Practical preimplantation genetic testing [3 ed.] 9783030431570, 3030431576

Table of contents :
Preface
Contents
1: Place of Preimplantation Genetic Testing (PGT) Among Available Options for Prevention of Genetic Disorders
1.1 Preconception Prevention of Congenital Anomalies by Folic Acid Containing Multivitamin Fortification Programs
1.2 Genetic History and Avoidance of Congenital Disorders by Prenatal and Preimplantation Genetic Testing
1.3 Prospective Carrier Screening as a Means for Improving PGT Uptake
References
2: Major Components of Preimplantation Genetic Testing
2.1 Introduction
2.2 Polar Body Sampling
2.2.1 Polar Body Testing as a Preconception Testing Strategy
2.2.2 Analysis of Sperm
2.3 Embryo Biopsy
2.4 Developments in Blastocyst Biopsy Procedures
2.4.1 Trophectoderm Biopsy Procedure Without Laser Assistance at Day 5
2.5 Noninvasive PGT (NIPGT)
2.6 PGT Without IVF
References
3: Major Components of Preimplantation Genetic Testing: Adjustment of Available Genetic Technology to PGT Practice
3.1 Adjustment of DNA Analysis to Avoid Misdiagnosis in Single-Cell PCR
3.2 Other Strategies to Avoid Misdiagnosis
3.3 Is There Still Place for FISH Analysis in Current PGT-A?
3.4 Microarray Analysis (Array CGH): Acceptable but Not up to Present Standards
3.5 Next-Generation Sequencing (NGS): Present Standard of PGT Practice
References
4: Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)
4.1 Traditional Indications and Strategies for PGT-M
4.1.1 Autosomal Recessive Disorders
4.1.2 Autosomal Dominant Disorders
4.1.3 X-Linked Disorders
4.2 Affected Homozygous or Compound Heterozygous Conditions
4.3 Concomitant PGT for Two or More Single-Gene Disorders
4.4 De Novo Mutations
4.5 Late-Onset Common Disorders with Genetic Predisposition
4.5.1 Cancer
4.5.1.1 Breast Cancer
4.5.1.2 Other Cancers
4.5.2 Neurodegenerative Diseases
4.5.2.1 Huntington Disease
4.5.2.2 Alzheimer Disease
4.5.3 Cardiac Disease
4.5.3.1 Conclusion
4.6 Other Genetic Conditions with Important Health-Related Implications
4.6.1 Blood Group Incompatibility
4.6.2 Congenital Malformations
4.6.2.1 Embryonic Lineage and Differentiation
4.6.3 Dynamic Mutations
4.6.4 Variants of Unknown Significance (VUS)
4.7 Contemporary Selection of Molecular Technologies
References
5: Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)
5.1 Thalassemia
5.2 Immunodeficiency
5.3 Preimplantation HLA Matching Without PGT-M
5.4 Limitations and Future Prospects of PGT- HLA
5.5 Practical Implications of PGT-HLA
References
6: Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic Testing for Chromosomal Disorders (PGT-A and PGT-SR)
6.1 Meiotic and Mitotic Abnormalities
6.2 Chromosome-Specific Meiotic Error Origin and Its Impact on Embryo Viability
6.3 Mitotic Errors in Embryos in Relation to Meiosis Errors
6.4 Practical Relevance of Autosomal Monosomy Detection
6.5 Uniparental Disomies
6.6 Mosaicism and Sub-chromosomal Variations
6.7 Possible Additional Tests to Preselect Euploid Embryos with Higher Potential for Pregnancy
6.7.1 Mitochondrial DNA
6.7.2 Morphokinetic Assessment
6.8 Characterizing Chromosomal Rearrangements
6.8.1 Polar Body Approach
6.8.2 Karyotyping of Embryos via Nuclear Conversion
6.8.3 PGT-SR Using Next-Generation Technologies
6.8.4 PGT-SR Developments to Distinguishing Noncarrier Balanced Embryos from Normal
6.8.5 Reproductive Outcome of PGT-SR
References
7: Clinical Outcome of Preimplantation Genetic Testing
7.1 Ovulation Stimulation for a PGT Cycle
7.1.1 Ovulation Stimulation and Aneuploidy
7.1.2 Specific Disorder Characterized by Poor Ovulation Response
7.1.2.1 Fragile X Syndrome
7.1.2.2 Myotonic Dystrophy
7.1.2.3 BRCA 1/2
7.1.2.4 Carriers of Balanced Translocations
7.2 Anomalies in ART/ PGT Pregnancies and in ART Pregnancies Without PGT
7.2.1 Birth Defects in ART/PGT Pregnancies
7.2.2 Birth Defects in ART without PGT
7.2.3 Pitfalls in Assessing Birth Defects in ART Offspring
7.2.4 Birth Defects in Offspring of Subfertile Couples Not Requiring ART
7.2.5 Genetic Counseling for Birth Defects in ART and PGT
7.3 Diagnostic Accuracy of PGT-M
7.4 Diagnostic Accuracy and Predictive Value of PGT-A
7.4.1 Predictive Value of PGT-A
7.4.2 Aneuploidy and Error Origin of Chromosomal Abnormalities
7.5 Mosaicism in PGT-A
7.5.1 Numerical Mosaicism
7.5.2 Segmental Mosaicism
7.5.3 Priority of Mosaic Embryos for Transfer
7.6 Diagnostic Accuracy of PGT-SR
7.7 Clinical Utility of PGT-A: Randomized Clinical Trials Involving 24-Chromosome Aneuploid Testing
7.7.1 RCTs Involving Polar Body Biopsy
7.7.2 RCTs Involving 24-Chromosome Testing of Blastocysts
7.7.3 Confirming Clinical Utility of a PGT-A
7.8 Ancillary Tests to Enhance PGT-A Pregnancy Rates
7.9 PGT-A for Recurrent Miscarriages
7.10 Why Did Cleavage-Stage RCTs Before 2010 Fail to Show Benefit of PGT-A?
7.10.1 Biopsy Damage in Cleavage-Stage Embryos
7.10.2 Observational Studies and RCTs Involving ­Cleavage-Stage Embryos That Showed Benefit
7.10.3 RCTs in Cleavage-Stage Embryos Not Showing Benefit
7.11 Universal Single Embryo Transfer to Avoid Multiple Gestations
References
8: Social, Ethical, and Legal Aspects
8.1 Legality and Regulation of PGT
8.2 PGT in Venues Having Restrictions
8.3 Ethical Issues Unique to PGT
8.4 PGT for Disorders with Genetic Predisposition
8.5 PGT for HLA Prediction and Stem Cell Transplantation
8.6 PGT for Sex Selection
8.7 Conclusions
References
Index

Citation preview

Practical Preimplantation Genetic Testing Anver Kuliev Svetlana Rechitsky Joe Leigh Simpson Third Edition

123

Practical Preimplantation Genetic Testing

Anver Kuliev • Svetlana Rechitsky  Joe Leigh Simpson

Practical Preimplantation Genetic Testing Third Edition

Anver Kuliev Reproductive Genetic Innovations Chicago, IL USA

Svetlana Rechitsky Reproductive Genetic Innovations Chicago, IL USA

Florida International University

Florida International University

Miami, FL USA

Miami, FL USA

Joe Leigh Simpson Florida International University Miami, FL USA Reproductive Genetic Innovations

Chicago, IL USA

ISBN 978-3-030-43156-3    ISBN 978-3-030-43157-0 (eBook) https://doi.org/10.1007/978-3-030-43157-0 © Springer Nature Switzerland AG 2020 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Switzerland AG The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

Preface

Before introducing the present edition of the book, we have to mention the changes of standardized terminology*, according to which the term “preimplantation genetic testing” (PGT) replaces the previous term “preimplantation genetic diagnosis” (PGD) and a quite inadequate “jargon” “preimplantation genetic screening” (PGS) used as an alternative to PGD for aneuploidy. Accordingly, “PGT-M” will be used as an abbreviation for preimplantation genetic testing for monogenic disorders, “PGT-A” for preimplantation genetic testing for aneuploidy, “PGT-SR” for preimplantation genetic testing for structural rearrangements, and “PGT-HLA” for preimplantation genetic testing for human leukocyte antigens (HLA). So, the title of the third edition is changed from “Practical Preimplantation Genetic Diagnosis” to “Practical Preimplantation Genetic Testing” to comply with the standardized international terminology for assisted reproductive technology (ART) and preimplantation genetics. It should be mentioned that although PGT has become an established procedure for genetics and ART practices already for the last decade, its wider application has been obvious only after the introduction of the next-­generation technologies in the last few years. At the present time, more than one third of ART centers and the majority of genetic practices in the United States have already been utilizing PGT services to allow at-risk couples to reproduce normally, without fear of having an affected offspring. The practical PGT experience at present may be estimated in hundreds of thousands, what makes the updating of this experience of practical utility to both medical profession and patients. This will include, first of all, an update of PGT accuracy, reliability, and safety to ensure an improved access to PGT of those who may benefit greatly from this technology. It is of note that PGT has now been applied in as many as 581 different conditions, with the accuracy in the leading PGT centers, such as ours, approaching almost 100%. In fact, PGT may now be performed for any genetic condition, even if it was identified in one of the parents or in the affected child de novo, with also a possibility of concomitant testing of a number of disorders in one test. Updating will include also the progress in the primary prevention of genetic disorders described in the introductory section, which will now Zegers-Hochschild F,, David Adamson G., Dyer S., Racowsky C., de Mouzon J., Sokol R., Rienzi L., Sunde A., Schmidt L., Cooke I.D., Simpson JL,. Van der Poel S. The international glossary on Infertility and Fertility Care. Fertil Steril. 2017; 108(3):393–406 * 

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include the approaches for prospective identification of at-risk PGT couples, through the application of the expanded carrier screening using an increasing number of gene in the panels, as a means for an improved prospective access to PGT.  In fact, because of dramatic technological improvements in all aspects of PGT, most of the sections will be updated, with addition of also new sections on the next-generation technologies and universal PGT with combined testing for single gene and chromosomal disorders, which has previously presented a real challenge. As we have presently accumulated the world’s largest experience in this area, the guiding PGT strategies for different genetic disorders, will be presented, with emphasis on the most complicated cases that might be of special utility in the wider application of PGT technologies worldwide. PGT indications continue to expand for those that have never been even predicted, so the new section will be devoted to borderline indications, which will include common adult-onset conditions with genetic predisposition and nongenetic indications. This section will include the expanding PGT application to heart disease and cancer, for which the number of requests has been increasing gradually, such as for breast cancer, with more than a hundred predisposing gene mutations tested by the present time. A unique experience on PGT-HLA for stem cell transplantation treatment of congenital and acquired disorders will be addressed in a separate section, for which the increasing outcome data have become available. Started with our pioneering experience, still representing one of the largest in the world, PGT for HLA typing has become a method of choice in considering the treatment regiments for bone marrow failures, requiring HLA-compatible stem cell transplantation treatment. As the majority of couples requesting PGTHLA are of advance reproductive age, our experience in overcoming this problem will be presented, which will help to avoid the potential problems in older PGT-HLA patients. As there is still some controversy in the utility of PGT for aneuploidy, mainly due to the procedure accuracy in the past based on the use of FISH technique, a special section will be devoted to the outcome data that have been obtained since the shift of embryo biopsy to the blastocyst stage and the introduction of NGS-based 24-chromosome aneuploidy testing. This will also include the available RCT data, showing the obvious clinical impact of preselection of aneuploidy-free embryos. However, it is also obvious that not all the preselected euploid embryos have a potential to implant, so additional contributing factors will be addressed, such as cytoplasmic DNA contents, time-lapse parameters, genetic expression profile related to the euploid embryo competence, endometrial receptivity, and possible epigenetic influences of IVF-related procedures. The special emphasis will be on sub-­ chromosomal variations, with additional sections on segmental aneuploidies and mosaicism, which were not detected previously with the previously available inferior technologies but were shown to affect implantation and pregnancy outcome. Significant improvement has been also achieved in PGT for chromosomal rearrangements, involving the application of next-generation technologies, which requires a significant updating of the corresponding section as well. In

Preface

Preface

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addition, special emerging technologies for distinguishing normal from balanced rearrangements will be described, such as mate-pair sequencing, with presentation of the available practical experience. As the application of NGS allows PGT for both chromosomal rearrangements and aneuploidy, the reproductive impact of such a combined PGT will be evaluated. It should be, however, noted that PGT is still an invasive procedure, and despite the lack of any detectable damage and considerable improvement of biopsy techniques with the shift from cleavage to blastocyst stage, the potential negative effect of embryo biopsy-based PGT cannot be totally excluded. So the current experience for attempting a noninvasive PGT will be described, including approaches with the use of blastocoel fluid and spent culture medium, as well as a possibility of PGT without IVF, with their limitations and possible practical applications in the future. Also, a recent progress in the development of noninvasive prenatal tests (NIPT) for the application to PGT will be demonstrated, as a realistic follow-up procedure after PGT. Thus, a dramatic progress above in PGT and related areas requires an update in all sections, with substantial revision in PGT for both single-gene and chromosomal disorders. These updates are illustrated by our 30  years pioneering experience, including 6,204 PGT-M cycles for as many as 581 different monogenic disorders, currently performed together with PGT-A in the majority of cases. This and other large experiences in PGT-A, PGT-SR, and PGT-HLA, based on the use of the next-generation technologies, demonstrated an extremely high accuracy, safety, and reliability of PGT technology, which represents a practical PGT for improving the standard of ART and genetic practices. Chicago, IL, USA Chicago, IL, USA Miami, FL, USA

Anver Kuliev Svetlana Rechitsky Joe Leigh Simpson

Contents

1 Place of Preimplantation Genetic Testing (PGT) Among Available Options for Prevention of Genetic Disorders����������������   1 1.1 Preconception Prevention of Congenital Anomalies by Folic Acid Containing Multivitamin Fortification Programs������   2 1.2 Genetic History and Avoidance of Congenital Disorders by Prenatal and Preimplantation Genetic Testing ��������������������   4 1.3 Prospective Carrier Screening as a Means for Improving PGT Uptake������������������������������������������������������������������������������   6 References������������������������������������������������������������������������������������������  10 2 Major Components of Preimplantation Genetic Testing��������������  13 2.1 Introduction������������������������������������������������������������������������������  13 2.2 Polar Body Sampling����������������������������������������������������������������  15 2.2.1 Polar Body Testing as a Preconception Testing Strategy ������������������������������������������������������������������������  16 2.2.2 Analysis of Sperm��������������������������������������������������������  20 2.3 Embryo Biopsy ������������������������������������������������������������������������  21 2.4 Developments in Blastocyst Biopsy Procedures����������������������  23 2.4.1 Trophectoderm Biopsy Procedure Without Laser Assistance at Day 5 ��������������������������������������������   24 2.5 Noninvasive PGT (NIPGT)������������������������������������������������������  26 2.6 PGT Without IVF����������������������������������������������������������������������  27 References������������������������������������������������������������������������������������������  27 3 Major Components of Preimplantation Genetic Testing: Adjustment of Available Genetic Technology to PGT Practice��������������������������������������������������������������������������������  31 3.1 Adjustment of DNA Analysis to Avoid Misdiagnosis in Single-­Cell PCR��������������������������������������������������������������������  31 3.2 Other Strategies to Avoid Misdiagnosis������������������������������������  35 3.3 Is There Still Place for FISH Analysis in Current PGT-A?������  38 3.4 Microarray Analysis (Array CGH): Acceptable but Not up to Present Standards������������������������������������������������������������  40 3.5 Next-Generation Sequencing (NGS): Present Standard of PGT Practice������������������������������������������������������������������������  42 References������������������������������������������������������������������������������������������  45

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4 Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M) ����������������������������������  49 4.1 Traditional Indications and Strategies for PGT-M��������������������  75 4.1.1 Autosomal Recessive Disorders������������������������������������  75 4.1.2 Autosomal Dominant Disorders ����������������������������������  88 4.1.3 X-Linked Disorders������������������������������������������������������  95 4.2 Affected Homozygous or Compound Heterozygous Conditions �������������������������������������������������������������������������������� 101 4.3 Concomitant PGT for Two or More Single-Gene Disorders������ 107 4.4 De Novo Mutations������������������������������������������������������������������ 113 4.5 Late-Onset Common Disorders with Genetic Predisposition���������������������������������������������������������������������������� 126 4.5.1 Cancer �������������������������������������������������������������������������� 126 4.5.2 Neurodegenerative Diseases ���������������������������������������� 140 4.5.3 Cardiac Disease������������������������������������������������������������ 147 4.6 Other Genetic Conditions with Important Health-­Related Implications������������������������������������������������������������������������������ 155 4.6.1 Blood Group Incompatibility���������������������������������������� 155 4.6.2 Congenital Malformations�������������������������������������������� 158 4.6.3 Dynamic Mutations������������������������������������������������������ 167 4.6.4 Variants of Unknown Significance (VUS)�������������������� 174 4.7 Contemporary Selection of Molecular Technologies���������������� 178 References������������������������������������������������������������������������������������������ 178 5 Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)���������������������������������������� 183 5.1 Thalassemia������������������������������������������������������������������������������ 183 5.2 Immunodeficiency�������������������������������������������������������������������� 190 5.3 Preimplantation HLA Matching Without PGT-M�������������������� 199 5.4 Limitations and Future Prospects of PGT- HLA���������������������� 202 5.5 Practical Implications of PGT-HLA ���������������������������������������� 206 References������������������������������������������������������������������������������������������ 210 6 Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic Testing for Chromosomal Disorders (PGT-A and PGT-SR) �������������������������� 213 6.1 Meiotic and Mitotic Abnormalities������������������������������������������ 213 6.2 Chromosome-Specific Meiotic Error Origin and Its Impact on Embryo Viability������������������������������������������ 220 6.3 Mitotic Errors in Embryos in Relation to Meiosis Errors�������� 222 6.4 Practical Relevance of Autosomal Monosomy Detection�������� 223 6.5 Uniparental Disomies���������������������������������������������������������������� 224 6.6 Mosaicism and Sub-­chromosomal Variations�������������������������� 226 6.7 Possible Additional Tests to Preselect Euploid Embryos with Higher Potential for Pregnancy���������������������������������������� 229 6.7.1 Mitochondrial DNA������������������������������������������������������ 229 6.7.2 Morphokinetic Assessment ������������������������������������������ 232 6.8 Characterizing Chromosomal Rearrangements������������������������ 233 6.8.1 Polar Body Approach���������������������������������������������������� 233 6.8.2 Karyotyping of Embryos via Nuclear Conversion�������� 234

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6.8.3 PGT-SR Using Next-­Generation Technologies������������ 234 6.8.4 PGT-SR Developments to Distinguishing Noncarrier Balanced Embryos from Normal���������������� 236 6.8.5 Reproductive Outcome of PGT-SR������������������������������ 242 References������������������������������������������������������������������������������������������ 248 7 Clinical Outcome of Preimplantation Genetic Testing ���������������� 253 7.1 Ovulation Stimulation for a PGT Cycle������������������������������������ 253 7.1.1 Ovulation Stimulation and Aneuploidy������������������������ 253 7.1.2 Specific Disorder Characterized by Poor Ovulation Response���������������������������������������� 254 7.2 Anomalies in ART/ PGT Pregnancies and in ART Pregnancies Without PGT�������������������������������������������� 255 7.2.1 Birth Defects in ART/PGT Pregnancies ���������������������� 255 7.2.2 Birth Defects in ART without PGT������������������������������ 255 7.2.3 Pitfalls in Assessing Birth Defects in ART Offspring���������������������������������������������������������� 256 7.2.4 Birth Defects in Offspring of Subfertile Couples Not Requiring ART���������������������������������������� 256 7.2.5 Genetic Counseling for Birth Defects in ART and PGT����������������������������������������������������������� 257 7.3 Diagnostic Accuracy of PGT-M������������������������������������������������ 257 7.4 Diagnostic Accuracy and Predictive Value of PGT-A�������������� 258 7.4.1 Predictive Value of PGT-A�������������������������������������������� 258 7.4.2 Aneuploidy and Error Origin of Chromosomal Abnormalities���������������������������������������������������������������� 258 7.5 Mosaicism in PGT-A���������������������������������������������������������������� 259 7.5.1 Numerical Mosaicism �������������������������������������������������� 259 7.5.2 Segmental Mosaicism �������������������������������������������������� 260 7.5.3 Priority of Mosaic Embryos for Transfer���������������������� 261 7.6 Diagnostic Accuracy of PGT-SR���������������������������������������������� 262 7.7 Clinical Utility of PGT-A: Randomized Clinical Trials Involving 24-Chromosome Aneuploid Testing�������������� 262 7.7.1 RCTs Involving Polar Body Biopsy ���������������������������� 263 7.7.2 RCTs Involving 24-Chromosome Testing of Blastocysts���������������������������������������������������������������� 263 7.7.3 Confirming Clinical Utility of a PGT-A ���������������������� 265 7.8 Ancillary Tests to Enhance PGT-A Pregnancy Rates �������������� 265 7.9 PGT-A for Recurrent Miscarriages ������������������������������������������ 267 7.10 Why Did Cleavage-Stage RCTs Before 2010 Fail to Show Benefit of PGT-A?������������������������������������������������������ 267 7.10.1 Biopsy Damage in Cleavage-­Stage Embryos �������������� 267 7.10.2 Observational Studies and RCTs Involving Cleavage-­­Stage Embryos That Showed Benefit����������� 268 7.10.3 RCTs in Cleavage-Stage Embryos Not Showing Benefit���������������������������������������������������� 268 7.11 Universal Single Embryo Transfer to Avoid Multiple Gestations���������������������������������������������������������������������������������� 269 References������������������������������������������������������������������������������������������ 269

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8 Social, Ethical, and Legal Aspects�������������������������������������������������� 275 8.1 Legality and Regulation of PGT ���������������������������������������������� 275 8.2 PGT in Venues Having Restrictions������������������������������������������ 276 8.3 Ethical Issues Unique to PGT �������������������������������������������������� 277 8.4 PGT for Disorders with Genetic Predisposition ���������������������� 277 8.5 PGT for HLA Prediction and Stem Cell Transplantation �������� 279 8.6 PGT for Sex Selection�������������������������������������������������������������� 280 8.7 Conclusions������������������������������������������������������������������������������ 280 References������������������������������������������������������������������������������������������ 281 Index���������������������������������������������������������������������������������������������������������� 283

