers, health-related genetic variability and chromosomal abnormalities in the same biopsied PB or blastomere.
However, the above approaches cannot be applied in cases of de-novo mutations (DNM) in parent(s) or affected children, as neither origin nor relevant haplotypes are available for tracing the inheritance of this DNM in single cells biopsied from embryos or in oocytes. On the other hand, with the improved awareness of PGD, an increasing number of couples request PGD, without any family history of the genetic disease that has been first diagnosed in one of the parents or in their affected children (
). So the present report describes the development of PGD strategies for the genetic conditions determined by DNM and presents the first systematic experience of PGD for 151 cycles involving DNM.
Materials and methods
PGD strategies were developed for a total of 80 families with 38 different genetic disorders, determined by 33 dominant, three recessive and two X-linked DNM (the list of DNM for which the PGD strategy was performed is presented in Table 1).
Table 1Outcome of preimplantation genetic diagnosis (PGD) for de-novo mutations.
The majority of these families (71/80) were with dominant mutations, of which 40 were of paternal origin, including two cases of gonadal mosaicism, 46 of maternal origin, including one with gonadal mosaicism, and nine detected for the first time only in the affected children. All three couples with DNM of autosomal recessive inheritance were of paternal origin, including cystic fibrosis (CF), spinal muscular atrophy (SMA) and Fanconi anaemia. PGD for two X-linked DNM included PGD for chronic granulomatosis and incontinentia pigmenti.
As seen in Figure 1, the PGD strategies for these families were different depending on the origin of DNM and included DNA analysis of the parents and affected children prior to PGD, including the mutation verification, polymorphic marker evaluation, whole and single sperm testing and PB analysis in order to establish the normal and mutant haplotypes, without which PGD cannot always be performed.
Figure 1Preimplantation genetic diagnosis (PGD) strategies for de-novo mutations (DNM) of different origin. (A) Case work-out for DNM detected in father (partner): (1) pedigree in two generations; (2) mutation verification in DNA extracted from blood and total spermatozoa; (3) amplification of partner’s single spermatozoa to establish normal and affected haplotypes required for PGD cycle preparation; (4) amplification of patient’s DNA to identify the most informative markers for PGD; and (5) PGD by blastomere or blastocyst biopsy for combined mutation and linkage analysis. (B) Case work-out for DNM detected in mother (patient): (1) pedigree in two generations; (2) mutation verification in DNA extracted from whole blood or cheek swabs and single lymphocytes (paternal haplotypes are analysed on single spermatozoa for more accurate prediction of embryos’ genotypes); (3) PGD by PB1 and PB2 analysis to identify DNM-free oocytes and establish maternal haplotypes; and (4) blastomere or blastocyst analysis to confirm the diagnosis. (C) Case work-out for DNM detected first in affected offspring: (1) pedigree in three generations; (2) verification of DNM in child’s DNA extracted from blood or cheek swabs and mutation testing of DNA extracted from parents’ whole blood and total spermatozoa; (3) mutation evaluation on single lymphocytes and single sperm testing to rule out paternal gonadal mosaicism; (4) PGD by polar body analysis to detect potential maternal gonadal mosaicism; and (5) blastomere or blastocyst analysis to confirm the absence of the mutation.
In cases of DNM of paternal origin (Figure 1A), the DNM was first confirmed on the paternal DNA from blood and total spermatozoa, followed by single sperm typing to determine the proportion of spermatozoa with DNM and relevant normal and mutant haplotypes, as described earlier (
). For a higher reliability of testing, the relevant maternal linked markers were also detected, to be able to trace for possible shared maternal and paternal markers. To exclude misdiagnosis, PGD involved simultaneous detection of the causative gene and at least three highly polymorphic markers, closely linked to the gene tested, to ensure the detection of preferential amplification and allele-specific amplification failure (allele drop out, ADO), the main potential causes of PGD misdiagnoses (
). This involved a multiplex nested PCR analysis, with the initial 30 cycles of the first round of PCR containing all the pairs of outside primers, followed by 25 cycles of amplification of separate aliquots of the resulting PCR product with the inside primers specific for each site. Following the nested amplification, PCR products were analysed either by restriction digestion or direct fragment size analysis.