Contents

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Place of Preimplantation Genetic Testing (PGT) Among Available Options for Prevention of Genetic Disorders

Preventing genetic disorders and birth defects is a universal goal. Dramatic progress has been made in corrective gene therapy in recent years using CRISPR, but prevention of genetic disorders remains the main approach. Primary preventive measures are ideally applied at the community level. Examples include suitable dietary intake or avoiding toxicants that can result in new mutations. Prevention is provided also by preconception and prenatal predictive testing for genetic and complex disorders and by prospective screening for genetic disorders, including that for common conditions specific for each ethnic group or expanding carrier screening [1–2]. The ideal time for offering many preventive measures is, in fact, the preconception or preimplantation stage because detection thereafter will involve the decision either to keep the pregnancy and confront long-term social, familial, and financial consequences that arise with a seriously affected child or to terminate a planned and wanted pregnancy. The most relevant approaches for primary prevention of congenital disorders include (1) avoidance of new mutations through environmental programs, (2) reduction of maternal age related conditions through community education and family planning, (3) reduction of neural tube defects and a few other congenital malformations by periconceptional folic acid supplementation or multivitamin fortification of basic foodstuffs, (4) avoidance of alcohol and smoking during © Springer Nature Switzerland AG 2020 A. Kuliev et al., Practical Preimplantation Genetic Testing, https://doi.org/10.1007/978-3-030-43157-0_1

pregnancy, and (5) rubella vaccination. These actions can reduce congenital disorders of environmental origin through public health measures and those of biological origin through sophisticated approaches for molecularly detecting and managing individuals at genetic risk. In fact, most congenital and other complex conditions (e.g., NTD) have both genetic and environmental components; thus, the actions are first addressed to environmental causes, through the finding of the key components to modify the occurrence of congenital disorders. The decision to adopt any of the available preventive programs depends on health services development, ethnic distribution of certain congenital diseases, and local attitudes to genetic testing and termination of pregnancy. For example, induced abortions are still not permissible in many countries. The number of countries permitting prenatal genetic diagnosis and termination of pregnancies for medical indications is steadily increasing, but there is still restriction on the stage of pregnancy when termination for medical reasons can be performed. The impact of community-based preventive approaches is obvious from Down syndrome preventative programs in industrialized countries of Europe, South America, Asia, Oceania, and the United States. Pregnancies at increased risk are offered invasive testing and, if appropriate, selective pregnancy termination. Prenatal diagnosis offered to all women of advanced maternal age can result in the reduction of the birth 1

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1  Place of Preimplantation Genetic Testing (PGT) Among Available Options for Prevention of Genetic…

prevalence of Down syndrome by more than 50% [3]. These programs were initially based solely on testing by invasive prenatal procedures (amniocentesis; chorionic villus sampling), but later women of all ages were screened based on maternal serum analysis and ultrasound criteria. Currently screening increasingly involves cellfree DNA in maternal plasma (NIPT). All these actions decrease number of invasive procedures required. However, this reduction is positively correlated with the number of pregnancy terminations. In some countries the effect of such program is still growing, while in others it seems to be reaching a plateau, reflecting differences in the development of the services as well as social and religious differences. That this reduction is achieved only through pregnancy terminations is, however, a cause for serious concern [3–4]. This is particularly relevant for high-income countries in which women use family planning to postpone childbearing, leading to a rebound in a higher proportion of older mothers. The paradigmatic example of a highly effective population-based preventive measure at the primary prevention preconception stage is, as noted, prevention of neural tube defects (NTD) by folic acid or folic acid-containing multivitamins. Preimplantation genetic testing has not typically been considered analogous to traditional preventive strategies like NTD prevention by folic acid supplementation, of course, the reason being the necessity for in vitro fertilization. However, PGT is genuinely an established and realistic option for preconception prevention of genetic disorders. The role PGT plays in primary prevention for genetic disorders will be described in detail in this book.

1.1

Preconception Prevention of Congenital Anomalies by Folic Acid Containing Multivitamin Fortification Programs

Despite the need for integrating programs and combining all feasible approaches maximizing the benefits and minimizing the negative aspects

of preventive programs for congenital malformations, the ideal is a primary preventive measure. The paradigmatic example is represented by vitamin/folic acid supplementation to prevent NTD and congenital disorders [5–14]. The success of this effect deserves attention. The strategy is folic acid dietary supplementation and folic acid food fortification (FFI (flour fortification initiative), www.Sph.emory.edu/wheatflour). Application of these approaches has resulted in the overall reduction of NTD by as much as by half (from 40.6 per 1000 to 20.6 per 1000). Supplementation or fortification with folic acid or folic acid-containing multivitamins not only reduces the frequency of NTD by 75% but may reduce the population prevalence of other congenital disorders: cardiovascular, urinary tract, and limb deficiencies. More data are needed to determine efficacy of reducing birth defects other than NTD, such as pyloric stenosis. Still, the positive impact of folic acid in reducing congenital anomalies is in agreement with the following: 1. Mothers who give birth to a child with neural tube defects have mildly elevated blood and amniotic fluid levels of homocysteine. 2. Hyperhomocysteinemia and/or lack of methionine can induce neural tube defects in animal experiments. 3. Low maternal folate status is associated with increased risk for neural tube defects. 4. Vitamins of B group, including folate/folic acid, are important in homocysteine metabolism. 5. Vitamins B6, B11, and B12 are also able to reduce hyperhomocysteinemia. 6. Homocysteine accumulates if its conversion to methionine is slowed with a shortage of folate or vitamin B12 or both. Elevated plasma homocysteine indicates suboptimal nucleic acid and amino acid metabolism and results in increased risk of cardiovascular disease through thickening the lining of blood vessels [15–18]. Folate deficiency is usually related to genetic factors. In many populations, approximately 20% are homozygous for a common polymor-

1.1  Preconception Prevention of Congenital Anomalies by Folic Acid Containing Multivitamin…

phism of the enzyme methyl tetrahydrofolate reductase (MTHFR). Valine replaces alanine at codon 677, reducing the enzyme activity in homozygotes by 50–70% and increasing risk of neural tube defects. Folic acid supplementation increases the supply of tetrahydrofolate, accelerates most folate-dependent metabolic reactions, and reduces plasma homocysteine levels [19–22]. In over 90% of pregnancies in which the fetus has a neural tube defect, there is no previous indication of increased risk. The only identifiable risk group consists of (a) women with a prior affected pregnancy, who have a 3–4% recurrence risk, and (b) women who are heterozygous for the MTHFR mutation. However, these groups account for only a small proportion of affected pregnancies. Trials of the effect of folic acid supplementation on the prevalence of neural tube defects were needed to generate conclusive scientific evidence for its preventive effect [5–14]. Recommendation evolved to recommend that (1) dietary supplementation with folic acid or with multivitamin preparations containing folic acid, before and during early pregnancy (periconceptional supplementation), markedly reduces both the first occurrence of neural tube defects and recurrence among women having had a previously affected pregnancy (3–4% risk); (2) benefit is greatest in regions in which a high baseline prevalence of neural tube defects is present, but efficacy exists also in lower prevalence areas; and (3) no harmful effects have been observed, based on levels of supplementation ranging from 360 μg to 5 mg of folic acid daily. Most recent data show that no harmful effects have been seen even with the extreme doses of folic acid [23]. The recommended current dietary folate intake for adults in the United States is 400 μg,

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representing a daily intake of 200 μg folic acid equivalents. The additional intake needed for pregnant women is recommended to be 600 μg [24]. The UK Committee on Medical Aspects of Food and Nutrition Policy recommendation of 240  μg of folic acid per 100  g flour [25] is approximately equivalent to an additional folic acid intake of 200 μg a day. To obtain adequate protection against risk of neural tube defects, the mean plasma folate should approximate 10 ng/ml. The mean plasma folate level in most populations is only 5  ng/ml [24–27]. There is considerable variation within and between populations, but a few individuals have a mean plasma folate in the recommended (IU  ng/ml) range. Thus, food fortification alone needs to be provided as well as folic acid supplements. Groups of greatest likelihood for folate intakes and plasma folate being at the lower end of the range are those residing in regions lacking food fortification. Folic acid fortification of all cereal grain products at a level of 140  μg/100  g flour has been mandatory in the United States and Canada since 1998 (Table 1.1) [28–31]. Since introduction, birth prevalence of neural tube defects has fallen by about 19–32%, with no adverse effects reported [28–33]. In Hungary, folic acid, vitamin B12, and vitamin B6 were added to bread in 1993, with average daily intake of folic acid, vitamin B12, and B6 from this source approximately 200 μg, 1 μg, and 1080 μg [7]. The prevalence of neural tube defects in Hungary has fallen by 41%. Mandatory fortification of wheat flour with folic acid are in place in 53 countries, with maximum and minimum level of folic acid set according to the World Health Organization guidelines [28–29]. The potential global estimate of reduction of congenital disorders is presented

Table 1.1  Global estimate of reduction of neural tube defects (NTD), congenital heart disease (CHD), and limb reduction defects (LRD) by folic acid (FA) food fortification Potential annual affected births Est’d annual affected births with FA Born malformation-free with FA Based on Crider et al. [31]

All three conditions 1,035,604 659,090 376,513

NTDs 388,442 137,402 251,040

CHD /LRD 647,162 521,689 125,473

NTDs % 37.5 20.8 66.7

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1  Place of Preimplantation Genetic Testing (PGT) Among Available Options for Prevention of Genetic…

Table 1.2  Reported fall in prevalence of neural tube defects following folic acid food fortification Country/region The United States and Canada Hungary Costa Rica Chile South Africa (urban) Oman

Year fortification started 1998

% fall in NTD 19–32

1998 1998 2000 2003 1996

41 19–35 19–35 19–35 44

Based on Crider et al. [31]

in Table 1.2. The available actual figures support these estimates (see Table  1.1), although there were also conflicting data from updated US data, which may be due to potential misclassification and bias that can attenuate the measured association of folic acid supplementation with neural tube defects [30–32]. Overall, the data suggest at least 5–6 per thousand of NTDs and other malformations mentioned are the lowest achievable in these primary preventive measures, with ­conservative estimate of up to 300,000 NTDs preventable worldwide [31]. In addition, folic acid fortification is highly cost-effective [28, 33]. The cost is only about $1 per metric ton of flour, so low that the extra cost is insufficient to change the price of a loaf of bread. Again, however, the multivitamin food fortification does not substitute completely for periconception supplementation nor for multivitamin supplementation during pregnancy. Taking into consideration the estimated lifetime cost for a single patient with spina bifida ($250,000), complementary periconception supplementation program is highly costeffective, the major benefit being avoidance of an affected child. Available experience from those countries which have implemented a national food staff fortification program is presented in the abovementioned Table 1.1. In summary, food fortification and preconception vitamin supplementation programs are a paradigmatic prevention that has maximized numbers of healthy babies.

1.2

Genetic History and Avoidance of Congenital Disorders by Prenatal and Preimplantation Genetic Testing

Prevention requires one to inquire into the health status of first-degree relatives (siblings, parents, offspring), second-degree relatives (nephews, nieces, aunts, uncles, grandparents, or grandchildren), and third-degree relatives (maternal and paternal first cousins). Similar family history taking is required for sperm or oocyte donors and their family members. A positive family history for a genetic disorder may indicate need for prenatal or preimplantation testing. Most couples undergoing PGT for a monogenic disorder (PGT-M) will have been ascertained in this fashion. On the other hand, the couples undergoing PGT because of a prior affected child is low compared to the general population unaware of their 25% risk. Few will have undergone genetic screening. This gap is a major deficiency, to be discussed in this book. Family history may reveal at-risk individuals in other ways. For example, a seconddegree relative (e.g., grandparent) may have an autosomal dominant disorder. This should warrant physical examination of the potential parents (grandchildren) presenting for prenatal or preimplantation testing. Subtle and previously unappreciated clinical features in the couple could indicate presence of a transmitted mutant gene. For example, multiple café au lait spots connote neurofibromatosis, an autosomal dominant disorder. A history of adverse reproductive outcomes should be sought, namely, queries for miscarriages, stillbirths, or as already noted liveborn infants with anomalies. This may require genetic tests. A woman having recurrent miscarriages carries an increased likelihood for a balanced chromosomal translocation if she is 95%) but not necessarily 100%. Still, a negative screening test reduces considerably the likelihood that an individual is a carrier and, hence, lowers risk for having an affected offspring. The American College of Obstetricians and Gynecologists (ACOG) has long recommended a selected number of disorders for ethnicity-based carrier screening [34]. American College of Medical Genetics and Genomics (ACMG) offers recommendations that are similar, but not identical recommendations. Disorders recommended for screening before conception or early in pregnancy were traditionally restricted to specific ethnic groups and based on gene products (protein), as DNA methodology did not exist at that time. At present, panethnic carrier screening intermingles protein- and DNA-based methods for greatest efficiency in carrier detection. Updated recommendations have been summarized in joint statements by multiple organizations: ACOG, National Society of Genetic Counselors (NSGC), ACMG, and Perinatal Quality Foundation [35–36]. Carrier screening may concurrently involve both potential parents. However, cost-effective approach is to test the partner of greater risk (e.g., based on family history of the disease of interest). If one partner has a mutation that could result in an autosomal recessive disorder, the next step is to test the other partner. It is perhaps preferable to test both partners concurrently if they are of equal risk. Any couple being screened should be informed of their options: invasive prenatal

diagnosis (chorionic villus sampling (CVS) or amniocentesis), PGT-M, donor gametes (eggs or sperm), or adoption. All would avoid offspring with the at-risk disorder. Among the original carrier screening programs were those for Tay-Sachs disease in individuals of Ashkenazi Jewish ancestry. Screening for Tay-Sachs disease initially was based on ratios of the gene products hexosaminidase A and B, the former deficient in carriers and almost absent in affected individuals. Proteinbased functional assays are still applicable and sometimes essential if one partner is not Jewish. However, DNA-based screening in the Jewish populations is highly efficient because only a few mutant alleles account for most heterozygotes. Over 95% of heterozygotes in the Jewish population are accounted for by: ΔF508, W1282X, G542X, N1303K, and 3849  +  10kbC  →  T.  In non-Jewish individuals, the same mutations are not far less predictable. ACOG later recommended carrier screening in Jewish populations for Canavan disease and familial dysautonomia (IKBKAP). “Consideration” was recommended for other conditions prevalent in this population: mucolipidosis IV; Niemann-Pick disease type A; Fanconi anemia types A, B, C; Bloom syndrome; Gaucher disease (GBA); Jourbert syndrome; familial hyperinsulinemia (ABCC8); maple syrup urine integrated (BCKDHA, BCKDHB, DBT) disease; and Usher syndrome. By contrast with ACOG, ACMGG explicitly recommends offering Niemann-Pick (type A), Bloom syndrome, Fanconi anemia type C, mucolipidosis IV, and Gaucher disease. DNA panels are utilized for all these conditions, with carrier detection rates of 95–99% in the Jewish population. Carrier frequency and detection for non-Jewish partners are less established in these Ashkenazi disorders. Unlike Tay-Sachs disease, there are no suitable protein-based methods, for which reason screening is less efficient in couples of mixed ethnicities. ACOG Committee Opinions 691 and 690 detail recommendations [35–36]. Carrier screening programs for the hemoglobinopathies were developed in the 1970s [37]. Sickle cell disease is prevalent among individuals of African or African-American ori-

1.3  Prospective Carrier Screening as a Means for Improving PGT Uptake

gin, 1 in 12 being carrier for sickle cell anemia. β-Thalassemias are prevalent among Greeks, Italians (Sicilians), Turks, Arabs, Southern Iranians, Azerbaijanis, and Asian Indians. β-Thalassemia carrier detection is based on anemia that is not due to iron deficiency origin: mean corpuscular volume (MCV) is less than 80  fL, and iron saturation levels are normal. Diagnosis is confirmed by hemoglobin electrophoresis. Hemoglobin electrophoresis will reveal diminished hemoglobin B and increased hemoglobin F.  Detection of α-thalassemia requires DNA testing. Cystic fibrosis was the first disorder screened solely by DNA testing. In 2001 American College of Medical Genetics (ACMG) and ACOG proposed a DNA panel consisting of 23 mutations [38]. Screening was “offered” only to Caucasians and Ashkenazi Jews on grounds of prevalence being higher than in Hispanic, African-American, and Asian populations. For these ethnicities, screening was “made available.” The rationale was that carrier frequencies and, hence, detection rates were much lower in the latter ethnic groups. Lower heterozygote rates and lower detection rates meant that likelihood of heterozygosity not only was lower, but likelihood of being a heterozygote despite a negative screen was also lower. In other words, screening was less efficient. Recommendations were later updated later by ACOG [39], ACMG, and NSGC [40]. At present all these organizations now recommend carrier screening for all ethnicities [35, 36]. Spinal muscular atrophy (SMA) has long been under discussion. Carrier frequencies for autosomal recessive SMA are 1 in 40 to 1 in 60 in most populations [41], albeit lower in Hispanics [42]. The causative gene undergoing mutations causing SMA is survival motor neuron 1 (SMN1). Based on carrier rates being similar to cystic fibrosis, ACMG has long recommended population screening for SMA. Initially ACOG did not recommend population screening, but the 2017 Committee Opinion does so [35]. Fragile X syndrome (FMR1) is an X-linked disorder that is the most common inherited form of intellectual disability in males (1 in 3600). In

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the United States, the carrier frequency in women with no known risk factors is approximately 1 in 250. The molecular basis is expansion of trinucleotide CGG repeats in the FMR1 gene. Affected individuals have >200 GGT repeats. In females with 55–200 repeats, gametes may expand during meiosis to result in >200 and thus offspring with FMR1. Such individuals are said to have a “premutation.” Population-based carrier screening for fragile X is recommended by neither ACOG, ACMG, nor NSGC. Screening in Europe and North America is recommended for women having a family history of fragile X, but not in the general population [43]. In Israel screening is more common. How effective is genetic screening in preventing birth of offspring with genetic disorders? Prenatal genetic diagnosis and PGT-M as traditionally practiced have marginally reduced the number of births with these monogenic disorders in the population. However, many couples are at risk but do not realize their status. Approximately half of all individuals heterozygous for an Ashkenazic specific mutant had not self-reported themselves as Jewish [1]. In 2017 ACOG explicitly codified that “ethnic-specific, panethnic, and expanded carrier screening are (all) acceptable strategies for pre-pregnancy and prenatal carrier screening” [35]. Expanded carrier screening means all ethnicities, more genes, and increased information on genes screened. In 2015 joint counseling recommendations were published by ACOG, ACMG, NGSC, SMFM, and Perinatal Quality Foundation. Such a strategy has become feasible by availability of the broad spectrum of sequencing methodology. This approach is panethnic. Panethnic screening entails Expanded carrier screening. ACOG explicitly states that “ethnic-­ specific, panethnic, and expanded carrier screening are acceptable strategies for prepregnancy and prenatal carrier screening” [35, 36]. Many more conditions are screened and the depth of screening is greater. In expanded carrier screening, several hundred additional genes are considered plausible candidates [44, 45]. Costs are no longer a major impediment. Copy number variants (CNV) are also

1  Place of Preimplantation Genetic Testing (PGT) Among Available Options for Prevention of Genetic…

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sought in certain genes traditionally difficult to sequence: FMR1, SMN1, congenital adrenal hyperplasia, and α-thalassemia. Detection rates are thus increased. Interrogating for CNV in cystic fibrosis and Duchenne/Becker muscular dystrophy has increased heterozygote detection to almost 100%. In the 2018 reported results of the Counsyl, using expanded (Foresight™) panel consisting of 254 genes, 1 in 22 c­ ouples was at risk for a detectable disorder; 1  in 300 fetuses was affected [45]. These advances have impacted PGT-M, increasingly following expanded carrier screening programs, as was demonstrated by our ongoing PGT-M experience, which represents the world’s largest series in one center. In 2016, 38% of PGT-M cases at our center were ascertained by carrier screening. That is, the majority were ascertained following an affected proband. Two years later in 2018, the majority (63%) of PGT-M cases were ascertained by carrier screening (Fig.  1.1) [46]. The dynamics of increase of at-­risk couples presenting for PGT-M through expanded carrier screening was observed for each genetic condition tested, compared to the baseline referrals through the traditional approach. Among the

most frequently referred conditions were cystic fibrosis (CFTR), the uptake of which increased from 50% in 2016 to 82.7% in 2018 (Fig. 1.2); deafness (GLB2) with the uptake growing from 31.2% in 2015 to 86.4% in 2018 fragile X (FMR1), the uptake growing from 73% in 2016 to 84% in 2018; and thalassemia and sickle cell disease (HBB), increasing from 45.8% in 2016 to 60.9% in 2018. Thus, the overall number of prospective PGT-M cases for the last 3 years more than doubled after referral through expanded screening, with a similar dynamics for each condition tested. This may become the major source for performing PGT-M in the near future, allowing to offer PGT-M prospectively before the birth of an affected child. The data show significant increase of the PGT-­ M uptake following expanded carrier screening, demonstrating the utility for offering PGT-M prospectively to the couples at risk. Gene panels for various adult-onset heritable disorders are also becoming common. This has led to at-risk couples wishing gamete and embryo to be tested, resulting in increased PGT-M. Greatest applicability of PGT-M for adult-­ onset heritable disorders lies in heritable cancers.