In cases of DNM of maternal origin (Figure 1B), DNM was first confirmed in maternal blood and PGD was performed, when possible, by PB analysis, to identify the normal and mutant maternal haplotypes. Also, in order to trace the relevant paternal haplotypes, single sperm typing was performed, whenever possible, for avoiding misdiagnosis caused by possible shared paternal and maternal markers, particularly when no relatives are available to obtain the paternal haplotypes.
In cases of DNM detected first in children (Figure 1C), the mutation was verified in their whole blood DNA, followed by testing for the mutation in paternal DNA from blood, total and single spermatozoa, if the DNM appeared to be of paternal origin. In DNM of maternal origin, PGD was performed by the PB approach, with confirmation of the diagnosis by embryo biopsy, if necessary.
So in contrast to previous PGD practice, performing PGD for DNM required additional preparatory DNA work before performing the actual PGD, including single sperm analysis and the requirement of performing sequential PB1 and PB2, followed by blastomere or blastocyst analysis, described in detail elsewhere (
). Single spermatozoa and PBs were removed using micromanipulation and placed in lysis solution, which contained proteinase K for PB and KOH and dithiothreitol for sperm samples.
As in previous PGD protocols, the embryos without DNM were transferred in the same cycles, while the affected ones were used for confirmation of diagnosis, at least for in-house cases. Predicted diagnoses were followed up after delivery, while the spare unaffected embryos were frozen for future use by the families.
Results
Overall, 151 PGD cycles for DNM were performed for 80 families under study, resulting in pre-selection and transfer of 219 (1.72 per cycle) DNM-free embryos in 127 (84.1%) PGD cycles. This resulted in 63 (49.6%) unaffected pregnancies and the birth of 59 (46.5%) healthy children, confirmed to free of the DNM tested, and five pregnancies are still ongoing (Table 1). No misdiagnosis has been observed in the follow-up analysis.
DNM of dominant inheritance
As mentioned, the largest group was PGD for DNM of autosomal dominant type, including 136 cycles from 71 patients, which resulted in the transfer of 201 DNM-free embryos in 115 cycles, yielding 57 pregnancies and birth of 53 unaffected children, with four ongoing pregnancies at the present time. The most frequent conditions in this group were neurofibromatosis (NF) type 1 (24 cycles), osteogenesis imperfecta (19 cycles), Marfan syndrome (13 cycles), facioscapularhumoral muscular dystrophy and Blackfan Diamond anaemia (nine cycles each), familial adenomatous polyposis, tuberous sclerosis type 1 and retinoblastoma (seven cycles each) and Gorlin and Crouson syndromes (five cycles each). Between one and three cycles were performed for the remaining conditions, listed in Table 1.
Dominant DNM of paternal origin
The example of PGD design for DNM of dominant inheritance is presented below for a couple with NF2 splicing mutation (c114+2 T–C) detected in the husband with no previous family history of the disease (Figure 2). DNA analysis in paternal blood confirmed the presence of NF2 splicing mutation (c114+2 T–C), while testing of single spermatozoa showed a gonadal mosaicism, represented by three types of spermatozoa corresponding to three different haplotypes. Only 30% of spermatozoa were represented by actual mutant haplotype, while 36% were normal, characterized by normal haplotype, and 34% contained a normal allele in the mutant haplotype.
Figure 2Preimplantation genetic diagnosis (PGD) for de-novo mutations (DNM) in neurofibromatosis type II gene (c114+2 T–C splicing mutation) of paternal origin. (A) Family pedigree showing that that DNM in NF2 gene was first detected in father. Single sperm analysis via multiplex heminested PCR revealed gonadal mosaicism with three different haplotypes: haplotype a (normal), haplotype b (mutant containing c.114+2 T–C allele) and haplotype c (mutant without c.114+2 T–C allele in NF2). Maternal linkage was based on DNA amplification of blastomeres in PGD cycle. (B) Outcome of the first PGD cycle. Blastomeres from five embryos were subjected to combined mutation and linkage analysis by multiplex heminested PCR. Three embryos (4, 5 and 7) were predicted to be free from the paternal mutation based on the presence of normal sequence (N) in NF2 gene and confirmed by linked markers (haplotype a). Two embryos (6 and 8) were predicted to have normal sequence (N∗) on haplotype c. The accuracy of this prediction was decreased due to a potential allele drop out of the dominant mutation in single blastomere. Trophectoderm (TE) biopsy from these embryos confirmed the presence of the normal sequence of NF2 gene. Embryos 5 and 7 were transferred (ET) resulting in unaffected pregnancy and birth of a healthy boy and girl confirmed by postnatal testing. (C) Outcome of the second PGD cycle. Combined mutation and linkage analysis by multiplex heminested PCR was performed on blastomeres from 10 embryos. Mutant haplotype b was detected only in embryo 13. Embryo 6 was missing all the maternal markers suggesting monosomy of chromosome 22, in which the gene is localized. Although all the remaining embryos were predicted to be normal and free of mutation, only four (embryos 3, 4, 9 and 12) were with normal (N) paternal haplotype a, while embryos 2, 5, 7 and 11 were predicted to have normal sequence (N∗) on the mutant haplotype c. Blastocyst biopsy confirmed normal genotypes predicted on blastomeres. Two normal embryos (3 and 4) were transferred, resulting in clinical pregnancy and delivery of a healthy girl confirmed by postnatal analysis.
PGD was based on detecting and avoiding the transfer of embryos with the mutant haplotype with or without the mutant gene, while the embryos with normal haplotypes of paternal and maternal origin were transferred. As can be seen in Figure 2, all the five tested embryos from the first PGD cycle may appear unaffected, despite the finding of the mutant haplotype in two of them (embryos 6 and 8), which, however, were missing the mutant gene. The remaining three embryos were with normal paternal and maternal haplotypes, of which two (embryos 5 and 7) were transferred, resulting in a twin pregnancy and birth of two unaffected children. In the second PGD cycle for this couple, 10 embryos were examined, of which only one contained the actual mutant haplotype, while three had the mutant haplotype without the mutant gene, and the remaining six had the normal haplotype. Two of these embryos (3 and 4) were transferred, resulting in a singleton pregnancy and birth of unaffected child.
Dominant DNM of maternal origin
The example of dominant DNM of maternal origin is presented in Figure 3, in which gonadal mosaicism was also detected. DNM in NF1 gene (intron 17–38 deletion) was first presented in the affected child and appeared to be originated from the mother, who had three cell populations, represented by three haplotypes, including the normal, mutant with intron 17–38 deletion and mutant without deletion. So PGD was based on pre-selection and transfer of the embryos with either normal maternal haplotype or with mutant maternal haplotype lacking intron 17–38 deletion. As seen in Figure 3, of 11 embryos examined, despite the presence of six embryos with mutant maternal haplotypes, actually only two were affected (embryos 6 and 8), the other four (embryos 1, 2, 10 and 12) had no intron 17–38 deletion so were unaffected. Of the remaining five embryos, two were monosomic for maternal chromosome (embryos 7 and 11), and three (embryos 3, 4 and 5) contained only the normal parental haplotypes. One of these embryos (embryo 3) and the other demonstrating mutant maternal haplotype without deletion (embryo 2) were transferred, resulting in a biochemical pregnancy (Figure 3).
Figure 3Preimplantation genetic diagnosis (PGD) for de-novo mutations (DNM) in NF1 gene (intron 27–38 deletion) of maternal origin. (A) Family pedigree of a couple with affected son carrying deletion of intron 27–38 in the NF1 gene. This deletion was not detected in maternal DNA from whole blood, although two haplotypes (c and d) were present, the latter corresponding to mutant haplotype corresponding to affected son’s haplotype, but with no deletion. The expected deleted area on this ‘benign’ chromosome (same haplotype as affected son received from the mother but without deletion) is framed. The actual mutant haplotype (e) with deletion was detected on maternal single lymphocytes. Paternal normal haplotypes (a) and (b) were established based on markers detected on the son’s normal chromosome. (B) Outcome of PGD cycle, performed by multiplex heminested PCR on blastomeres from 11 embryos. Three embryos (3, 4 and 5) were predicted normal (N) based on the presence of maternal normal haplotype (c) and suitable for embryo transfer (ET). Four embryos (1, 2, 10 and 12) inherited the ‘benign’ mutant maternal haplotype (d) and were also predicted normal (N∗) and suitable for ET. Of the remaining four embryos, embryos 7 and 11 were predicted to have monosomy of chromosome 17, based to the absence of maternal alleles, while the other two (embryos 6 and 8) were predicted to be affected, based on the absence of maternal markers in deleted area (DEL). Two embryos (2 and 3) were transferred and resulted in a biochemical pregnancy.