500 450 400 350 63%

300 250

53%

200 39%

150 100 50 0 2016

2017

total requests for PGT-M

Fig. 1.1  Impact of expanded carrier screening on PGT-M uptake

2018 referred after carrier screening

1.3  Prospective Carrier Screening as a Means for Improving PGT Uptake

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60

50 82% 76%

40

70%

30

20

50%

10

0 2015

2016 total requests for CF PGT-M

2017

2018

referred after carrier screening

Fig. 1.2  Increase in PGT-M requests for cystic fibrosis (CFTR gene) after expanded carrier screening

The most recently reported population genomic screening for multiple conditions involved 2,688,192 young adults aged 18–25 years with the purpose of disease prevention [47]. This appeared to be highly cost-effective in significantly reducing the incidence and mortality of hereditary cancers and the burden of severe childhood-­onset genetic diseases, compared with targeted testing. The NIH National Cancer Institute lists of Familial Cancer Susceptibility Syndromes numbers 39. Many are of adult-onset, autosomal dominant in inheritance. Interest is often initiated when a family member is found to have a heritable cancer and the mutation is identified. Unaffected relatives of reproductive age naturally seek to learn if they themselves have the same cancer-susceptible mutation. If this proves to be the case, avoiding transmission of their mutant alleles to offspring is sought through PGT-M.  PGT-M is preferable to invasive prenatal genetic diagnosis because an ongoing pregnancy monitored by CVS or amniocentesis carries a 50% likelihood for an affected fetus; repeated clinical pregnancy terminations may be needed to achieve a normal offspring. By contrast, PGT-M allows selection of an unaffected embryo to transfer without fear of an affected

clinical pregnancy. Despite obvious benefit, PGT-M for adult-­onset cancers is diagnostically and emotionally complex. Chapter 4 will provide this strategy. Heritable cancers most often subjected to PGT include breast cancer, familial adenomatous polyposis 1 (FAP1), Fanconi anemia, neurofibromatosis, tuberous sclerosis, and hereditary nonpolyposis colon cancer (HNPCC) (see Chap. 4). Adult-onset PGT-M is also increasingly utilized for adult-onset autosomal dominant cardiac disorders. Most common are Long QT syndrome (LGT1, LGT2, LGT8), hypertrophic cardiomyopathy (CMH1, CMH4, CMH8), and dilated cardiomyopathy (Type 1A, 1DD, 1E, 1G). Monogenic cardiac disorders also commonly subjected to PGT-M include Holt-Oram syndrome and Noonan syndrome, to mention only a few, described in Chap. 4). In conclusion, the main objective of screening or prenatal genetic diagnosis is to assist couples to have an unaffected child of their own. PGT does this by embryo selection. We have noted that prenatal genetic diagnosis during an ongoing pregnancy is efficacious but requires termination of affected pregnancies. That is, prevention is

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1  Place of Preimplantation Genetic Testing (PGT) Among Available Options for Prevention of Genetic…

secondary. Pregnancy termination is not tolerated in many communities nor ethnic groups and is undesirable by all. Providing an option for couples at risk assuring that nonclinical pregnancy is affected is the goal. For this reason PGT has already become an integral part of preventive services for congenital disorders, providing a choice for those couples who are unable to accept prenatal screening and termination of pregnancy. PGT as the preferred form of prevention is performed under this assumption. Less appreciated is that PGT can accomplish primary prevention in certain circumstances, i.e., preconception PGT for primary prevention; PGT represents an important component of ART and genetic practices. PGT management usually includes embryo selection with discarding affected embryos. However, in exceptional cases PGT can identify a transferable embryo without even having to discard or permanently cryopreserve an affected embryo. Examples will be provided in Chap. 2.

References 1. Edwards JG, Feldman G, Goldberg J, et al. Expanded carrier screening in reproductive medicine-points to consider. Obstet Gynecol. 2015;125:653–62. 2. Ben-Shachar R, Svenson A, Goldberg JD, Mussey D.  A data –driven evaluation of size and content of expanded carrier screening panels. Genet Med. 2019; https://doi.org/10.1038/s41436-019-0466-5. 3. Stoll C, Alembik Y, Dott B, Roth MP. Impact of prenatal diagnosis on livebirth prevalence of children with congenital anomalies. Annales de Genetique. 2002;45:115–21. 4. EUROCAT report No 8. Surveillance of congenital anomalies in Europe 1980–1999. Edited by EUROCAT Working Group. EUROCAT Central Registry, Room 1F08, University of Ulster Newtownabbey, County Antrim, Northern Ireland BT37 0QB. Email eurocat@ ulster.ac.uk. 5. MRC Vitamin Study Research Group. Prevention of neural tube defects: results of the Medical Research Council vitamin study. Lancet. 1991;338:131–7. 6. Center for Disease Control and Prevention. Use of folic acid for prevention of spina bifida and other neural tube defects- 1983–1991. MMWR. 1991;40:513–6. 7. Czeizel AE, Dudás I. Prevention of the first occurrence of neural tube defects by periconceptional vitamin supplementation. N Engl J Med. 1992;327:1832–5.

8. US Department of Health and Human Services, Food and Drug Administration. Food standards: amendment of standards of identity for enriched grain products to require addition of folic acid. Fed Regist. 1996;61:8781–807. 9. Czeizel AE.  Primary prevention of neural-tube defects and some other major congenital abnormalities. Pediatr Drugs. 2000;2:437–49. 10. Wald NJ, Noble J. Primary prevention of neural tube defects. In: Rodeck CH, Whittle MJ, editors. In fetal medicine: basic science and clinical practice. London: Churchill Livingstone; 1999. p. 283–90. 11. Berry CJ, Li Z, Erickson JD.  Prevention of neural-­ tube defects with folic acid in China. N Engl J Med. 1999;341:1485–90. 12. Selhub J, Jacques PF, Rosenberg IH, et  al. Serum total homocystein concentrations in the third National Health and Nutrition Examination Survey (1991– 1994): population reference ranges and contribution of vitamin status to high serum concentrations. Ann Intern Med. 1999;131:331–9. 13. Ruddell LJ, Chisholm A, Williams S, Mann JI. Dietary strategies for lowering homocystein concentrations. Am J Clin Nutr. 2000;71:1448–54. 14. Olney RS, Mulinare J.  Trends in neural tube defect prevalence, folic acid fortification, and vitamin supplement use. Semin Perinatol. 2002;26:277–85. 15. Wald DS, Law M, Morris JK.  Homocysteine and cardiovascular disease: evidence on causality from a meta-analysis. BMJ. 2002;325:1202. 16. Schnyder G, Roffi M, Pin R, Flammer Y, Lange H, Eberli FR, Meier B, Turi ZG, Hess OM. Decreased rate of coronary restenosis after lowering of plasma homocysteine levels. N Engl J Med. 2001;345:1593–600. 17. La Vecchia C, Negri E, Pelucchi C, Franceschi S. Dietary folate and colorectal cancer. Int J Cancer. 2002;102:545–7. 18. McIlroy SP, Dynan KB, Lawson JT, Patterson CC, Passmore AP. Moderately elevated plasma homocysteine, methylenetetrahydrofolate reductase genotype, and risk for stroke, vascular dementia, and Alzheimer disease in Northern Ireland. Stroke. 2002;33:2351–6. 19. Klerk M, Verhoef P, Clarke R, Blom HJ, Kok FJ, Schouten EG, MTHFR Studies Collaboration Group. MTHFR 677C-->T polymorphism and risk of coronary heart disease: a meta-analysis. JAMA. 2002;288(16):2023–31. 20. Shields DC, Kirke PN, Mills JL, Ramsbottom D, Molloy AM, Burke H, Weir DG, Scott JM, Whitehead AS. The “thermolabile” variant of methylenetetrahydrofolate reductase and neural tube defects: an evaluation of genetic risk and the relative importance of the genotypes of the embryo and the mother. Am J Hum Genet. 1999;64:1045–55. 21. Brody LC, Conley M, Cox C, Kirke PN, McKeever MP, Mills JL, Molloy AM, O’Leary VB, Parle-McDermott A, Scott JM, Swanson DA. A polymorphism, R653Q, in the trifunctional enzyme methylenetetrahydrofolate dehydrogenase/methenyltetrahydrofolate cyclohydro-

References lase/formyltetrahydrofolate synthetase is a maternal genetic risk factor for neural tube defects: report of the Birth Defects Research Group. Am J Hum Genet. 2002;71:1207–15. 22. Moat SJ, Ashfield-Watt PA, Powers HJ, Newcombe RG, McDowell IF.  Effect of riboflavin status on the homocysteine-lowering effect of folate in relation to the MTHFR (C677T) genotype. Clin Chem. 2003;49:295–302. 23. Wald NJ. Folic acid and neural tube defects. In: Walter P, Hornig D, Moser U, editors. Functions of vitamins beyond recommended dietary allowances. Bibl. Nutr Dieta. Basel: Karger; 2001. No 55. p. 22–33. 24. Sheehy TW.  Folic acid: lack of toxicity. Lancet. 1973;1:37. 25. Department of Health. Report on Health and Social Subjects. 50. Folic Acid and the Prevention of Disease: Report of Committee on Medical Aspects of Food and Nutrition Policy. The Stationery Office, London. 2000. 26. Moore LL, Bradlee ML, Singer MR, Rothman KJ, Milunsky A.  Folate intake and the risk of neural tube defects: an estimation of dose-response. Epidemiology. 2003;14:200–5. 27. Raats M, Thorpe L, Hurren C, Elliott K.  Changing preconceptions: the HFEA folic acid campaign 1995– 98. London: Health Education Authority 2; 1998. 28. Report of WHO\EURO Meeting on Development of EURO Strategy on Congenital Disorders Minsk, November 29–30, 2001 Unpublished WHO/EURO Document (#51203630). WHO Copenhagen, Denmark. 29. Allen L, de Benoist B, Dary O, Hurrell R, editors. Guidelines on Food Fortification with Micronutrients. 1st. World Health Organization; Portland, OR, USA: 2006. Annex D-A procedure for estimating feasible fortification levels for a mass fortification programme; p. 294–312. 30. Viswanathan M, Treiman KA, Kish-Doto J, et  al. Folic acid supplementation for the prevention of neural tube defects. An updated evidence report and systematic review for the US Preventive Services Task Force. JAMA. 2017;317:190–203. 31. Crider KS, Bailey LB, Berry BJ. Folic acid food fortification—its history, effect, concerns, and future directions. Nutrients. 2011;3:370–84. 32. Centers for Disease Control and Prevention, authors. CDC grand rounds: additional opportunities to prevent neural tube defects with folic acid fortification. MMWR Morb Mortal Wkly Rep. 2010;59: 980–4. 33. Report of the WHO/EURO Meeting on the Regional Policy for Prevention of Congenital Disorders. Folic acid: from research to public health practice. Rome, Italy, 11–12 November 2002. Instituto Superiore di Sanita, Rome, Italy. 34. Driscoll DA, Simpson JL, Holzgreve W, Otaño L.  Genetic screening and prenatal genetic diagno-

11 sis. In: Gabbe SG, Niebyl JR, Simpson JL, Landon MB, Galan HL, ERM J, Driscoll DA, Berghella V, Grobman WA, editors. Obstetrics: normal and problem pregnancies. 7th ed. Philadelphia: Elsevier; 2017. p. 193–218. 35. American College of Obstetricians and Gynecologists. Carrier screening for genetic conditions. Committee opinion no. 691. Obstet Gynecol. 2017;129(3): e41–55. 36. American College of Obstetricians and Gynecologists. Carrier screening in the age of genomic medicine. Committee opinion no. 690. Obstet Gynecol. 2017;129(3):e35–40. 37. American College of Obstetricians and Gynecologists. Hemoglobinopathies in pregnancy. ACOG practice bulletin no. 78. Obstet Gynecol. 2007;109:229–37. 38. Grody WW, Cutting GR, Klinger KW, et al. Laboratory standards and guidelines for population based cystic fibrosis screening. Genet Med. 2001;3:149–54. 39. American College of Obstetricians and Gynecologists. Update on carrier screening for cystic fibrosis. Committee opinion no. 486. Obstet Gynecol. 2011;117:1028–31. 40. Langfelder-Schwind E, Karczeki B, Strecker MN, Redman J, Sugarman E, Zaleski C, et  al. Molecular testing for cystic fibrosis carrier status practice guidelines: recommendations of the National Society of Genetic Counselors. J Genet Couns. 2014;23: 5–15. 41. Prior TW, Professional Practice and Guidelines Committee. Carrier screening for spinal muscular atrophy. Genet Med. 2008;10:840–2. 42. ACOG Committee on Genetics. ACOG Committee opinion no. 432: spinal muscular atrophy. Obstet Gynecol. 2009;113:1194–6. 43. American College of Obstetricians and Gynaecologists Committee on Genetics. ACOG Committee opinion no. 469: carrier screening for fragile X syndrome. Obsetet Gynecol. 2010;116:1008–10. 44. Haque IS, Lazarin GA, Kang P, Evans EA, Goldberg JD, Wapner RJ. Modeled fetal risk of genetic diseases identified by expanded carrier screening. JAMA. 2016;3016:734–42. 45. Hogan GJ, Vysotskaia VS, Beauchamp KA, Seisenberger S, Grauman PV, Haas KR, et  al. Validation of an expanded carrier screen that optimizes sensitivity via full-exon sequencing and panel-wide copy number variant identification. Clin Chem. 2018;64:1063–73. https://doi.org/10.1373/ clinchem.2018.286823. 46. Simpson JL, Rechitsky S, Kuliev A. Before the beginning: the genetic risk of a couple aiming to conceive. Fertil Steril. 2019;112:622–30. 47. Zang L, Bao Y, Riaz M, et  al. Population genomic screening of all young adults in a health-care system: a cost-effectiveness analysis. Genet Med. 2019; https://doi.org/10.1038/s41436-019-0457-6.

2

Major Components of Preimplantation Genetic Testing

2.1

Introduction

tural rearrangements (PGT-SR) has a clear advantage over the traditional prenatal diagnosis in Preimplantation genetic testing (PGT) is now an assisting these couples to establish an unaffected established clinical option in reproductive medi- pregnancy and deliver a child free from unbalcine [1–3]. Tens of thousands of PGT cases have anced translocation [1, 30–33]. Reproductive been performed in hundreds of centers around outcomes depend in turn governing efficiency of the world, allowing at-risk couples to avoid pro- achieving an unaffected pregnancy on the origin ducing offspring with genetic disorders. More and type of translocation. The majority result in importantly, children have been healthy, validat- early fetal loss and rarely in an affected birth; ing no ostensible evidence incurred by embryo thus, it may take years until the translocation carbiopsy or embryo culture (see Chap. 7). riers are fortunate enough to have an unaffected Applied first in 1990 for preexisting Mendelian offspring; thus, current recommendations of diseases [4, 5], namely, cystic fibrosis (CF) and PGDIS, ESHRE and ASRM Practice Committee X-linked disorders, PGT initially did not seem to include chromosomal rearrangements as one of be practical. Only a few babies were born during the main indications for PGT [34]. The experithe first 3 years, and several misdiagnoses were ence of thousands of PGT-SR cycles accumureported [6, 7]. After the introduction of fluores- lated to date demonstrates a fourfold reduction of cent in situ hybridization (FISH) analysis in spontaneous abortions in these couples, com1993–1994 for PGT of chromosomal disorders pared to their experience before PGT [32–37]. In [8–13] (Chap. 6), the number of PGT cycles addition, carrier couples can avoid transfer of began to double annually, yielding more than 100 translocation carrier embryos; approaches develunaffected children by the year 1996 [14, 15]. oped to distinguish carrier embryos from the norApplication of PGT increased further when mal ones may currently be offered to carriers of the ability to detect chromosomal rearrangements balanced translocations (Chap. 6). became possible in 1996, first using locus-­ The natural extension of PGT’s ability to specific FISH probes, then more widely available allow transfer of euploid embryos (PGT-A) sub-telomeric probes [16, 17] (Chap. 6), haplo- would be expected to have positive impact on the typing and microarray technology (array CGH) liveborn pregnancy outcome, especially in poor [18–29], and presently next-generation sequenc- prognosis IVF patients (prior IVF failures, matering (NGS) (Chap. 3). Because many carriers of nal age over 37, repeated miscarriages). balanced translocations have a low likelihood of Introduction of commercially available FISH having an unaffected pregnancy, PGT for struc- probes in 1998–1999, followed a decade later by © Springer Nature Switzerland AG 2020 A. Kuliev et al., Practical Preimplantation Genetic Testing, https://doi.org/10.1007/978-3-030-43157-0_2

13

14

2  Major Components of Preimplantation Genetic Testing

current 24-chromosome testing by microarray analysis and NGS, have led to the accumulated experience of tens of thousands of PGT-A cycles worldwide [35, 37–41] (Chaps. 6 and 7), demonstrating the usefulness of PGT-A in assisted reproduction practices. According to the experience of centers, the overall pregnancy rate per transfer is higher than that in non-PGT IVF patients of comparable age groups, although the details of its application to different patient groups are still debated (see Chap. 7). The current IVF practice of transferring embryos based solely on morphological criteria is inefficient, given that half of these embryos are chromosomally abnormal and would compro­ mise the reproductive outcome (Chap. 7). Introduction of 24-chromosome testing combined with blastocyst biopsy and current strategy of single embryo transfer further improves reproductive outcome in poor prognosis IVF patients, confirming the need for preselection of euploid embryos for transfer [18–29]. The application of PGT has further expanded with its introduction to late-onset diseases with genetic predisposition [42] (Chap. 4), an indication that had never been considered for the traditional prenatal diagnosis. For patients with inherited pathological adult-onset predisposition, PGT provides a realistic reason for undertaking pregnancy. Despite 50% risk, offspring without genetic predisposition to the disease can be obtained. Prospective parents at such risk and their physicians should be aware of this option, especially when there is no opportunity to diagnose the disease until it is fully manifested (Chap. 4). Another unique option of PGT is HLA typing as a component of PGT (PGT-HLA) [43] (Chap. 5). In this application PGT offers not only preventative technology to avoid an affected offspring but also an approach for treating (older) siblings with congenital or acquired bone marrow diseases for which there is still no other therapy. This may in future be applied to any condition that can be treated by embryonic stem cell transplantation.

PGT- HLA was first applied to couples desiring of having an unaffected (younger) child free from the genetic disorder in the older sibling. In addition to diagnosis to assure a genetically normal embryo, HLA-matched, unaffected embryos were chosen. At delivery cord blood (otherwise to be discarded) was gathered for stem cell transplantation. As will be described, this approach has been also used without testing of the causative gene, with the sole purpose of finding a matching HLA progeny for a source of stem cell transplantation for affected siblings with congenital or acquired bone marrow disease or cancer [44] (see Chap. 5). As will be described in this book, 30 years of PGT experience has demonstrated considerable progress. Hundreds of thousands of PGT attempts worldwide have resulted in birth of a large cohort of apparently unaffected children, with no detrimental effect on embryo development. There are no significant differences in the overall congenital malformation rate after PGT compared to population prevalence [45–47] (Chap. 7). With the highly improved accuracy of genetic analysis and indications expanding well beyond those for prenatal diagnosis, up to hundred thousand PGT cycles are now performed annually. This reflects PGT offering a special attraction not possible with traditional prenatal diagnosis, namely, avoiding clinical pregnancy termination. As mentioned, this is extremely useful for translocation carriers, couples at risk for producing offspring with common diseases of autosomal dominant or recessive etiology, and couples wishing to have not only an unaffected child but also an HLA-­compatible stem cell donor for treatment of an older moribund sib with a congenital disorder. Yet the greatest numerical impact of PGT involves assisted reproduction practices to increase pregnancy rates (Chaps. 6 and 7). The estimated number of ART centers using PGT solely for this purpose is well over one-third of cycles in the United States. Thus, improved IVF efficiency through aneuploidy testing has become standard, despite the technology being quite sophisticated.

2.2  Polar Body Sampling

Special expertise and training in the main components of PGT is required. Pivotal to PGT is obtaining biopsy material from oocytes and embryos. Biopsy material for performing PGT may be obtained from three major sources: 1. Matured and fertilized oocytes from which the first and second polar body (PB1 and PB2) are removed 2. Eight-cell cleavage-stage embryo, from which a single blastomere is removed 3. Blastocyst-stage embryo, from which not less than 5, but not more than 10, cells are removed Material obtained is tested for single-gene disorders using PCR analysis, or used for chromosomal abnormalities, previously done by fluorescent in situ hybridization (FISH) and now by microarray (array CGH) or next-generation sequencing (NGS) (Chaps. 3  and 6). Each of these PGT methods has its advantages and disadvantages, with method selected depending on circumstances; in some cases combination of these methods may be required. Despite reduction in embryo cell number after biopsy, having a potential deleterious influence on embryo viability, blastomere or blastocyst biopsy allows detection of paternally derived abnormalities. On the other hand, removal of PB1 and PB2 should not have much effect on the embryo viability as polar bodies are naturally extruded from oocytes as result of maturation and fertilization. Polar bodies provide no information on the paternally derived anomalies, even if this constitutes less than 5% of chromosomal errors in preimplantation embryos.

2.2

Polar Body Sampling

Introduced almost 30 years ago [5], PB biopsy is still one of the alternative approaches in PGT, although its application has been limited to ethnic and social groups that cannot accept embryo biopsy or wish to avoid technical problems in certain PGT indications. The rationale for per-

15

forming PGT by the use of PB is based on the fact that PBs are the by-products of female meiosis and allow predicting by deduction of the resulting genotype of the maternal contribution to the embryo. Neither PB1, extruded as a result of the first meiotic division, nor PB2, extruded following the second meiotic division, has any known biological value for pre- and postimplantation development of the embryo. Initially, only PB1 was tested, based on the fact that in the absence of crossing over, PB1 will be homozygous for the allele not contained in the oocyte and PB2 [48, 49]. However, the PB1 approach was not applicable for predicting the eventual genotype of the oocytes if crossing over had occurred, because the primary oocyte in this case would be heterozygous for the mutant gene. Frequency of crossing over varies with the distance between the locus and the centromere, approaching as much as 50% for telomeric genes. Thus, PB1 approach is of limited value, unless the status of the oocyte can be deduced by PB2, which allows detecting hemizygous normal oocytes resulting after the second meiotic division. As will be described below, this PGT technique involves a two-step oocyte analysis, with a sequential testing of PB1 and PB2 (see details of micromanipulation setup and procedure steps elsewhere [48]). In brief, PB1 and PB2 are removed following stimulation and oocyte retrieval using a standard IVF protocol. Following extrusion of PB1, the zona pellucida (ZP) is opened mechanically using a microneedle, or laser, and PB1 aspirated into a blunt micropipette. Oocytes are then inseminated with motile sperm, or using intracytoplasmic sperm injection (ICSI), and examined for the presence of pronuclei and extrusion of PB2, which is removed in the same manner as PB1. To avoid an additional invasive procedure, both PB1 and PB2 may be removed simultaneously, fixed, and analyzed on the same slide (acceptable only for FISH analysis). However, for PGT-M, PGT-SR, and PGT-A by microarray and NGS analysis, PB1 and PB2 are removed sequentially as mentioned above. The biopsied oocytes  are then fertilized,  returned to culture,

16

2  Major Components of Preimplantation Genetic Testing

checked for cleavage, and transferred, depending on the genotype of the corresponding PB1 and PB2 [48]. As mentioned, PB1 and PB2 have no any known biological significance in pre- and postimplantation development to affect embryo viability, as also shown by the follow-up study. In this study, it was demonstrated that following the procedure, zygotes with two pronuclei were observed in 1192 (81.8%) of 1458 oocytes, compared to 30,972 (77.3%) of 40,092 in a routine non-PGT cycles, suggesting no difference in fertilization rate observed after PB1 removal in comparison with non-biopsied oocytes. There was also no difference in blastocyst formation of the embryos resulting from the biopsied oocytes. Blastocyst formation of embryos resulting from biopsied oocytes was observed in 1653 (50.2%) of 3293 embryos, not different from 49.8% (9726 of 19,529) non-biopsied embryos observed in routine IVF. Similarly, no detrimental effect was noted after PB2 removal, which was evident from cleavage rate, blastocyst formation, and the ­number of cells in the respective blastocysts [50]. As will be seen below, there was no difference after a sequential PB1–PB2 and embryo biopsy.

combined with freezing of the oocytes at the pronuclear stage. After analysis, the oocytes predicted of having the normal maternal allele may be thawed and cultured to allow the pronuclear fusion, embryo development, and transfer  in a subsequent menstrual cycle. In fact, it is possible to complete the testing of PB2 in approximately 9 hours after removal. This avoids the need for freezing of the mutation or aneuploidy-free oocytes, allowing continued culture as usual and replacement on day 3 or day 5; abnormal oocytes are frozen at the pronuclear stage or discarded [51]. Because zygotes are not considered to be embryo until pronuclear fusion, and no abnormal oocytes are thawed and cultured, the establishment of the affected embryos is obviated; thus, this technique may be ethically more acceptable to many couples. This technique creates a new class of genetic testing, which may be called pre-embryonic genetic testing (PEGT), pushing the frontier of genotyping to an even earlier stage. The first attempt of PEGT in testing the feasibility of the approach included sickle cell anemia and Sandhoff disease (SHD). PEGT was performed for a 33-year-old woman and her spouse at risk for producing a child with sickle cell disease. The couple could not accept neither a possible termination of a pregnancy following prenatal diagnosis nor any 2.2.1 Polar Body Testing manipulation of the embryo. A standard IVF proas a Preconception Testing tocol was used, but the patient suffered hyperStrategy stimulation syndrome, which precluded transfer of embryos from that cycle. Twenty-eight mature Although not used on a large scale, PB as a pre- oocytes were aspirated and placed in culture conception strategy may have selected utility. In medium. Of the 28 aspirated oocytes, 14 extruded certain venues  – Austria, Germany, Switzerland PB1s, which were removed. The oocytes were (until recently), and Malta  – PGT has been then fertilized by intracytoplasmic sperm injecrestricted to micromanipulations only prior to tion. As soon as the PB2s were removed and prior fertilization. This strategy is also applicable to to the fusion of the male and female pronuclei, all certain religious groups. While laws are being the oocytes were frozen. PB1 and PB2 were anaevolved in some of these communities, PB1-­ lyzed by multiplex nested PCR to ensure detectbased testing still remains an option for haplo- ing a potential allele dropout (ADO), which type analysis in PGT for de novo mutations of occurs in approximately 5–10% of cases in the maternal origin (Chap. 4). PB1 testing is not suf- PB analyses (see Chap. 3). This involved a nested, ficient to predict embryo genotype, unless PB2 is multiplex PCR with primer sets for the sickle cell tested before pronuclear fusion. This may be mutation and two linked short tandem repeat

2.2  Polar Body Sampling

(STR) markers, one located at the 5′ end of the beta-globin gene (5′ STR) and the other in the human tyrosine hydroxylase gene (THO-STR), for both of which the mother was heterozygous. To detect potential contamination with extraneous DNA and identify the embryo that implanted and established a pregnancy, additional non-­ linked STRs were amplified (the details of this first preconception PGT case were described elsewhere [48, 52]). The pronuclear-stage oocytes predicted to be normal were thawed, cultured to develop into the cleaving embryos, and transferred back to the patient in the two subsequent clinical cycles. The oocytes predicted to contain the mutant maternal gene were not thawed, but analyzed directly at the pronuclear stage for the confirmation of PB diagnosis. Following intracytoplasmic sperm injection of 14 oocytes with extruded PB1, PB2s were extruded from 13 of them, with the results of both PB1 and PB2 available in 12 of these 13 oocytes. Overall, six oocytes were predicted to contain a normal allele, based on heterozygous status of PB1 and hemizygous mutant status of PB2.The frozen cycle with the transfer of two unaffected embryos resulted in a singleton pregnancy and birth of unaffected child, following confirmation of PB diagnosis by chorionic villus sampling (CVS). The second case of PEGT was done without pronuclear stage freezing, based on the technological possibilities to complete testing before pronuclear fusion. It was offered to a 32-year-old woman and her spouse, who were at risk for producing a child with Sandhoff disease (SHD) and specifically requested PGT to be performed without any possible discard of embryos even if affected [51]. As seen from the pedigree shown in Fig.  2.1, the couple had one affected son with classical features of SHD, who died at the age of 1  year and 3  months despite bone marrow transplantation. SHD results from the defect in the beta chain of hexosaminidase B gene (HEXB) on chromosome 5, which consists of 14 exons distributed over 40 Kb of DNA (MIM 268800; 606,873).

17

Mutation in this gene causes beta-­hexosaminidase deficiency, resulting in the lysosomal storage disease GM2-gangliosidosis. The same condition is caused by Tay–Sachs disease resulting from the defect of hexosaminidase A gene (HEXA). The child inherited two different mutations from his parents: the paternally derived I 270 V mutation in exon 5 of HEXB gene, resulting from ATT to GTT substitution, and a large maternal 16Kb deletion (16Kb Del), involving as many as 5 exons, from exon 1 to exon 5. The paternal mutation was identified by the Hinf I restriction digestion, which cuts the normal allele into two fragments of 32 and 25  bp, leaving the mutant allele uncut, and the maternal 16Kb Del detected by a fragment size analysis. Five closely linked polymorphic markers, D5S1982, D5S1988, D5S2003, D5S349, and D5S1404, were tested simultaneously with the HEXB gene in a multiplex heminested PCR system. A single PGT cycle was initiated, which was performed according to the modified timetable of the applied procedures of sequential PB1 and PB2 analysis. PB1 was removed as usual 3.5 hrs after aspiration, followed by ICSI. PB2 was removed soon after it was extruded, approximately within 6.5 hrs after ICSI, to allow sufficient time for the completion of the DNA analysis before pronuclear fusion (see Fig. 2.2). DNA analysis was done in less than 9 hrs overall, making it realistic to freeze the oocytes predicted to contain the deleted HEXB allele before syngamy (within 24  hrs after aspiration or 12  hrs after PB2 removal), and culture the HEXB deletion-free oocytes to blastocyst and transfer at day 5, following confirmation of the maternal mutation-­free status of the embryos by the embryo biopsy. Of 18 oocytes available for testing in a single PEGT cycle, 16 showed conclusive PB1 and PB2 results, of which 8 contained the maternal 16Kb deletion and frozen at the pronuclear stage (Fig.  2.1). Four of these oocytes contained heterozygous PB1 and normal PB2 (oocytes #3, # 9, #11, and #14) and four homozygous normal PB1 and mutant PB2 (Fig. 2.1b). The remaining eight

2  Major Components of Preimplantation Genetic Testing

18

oocytes were free of the deletion, two originating from the oocytes with heterozygous PB1 and mutant PB2 (oocytes # 1 and # 5) and the others from the oocytes with homozygous mutant PB1 and normal PB2. As the predicted genotypes in these oocytes may erroneously appear opposite, due to a possible undetected ADO of one of the alleles in the actually heterozygous PB1, similar to the four mutant oocytes predicted on the basis of homozygous normal PB1 and mutant PB2, the testing for five closely linked polymorphic markers was essential, confirming all the predicted oocyte genotypes mentioned.

a

117 206 I 207V 130 102 165

Family pedigree

A follow-up blastomere analysis of the embryos deriving from the oocytes predicted to be free of maternal deletion showed complete correspondence to the PB testing. Six of these embryos appeared to contain also a normal paternal allele (embryos #1, 4, 5, 6, 8, and 10), while only two (embryos #16 and 18) inherited the paternally derived mutant allele, confirmed by all five linked polymorphic markers tested (Fig. 2.1). Results showed that PEGT is a realistic option for couples who cannot accept traditional PGT, because of their objection to micromanipulation

115 117 203 201 16 Kb Del N 132 128 98 98 185 185

115 194 N 116 104 193

1.1 115 117 203 206 I 207V 16 Kb Del 132 130 98 102 185 165

b

1.2

PGT

2.1

2.2

32 y.o. 115 194 N 116 104 193

Markers order: D5S1982 D5S1988 HEXB 16Kb Del HEXB I207V D5S2003 D5S349 D5S1404

117 201 N 128 98 185

Sequential Polar Body Analysis 1

3

117 201 N 128 98 185

115 203 Del 132 98 185

N

c

4

5

6

7

117 201 N 128 98 185

117 201 N 128 98 185

117 201 N 128 98 185

115 203 Del 132 98 185

117 201 N 128 98 185

115 203 Del 132 98 185

M

N

M

N

M

N

N

5

6

8

9

10

11

13

14

15

117 201 N 128 98 185

115 203 Del 132 98 185

115 203 Del 132 98 185

115 203 Del 132 98 185

115 203 Del 132 98 185

N

M

M

M

M

16

17

18

117 201 N 128 98 185

115 203 Del 132 98 185

117 201 N 128 98 185

N

M

N

Blstomeres Analysis 1

115 117 194 201 N N 116 128 104 98 193 185 Normal

ET

4

115 117 194 201 N N 116 128 104 98 193 185 Normal

Fr

115 117 194 201 N N 116 128 104 98 193 185 Normal

Fr

115 194 N 116 104 193

117 201 N 128 98 185

Normal

Fr

8

115 117 194 201 N N 116 128 104 98 193 185 Normal

Fr

Fig. 2.1  Pre-embryonic testing for Sandhoff disease. Upper portion, showing pedigree (a) results of polar bodies (b) and blastomeres (c); and Figure 2.1. Bottom portion, showing the actual DNA analysis used: schematic presentation of the mutation and linked polymorphic markers (A); polar body analysis of the maternal 16 Kb

10

115 194 N 116 104 193

117 201 N 128 98 185

Normal

ET

16

117 206 I 207V 130 102 165

117 201 N 128 98 185

Carrier

Fr

18

117 206 I 207V 130 102 165

117 201 N 128 98 185

Carrier

Fr

deletion (N – normal; D – deletion) (B); Restriction map: HhaI enzyme created 2 fragments in normal gene, leaving the paternal mutation I 207V uncut (C); and Blastomere analysis for maternal deletion and paternal mutation, confirming the PB diagnosis (D)

2.2  Polar Body Sampling

19

HEXB

A

I 207 V

D5S1982

1

D5s1988

0.72 cM

3

2

4 5

6

0.52 cM

Exon

D5S349

1.08 cM

D5S1404 1.26 cM

16 Kb Deletion

Intron

C

87

Normal allele Mutant allele

32

87

L

PB1 PB2 PB1 PB2 PB1 PB2 PB1 PB2 F M C

L

L

PB1 PB2 PB1 PB2 PB1 PB2 PB1 PB2 PB1 PB2

Sequential Polar Body Analysis for 16 Kb Deletion in HEXB gene PB1 PB2 PB1 PB2 PB1 PB2 PB1 PB2 PB1 PB2 PB1 PB2

Paternal mutation I 207V (ATT -GTT) Mutant Allele 5’ TGGTTGATA…. 3’ Normal Allele 5’ TGATTGATA..... 3’ 3’ end of 4R primer 3’ AGTAT…5’ Hinf I restriction site G’ANTC

513 bp Deletion allele

B

D5S2003

0.55 cM

25

15

57

15

Deletion (513 bp) Normal (117bp)

1 N

3 D

4 N

5 N

6 N

7 D

8 N

9 10 11 13 14 D N D D D

analysis D ConfirmationalforBlastomere the

15 16 17 18 D N D N

Blastomere analysis for the paternal mutation I 207 V

maternal 16 Kb deletion L

L

B1 B4 B5 B6 B8 B10 B16 B18 F M C

Predicted Genotype

B1 B4 B5 B6 B8 B10 B16 B18 F M C Uncut

Oocyte No.

Del

Invariant N

(87 bp)

Mutant (57 bp)

Fig. 2.1 (continued)

Fig. 2.2  Timeframe for pre-embryonic diagnosis of Sandhoff disease (see explanation in the figure)

hCG

–

8 p.m.

Aspiration

–

7 a.m. (on the second day)

PB1 removal

–

11.30 a.m.

ICSI

–

12 p.m.

PB2 removal

–

6.30 p.m.

DNA testing

–

7 p.m. – 4 a.m.

Fertilization control

–

12.30 a.m. & 6 a.m.

Freezing (of mutant oocytes at pronuclear stage)

–

6.30 a.m.

Blastomere biopsy (of embryosfree of maternal mutation)

–

day 3

Transfer of unaffected embryos

–

day 5

35 h 35 h

65 h 24 h 18 h 9h

20

2  Major Components of Preimplantation Genetic Testing

and potential discard of the tested embryos. The previous PEGT case described involved freezing of all the tested oocytes at the pronuclear stage immediately after ICSI and extrusion of PB2. The present case is proceeded without freezing of the mutation-free oocytes, which were detected well before the pronuclear fusion and prior to when a decision to discard could not be avoided. Although all oocytes could have been frozen irrespective of DNA diagnosis (as in the previous case), recovering all frozen pronuclear stage oocytes might not be possible. Those not recovered may have included preselected unaffected embryos that if not transferred could have negatively affected the PEGT outcome. PEGT in the same clinical cycle is clearly an important practical step, which has become realistic because of DNA analysis being completed within less than 9 hrs. PEGT may be also applied for aneuploidy, as given the great majority of chromosomal disorders deriving from the female meiosis and testable by PB analysis. Available experience is presently limited to translocation or aneuploidy testing by PB1 analysis, which, as mentioned, leaves meiosis II errors undetected. As seen from the present results, detection of the second meiosis errors is currently feasible within the timeframe available prior to pronuclear fusion; thus, PEGT for chromosomal disorders may in future be also applied in those countries where PGT is still not acceptable because of the potential discard of the affected embryos with the currently used methods. Presented data demonstrate feasibility of performing PEGT for single-gene disorders, which resulted in obtaining unaffected pregnancies and birth of healthy children. Of course, PGT-M may be performed by the use of PB1 analysis alone, as described in the first case of PGT by PB1 [5]. Although this allowed preselection of a few mutation-free oocytes inferred from the homozygous abnormal status of PB1, the majority of oocytes were heterozygous after the first meiotic division, so the genotype of the resulting embryos could not be predicted, thus limiting the number of normal embryos for transfer. Data also show that to avoid discard of preimplantation embryos reaching the cleavage stage

by the time the PB genotyping results were obtained, freezing of oocytes may be applied immediately after ICSI and extrusion of PB2, as well as prior to fusion of the male and female pronuclei (the actual point considered to be the beginning of the embryonic period of development [53]). In fact, freezing may be omitted entirely, as developments in PCR analysis allow completing the genetic diagnosis before pronuclei fusion. This opens a possibility for application of PGT for couples who are unable to accept any intervention and discard of the human embryos.

2.2.2 Analysis of Sperm No method has yet become available for testing the outcome of male meiosis because genetic analysis destroys the sperm, rendering it useless for fertilization. To overcome this problem, an original technique has been introduced, allowing duplicating a sperm before genetic analysis. One of the duplicated sperms can be used for testing whereas the other for fertilization and consequent transfer of the resulting embryos, i.e., provided genetic analysis of the corresponding duplicate shows normal genotype [54, 55]. In this way the establishment and discard of any embryo containing paternal mutation may be avoided. However, more data are necessary to define special conditions required for the faithful replication of human sperm genome, i.e., assuring that the haploid cell pairs obtained from sperm duplication are identical. Still lacking is validation for preconception testing and application to exclude paternally derived mutations. The genotype of sperm may be also tested following the testicular biopsy culture and tracking the developmental progression of spermatocytes through meiosis in  vitro. This provides the possibility of meiosis outcome analysis to infer the genotype of the resulting sperm to be used for fertilization. However, this is still not practical. There have also been attempts to approach preconception testing through development of artificial gametes, using techniques of somatic cell haploidization

2.3  Embryo Biopsy

induced by introduction of somatic cells into enucleated oocytes (ooplast) [56–59]. However, 90% of the resulting haploid nuclei have chromosomal aneuploidies. Haploidization of somatic cells may be achieved using MII oocyte cytoplasm, but the aneuploidy rate is much higher than in the natural meiosis process; thus, these techniques are not acceptable at present for clinical practice.

2.3

Embryo Biopsy

Embryo biopsy may be performed as soon as the embryo reaches a minimum of 6–8 cells, thus avoiding a decrease in cell number at the later stages of development. A mechanical opening of zona pellucida has been developed, called 3D-PZD, allowing the creation of a V-shaped triangular flap or square flap opening, sufficient in size for a micropipette to pass through in order to remove a single blastomere. Micromanipulation dishes are prepared the same way as for PB1 removal, except sucrose is eliminated. The micromanipulation setup is also the same as for PB1 removal with one exception that a micropipette of larger diameter (25–30  μm) is needed, as detailed elsewhere [48]. With the current tendency of shifting from the cleavage stage to blastocyst transfer [60], increasing numbers of PGT cycles are performed by blastocyst biopsy for genetic and chromosomal disorders, resulting in thousands of unaffected clinical pregnancies and births of apparently healthy children. Blastocyst biopsy was initially performed by mechanical methods, involving a smooth aspiration of several trophectoderm cells herniated through the zona pellucida into biopsy pipette with internal diameter of 30  μm [48]. However, most centers use a laser procedure, which is applied to break down the tight junctions between trophectoderm cells, followed by aspiration of 5–10 trophectoderm cells (see below). Blastocyst biopsy is increasingly accompanied by improved freezing techniques, particularly vitrification. Blastocyst (trophectoderm) biopsy also has important implications with the

21

application of the next-generation technologies. Given sufficient DNA, microarray [23, 28, 29, 62] and NGS provide much more accurate and reliable results when performed on blastocyst biopsy sample (Chap. 3). Follow-up studies of embryos after embryo biopsy have not shown any detrimental effect. No increase in congenital malformation has been reported among thousands of children born either following PB or embryo biopsy, although further systematic studies are necessary to monitor the clinical outcomes of PGT using different sampling procedures (Chap. 7). According to the data presented in Fig.  2.3, there seems to be no difference in developmental potential of embryos following single, double, or triple biopsy procedures, compared to the rate of blastocyst formation following ICSI (control group). No significant differences were observed either in blastocyst formation between the embryos with one or two biopsy procedures, even with prior PB1 and PB2 removal on day 1. There was no decrease, and perhaps even an increase, in the rate of blastocyst formation when embryos with three biopsy procedures (PB1 and PB2 sampling, folowed by embryo biopsy on day 3) were compared to a control group. The blastocyst formation rate after single or double biopsy was 68%. This is in agreement with pregnancy outcome data (see below), showing a pregnancy rate of PGT-M cycles by sequential PB1 and PB2 analysis, followed by blastomere biopsy being similar or even higher than in non-PGT cycles. Even safer than blastomere biopsy is blastocyst biopsy, as seen from a well-designed randomized control study, comparing embryo viability after blastomere and blastocyst biopsy [61]. The current shift to blastocyst transfer has resulted in an improvement in outcomes of PGT-M and PGT-A. Blastocyst biopsy may be performed prior to the expanded blastocyst stage by mechanical methods, namely, after trophectoderm starts herniating through the zona pellucida [48]. Several trophectoderm cells (5–10 cells) are removed mechanically by aspiration with a biopsy pipette, or after breaking down the tight junctions between trophectoderm cells by laser. An average blastocyst contains more than 100 cells, so

2  Major Components of Preimplantation Genetic Testing

22

2003−2004 70 60 50 40

a

d 51.4

30 20 10 0

n = 12,022

b

c 44.6 n = 56

58.3

49.9

n = 2,341

n = 547

34.3 ±4.6 34.6 ±6.1 37.8 ±3.8 32.8 ±4.3 Blastocyst rate

Mean age

ICSI ICSI / embryo biopsy ICSI / PB1 & PB2, embryo biopsy ICSI / PB1, PB2, embryo biopsy

Fig. 2.3  Effect of micromanipulations on embryo development. Bar graph demonstrating nondetrimental effect of biopsy procedures on the rate of blastocyst formation following intracytoplasmic sperm injection (control group). No significant difference was seen between embryos after (1) embryo biopsy procedure of a single blastomere and also embryos in which (2) biopsy procedures were performed in which the first and second polar bodies were removed simultaneously on day 1 (pronu-

clear stage) followed by the removal of a single blastomere on day 3, when compared to the control group. There was no decrease, but a significant increase in the rate of blastocyst formation when compared to the control group, for the group of embryos in which three biopsy procedures were performed. The first polar body was removed prior to ICSI, the second polar body was removed at the time of fertilization assessment, and a single blastomere was removed on day 3

removal of 5–10 cells from the trophectoderm is unlikely to have a detrimental effect on the blastocyst development and particularly on the development of the fetus originating from the inner cell mass. Still, removal of more than ten cells is avoided to exclude any potential damage. Of course, a major advantage of this method is that more than one cell provides amounts of genetic material more suitable for testing. Having DNA from more cells to work with prevents hybridization and amplification failures. It also detects mosaicism and segmental variations if coupled with a high-resolution NGS, thereby increasing the reliability and accuracy of the testing. However, time for analysis is limited by the implantation window of the blastocyst, which is less than 24 hours. This requires freezing embryos with subsequent transfer after thawing. Another limitation is the need for an optimum laboratory setup with culture conditions providing an acceptable rate of blastocyst development; thus, proper laboratory standards are required to pro-

duce maximum embryos reaching the blastocyst stage. Success rate varies greatly among different laboratories. Several culture systems and media have been developed for culturing embryos to the blastocyst stage in  vitro, involving a defined sequential culture media [62]. Sequential media pairs consist of one medium for culture from day 1 to day 3 and a second medium for culture from day 3 onward to blastocyst. The media are purported to mimic the in vivo environment (oviduct and uterus, respectively) and also to reflect changes in the metabolic patterns of the embryo. However, a special culture medium was then designed (Global media  – modified KSOMAA media for human embryos) that is capable of supporting human embryos to develop from zygote to the blastocyst stage [63, 64]. The latter approach, according to our experience, provides an average success rate of over 60% in culturing embryos to blastocyst stage. One of the advantages of culturing embryos to blastocyst stage is that the embryo passes the

2.4  Developments in Blastocyst Biopsy Procedures

natural self-correction mechanisms, overcoming the natural errors of the cleavage stage; thus, testing is limited only to those achieving the blastocyst stage. Although usually day 5 or day 6 blastocysts are sampled, available experience suggests that day 7 blastocysts can also be utilized, albeit with a lower success rate. In our current experience, 72.7% of blastocysts were biopsied on day 5, 27% on day 6, and only a few on day 7. The rates of aneuploid embryos were 45.3% on day 5 and 57.4% on days 6 and 7. The other important development in the PGT arrangements is that all the biopsied blastocysts are frozen, using the progress in vitrification to overcome the timeframe limitation for shipping samples for genetic analysis, with the transfer of tested blastocysts in a subsequent frozen–thawed cycle. In contrast to the traditional slow freezing procedure used previously as a standard, the currently applied vitrification procedure is a very rapid freezing approach with use of high concentration of cryoprotectant to solidify without ice crystal formation. Vitrified embryos seem almost unchanged morphologically after warming, but sufficient experience is required to achieve acceptable embryo development after thawing. For vitrification the embryos are placed in a loading device surrounded by vitrification media, which is placed directly into liquid nitrogen where it is stored. There are a variety of loading devices available, including the Cryotop, Cryoloop, CryoTip, Cryoleaf, High Security straws, and Rapid-I.  Success of vitrification  – measured by survival, implantation, and pregnancy rate  – depends not only on the loading devices used but also on blastocyst selection, media protocol, freezing and warming rates, and operator-dependent factors [65]. According to our experience, blastocyst survival and pregnancy rate after vitrification is over 60%, similar for different age groups.