Only four PGD cycles were performed for DNM of autosomal recessive type: one cycle for CF, one for SMA and two for Fanconi anaemia, which resulted in the transfer of six embryos, yielding two pregnancies and the birth of unaffected twins from the couple tested for CF, and the other pregnancy is ongoing (Table 1). In case of PGD for SMA, the maternal mutation was inherited from parents, while the paternal one was not found in relatives, so the paternal haplotypes were identified by single sperm testing. In PGD for CF, the affected baby was double-heterozygous for CFTR DF508/R75X, the latter representing DNM, not identified in family testing.
As presented in Figure 4, DNM for FANCI was first detected in the child, who was compound heterozygous for C750G/E837X mutation. Testing of both parents for the presence of these mutations showed that the mother was a carrier of E837X mutation, and characterized by two relevant haplotypes, while no mutation was found either in the paternal blood or whole spermatozoa, despite the presence of both normal and mutant haplotypes in single spermatozoa, which however was lacking C750G mutation. Because the couple also requested HLA typing for possible stem cell transplantation required for the affected sibling, the embryos were also tested for HLA haplotypes. As can be seen in Figure 4, of the six embryos tested, only one (embryo 2) inherited both maternal and paternal normal haplotypes, while two were with both paternal and maternal mutant haplotypes (embryos 3 and 6), but were unaffected heterozygous carriers, because the paternal mutant haplotype was missing the expected paternal FANC750G mutation. The remaining three embryos had normal maternal and mutant paternal haplotype, without a mutant gene involved. So all the embryos were actually unaffected, of which two heterozygous carriers also appeared to be HLA matched to the affected child (embryos 3 and 6) and these embryos were transferred, resulting in no pregnancy.
Figure 4Preimplantation genetic diagnosis (PGD) for autosomal recessive de-novo mutations (DNM) detected first in an affected child who was compound heterozygous for C750G/E837X mutations in the FANCI gene, combined with human leukocyte antigen (HLA) genotyping. (A) Family pedigree, showing HLA and mutation haplotypes, based on parental and affected child’s genomic DNA testing. E837X mutation was detected in the carrier mother, but C750G mutation was absent in DNA extracted from paternal blood or whole sperm samples. However, both normal and mutant haplotypes were detected in testing of single spermatozoa. (B) PGD cycle combined with HLA testing: (1b) Multiplex heminested and fully nested amplification performed on blastomeres from six embryos did not reveal the paternal mutation. Four embryos (1, 2, 4 and 5) were predicted normal based on the absence of both mutations, of which embryos 1, 4 and 5 inherited ‘benign’ paternal haplotype a, similar to one of the mutant haplotypes in the affected child, and embryo 2 inherited the normal haplotype b. The remaining embryos (3 and 6) were predicted to be carriers of the maternal mutation E837X, but inherited the paternal haplotype a; and (2b) HLA marker analysis demonstrated the presence of two HLA-matched embryos (3 and 6), which were transferred but did not achieve pregnancy. Normal∗ = benign paternal haplotype a similar to the mutant haplotype of the affected child; white bars = HLA-match genotypes; grey bars = non-HLA-match haplotypes.
The results and outcome of 11 PGD cycles for X-linked DNM are presented and Table 1, and the example of PGD for chronic granulomatous disease, determined by DNM IVS9+5 G–A in CYBB gene is shown in Figure 5. DNM in this case was first detected in the affected child, who also required HLA-matched stem cell transplantation. So in addition to mutation analysis, HLA typing was performed, together with aneuploidy testing, because the mother was 36 years old. DNA analysis in maternal blood failed to detect the mutant gene, while both normal and mutant haplotypes were present, despite the latter missing the mutant gene. PGD was performed by sequential PB1 and PB2 analysis in nine oocytes, showing that all the oocytes were normal, although four of them (oocytes 2, 3, 7 and 11) contained the maternal mutant haplotype, without the mutant gene. The testing of the embryos resulting from each of these oocytes confirmed the PB haplotype analysis, showing the lack of the mutant gene. All the embryos were also found to be aneuploidy free, of which four (embryos 7, 8, 9 and 11) also appeared to be the exact HLA match to the affected sibling. Two of these (8 and 9) were transferred, resulting in a clinical pregnancy, which spontaneously aborted in the first trimester.