2.4

Developments in Blastocyst Biopsy Procedures

The conventional trophectoderm biopsy procedure is demonstrated in Fig. 2.4. In brief, trophectoderm biopsy is performed at day 5 or 6

23

Fig. 2.4 Standard procedure

laser-assisted

blastocyst

biopsy

(rarely at day 7) of development, after the trophectoderm starts to herniate through the zona pellucida. Several trophectoderm cells are removed by aspiration of herniated trophectoderm cells into a biopsy pipette of internal diameter 25–27 μm passed through zona pellucida, opposite the inner cell mass. To break down the tight junctions between trophectoderm cells, 2–3 laser blasts are applied (duration of 0.4 mSec pulse each at 100% power) (Fig. 2.4). In selection of embryos for blastocyst biopsy, poor-quality blastocysts and those with early stage of herniation are avoided. A detailed description of the laserassisted hatching and blastocyst biopsy is described in detail in our Atlas of Preimplantation Genetic Diagnosis [48]. An important step for blastocyst biopsy is assisted hatching, performed by laser to create a hole or trench in the embryo zona pellucida. Laser-assisted hatching (laser zona drilling) is performed on day 3 or 4 of embryo development (at the 8- to 32-cell stage) inside of the lead of 35  ×  10  mm Petri dish in a 5  μl drop of 10% Global media, covered by 2  ml of mineral oil (LifeGlobal). A 10–25 μm hole is made, using a laser pulse width of 400–500 μs, to create a hole wide enough to breach the zona pellucida (as seen at the focal plane, when the embryo is observed at its largest diameter). The optimal diameter of the hole is determined by the thickness of the zona pellucida. Holes of wide diameter are necessary to accommodate a thick zona, requiring one or more laser pulses. Holes of smaller diameter are preferred for thin zona. To

24

2  Major Components of Preimplantation Genetic Testing

minimize the risk of damage, as few laser pulses as possible are administered at the lowest power levels possible to achieve safe zona drilling or thinning effects. The main components of the procedure are: 1. Laser-assisted hatching is performed at the cleavage or morula stage on day 3 or 4 of embryo development. The embryo zona pellucida is drilled with 1 or 2–3 ZILOS-tk pulses at the high setting (laser pulse width of 500 μs), with the diameter of the drilled hole at 25–30 μm. 2. At the expanded blastocyst stage, day 5, 6, or 7 of embryo development, when trophectoderm is herniating through the hole, the embryo is transferred into the lead of 35x10 mm Petri dish with 5 μl drops of 20% Global media covered with 2 ml of oil. Each blastocyst is separated – one per biopsy dish – and placed into appropriately numbered (at the bottom) drop in the Petri dish. 3. The blastocyst is supported on a holding pipette and held at the floor of the Petri dish. Using a polished thin-walled biopsy pipette with internal diameter 25–27 μm, 5–10 cells are drawn into the pipette and pulled away from the embryo. Separation from the rest of the trophectoderm is achieved by 2–5 pulses of the laser at the medium setting (laser pulse width 350–500  μs), aimed at the junction of the cells near the biopsy pipette, with biopsy time not exceeding 3 minutes. 4. The biopsied cells are transferred into the empty 5 μl drop of 20% Global media in the same Petri dish at 3 o’clock position. 5. After the blastocyst has been biopsied, the dish is taken back to the dissecting scope and, under supervision of a witness, individually placed into the appropriately numbered culture drop in the separate growth dish which had been equilibrated at least 2 hours. 6. Trophectoderm samples are transferred into properly labelled micro-centrifuge tubes containing 2  μl of buffer using an individual attenuated glass pipettes with inner diameter of 120–140 μm, or plastic stripper tip with inner diameter of 135 μm. Samples

are transferred within a minimum possible volume of media, which does not exceed 0.5  μl, the transfer pipettes being changed between every sample. Micro-centrifuge tubes are handled the same way as for PCR testing. As mentioned above, the follow-up studies of embryos after embryo biopsy did not show any detrimental effect, in agreement with the outcome of thousands of PGT-M cycles.

2.4.1 Trophectoderm Biopsy Procedure Without Laser Assistance at Day 5 This technique was developed to overcome the major disadvantages of a regular laser-assisted trophectoderm biopsy, which include: (1) potential damage due to removal of excess trophectoderm cells; (2) stickiness (adherence) of the resulting blastocyst edge in the obtained biopsied material, leading to an increased chance of a loss during the sample transfer into the testing tube; and (3) possibly damaged nuclei by laser bean, interfering with correct interpretation of embryo genetic status, which may contribute to a mosaicism rate reported in PGT-A. The technique depends on use of the force of surface tension on the boundary of culture media of the biopsy drop with mineral oil, in lieu of laser pulses (Fig. 2.5). The method can be applied only on grade 2 to 2–3 of day 5 blastocysts, given the tight junction between trophectoderm cells of early blastocyst (grades 2 and 2–3) is not as strong as cell connection in more advanced blastocyst (grades 3–4, 4, and higher). Grade 2 and 2–3 blastocysts are placed into 5 mkl equilibrated culture medium drops covered by 2  ml of equilibrated mineral oil, held by holding pipette on the left side, while the biopsy pipette (ID 25 mkm) on the right side is used to suck 5–10 extruded trophectoderm cells into the pipette. The embryo is then moved to the right edge of the biopsy drop with the biopsy pipette, moving slowly toward the inside of the oil environment, making cytoplasm bridge

2.4  Developments in Blastocyst Biopsy Procedures

25

Fig. 2.5  Non-laser-assisted blastocyst biopsy procedure (see explanation in the text)

between blastocyst and sample thinner, until it is finally removed and placed into a separate 5 mkl culture medium drop in the same dish. The usefulness of the method was evaluated by amplification efficiency of biopsied samples that totaled 55 early day 5 blastocysts, compared to the results of 87 conventional laserassisted biopsies on day 5–6 blastocysts. Significant differences were observed in the amplification efficiency, from 84% to 96.4%. In addition, the described biopsy procedure results in a non-­sticky TE samples with no evidence of damaged nuclei. To apply this technique to day 6 or 7 blastocysts, this non-laser-assisted trophectoderm biopsy is combined by laser assistance. Advanced grade 3–6 blastocysts are placed into 5 mkl equilibrated culture medium drops covered by 2 ml of equilibrated mineral oil, held by holding pipette on the left side; the biopsy pipette (ID 25–27

mkm), held on the right side, aspirates 5–10 cells of the herniated trophectoderm into the pipette. The biopsied embryo is then first moved to the right edge of the biopsy drop with the biopsy pipette and then slowly toward the inside of the oil environment to make a cytoplasm bridge between blastocyst and sample thinner. In this position separation from the rest of the trophectoderm requires only 1–2 pulses of the laser at the 400  μs pulse length (versus 2–5 pulses when doing conventional laser-assisted trophectoderm biopsy)  to minimize biopsied trophectoderm cells damage. The described biopsy procedure also resulted in obtaining non-sticky trophectoderm samples with no evidence of damaged nuclei, with the improved amplification efficiency, and avoiding a potential impact on technically derived mosaicism, possibly  contributing to different mosaicism rates between various PGT centers.

2  Major Components of Preimplantation Genetic Testing

26

2.5

Noninvasive PGT (NIPGT)

Although there is no evidence for detrimental effect of biopsy procedures on embryo viability, prudence dictates that any potential damage be minimized. Thus, research into truly noninvasive approaches to PGT procedures is pursued, analogous to options for prenatal genetic diagnosis in ongoing clinical pregnancies. Aspiration of Blastocoel Fluid  Aspiration of blastocoel fluid has been proposed as in theory being less invasive than trophectoderm biopsy. However, this is not proved. Accuracy in particular yet has to be demonstrated [66]. Although usefulness of blastocoel fluid for PGT by different groups lacks consensus, its use has been proposed to identify at-risk embryos from younger patients, who otherwise have no accessible indication for PGT-A [66]. The analogy to amniocentesis is attractive but very arguable. Insufficient data on the concordance of blastocoel-derived DNA and trophectoderm sample is available to justify diagnostic application of blastocoel fluid.

Spent Culture Medium  A noninvasive approach for PGT could use analysis of “spent” culture medium [66–69]. This genuinely noninvasive approach [67, 68] is based on cell-free DNA in culture media and is analogous to monitoring cell-free DNA in maternal plasma during pregnancy. Although attractive, data are preliminary and show limitations. One of the first reports was based on testing of 55 samples, of which only six had sufficient DNA levels to be tested for aneuploidy by whole-genome amplification (WGA) and oligonucleotide array comparative genomic hybridization (array CGH). The authors claimed the agreement between invasive and noninvasive tests, based on only a few cases, one of which demonstrated monosomy 13 [67]. Another early report was based on aneuploidy concordance testing between results of spent culture media and results of embryo biopsy from couples having a balanced translocation, azoospermia, or recurrent pregnancy loss [68]. Reproductive outcome was available in six

clinical pregnancies that yielded five healthy live births. Feasibility was suggested using the consistencey of  free embryonic DNA in spent media  results, with results of trophectoderm biopsy. However, another early study that involved testing nuclear and mitochondrial DNA in spent embryo culture medium concluded that DNA from culture medium cannot yet be used for PGT because contamination arises from DNA from other origins, such as cumulus cells [69]. The concordance in spent medium and embryo biopsy was reported for IVSII654 thalassemia mutation [70]. Overall, these early data demonstrated the complexity of the methodology of applying cell-free DNA to perform PGT without biopsy. Following initial results and pilot studies, larger systematic concordance studies were initiated in several active PGT centers. In one of the studies, performed in the format of clinical trial, the test was offered to infertility patients presenting for PGT-A. A provisional study of 91 samples showed enough DNA to test in 88% of cases; concordance with biopsy results was in 80% of cases [71]. Study design requires liquid and biopsy tests to be compared to determine concordance with biopsy sample results. In another prospective study, 92 blastocoel samples and 72 spent culture medium samples were compared to the diagnostic biopsy samples, which were processed for PGT-M and PGT-­ A. Overall results demonstrated that neither blastocoel samples nor spent media were sufficiently robust approaches for aneuploidy or single-gene disorders. NIPGT cannot in particular be applied clinically until the risk of maternal contamination can be excluded. This is particularly high in spent culture medium samples due to maternal cumulus DNA. Further work is needed to understand the origin of DNA in the blastocoel fluid and spent culture medium [72]. In a further recent study, spent media from donated embryos were collected on days 3 and 5, one-half of the embryos undergoing assisted hatching and the other without [73]. Spent media were collected on day 5 before blastocyst biopsy for PGT-A by next-generation

References

sequencing; PGT-A results were compared with those from spent medium. Of 141 samples, amplification efficiency was 39% and 80.4% of days 3 and 5 in spent culture medium samples, with concordances of 56.3% and 81.3% between day 3 spent medium and whole embryos. Concordances were 65% and 70% between day 5 spent medium and biopsy samples. Day 5 medium sensitivity and specificity for aneuploidy were 0.8 and 0.61, respectively. Positive predictive value was 0.47 and negative predictive value 0.88, whereas neither was influenced by timing of assisted hatching, nor the morphology influenced DNA concentration or accuracy. Thus, DNA in spent medium is detectable on day 3 but more reproducibly on day 5. Overall, concordance rates are not high enough for practical application. The origin of the DNA in spent culture medium is not clear. It has been found that results of the test are more likely to have a prognostic utility and better prediction of reproductive outcome, if spent  culture medium results of the transferred euploid embryo also showed euploidy. This is in contrast to the euploid embryo transfer outcome with spent media showing imbalance results [74]. This is also in agreement with the blastocoel data, which suggested a significantly improved embryo transfer outcome for those euploid embryos whose blastocoel fluid has a higher quantity of DNA based on WGA [75]. Better understanding of the origin of DNA both in spent media and blastocoel fluid is underway, using novel technologies to identify the exact sources of tested DNA [76, 77]. Overall, NIPGT is not yet ready for clinical application due to still quite low amplification efficiency, insufficient concordance to the biopsy samples, and high rate of contamination.

2.6

PGT Without IVF

Progress has occurred in the development of PGT without IVF, which is based on the recovery of embryos from the uteri using uterine lavage [78]. In an ongoing study of 134 lavage cycles, 136 embryos were recovered in 56 cycles; 96 embryos

27

were at the blastocyst stage. Of these in vivo conceived embryos, 63% were euploid or low-­ mosaic, similar to that of IVF-conceived embryos (64%). High mosaicism rate (11% vs 9%), complex abnormality rates (8% vs 10%), and ­aneuploidy rates (19% vs 17%) were also similar in vivo to in vitro. This procedure has the potential to provide a low-cost, minimally invasive, reproducible, and effective way to acquire in vivo conceived embryos for PGT-A.

References 1. Preimplantation Genetic Diagnosis International Society (PGDIS). Guidelines for good practice in PGD: program requirements and laboratory quality assurance. Reprod Biomed Online. 2008;16:134–47. 2. Preimplantation Genetic Diagnosis International Society (PGDIS). 10th international congress on preimplantation genetic diagnosis. Reprod Biomed Online. 2010;20:S1–S42. 3. Kuliev AM.  Expanding indications for preimplantation genetic diagnosis. Expert Rev Obstet Gynecol. 2011;6:599–607. 4. Handyside AH, Kontogiani EH, Hardy K, Winston RML. Pregnancies from biopsied human preimplantation embryos sexed by Y-specific DNA amplification. Nature. 1990;344:768–70. 5. Verlinsky Y, Ginsberg N, Lifchez A, Valle J, Moise J, Strom CM. Analysis of the first polar body: preconception genetic diagnosis. Hum Reprod. 1990;5:826–9. 6. International Working Group on Preimplantation Genetics. Current progress in preimplantation genetic diagnosis. J Assist Reprod Genet. 1993;10:353–60. 7. International Working Group on Preimplantation Genetics. Preimplantation diagnosis of genetic and chromosomal disorders. J Assist Reprod Genet. 1994;11:236–43. 8. Griffin DK, Handyside AH, Penketh RJA, Winston RML, Delhanty JDA.  Fluorescent in-situ hybridization to interphase nuclei of human preimplantation embryos with X and Y chromosome specific probes. Hum Reprod. 1991;6:101–5. 9. Griffin DK, Wilton LJ, Handyside AH, Winston RML, Delhanty JDA. Dual fluorescent in-situ hybridization for the simultaneous detection of X and Y chromosome specific probes for the sexing of human preimplantation embryonic nuclei. Hum Genet. 1992;89:18–22. 10. Munne S, Lee A, Rozenwaks Z, Grifo J, Cohen J.  Diagnosis of major chromosome aneuploidies in human preimplantation embryos. Hum Reprod. 1993;8:2185–91. 11. Munne S, Weier HUG, Stein J, Grifo J, Cohen J.  A fast and efficient method for simultaneous X and Y

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2  Major Components of Preimplantation Genetic Testing

in-situ hybridization of human blastomeres. J Assisted Reprod Genet. 1993;10:82–90. 12. Delhanty JDA, Griffin DK, Handyside AH. Detection of aneuploidy and chromosomal mosaicism in human embryos during preimplantation sex determination by fluorescent in situ hybridisation (FISH). Hum Molec Genet. 1993;2:1183–5. 13. Munné S, Weier HUG, Grifo J, Cohen J. Chromosome mosaicism in human embryos. Biol Reprod. 1994;51:373–9. 14. Verlinsky Y.  Preimplantation genetic diagnosis. J Assist Reprod Genet. 1996;13:87–9. 15. International Working Group on Preimplantation Genetics. Current status of preimplantation diagnosis. J Assist Reprod Genet. 1997;14:72–5. 16. Munne S, Morrison L, Fung J, et  al. Spontaneous abortions are reduced after preconception diagnosis of translocations. J Assit Reprod Genet. 1998;15:290–6. 17. Verlinsky Y, Kuliev A.  Preimplantation genetics. J Assist Reprod Genet. 1998;15:215–8. 18. Munne S, Wells D, Cohen J.  Technology requirements for preimplantation genetic diagnosis to improve assisted reproduction outcomes. Fertil Steril. 2010;92:408–30. 19. Treff NT, Levy B, Su J, Northrop LE, Tao X, Scott RT.  SNP microarray-based 24 chromosome aneuploidy screening is significantly more consistent that FISH. Mol Hum Reprod. 2010;16:583–9. 20. Johnson DS, Gemelos G, Ryan A, et  al. Preclinical validation of a microarray method for full molecular karyotyping of blastomeres in a 24-h protocol. Hum Reprod. 2010;25:1066–75. 21. Treff NT, Sue G, Tao X, Levy B, Scott RT. Accurate single cell 24 chromosome aneuploidy screening using whole genome amplification and single nucleotide polymorphism microarrays. Fertil Steril. 2010;94:2017–21. 22. Gutierrez-Mateo C, Colls P, Sanchez-Garcia J, Escudero T, Prates R, Wells D, Munne S. Validation of microarray comparative genomic hybridization for comprehensive chromosome analysis of embryos. Fertil Steril. 2011;95:953–8. 23. Fragouli E, Alfarawati S, Daphnis DD, et  al. Cytogenetic analysis of human blastocyst with the use of FISH, CGH, and aCGH: scientific data and technical evaluation. Hum Reprod. 2011;26:480–90. 24. Kuliev A, Zlatopolsky Z, Kirillova I, Spivakova J, Cieslak-Janzen G.  Meiosis errors in over 20,000 oocytes studied in the practice of preimplantation aneuploidy testing. Reprod Biomed Online. 2011;22:2–8. 25. Gabriel AS, Thornhill AR, Ottolini CS, et  al. Array comparative genomic hybridization on first polar bodies suggests that non-disjunction is not the predominant mechanism leading to aneuploidy in humans. J Med Genet. 2011;48(7):433–7. 26. Geraedts J, Collins J, Gianaroli L, et  al. What next for preimplantation genetic screening? A polar body approach. Hum Reprod. 2010;25:575–7.

27. European Society of Human Reproduction and Embryology (ESHRE). Abstracts of the 26th annual meeting of European Society of Human Reproduction and Embryology. Hum Reprod. 2010;25(Supplement 1) 28. Schoolcraft WB, Fragouli E, Stevens J, Munne S, Katz-Jaffe MG, Wells D. Clinical application of comprehensive chromosomal screening in the blastocyst stage. Fertil Steril. 2010;94:1700–6. 29. Scott RT, Tao X, Ferry KM, Treff NR. A prospective randomized controlled trial demonstrating significantly increased clinical pregnancy rates following 24 chromosome aneuploidy screening: biopsy on day 5 with fresh transfer. Fertil Steril. 2010;94(Supplement S2):O-05. 30. Munné S, Sandalinas M, Escudero T.  Outcome of premplantation genetic diagnosis of translocations. Fertil Steril. 2000;73:1209–18. 31. ESHRE Preimplantation Genetic Diagnosis (PGD) Consortium. Best practice guidelines for preimplantation genetic diagnosis/screening (PGD/PGS). Hum Reprod. 2011;26:14–46. 32. Verlinsky Y, Tur-Kaspa I, Cieslak J, Bernal A, Morris R, Taranissi M, Kaplan B, Kuliev A. Preimplantation testing for chromosomal disorders improves reproductive outcome of poor prognosis patients. Reprod Biomed Online. 2005;11:219–25. 33. Kuliev A, Jansen JC, Zlatopolski Z, et al. Conversion and non-conversion approach to preimplantation diagnosis for chromosomal rearrangements in 475 cycles. Reprod Biomed Online. 2010;21:93–9. 34. The Practice Committee of the Society for Assisted Reproductive Technology and Practice Committee of the American Society for Reproductive Medecine. Preimplantation genetic testing: a practice committee opinion. Feril Steril. 2007;88:1497–504. 35. Munne S.  Preimplantation genetic diagnosis of numerical and structural chromosome abnormalities. Reprod Biomed Online. 2002;4:183–96. 36. Verlinsky Y, Cieslak J, Evsikov S, Galat V, Kuliev A.  Nuclear transfer for full karyotyping and preimplantation diagnosis of translocations. Reprod Biomed Online. 2002;5:302–7. 37. Kuliev A, Verlinsky Y. Thirteen years’ experience of preimplantation diagnosis: report of the fifth international symposium on preimplantation genetics. Reprod Biomed Online. 2004;8:229–35. 38. International Working Group on Preimplantation Genetics. Preimplantation diagnosis: an integral part of assisted reproduction. Report of the 9th annual meeting international working group on preimlantation genetics, in association with the 11th IVF congress, Sydney, May 10, 1999. J Assist Reprod Genet. 2000;16:161–4. 39. International Working Group on Preimplantation Genetics. 10th anniversary of preimplantation genetic diagnosis. Report of the 10th annual meeting of international working group on preimlantation genetics, in conjunction with 3rd international symposium on

References preimplantation genetics, Bologna June 23, 2000. J Assist Reprod Genet. 2001;18:66–72. 40. International Working Group on Preimplantation Genetics. Preimplantation genetic diagnosis – experience of three thousand clinical cycles. Report of the 11th annual meeting international working group on Preimlantation genetics, in conjunction with 10th international congress of human genetics, Vienna, May 15, 2001. Reprod Biomed Online. 2001;3:49–53. 41. Verlinsky Y, Munne S, Cohen J, et al. Over a decade of preimplantation genetic diagnosis experience  – a multi-center report. Fertil Steril. 2004;82:292–4. 42. Verlinsky Y, Rechitsky S, Verlinsky O, et  al. Preimplantation diagnosis for p53 tumor suppressor gene mutations. Reprod Biomed Online. 2001;2:102–5. 43. Verlinsky Y, Rechitsky S, Schoolcraft W, Strom C, Kuliev A.  Preimplantation diagnosis for Fanconi anemia combined with HLA matching. JAMA. 2001;285:3130–3. 44. Verlinsky Y, Rechitsky S, Sharapova T, Morris R, Tharanissi M, Kuliev A. Preimplantation HLA typing. JAMA. 2004;291:2079–208. 45. Kuliev A, Verlinsky Y. Current feature of preimplantation genetic disgnosis. Reprod Biomed Online. 2002;5:296–301. 46. ESHRE Preimplantation Genetic Diagnosis (PGD) Consortium. Data Collecttion II (May 2002). Hum Reprod. 2002;15:2673–83. 47. Liebaers I, Desmyttere S, Verpoest W, De Rycke M, Staessen C, Sermon K, Devroey P, Haentjens P, Bonduelle M. Report on a consecutive series of 581 children born after blastomere biopsy for preimplantation genetic diagnosis. Hum Reprod. 2010;25:275–82. 48. Kuliev A, Rechitsky S, Verlinsky O. 2014 atlas of preimplantation genetic diagnosis. 3rd ed. London: CRC Press, Taylor & Francis; 2014. 49. Verlinsky Y, Milayeva S, Evsikov S, et  al. Preconception and preimplantation diagnosis for cystic fibrosis. Prenat Diagnosis. 1992;12:103–10. 50. Kaplan B, Wolf G, Kovalinskaya L, Verlinsky Y, et al. Viability of embryos following second polar body removal in a mouse model. J Assist Reprod Genet. 1995;12:747–9. 51. Kuliev A, Rechitsky S, Laziuk K, Verlinsky O, Tur-Kaspa I, Verlinsky Y.  Pre-embryonic diagnosis for Sandhoff disease. Reprod Biomed Online. 2006;12:328–33. 52. Kuliev A, Rechitsky S, Verlinsky O, Strom S, Verlinsky Y.  Preembryonic diagnosis for sickle cell disease. Mol Cell Endocrinol. 2001;183:S19–22. 53. Larsen WJ.  Human embryology. Edinburgh: Churchill Livingstone; 1994. 54. Willadsen S, Munne S, Schmmel T, Cohen J.  Applications of nuclear sperm duplication. Fifth international symposium on preimplantation genetics, 5–7 June, Antalya, Turkey; 2003. p. 35. 55. Munne S, Willadsen S, Schmmel T, Cohen J. Nuclear sperm duplication as a tool to study mosaicism. Fifth

29 international symposium on preimplantation genetics, 5–7 June, Antalya, Turkey; 2003a. p. 55–6. 56. Lacham-Kaplan O, Daniels R, Trounson A.  Fertilization of mouse oocytes using somatic cells as male germ cells. Reprod Biomed Online. 2001;3:205–11. 57. Tesarik J, Nagy ZP, Sousa M, et al. Fertilized oocytes reconstructed from patient’s somatic cell nuclei and donor ooplast. Reprod Biomed Online. 2001;2:160–4. 58. Galat V, Verlinsky Y.  Haploidization of somatic cell nuclei by cytoplasm of human oocytes. Reprod Biomed Online. 2002;4(Supplement 2):48–9. 59. Galat V, Ozen S, Rechitsky L, Verlinsky Y.  Is haploidization by human mature oocytes real? Fifth international symposium on preimplantation genetics, 5–7 June, Antalya, Turkey; 2003. p. 36–7. 60. McArthur S, Marshall J, Wright D, de Boer K. Successful Pregnancies following blastocyst (day 5) biopsy and analysis for reciprocal and Robertsonial translocations. Fifth international symposium on preimplantation genetics, 5–7 June, Antalya, Turkey; 2003. p. 33. 61. Scott RT, Upham KM, Forman EJ, Zhao T, Treff NR.  Cleavage –stage biopsy significantly impairs human embryonic implantation implantation potential while blastocyst biopsy does not: a randomized and pared clinical trial. Fertil Steril. 2013;100:624–30. 62. Gardner DK, Lane M. Culture of viable human blastocyscts in defined sequential serum-free media. Hum Reprod. 1998;13(Suppl. 3):148–59. 63. Bigger JD, Racowsky C.  The development of fertilized human ova to the blastocyst stage in KSOM (AA) medium: is a two-step protocol necessary? Reprod Biomed Online. 2002;5:133–40. 64. Rieger D. The use of global for 1-step (uninterrupted) culture from zygote to blastocyst. Fertility Magazine. 2012;14:5–11. 65. Kader A, Choi A, Orief Y, Agrawal A. Factors affecting the outcome of human blastocyst vitrification. Reprod Biol Endocrinol. 2009;7:99. 66. Gianaroli L, Magli MC, Pomante A, et  al. Blastocentesis: a source of DNA for preimplantation genetic testing. Results from a pilot study. Fertil Steril. 2014;102:1692–1699.e6. 67. Shamonki MI, Jin H, Haimowitz Z, Liu L.  Proof of concept: preimplantation genetic screening without embryo biopsy through analysis of cell-free DNA in spent embryo culture media. Fertil Steril. 2016;106:1312–8. 68. Xu J, Fang R, Chen L, et al. Noninvasive chromosome screening of human embryos by genome sequencing of embryo culture medium for in  vitro fertilization. PNAS. 2016;113:11907–12. 69. Hammond ER, McGillivray BC, Wicker SM, et  al. Characterizing nuclear and mitochondrial DA in spent embryo culture media: genetic contamination identified. Feril Steril. 2017;107:220–8. 70. Liu W, Liu J, Zi H, Du HZ, Ling J, Sun X, Chen D. Non-invasive pre-implantation aneuploidy screen-

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ing and diagnosis of beta thalassemia IVSII654 mutation using spent embryo culture medium. Ann Med. 2017;49(4):319–28. 71. Katz-Jaffe M, McCallie B, McReynolds S, Henry L, McCormick S, Schoolcraft WB.  Analysis of the embryonic media drop as a non-invasive alternative to blastocyst biopsy in preimplantation genetic testing for aneuploidy. ASRM 2018, Abstract #P786. 72. Capablo A, Romanelli V, Patassini C, et al. Diagnostic accuracy of blastocoelic fluid and spent media as source of DNA for preimplantation genetic testing in standard clinical conditions. Fertil Steril. 2018;110:870–9. 73. Ho JR, Arrach N, Rhodes K, et al. Pushing the limits of detection: investigation of cell-free DNA for aneuploidy screening in embryos. Fertil Steril. 2018;110:467–47. 74. Rubio C.  Non-invasive PGT-A with the analysis of embryonic cell free DNA in spent blastocyst media. PGDIS 2019. Reprod Biomed Online. 2019;39:E12.

75. Magli C, Andor C, Crippa A, Tabanelli C, Ferraretti AP, Gianaroli L. Deoxyribonucleic acid detection in blastocoelic fluid: a new predictor of embryo ploidy and viable pregnancy. Fertil Steril. 2019;111:77–85. 76. Vera-Rodriguez M, Diez-Juan A, Jimenez-Almazan J, et  al. Origin and composition of cell-free DNA in spent medium from human embryo culture during preimplantation development. Hum Reprod. 2018;1(33):745–56. 77. Liang B, Gao Y, Xu J, et  al. Raman profiling of embryo culture medium to identify aneuploid and euploid embryos. Fertil Steril. 2019;111:753–62. 78. Munné S, Najmabadi S, Rivas J, Nakajima ST, Buster JE. Advances in in-vivo fertilized and matured human embryos and characterization by preimplantation genetic testing (PGT-A). PGDIS 2019. Reprod Biomed Online. 2019;39:e12–3.

3

Major Components of Preimplantation Genetic Testing: Adjustment of Available Genetic Technology to PGT Practice

PGT became realistic only after the discovery of PCR, which made a single-cell genetic analysis a reality and served as the basis for performing PGT in assisted reproduction and genetic practices. This provided an important option for couples at the genetic risk to avoid the birth of an affected offspring and have a healthy child of their own. Still, accuracy remained to be verified.

3.1

 djustment of DNA Analysis A to Avoid Misdiagnosis in Single-Cell PCR

Because PGT for monogenic disorders (PGT-M) is based on a single or few cells, accuracy depends heavily on addressing limitations of single-cell DNA analysis to avoid the causes of misdiagnosis (Table  3.1). One of the key contributors to misdiagnosis is the phenomenon of preferential amplification, also known as allele-specific amplification failure (allele dropout  – ADO). Avoiding ADO requires the application of special protocols to ensure detecting ADO [1–3]. In the 1990s, misdiagnoses at the initial stage of PGT-M application were reported for myotonic dystrophy (DM), fragile X syndrome (XMR1), and cystic fibrosis (CF). These might have been caused by ADO or preference amplification, but were not fully recognized [4–8]. It has since been demonstrated that ADO rates in single cells differ for different types of hetero© Springer Nature Switzerland AG 2020 A. Kuliev et al., Practical Preimplantation Genetic Testing, https://doi.org/10.1007/978-3-030-43157-0_3

zygous cells [9]. Rates exceed 20% in blastomeres, compared to the ADO rates in single fibroblasts and PB1 of under 10% (Fig.  3.1). A high rate of ADO in blastomeres may potentially lead to a misdiagnosis, especially in compound heterozygous embryos. This was likely the case in some of the abovementioned misdiagnoses observed in PGT when performed by blastomere biopsy. Accuracy of PGT-M depends on approaches that control, detect, and prevent ADO. ADO rates vary by different lysis procedures, cell types, and loci analyzed [1–2, 10]. ADO must be excluded, being the key to avoiding misdiagnosis in PGT-­ M. Our own experience shows that ADO can be detected reliably by sequential analysis of oocytes, using PB1 and PB2, or embryos. The strategy involves simultaneous amplification of mutant genes with linked polymorphic markers, described in detail elsewhere and presented in brief below [1, 3, 10]. Biopsied single cells are placed directly into a lysis solution, consisting of 0.5 mcl 10 × PCR buffer, 0.5  mcl 1% Tween 20, 0.5  mcl 1% Triton  ×  100, 3.5  mcl H2O, and 0.05  mcl Proteinase K (20 mg/ml in a 0.5 ml PCR tube). After spinning down at a low speed in a microfuge for a few seconds, samples are covered with 1 drop of mineral oil and incubated at 45 °C for 15 min in a thermal cycler. Proteinase K is then inactivated at 96 °C for 20 min, which is also the beginning of the hot start of the first31

32

3  Major Components of Preimplantation Genetic Testing: Adjustment of Available Genetic Technology…

round PCR.  Lower stringency and longer annealing time are used in the first-round PCR, with the introduction of the mixture of all outside primers for both mutant genes and polymorphic markers. Following the first-round PCR, separate aliquots are amplified in the second-round PCR with specific inside primers for each site, using a higher stringency. Such a dual or multiple amplification reaction allows recognition of most ADO cases. If pseudogenes are involved, false priming is eliminated by use of first-round primers designed to anneal to the regions of nonidentity with pseudogenes [1]. In addition to short tandem repeats (STR) or single Table 3.1  Expected problems in PGT-M Preferential amplification Allele dropout – ADO Contamination Recombination Pseudogene presence Aneuploidy Uniparental disomy

27.70%

30.0% 25.0%

nucleotide polymorphisms (SNPs) linked to the genes studied, STRs or SNPs located on other chromosomes may be also tested to exclude possible contamination by extraneous DNA. Identification of the origin of individual embryos in established pregnancies can be determined. A list of linked markers, their identified sequences, and requisite PCR conditions are presented in each respective section. Fluorescent PCR (F-PCR) is also an option for direct fragment size analysis of PCR products [1] and may be useful also for a direct sequencing of the PCR product in the detection of point mutations and for distinguishing preferential amplification from ADO.  Considerable proportions of ADO can be detected by sequential analysis of PB1 and PB2, as initially demonstrated in sequential PCR analysis of 26 alleles in PB1 and PB2 from 1047 oocytes [1]. A total of 32 of 53 ADOs in mutation analysis were detected by sequential analysis of PB1 and PB2, even when no informative polymorphic markers were available.

21.70%

27%

19.90% Corona cells Polar Body 1

20.0%

Blastomere Blastomere WGA

15.0% 7.40% 10.0%

5.50%

7.00% 3.60% 6.20%

6.50% 6.80% 8.00% 6.00% 6%

Trophectoderm Trophectoderm WGA Trophectoderm MDA

5.0% 0.0% CFTR

Fig. 3.1  ADO rate in different types of cells with cystic fibrosis (CFTR) (on the left) and thalassemia (HBB), increasing following WGA allele dropout (ADO) rates in different types of cells heterozygous for cystic fibrosis (CFTR gene) and beta-globin gene mutations with or

HBB

without whole-genome amplification (WGA) (see description in the text). In both mutations a significantly higher ADO rate is seen following WGA, which is much lower in blastocyst samples, than in blastomere samples

3.1 Adjustment of DNA Analysis to Avoid Misdiagnosis in Single-Cell PCR

The most universal approach for detection of potential ADO in PGT-M is the testing for presence or absence of linked polymorphic markers. This can be performed on any type of biopsied single or small number of cells. Analysis of linked polymorphic markers reduces the undetected ADO rate considerably, irrespective of use of conventional or fluorescent PCR.  With each additional linked marker in a multiplex PCR, the rate of undetected ADO may be expected to be reduced by half, overall reducing misdiagnosis to practically zero (Fig.  3.2). Contrary to expecta-

33

tion, F-PCR does not sufficiently improve detection of potential misdiagnoses in PGT-M, as demonstrated by testing of 148 single fibroblasts by both conventional and fluorescent PCR. The multiplex-nested PCR analysis is performed with the initial PCR containing all the pairs of outside primers; thus, following the first-round PCR, separate aliquots of the resulting PCR product may be amplified using the inside primers specific for each site. Only when the polymorphic sites and the mutation agree are embryos transferred, and in this way multiplex amplification

7.1 9

7.9

8.1

8 6.8 7

6 5 3.5 4 3 1.5 2

0

1 0

e

on

Al

M

+1

8 50

F

Delta F508

M

+2

8 50

F

Intron 6

Fig. 3.2  Use of polymorphic markers in multiplex-nested PCR analysis to detect and avoid ADO in PGT-M. Testing for Delta F508 mutation alone or each linked marker (Intron 6, Intron 8, or Intron 17) separately results in ADO rates of 6.8–8.1% while with addition of each linked

M

+3

8 50

F

Intron 8

Intron 17

marker in multiplex PCR reduces the risk for ADO almost by half with each additional marker (by 3.5% with one, by 1.5% with two, and by almost zero rate with addition of three markers)

3  Major Components of Preimplantation Genetic Testing: Adjustment of Available Genetic Technology…

34

allows detecting and excluding the impact of preferential amplification and ADO on misdiagnosis, that is, the major part of excluding from transfer the majority of misdiagnosed affected embryos. (Example of detection of ADO of dominant mutation by linked markers is demonstrated in the case of PGT for breast cancer, presented in Fig. 3.3.) The usefulness of linkage analysis in polar body analysis was demonstrated in a series of 114 PGT cycles for couples at high risk of having children with single-gene disorders. Preselection and transfer of mutation-free oocytes were possible in almost all cycles. Of 1047 oocytes with

DNA results, 672 (64.1%) had heterozygous PB1, i.e., with both normal and mutant genes amplified. This was ideal for further testing with selection of an embryo to transfer after identification of mutant gene in the sequential analysis of PB2. Thus, priority in preselection of embryos for transfer was given to embryos resulting from the oocytes with heterozygous PB1 because in the absence of DNA contamination, this could have indicated absence of ADO of either normal or mutant allele. Although most of the transferred embryos were preselected using this particular strategy, some preselected embryos originated from homozygous normal oocytes, inferred from

a N / 5385insC

PGT

b

PGT-M for BRCA1 mutation

Embryo #

D17S1801

D17S932

BRCA1 gene

8(CA)

3’end

D17S934

01

FA

99 / 118

N / 5385insC

84 / 104

177 / 173

159 / 167

02

ADO/ 129

110 / 120

N

96 / 106

173 / 175

161 / 152

NORMAL

03

137 / 127

110 / 118

N / ADO

96 /104

173/173

161 / ADO

AFFECTED

04

125 / 129

99 / 120

N

84 / 106

177 / 175

159 / 152

NORMAL

05

125 / 129

99 / 120

N

84 / 106

177 / 175

159 / 152

NORMAL

06

137 / 127

110 / 118

N / 5385insC

96 / 104

173/173

ADO / 167

AFFECTED

08

ADO/ 127

110 / 118

N / 5385insC

96 / 104

173/173

161 / 167

AFFECTED

09

137 / 127

110 / 118

N / 5385insC

96 / 104

173/173

161 / 167

AFFECTED

10

137 / 127

110 / 118

N / 5385insC

96 / 104

173/173

161 / 167

AFFECTED

PARTNER

125 / 137

99 / 110

N/N

84 / 96

173 / 177

159 / 161

NORMAL

PATIENT

129 / 127

120 / 118

N / 5385insC

106 / 104

175 / 173

152 / 167

AT RISK

Fig. 3.3  ADO of dominant mutant allele detected by linked markers. (a) Pedigree indicated affected family members and patient at risk of developing breast or ovarian cancer. (b) PGT-M was performed for breast cancer predisposition caused by 5385 insertion “C” in BRCA1 gene. Table contains the outcome of the cycle. Left vertical column contains embryo numbers followed by informative markers surrounding BRCA1 gene and genotype predictions for each embryo. Out of nine tested embryos, three were predicted to be normal and recommended for

Predicted Genotype

AFFECTED

transfer. Six other embryos were predicted to have a mutation in BRCA1 gene. ADO was detected in embryo #2 (D17S1801 marker), embryo #3 (mutation site and D17S934), embryo #6 (D17S934), and embryo #8 (D17S1801). Without linkage analysis embryo #3 with ADO at a mutation site would be predicted normal, and transfer of this embryo would lead to a misdiagnosis. One normal embryo #04 was transferred, resulting in ­pregnancy and delivery of a healthy baby free from from predisposition to breast cancer

3.2 Other Strategies to Avoid Misdiagnosis

homozygous mutant status of PB1 and hemizygous normal status of PB2. Resulting embryos from these oocytes were accepted for transfer only if ADO could be excluded using linked polymorphic marker analysis. Otherwise, such embryos were excluded from transfer or further investigated following embryo biopsy. To determine the proportion of undetected ADO in mutation analysis, whole genomic DNA analysis of donated non-transferred embryos was undertaken, including the affected ones or those with insufficient information available for their transfer. Overall, of 1052 embryos tested, 82 (7.8%) ADOs were observed, of which 75 were detected and only 7 were undetected by the strategy used, suggesting an extremely high accuracy of PGT-M. As mentioned, one of the most efficient approaches for avoiding misdiagnosis is the sequential genetic analysis of PB1 and PB2  in PGT for maternally derived mutations. Detection of both mutant and normal alleles in the heterozygous PB1, together with the mutant allele in the corresponding PB2, leaves no doubt that the resulting maternal contribution to the embryo is normal, even without testing for the linked markers as a control. Ideally, one tests simultaneously at least one linked marker to confirm the diagnosis. Alternatively, mutation-free oocytes may be predicted when the corresponding PB1 is homozygous mutant, in which scenario the corresponding PB2 should be hemizygous normal and similar to the resulting maternal pronuclear. However, the genotype of the resulting maternal contribution may be quite opposite, i.e., mutant, if the corresponding PB1 is in fact heterozygous but erroneously diagnosed as homozygous mutant because of ADO involving the normal allele. In the above scenario, the extrusion of the normal allele with PB2 would mean that the mutant allele remains in the resulting oocyte. Therefore, the embryo resulting from the oocyte with homozygous mutant PB1 cannot be acceptable for transfer unless the heterozygous status of PB1 is excluded by the use of linked markers as described above. This example of misdiagnosis due to ADO of the normal allele in PB1 has been described in a PGT cycle performed for FMR1 (see Chap. 4). To avoid misdiagnosis, sequential

35

PB1 and PB2 analysis may be required to combine with multiplex PCR to exclude an undetected ADO in a heterozygous PB1. As described above [1–3], analysis of more than 1000 oocytes tested by sequential PB1 and PB2 analysis showed that more than half of ADOs were detected by sequential analysis of PB1 and PB2; the remaining cases were detected by multiplex PCR.  Accuracy of this approach was demonstrated by reports of PGT-M [11–12] and description below of our overall experience in Chap. 4. Another method with proved potential for detecting and avoiding misdiagnosis due to preferential amplification is fluorescence PCR (F-PCR). This allows detection of heterozygous PB1 or blastomeres misdiagnosed as homozygous in conventional PCR.  The method also allows simultaneous gender determination [13], DNA fingerprinting, and detection of common aneuploidies, the latter initially at a time when microarray and NGS technologies were not available. Application of F-PCR combined with a multiplex system and sequential PB1 and PB2 analysis excluded almost completely the risk for misdiagnosis due to preferential amplification in maternally derived mutations. Accuracy of PGT can further be improved by application of fluorescent PCR with expand long template (ELT) kit, which reduced the undetected ADO rate from as high as 30–35% in both conventional and fluorescent PCR to as low as 5% in testing for myotonic dystrophy [14]. Another development in improving the accuracy of singlecell PCR analysis involves application of realtime PCR, which reduces the undetected ADO rate by almost half compared to conventional or fluorescent PCR. Application of these approaches together with simultaneous testing for the causative mutation and one or two linked markers reliably avoids the risk for misdiagnosis.