Figure 5Preimplantation genetic diagnosis (PGD) for chronic granulomatous disease, determined by X-linked DNM IVS9+5 G–A, combined with human leukocyte antigen (HLA) genotyping and aneuploidy testing. (A) Family pedigree showing the mutation and closely linked to CYBB gene markers. (B) Sequential PB1 and PB2 analysis, showing that all the tested oocytes are normal, despite four of them containing the ‘benign’ mutant haplotype without IVS9+5 mutation (N∗). (C) (1c) multiplex heminested PCR for combined mutation analysis; (2c) HLA genotyping; and (3c) karyotyping, for six chromosomes by PCR on blastomeres. Two of four embryos (8 and 9), predicted to be HLA matched and free of mutation and aneuploidy, were transferred, resulting in a clinical pregnancy that spontaneously aborted at 9 weeks. White bars = HLA-match genotypes; grey bars = non-HLA-match haplotypes.
This is the first report of systematic PGD for DNM, which could not previously be performed, due to unavailability of family history and lack of any affected family member to identify the origin of a mutation and trace the inheritance of the mutant and normal alleles in oocytes and embryos. However, the presented data show that strategies may be developed in the search for the possible origin of DNM and relevant haplotypes as the basis for developing a PGD design for each particular couple with DNM, allowing a highly accurate pre-selection of oocytes and embryos free from the DNM in question.
Although the strategies may differ depending on the type of DNM inheritance, the general approach involves the identification of the DNM origin and search for a possible gonadal mosaicism and relevant parental haplotypes. As demonstrated in Figure 1, one of the important steps is single sperm typing, which was performed in 37/80 patients (46.3%). Overall, 964 single spermatozoa were tested, with the requirement for testing of at least 15 single spermatozoa per patient, and as many as 50 per patient to exclude a possible gonadal mosaicism. Even if a DNM is not identified, a single sperm typing may identify a ‘benign’ mutant haplotype, represented by a mutant haplotype without a DNM. The other important requirement is to identify the relevant linked markers in both parents even if only one is a DNM carrier. Although not always possible, PGD by the PB approach to detect or confirm the maternal normal and mutant haplotypes is always the method of choice, performed in this study in 54 (35.8%) of 151 cycles.
The implications of gonadal mosaicism for genetic counselling of dominant disorders, such NF1, tuberous sclerosis types 1 and 2, lethal osteogenesis inperfecta, familial adenomatous polyposis, retinoblastoma and X-linked dominant trait incontinentia pigmenti, have been recognized previously (
). Although germinal mosaicism is thought to be common, its presence in families is usually difficult to detect and depends on the gene penetrance. A majority of the newly mutant genes will have mutated during the development of the affected individual and not during development of one of the parents’ gonads, so testing of the affected child will often reveal mosaicism for the gene in question, but in many cases will remain undetected.
For example, all the oocytes and embryos tested from such cases in the study centre’s experience appeared to be unaffected, irrespective of the origin of the DMN (29 oocytes and 87 embryos). However, this does not mean that PGD is not justified in these cases, because the possibility of a low level of mosaicism in parents’ gonads cannot be completely excluded. In the cases of DNM detected first in children, although no mutation was identified in either parent, mutant haplotypes without the mutant gene were present in the parents, suggesting the possibility of very low levels of mosaicism. So PGD is indeed indicated in such cases to exclude any possibility of mutant oocyte and embryo production due to undetected germinal mosaicism.
As expected, the majority of cases involved DNM of dominant inheritance, in agreement with the high mutation rate of dominant disorders. However, almost a similar proportion of DNM of dominant type was either of paternal (43%) or maternal (49%) origin, requiring the testing for the presence of DNM in both parents. On the other hand, all the cases of DNM of recessive inheritance were of paternal origin, but the number of cases is not sufficient for conclusions.