3.2

 ther Strategies to Avoid O Misdiagnosis

Mosaicism exists in preimplantation genetic analysis, and its recognition is of obvious relevance for PGT-M. Lack of a mutant allele could

36

3  Major Components of Preimplantation Genetic Testing: Adjustment of Available Genetic Technology…

be due to monosomy for the chromosome on which the gene was located. Before the availability of 24-chromosome microarray or NGS technologies, aneuploidy testing to exclude this possibility was performed by adding primers for chromosome-specific microsatellite markers to the multiplex PCR protocols worked out for specific genetic disorder [15]. Development of multiplex-­nested PCR systems also allowed PGT for different conditions simultaneously, for example, PGT for CFTR together with FMR1 or gender determination (X,Y) [16]. Simultaneous PGT was recently performed for as many as five different conditions, namely, PGT-M for four autosomal recessive diseases and PGT-A, presented in Chap. 4. This resulted in birth of a child free of all four recessive conditions and aneuploidy, despite a very low probability of PGT outcome for the first-degree consanguineous ­couple [17]. Development of a custom-made PGT strategy for each mutation and for each couple has become an integral part of PGT-M excluding a potential misdiagnosis. For example, a particular set of outside primers has to be designed to eliminate false priming to a pseudogene, as applied in PGT for long-chain 3-hydroxyacyl-Coa dehydrogenase deficiency [18]. Single-sperm typing is needed to determine paternal haplotypes in PGT for de novo mutations, enabling linked marker analysis in addition to mutation testing. This is especially essential in cases of paternally derived dominant conditions (see below). The use of haplotyping for PGT without direct mutation testing is also applicable, referred to as preimplantation genetic haplotyping (PGH) [19], to improve the accuracy of PGT.  Availability of the parental haplotypes, irrespective if mother or father is a carrier, not only confirms the absence of the mutant gene but also the presence of maternal and paternal wild-­ type alleles. This is especially helpful when only one informative marker is available (see Chap. 4). Of special importance is detection of recombination, which will potentially contribute to accuracy of PGT-M, as demonstrated in Fig. 3.4. As will be described in Chap. 5, the failure to identify recombination in the HLA cluster may lead to failure in identification of HLA-matched

embryo in PGT-HLA and contribute to the effectiveness of stem cell transplantation therapy. One of the major developments used in PGT-M is the whole-genome amplification (WGA) using Super Plex Single Cell Whole Genome Amplification Kit. As a first step, this is a key component of the microarray and NGS technologies. The WGA product is currently used for combining PGT-M with aneuploidy testing for 24 chromosomes. The first series of conditions for which PGT-M was performed together with 24-chromosome aneuploidy involved testing of a total of 1054 embryos, including 320 by blastocyst biopsy in 78  cycles, 670 by blastomere biopsy in 53  cycles, and 64 by combined polar body and blastomere biopsy in 4 cycles [20]. Five of these cycles were performed with concomitant HLA typing, including two for chronic granulomatous disease, two for B-thalassemia, and one for Diamond–Blackfan anemia; in each, testing involved a set of chromosome 6 short tandem repeats throughout HLA region. Of 1054 embryos tested, 434 (41.2%) were predicted to be aneuploidy-­ free and also unaffected. Overall, embryos suitable for transfer were available in all cycles, transferred without freezing in 41 of 92 cycles performed by polar body and blastomere biopsy resulting in 21 (51.2%) clinical pregnancies and 24 unaffected children. Embryo freezing was performed in the remaining 51 transfer cycles performed by blastocyst biopsy, resulting in 40 (78.4%) clinical pregnancies and 45 unaffected births. Overall, concomitant 24-chromosome aneuploidy testing, together with analysis for single-gene disorders and HLA typing, resulted in transfer of 137 embryos in 92 cycles, yielding 61 (66.3%) clinical pregnancies and 69 unaffected children. Thus, combined comprehensive testing for 24 chromosomes with PGT for single-gene disorders and preimplantation HLA typing has become a practical application in the framework of ART to optimize the pregnancy success. On the other hand, WGA increases significantly the risk of ADO rate in all types of biopsied cells [20]. Even with the trend to the blastocyst biopsy that permits testing as many as 5–10 cells, ADO rate doubles after WGA [20], thus increasing the risk for misdiagnosis

3.2 Other Strategies to Avoid Misdiagnosis a

37 Chromosome crossover

Homologous chromosomes aligned

Recombinant chromatids

Non-recombinant chromatids

b

Normal

Mutant / Normal

Recombination as a potential source of misdiagnosis

Recombination not leading to misdiagnosis

K

A

K

a

L

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L

b

m

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m

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Normal

Mutant

O

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E

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q

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Fig. 3.4  Recombination as a source of misdiagnosis. (a) Schematic representation of meiotic recombination. (b) Recombination position and correct genotype prediction. Recombination is shown in the middle panel, resulting in

recombinant chromatids, shown in different colors in the long arms of each chromatid of homologous chromosomes, which is a potential source of misdiagnosis, as shown in the panel below on the right

(Fig. 3.1). However, no misdiagnosis has actually been observed in PGT-M by applying the approaches described above. Still, a challenge exists to devise more efficient WGA procedures that may help to completely avoid ADO-­ associated misdiagnosis. Development of a family-specific strategy appeared of particular importance in PGT for de novo mutations (DNM), detected in parent(s) or

affected children. Neither origin nor relevant haplotypes are available for tracing the inheritance of DNM in a single or in a number of cells biopsied from embryos or oocytes. The number of such cases is not rare among those requesting PGT-M, when there is no family history of the genetic disease, which is first being diagnosed in one of the parents or in one of their affected children. The first systematic PGT for de novo muta-

38

3  Major Components of Preimplantation Genetic Testing: Adjustment of Available Genetic Technology…

tions involving maternal or paternal origin demonstrated that PGT-M strategies for these families depend on the parental origin of DNM and must include an extensive DNA analysis of the parents and affected children prior to PGT-M [21]. Mutation verification, polymorphic marker evaluation, whole- and single-sperm testing, or PB analysis may be necessary to establish normal and mutant haplotypes, without which PGT for DNM cannot be performed. Despite the complexity of PGT for DNM, applied strategies appear to be highly accurate, making PGT-M feasible for any couple at risk including affected family members (see Chap. 4). Although PGT-M is mainly designed for specific conditions, a comprehensive approach has been also used to test simultaneously two or more conditions [22], with concomitant testing for chromosomal aneuploidy and translocations [20]. Preparatory work is still required to construct a special PGT-M design for each couple. As noted, PGT for paternally derived dominant conditions may involve single-sperm typing to establish paternal haplotypes based on linked marker analysis. As also mentioned above, a more universal approach by preimplantation genetic haplotyping (PGH) may possibly be required even without mutation testing. Another indirect approach applicable for PGT-M is karyomapping, which does not require testing for the causative gene and may be combined with PGT-A [23]. However, karyomapping is not universally applicable to all PGT-M cases because parental data may not be available, both parents may have identical recessive mutation in a highly consanguineous background, and the affected case may have arisen by a de novo mutation. Our data analyzing the outcome of PGT-M series of 2780 couples showed that karyomapping could not be applicable to as many as one-­ third of families referring for PGT-M [20].

years during which extensive use of PGT-A by FISH was performed, improved pregnancy rates in poor-prognosis IVF patients were shown [24– 26]. This application utilized commercially available chromosome-specific probes, still in use in many Eastern Mediterranean and Southeast Asian countries. FISH may also be applied as an additional test in circumstances in which no contemporary technology exists. The majority of FISH cycles were performed by blastomere biopsy; approximately one-fifth was done by PB1 and PB2 analysis. Overall, these resulted in thousands of unaffected pregnancies and healthy births. Follow-up confirmation studies utilizing abnormal embryos demonstrated acceptable accuracy [1, 24–27]. There were also limitations, in part due to mosaicism [28], which was particularly high in slowly growing embryos exhibiting an arrested development. As will be described below, mosaicism also persists to blastocyst stage. To recapitulate, recall that FISH has been extensively applied to PB analysis since 1994 [29]. In the initial work, 130 unfertilized MII oocytes were tested simultaneously with their PB1, using X chromosome- and chromosome 18-specific probes. It was demonstrated that PB1 data allows an exact prediction by deduction of the chromosome set in the corresponding oocytes [29–30]. Each chromosome in PB1 was represented by double sets (signals), corresponding to two chromatids in each univalent. The number of sets (chromatids) in PB1 reliably predicts the corresponding number of sets (chromatids) in the MII oocytes, therefore providing an excellent tool for the genetic preselection of oocytes. In addition to a normal distribution of signals in PB1 and the corresponding MII oocytes, meiotic errors were detected, confirming the accuracy of PB1 testing for predicting the genotype of the corresponding oocyte. For example, in one PB1 four copies for chromosome 18 were detected, 3.3 Is There Still Place for FISH perfectly in accordance with lack of chromosome 18  in the corresponding MII oocyte (chromoAnalysis in Current PGT-A? some 18 nondisjunction). The chromosomal FISH is no longer a standard technology for PGT-­ complements of the oocyte could thus be inferred A, but it was a method of choice for age-related from testing PB1, which can be removed followaneuploidies for many years. Over the dozen of ing its extrusion from the mature oocyte, with no

3.3 Is There Still Place for FISH Analysis in Current PGT-A?

potential influence on the embryo viability. Another interesting phenomenon was the observation of chromatid malsegregation as a possible cause of chromosomal aneuploidy in the resulting mature oocytes. In four oocytes, three instead of the expected two copies were found in the MII oocytes. This perfectly complemented a single set in the corresponding PB1. Similar results were reported by another group, confirming diagnostic significance of PB1 analysis for predicting the genotype of the preimplantation embryo [31]. PB1 testing was also one of the first approaches used for PGT of structural rearrangements (PGT-SR) [32], based on PB1, which never forms an interphase nucleus, but consists of metaphase chromosomes. PB1 chromosomes are recognizable when isolated 2–3  h after in  vitro culture, with degeneration beginning 6–7  h after extrusion. Therefore, whole chromosome painting or chromosome segment-specific probes were applied for testing maternally derived chromosomal translocations in PB1. However, malsegregation and/or recombination occurs between chromatids, requiring follow-up analysis of PB2  in order to predict accurately the meiotic outcome following the second meiotic division (see Chap. 6). In contrast to PB1, PB2 is the by-product of the second meiotic division, extruded following fertilization of oocyte. The need for PB2 analysis for PGT of chromosomal imbalances is obvious, given PB1 testing alone not allowing prediction of the resulting genotype of the oocyte [33]. In contrast to the paired copies in PB1, each chromosome in PB2 is represented by a single copy; thus, lack of or addition of a copy for a particular chromosome under interrogation provides evidence of a second meiotic division error. Although only 19 of 55 oocytes in the initial study were tested by both PB1 and PB2, evidence for errors was observed not only in meiosis I but also in meiosis II (PB2 testing). This suggested that some oocytes selected as normal based on PB1 analysis still could have been abnormal following nondisjunction in the second meiotic division. Therefore, analysis of both PB1 and PB2 has become the basic requirement for PGT-A, because this detects errors in both the first and

39

second meiotic divisions. Overall, more than 25,000 oocytes have been analyzed, showing accuracy and reliability of PB1 and PB2 testing for predicting the karyotype of the embryo resulting from the corresponding oocyte [34]. More than 50% oocytes from IVF patients of advanced reproductive age are abnormal, resulting from errors in both the first and second meiotic divisions. Prior to PB2 analysis, it had been believed that aneuploidies mainly originated from meiosis I. PB2 testing is now recognized as an important component of PGT for maternally derived translocations. Although this was initially done on interphase nuclei, progress in transforming PB2 into metaphase chromosomes had been achieved via electrofusion of a PB2 nucleus with a foreign one-cell human embryo; however, proportion of metaphase plates obtained was not sufficient to be useful in clinical practice [35]. Much higher efficiency was observed in conversion of interphase blastomere nuclei of blastomeres, as described below. Visualization of chromosomes of individual blastomere nuclei is possible but requires the application of nuclear transfer techniques. These were initially employed to convert a PB2 interphase nuclei into metaphase chromosomes [36]. The original experimental design was to fuse individual blastomeres with enucleated human oocytes. Metaphases were indeed obtained from two-thirds of blastomeres treated by this method, but efficiency was not high enough to be clinically applicable. This was due to inability of a replicating nucleus to form metaphase chromosomes after induction of premature chromosome condensation (PCC). Because biopsied blastomeres may be at any stage of the cell cycle at the time of biopsy, there was need to control the ­timing of mitosis of a blastomere nucleus. This can be in principle achieved by introduction of the biopsy blastomere into cytoplasm of a cell at a known cell cycle point. To achieve such reprogramming, individual blastomeres were fused with intact or enucleated mouse zygotes at pronuclear stage, i.e., the S-phase of the cell cycle. Overall success rate of the procedure was as high as 83%, efficiency improving with experience [1, 37]. Similar results were obtained using

40

3  Major Components of Preimplantation Genetic Testing: Adjustment of Available Genetic Technology…

bovine ooplasts for fusion with human blastomeres [38]. Failures occurred either because of absence of the nucleus in biopsied blastomeres or because heterokaryons were fixed after they have already cleaved. Blastomere biopsy was performed not earlier that day 3 or day 4, thus avoiding 2- and 4-cell embryos that could lead to accelerated heterokaryon cleavage. The success rate did not depend at all on whether mouse zygotes were enucleated before fusion with blastomeres; thus, simplifying the procedure should be possible using intact mouse zygotes. Thus, the procedure was quite simple and included the following components: mouse zygotes were thawed, freed of zonae pellucidae and PB2, approximately, 1–2 h before electrofusion with human blastomeres; 4  h after fusion heterokaryons were monitored for signs of the disappearance of pronuclei and fixed at mitosis following hypotonic treatment. To avoid monitoring and a possible failure to recognize mitosis, heterokaryons were also cultured in the presence of the microtubule inhibitors vinblastine or podophyllotoxin. All embryos left in the culture by the ninth hour after fusion were fixed following 1 h pre-treatment with OA.  The example of blastomere nucleus conversion applied for PGT of reciprocal translocation 46,XX, t(9;16) (q34.3;p13.1) is shown in Fig.  3.5. The method was applied for PGT of paternally derived reciprocal translocations and for confirmation of PGT of chromosomal abnormalities performed by PB1 and PB2 analysis. It was also reported that the blastomere metaphase can be also obtained without the conversion method [39]. To obtain analyzable chromosomes, embryos are monitored closely at the second day after ICSI in order to identify the blastomere with nuclear breakdown. This is then biopsied and fixed within 1 h. This method was later modified by 1 h culture of the biopsied blastomere in medium containing vinblastine, which resulted in harvest of metaphase chromosomes of good quality. To obtain more reproducible results, this method was further improved using chemical agents, such as caffeine, which facilitated an earlier nuclear envelope breakdown, premature

metaphase, PCC, and earlier onset of DNA synthesis that increases the rate of metaphase formation. For better results, it requires a morphological selection for biopsy of the largest blastomere, i.e., characterized by 1–2 large nucleoli within the cell nucleus. Upon embryo biopsy, each blastomere was placed in microdrops of Global culture medium (LifeGlobal, USA) supplemented with Plasmanate (Bayer Biological, USA) 10% vol:vol., containing caffeine (Sigma) (1 mmol/l) and a low dose of colcemid (Sigma) (2)

c 1

2

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Fig. 3.6  NGS-based PGT-A. Panel a shows examples of monosomies: monosomy 16 on the left and monosomy 21 on the right (circles in red). Panel b shows examples of trisomy: trisomy 16 on the left and trisomy 21 on the right

(circles in blue). Panel c demonstrates complex abnormalities, involving 3 or more chromosomal abnormalities in each embryo (circles in red and blue)

is, therefore, better able to identify presence of mosaic aneuploidy within the blastocyst. The most widely used NGS platform is MiSeq, developed by Illumina. The basis is a highly optimized amplification procedure and software. The WGA product produced by this amplification procedure is digested into millions of DNA fragments, which are used for library preparation as distinguished with a molecular barcode. This is followed by a specific PCR step, optic-based sequencing of the DNA fragments of the whole genome at a depth of 1X, and comparison with sequence data of a human hap map reference genome. MiSeq allows simultaneous analyses of 22 samples, with the final report of sequencing analysis available within 16  h (example of PGT-SR performed by NGS is presented in Fig. 3.7). Sequencing data output generally involves two rounds of quality assurance metrics, includ-

ing detailed analysis with BlueFuse software. VeriSeq genome analysis on the MiSeq detects whole-genome aneuploidy and mosaicism to a level of 50%. Different labs use different cutoff rates [55–59], but PGDIS recommendations currently recommend a 20% cutoff. That is, embryos are considered non-mosaic euploid if non-model DNA proportions are below 20%. Non-model DNA over 80% is considered non-mosaic aneuploid. Between 20 and 80% are considered mosaic [60]. Figure 3.8 presents examples of different mosaicism levels, shown as a gradient [61]. Although this methodology was originally designed to identify whole chromosome aneuploidy, MiSeq also detects structural variations. An option also exists to apply a low-density NGS technology, perhaps amplifying up to 10,000 genomic loci and providing sequencing data at each locus. This is too insensitive to detect mosaicism, but in certain circumstances is advanta-

44

3  Major Components of Preimplantation Genetic Testing: Adjustment of Available Genetic Technology…

Fig. 3.7  NGS-based PGT-SR. Embryo A in addition to derivative chromosome 9 has also shown mosaicism for monosomy 5 and mosaicism for trisomy 17 (pointed by

arrows). Embryo B has no other variations except for derivative chromosome 7, while embryo C in addition to derivative chromosome 7 has also trisomy 5 (circles in red)

3

20%

2

20%

Mosaicism level

Copy number

80%

80% 1 Chromosomes 1-24

Fig. 3.8  Gradient level of mosaicism for risk evaluation (see explanation in the text). PGDIS Position Statement on the Transfer of Mosaic Embryos. D.S. Cram, D. Leigh, A. Handyside, L. Rechitsky, K. Xu, G. Harton, J. Grifo, C.

Rubio, E. Fragouli, S. Kahraman, E. Forman, M. KatzJaffe, H. Tempest, A. Thornhill, C. Strom, T. Escudero, J. Qiao, S. Munne, J.L. Simpson, A. Kuliev. Reprod BioMed Online 2019; 39: e1–e4

References

geous because “false positive” results of mosaicism in embryos are minimized. The applicable commercially available kit for NGS is VeriSeq™ PGT Kit (Illumina). Karyomapping supplied by Illumina may also be used for PGT-A but requires different equipment and reagents than NGS. The primary application of karyomapping is PGT-M. An alternative NGS platform is Personal Genome Machine (PGM), developed by Thermo Fisher Scientific. It is based on the use of emulsion PCR which follows library preparation, leading to release of ions, detected during sequencing, that results in a shift of pH that is detected on the sensor. PGM simultaneously tests as many as 60 DNA samples, with the results in 12 h. It detects chromosome aneuploidy, segmental variation with resolution to 800 kb, and mosaicism to the 20% level. The commercially available kit for this platform is Ion ReproSeq™ PGS Kit (Thermo Fisher Scientific). NGS also can provide the option of combining aneuploidy testing for 24 chromosomes together with PGT-M. Combined PGT-M and PGT-A was first performed for GM1 gangliosidosis, the severe autosomal recessive lysosomal storage disorder [62]. Thereafter, application was extended to other genetic conditions and used together with preimplantation HLA typing. At present this combined approach is applied as a routine, together with HLA typing and translocations. Testing for multiple indications in a single comprehensive test avoids additional biopsy procedures. Our experience includes more than 2000 cycles in which such an approach was used, with 70% pregnancy rates per single embryo transfer, birth of up to 1000 unaffected children, and reduction of spontaneous abortion rate to under 10%. The pregnancy rate is increased by 15–20% if PGT-A accompanies PGT-M. Specifically designed platforms are being developed for a comprehensive testing of monogenetic and chromosomal disorders, with a commercial kit under development by PerkinElmer (PG-Seq™ NGS Kit (RHS Ltd), Agilent, and others, being validated for clinical use. In summary, advances in molecular technologies have changed current practices in PGT.  Traditional technologies are being

45

replaced by molecular genetic techniques that are more efficient, flexible, of higher throughput, potentially noninvasive, and of greater productivity. Use of NGS in PGT has dramatically increased our ability to detect aneuploid embryos and other genetic abnormalities. This naturally improves choices for embryo selection. NGS has also revolutionized the detection of point mutations, enabling parallel testing of many genes at the same time and again improving preselection of an embryo of highest potential for a viable pregnancy and birth of unaffected child.