It should be mentioned, that despite the complexity of PGD for DNM, the applied strategies appeared to be highly accurate. Based on testing of 631 oocytes by PB analysis and 1145 embryos by blastomere biopsy, 219 mutation-free embryos were transferred, resulting in 49.6% pregnancy rate and birth of 59 healthy children with no misdiagnosis detected, suggesting a 100% accuracy of the applied technique of PGD. This is in agreement with 99.4% accuracy rate per transfer cycle in the study centre’s overall experience of PGD for Mendelian disorders, which is the world’s largest PGD series for this indication, including 2028 PGD cycles performed for 227 genetic conditions, which resulted in 677 unaffected pregnancies (40.2% pregnancy rate per transfer cycle) and birth of 690 healthy children (
; Rechitsky and Kuliev, 2010). Of the four misdiagnoses observed, one was due to undetected ADO in PGD for CF, when the mutant double heterozygote embryo was erroneously diagnosed as an unaffected carrier (
) and the other three were due to transfer of embryos with predicted low accuracy in PGD for Fragile X, muscular dystrophy and β-thalassaemia, when the couple opted to transfer the embryo tested normal based on an insufficient number of markers, leaving the probability for ADO (
). There were also three cases of the wrong embryo transfer, which was obvious from the haplotypes of the resulting fetuses and children contrasting with the haplotypes of the embryos recommended for transfer.
The high accuracy has also recently been reported from the other second largest PGD experience for monogenic disorders (
). In this report only a 0.6% misdiagnosis rate was observed in PGD of 1443 PGD cycles, one in PGD for muscular dystrophy and three in PGD for CMT1A, the latter being due to error in linkage analysis in preparation for PGD (
ESHRE PGD consortium data collection VIII: cycles from January to December 2005 with pregnancy follow-up to October 2006 European Society of Human Reproduction and Embryology PGD Consortium.
In conclusion, the presented data show that PGD for DNM is an important addition to the practice of PGD for Mendelian diseases, as it now makes it possible to offer PGD to any couple at risk for producing offspring with a genetic disease, without the traditional requirement of family data, which is not always available even in cases with known family history for the disease. So the data demonstrate feasibility of PGD for DNM, which may now be routinely performed with the accuracy of over 99%, using the established PGD strategy.
Acknowledgement
This paper is dedicated to the memory of Yury Verlinsky, founder of Reproductive Genetics Institute.
References
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Brooks B.
Kaplan Y.
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Single sperm análisis for haplotype construction of de-novo paternal mutations: application to PGD for neufibromatosis type 1.
ESHRE PGD consortium data collection VIII: cycles from January to December 2005 with pregnancy follow-up to October 2006 European Society of Human Reproduction and Embryology PGD Consortium.
The Preimplantation Genetic Diagnosis International Society (PGDIS). 2008 Guidelines for good practice in PGD: program requirements and laboratory quality assurance. Reprod BioMed Online 16, 134–147.
The Preimplantation Genetic Diagnosis International Society (PGDIS). 2009 Ninth International Congress on Preimplantation Genetics: PGD and Stem Cells. Reprod BioMed Online 18, S1–S32.
The Preimplantation Genetic Diagnosis International Society (PGDIS). 2010 10th International Congress on Preimplantation Genetic Diagnosis. Reprod BioMed Online 20, S1–S42.
Rechitsky, S., Kuliev, A., 2010. Novel indications for preimplantation genetic diagnosis. 10th International Congress on Preimplantation Genetic Disgnosis, Montpellier, France, 5–8 May 2010. Reprod Biomed Online 20, Suppl. S1–S2.
Declaration: The authors report no financial or commercial conflicts of interest.
Footnotes
Dr. Svetlana Rechitsky is a graduate of Kharkov University’s Genetics Faculty, and received her PhD in experimental molecular embryology from the second Moscow Medical Institute in 1986. She moved to the Reproductive Genetics Institute in 1989, where she heads the DNA laboratory, which has performed the largest preimplantation genetic diagnosis (PGD) series for single gene disorders, with PGD design for the majority of them developed for the first time. She has published more than 40 papers in the field of PGD, including a key contribution to the Atlas of Preimplantation Genetic Diagnosis.
Dr. Svetlana Rechitsky is a graduate of Kharkov University’s Genetics Faculty, and received her PhD in experimental molecular embryology from the second Moscow Medical Institute in 1986. She moved to the Reproductive Genetics Institute in 1989, where she heads the DNA laboratory, which has performed the largest preimplantation genetic diagnosis (PGD) series for single gene disorders, with PGD design for the majority of them developed for the first time. She has published more than 40 papers in the field of PGD, including a key contribution to the Atlas of Preimplantation Genetic Diagnosis.
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