References 1. Kuliev A, Rechitsky S, Verlinsky O. Atlas of preimplantation genetic diagnosis. 3rd ed. Taylor & Francis, London: CRC Press; 2014. 2. Rechitsky S, Verlinsky O, Strom C, et al. Experience with single-cell PCR in preimplantation genetic diagnosis: how to avoid pitfalls. In: Hahn S, Holzgreve W, editors. Fetal cells in maternal blood. New developments for a new millennium. 11th fetal cell workshop, Basel. Basel: Karger; 2000. p. 8–15. 3. Rechitsky S, Verlinsky O, Amet T, et al. Reliability of preimplantation diagnosis for single gene disorders. Mol Cell Endocrinol. 2001;183(Suppl 1):S65–8. 4. International Working Group on Preimplantation Genetics. Tenth anniversary of preimplantation genetic diagnosis. Report of the 10th annual meeting of international working group on preimplantation genetics, in conjunction with 3rd international symposium on preimplantation genetics, Bologna, June 23, 2000. J Assist Reprod Genet. 2001;18:66–72. 5. International Working Group on Preimplantation Genetics. Preimplantation genetic diagnosis – experience of three thousand clinical cycles. Report of the 11th annual meeting international working group on preimplantation genetics, in conjunction with 10th international congress of human genetics, Vienna, May 15, 2001. Reprod Biomed Online. 2001;3:49–53. 6. Verlinsky Y, Munne S, Cohen J, et al. Over a decade of preimplantation genetic diagnosis experience  – a multi-center report. Fertil Steril. 2004;82:292–4. 7. ESHRE Preimplantation Genetic Diagnosis (PGD) Consortium. Data collection II (May 2002). Hum Reprod. 2002;15:2673–83. 8. Verlinsky Y, Rechitsky S, Verlinsky O, Strom C, Kuliev A. Polar body based preimplantation diagnosis for X-linked genetic disorders. Reprod Biomed Online. 2002;4:38–42. 9. Rechitsky S, Strom C, Verlinsky O, et al. Allele drop out in polar bodies and blastomeres. J Assist Reprod Genet. 1998;15:253–7.

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10. Rechitsky S, Strom C, Verlinsky O, et al. Accuracy of preimplantation diagnosis of single-gene disorders by polar body analysis of oocytes. J Assist Reprod Genet. 1999;16:192–8. 11. Kuliev A, Rechitsky S, Verlinsky O, et  al. Birth of healthy children following preimplantation diagnosis for thalassemias. J Assist Reprod Genet. 1999;16:219–25. 12. Rechitsky S, Verlinsky O, Kuliev A, Ozen S, Masciangelo C, Lifchez A, Verlinsky Y. Preimplantation genetic diagnosis for familial dysautonomia. Reprod Biomed Online. 2003;6:488–93. 13. Findlay I, Quirke P.  Fluorescent polymerase chain reaction: Part I. A new method allowing genetic diagnosis and DNA fingerprinting of single cells. Hum Reprod Update. 1996;2:137–52. 14. Sermon K, Seneca S, De Rycke M, et  al. PGD in the lab for triplet diseases-myotonic dystrophy, Huntington’s disease and Fragile-X syndrome. Mol Cell Endocrinol. 2001;183:S77–85. 15. Katz MG, Mansfield J, Gras L, Trounson A, Cram DS.  Diagnosis of trisomy 21  in preimplantation embryos by single-cell DNA fingerprinting. Reprod Biomed Online. 2002;4:43–50. 16. Rechitsky S, Verlinsky O, Kuliev A. PGD for cystic fibrosis (CF) patients and couples at risk for additional genetic disorders combined with 24 chromosome aneuploidy testing. Reprod Biomed Online. 2013;26:420–30. 17. Prokhorovich M, Rechitsky S, Pakhalchuk T, San Ramon G, Gershman R, Bond E, Kuliev A.  Simultaneous preimplantation genetic testing (PGT) for 5 different genetic conditions. Reprod Biomed Online. 2019;39:e51. 18. Verlinsky Y, Rechitsky S, Verlinsky O, Strom C, Kuliev A.  Preimplantation diagnosis for long-chain 3-hydroxiacyl-CoA dehydrogenase deficiency. Reprod Biomed Online. 2001;2:17–9. 19. Renwick P, Trussler J, Braude P, Ogilvie CM. Preimplantation genetic haplotyping: 127 diagnostic cycles demonstrating a robust, efficient alternative to direct mutation testing on single cells. Reprod Biomed Online. 2010;20:470–6. 20. Rechitsky S, Pakhalchuk G, San Ramos G, et al. First systematic experience of preimplantation human leukocyte antigen typing combined with 24-chromosome aneuploidy testing. Fertil Steril. 2016;103:503–12. 21. Rechitsky S, Pomerantseva K, Pakhalchuk T, Polling D, Verlinsky O, Kuliev A.  First systematic experience of preimplantation genetic diagnosis for de novo mutations. Reprod Biomed Online. 2011;22:350–61. 22. Kuliev A, Rechitsky S. Preimplantation genetic testing: current challenges and future prospects. Expert Rev Mol Diagn. 2017;17(12):1071–88. 23. Handyside AH, Thornhill AR, Harton GL, et  al. Karyomapping: a novel molecular karyotyping method based on mapping crossovers between parental haplotypes with broad applications for preimplantation genetic diagnosis of inherited disease. J Med Genet. 2010;47:651–65.

24. Gianaroli L, Magli MC, Ferraretti AP, Munne S.  Preimplantation diagnosis for aneuploidies in patients undergoing in  vitro fertilization with poor prognosis: identification of the categories for which it should be proposed. Fertil Steril. 1999;72:837–44. 25. Munne S, Magli C, Cohen J, et al. Positive outcome after preimplantation diagnosis of aneuploidy in human embryos. Hum Reprod. 1999;14:2191–9. 26. Munne S, Sandalinas M, Escudero T, et al. Improved implantation after preimplantation genetic diagnosis of aneuploidy. Reprod Biomed Online. 2003;7:91–7. 27. Munné S, Alikani M, Tomkin G, Grifo J, Cohen J.  Embryo morphology, developmental rates and maternal age are correlated with chromosome abnormalities. Fertil Steril. 1995;64:382–91. 28. Harper JC, Coonen E, Handyside AH, et al. Mosaicism of autosomes and sex chromosomes in morphologically normal monospermic preimplantation human embryos. Prenat Diagn. 1995;15:41–9. 29. Dyban A, Fredine M, Severova E, Cieslac J, Wolf G, Kuliev A, Verlinsky Y.  Detection of aneuploidy in human oocytes and corresponding first polar bodies using FISH. In: 7th international conference on early prenatal diagnosis. Jerusalem, Israel, May 22–27, 1994 (Abstract #97). 30. Dyban A, Fredine M, Severova E, et  al. Detection of aneuploidy in human oocytes and corresponding first polar bodies by FISH.  J Assist Reprod Genet. 1996;13:72–7. 31. Munné S, Daily T, Sultan KM, Grifo J, Cohen J. The use of first polar bodies for preimplantation diagnosis of aneuploidy. Hum Reprod. 1995;10:1014–120. 32. Munne S, Morrison L, Fung J, et  al. Spontaneous abortions are reduced after preconception diagnosis of translocations. J Assist Reprod Genet. 1998;15: 290–6. 33. Verlinsky Y, Cieslak J, Freidine M, Ivakhnenko V, Wolf G, Kovalinskaya L, White M, Lifchez A, Kaplan B, Moise J, Ginsberg N, Strom C, Kuliev A. Pregnancies following pre-conception diagnosis of common aneuploidies by fluorescent in-situ hybridization. Hum Reprod. 1995;10:1923–7. 34. Kuliev A, Zlatopolsky Z, Kirillova I, Spivakova J, Cieslak-Janzen G.  Meiosis errors in over 20,000 oocytes studied in the practice of preimplantation aneuploidy testing. Reprod Biomed Online. 2011;22:2–8. 35. Verlinsky Y, Evsikov S.  Karyotyping of human oocytes by chromosomal analysis of the second polar body. Mol Hum Reprod. 1999;5:89–95. 36. Verlinsky Y, Evsikov S.  A simplified and efficient method for obtaining metaphase chromosomes from individual human blastomeres. Fertil Steril. 1999;72:1–6. 37. Verlinsky Y, Cieslak J, Evsikov S, Galat V, Kuliev A.  Nuclear transfer for full karyotyping and preimplantation diagnosis of translocations. Reprod Biomed Online. 2002;5:302–7. 38. Willadsen S, Levron J, Munne S, et al. Rapid visualization of metaphase chromosomes in single human

References blastomeres after fusion with in-vitro matured bovine eggs. Hum Reprod. 1999;14:470–4. 39. Tanaka A, Nagayoshi M, Awata S, Mawatari Y, Tanaka I, Kusunoki H.  Preimplantation diagnosis of repeated miscarriage due to chromosomal translocations using metaphase chromosomes of a blastomere biopsied from 4-6 cell stage embryo. Fertil Steril. 2004;81:30–4. 40. Kuliev A, Jansen JC, Zlatopolski Z, et al. Conversion and non-conversion approach to preimplantation diagnosis for chromosomal rearrangements in 475 cycles. Reprod Biomed Online. 2010;21:93–9. 41. Shkumatov A, Kuznyetsov V, Cieslak J, Ilkevitch Y, Verlinsky Y.  Obtaining metaphase spreads from single blastomeres for PGD of chromosomal rearrangements. Reprod Biomed Online. 2007;14:498–503. 42. Munne S, Wells D, Cohen J.  Technology requirements for preimplantation genetic diagnosis to improve assisted reproduction outcomes. Fertil Steril. 2010;92:408–30. 43. Treff NT, Levy B, Su J, Northrop LE, Tao X, Scott RT.  SNP microarray-based 24 chromosome aneuploidy screening is significantly more consistent than FISH. Mol Hum Reprod. 2010;16:583–9. 44. Johnson DS, Gemelos G, Ryan A, et  al. Preclinical validation of a microarray method for full molecular karyotyping of blastomeres in a 24-h protocol. Hum Reprod. 2010;25:1066–75. 45. Treff NT, Sue G, Tao X, Levy B, Scott RT. Accurate single cell 24 chromosome aneuploidy screening using whole genome amplification and single nucleotide polymorphism microarrays. Fertil Steril. 2010;94:2017–21. 46. Gutierrez-Mateo C, Colls P, Sanchez-Garcia J, Escudero T, Prates R, Ketterson K, Wells D, Munne S.  Validation of microarray comparative genomic hybridization for comprehensive chromosome analysis of embryos. Fertil Steril. 2011;95:953–8. 47. Fragouli E, Alfarawati S, Daphnis DD, et  al. Cytogenetic analysis of human blastocyst with the use of FISH, CGH, and aCGH: scientific data and technical evaluation. Hum Reprod. 2011;26:480–90. 48. Gabriel AS, Thornhill AR, Ottolini CS, et  al. Array comparative genomic hybridization on first polar bodies suggests that non-disjunction is not the predominant mechanism leading to aneuploidy in humans. J Med Genet. 2011;48:433–7. 49. Geraedts J, Collins J, Gianaroli L, et  al. What next for preimplantation genetic screening? A polar body approach. Hum Reprod. 2010;25:575–7. 50. European Society of Human Reproduction and Embryology (ESHRE). Abstracts of the 26th annual meeting of European Society of Human

47 Reproduction and Embryology. Hum Reprod. 2010;25(Suppl. 1):17–8. 51. Schoolcraft WB, Fragouli E, Stevens J, Munne S, Katz-Jaffe MG, Wells D. Clinical application of comprehensive chromosomal screening in the blastocyst stage. Fertil Steril. 2010;94:1700–6. 52. Scott RT, Tao X, Ferry KM, Treff NR. A prospective randomized controlled trial demonstrating significantly increased clinical pregnancy rates following 24 chromosome aneuploidy screening: biopsy on day 5 with fresh transfer. Fertil Steril. 2010;94(Supplement S2):O-05. 53. Treff NR, Northrop LE, Kasabwala K, Su J, Levy B, Scott RT.  Single nucleotide polymorphism microarray-based concurrent screening of 24-­ chromosome aneuploidy and unbalanced translocations in preimplantation human embryos. Fertil Steril. 2011;95(5):1606. 54. Wells D, Kaur K, Grifo J, et al. Clinical utilisation of a rapid low-pass whole genome sequencing technique for the diagnosis of aneuploidy in human embryos prior to implantation. J Med Genet. 2014;51:553–62. 55. Fiorentino F, Bono S, Biricik A, et al. Application of next-generation sequencing technology for comprehensive aneuploidy screening of blastocysts in clinical preimplantation genetic screening cycles. Hum Reprod. 2014;29(12):2802–13. 56. Rubio C, Bellver J, Rodrigo L, et  al. In vitro fertilization with preimplantation genetic diagnosis for aneuploidies in advanced maternal age: a randomized, controlled study. Fertil Steril. 2017;107(5):1122–9. 57. Fragouli E, Alfarawati S, Spath K, et al. Analysis of implantation and ongoing pregnancy rates following the transfer of mosaic diploid-aneuploid blastocysts. Hum Genet. 2017;136(7):805–19. 58. Greco E, Minasi MG, Fiorentino F.  Healthy babies after intrauterine transfer of mosaic aneuploid blastocysts. N Engl J Med. 2015;373(21):2089–90. 59. Munne S, Blazek J, Large M, et al. Detailed investigation into the cytogenetic constitution and pregnancy outcome of replacing mosaic blastocysts detected with the use of high-resolution next-generation sequencing. Fertil Steril. 2017;108(1):62–71.e8. 60. PGDIS.  Position statement on mosaicism. 2016. www.pgdis.org. 61. PGDIS. Position statement on mosaic transfer. 2019. www.pgdis.org. 62. Brezina PR, Benner A, Rechitsky S, et al. Single-gene testing combined with single nucleotide polymorphism microarray preimplantation genetic diagnosis for aneuploidy: a novel approach in optimizing pregnancy outcome. Fertil Steril. 2011;95:1786.

4

Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Monogenic disorders were the first group of indications for which preimplantation genetic testing (PGT-M) was originally introduced almost 30  years ago. The purpose was performing genetic testing before clinical pregnancy in order to establish only unaffected pregnancies and to avoid the need for pregnancy termination [1, 2]. Despite the requirement for ovarian hyperstimulation and in  vitro fertilization (IVF) needed to perform genetic testing of oocytes or embryos prior to transfer, PGT-M has been accepted in most parts of the world [3, 4]. Hundreds of thousands of PGT cycles have now been performed for single-gene disorders and, as will be shown below, are presently offered for some indications that have never been offered in invasive prenatal diagnosis, such as late-onset disorders with genetic predisposition or preimplantation HLA typing, making PGT-M a real alternative to prenatal diagnosis [5–8]. This book will explain strategies needed for PGT-M for different circumstances as well as major indications compared to prenatal genetic diagnosis. Practical details needed to assure accuracy will be described. Initially, indications for PGT were similar to those practiced in prenatal diagnosis and applied for those at-risk couples which could not accept pregnancy termination, expected in 25–50% of cases following prenatal diagnosis, depending on the mode of inheritance. Indications have now extended beyond those for invasive prenatal diag© Springer Nature Switzerland AG 2020 A. Kuliev et al., Practical Preimplantation Genetic Testing, https://doi.org/10.1007/978-3-030-43157-0_4

nosis and currently include conditions with a low penetrance, late-onset disorders with genetic ­predisposition, and HLA typing with or without testing for causative genes [8]. The list of disorders for which we applied PGT-M now comprises 581 different conditions (Table  4.1), with the most frequent shifting to common conditions with genetic predisposition, such as breast cancer. Initially most frequent PGT-M was opted for cystic fibrosis (CFTR), hemoglobin disorders, and dynamic mutations such as myotonic dystrophy. The choice between invasive prenatal diagnosis and PGT-M mainly depended on the patient’s attitude to termination of pregnancy, which is strongly influenced by social and religious factors but steadily is becoming a part of family planning for couples at risk to ensure having only unaffected pregnancy. Risk of having offspring with severe late-onset common disorders of strong genetic predisposition is an increasingly accepted indication for PGT-M [8]. Spectrum of referral to PGT-M has also changed, with current shift to direct referral through the information in the Internet. As described in Chap. 1, extended preconception carrier screening is currently introduced to identify couples without a family history who can benefit from PGT-­ M. Still, there is a need for improving awareness of PGT-M both for couples at risk and for medical providers. Table 4.2 presents the outcome of the above experience through 2018, involving over 6204 49

Colorectal cancer, hereditary nonpolyposis, type 2; HNPCC2 Colorectal cancer, hereditary nonpolyposis, type 4; HNPCC4 Colorectal cancer, hereditary nonpolyposis, type 5; HNPCC5 Dyskeratosis congenita, autosomal dominant 3; DKCA3 Dyskeratosis congenita, autosomal dominant, 2; DKCA2 Dyskeratosis congenita; DKCA2 Dyskeratosis congenita, autosomal recessive, 5; DKCB5 Dyskeratosis congenita, X-linked; DKCX Epidermolysis bullosa dystrophica, autosomal dominant; DDEB

Disease Familial cancer susceptibility syndromes Albinism, oculocutaneous, type IA; OCA1A Albinism, oculocutaneous, type III; OCA3 Ataxia–telangiectasia; AT Basal cell nevus syndrome; BCNS (Gorlin) Birt–Hogg–Dube syndrome; BHD Bloom syndrome; BLM Breast and colorectal cancer, susceptibility Breast cancer Breast–ovarian cancer, familial, susceptibility to, 1; BROVCA1 Breast–ovarian cancer, familial, susceptibility to, 2; BROVCA2 Breast–ovarian cancer, familial, susceptibility to, 3; BROVCA3 Carney complex, type 1; CNC1 AR AR AR AD AD AR AD AD AD AD AR AD AD AD AD AD AD

AR XL AR

203100 203290 208900 109400 135150 210900 604373 114480 113705 612555 613399 160980 609310 614337 614350 613990 613989

615190 305000 131750

DKC1 COL7A1

RTEL1

TERT

TINF2

MSH6

PMS2

MLH1

PRKAR1A

RAD51C

BRCA2

TYR TYRP1 ATM PTCH1 FLCN RECQL3 CHEK2 PALB2 BRCA1

Gene name/ Inheritance symbol

OMIM number

Table 4.1  List of genetic conditions for which PGT was performed

H/ACA ribonucleoprotein complex subunit 4 Collagen alpha-1(VII) chain

Regulator of telomere elongation helicase 1

Telomerase reverse transcriptase

TERF1-interacting nuclear factor 2

DNA mismatch repair protein Msh6

Mismatch repair endonuclease PMS2

cAMP-dependent protein kinase type I-alpha regulatory subunit DNA mismatch repair protein Mlh1

DNA repair protein RAD51 homolog 3

Breast cancer type 2 susceptibility protein

Tyrosinase 5,6-Dihydroxyindole-2-carboxylic acid oxidase Serine–protein kinase ATM Protein patched homolog 1 Folliculin Bloom syndrome protein Serine/threonine-protein kinase Chk2 Partner and localizer of BRCA2 Breast cancer type 1 susceptibility protein

Protein name

Xq28 3p21.31

20q13.33

5p15.33

14q12

2p16.3

7p22.1

3p22.2

17q24.2

17q22

13q13.1

11q14.3 9p23 11q22.3 9q22.32 17p11.2 15q26.1 22q12.1 16p12.2 17q21.31

Location

50 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

AD AD AD AD

AD AD AD AD AR

175200 601399 601200 103580 603233 605074 180200 609322 187300 600376 191100 613254 606219

Peutz–Jeghers syndrome; PJS Platelet disorder, familial, with associated myeloid malignancy; FPDMM Pleuropulmonary blastoma; PPB Pseudohypoparathyroidism, type IA; PHP1A

Pseudohypoparathyroidism, type IB; PHP1B

Renal cell carcinoma, papillary, 1; RCCP1 Retinoblastoma; RB1 Rhabdoid tumor predisposition syndrome 1; RTPS1

Telangiectasia, hereditary hemorrhagic, of Rendu, Osler, and Weber; HHT Telangiectasia, hereditary hemorrhagic, type 2; HHT2 Tuberous sclerosis 1; TSC1 Tuberous sclerosis 2; TSC2 Tumor necrosis factor receptor-associated protein 1; TRAP1

AD AD AD

AD

AD

614165

Paragangliomas 5; PGL5

AD AD AR AD AD AD AD AD AD AD AD AD

175100 137215 203300 151623 120435 131100 171400 610755 614286 162200 101000 606864

Familial adenomatous polyposis 1; FAP1 Gastric cancer, hereditary diffuse; HDGC Hermansky–Pudlak Syndrome 1; HPS1 LI–Fraumeni syndrome 1; LFS1 Lynch syndrome I Multiple endocrine neoplasia, type I; MEN1 Multiple endocrine neoplasia, type IIA; MEN2A Multiple endocrine neoplasia, type IV; MEN4 Myelodysplastic syndrome; MDS Neurofibromatosis, type I; NF1 Neurofibromatosis, type II; NF2 Paraganglioma and gastric stromal sarcoma

ACVRL1 TSC1 TSC2 TRAP1

ENG

MET RB1 SMARCB1

GNAS

DICER1 GNAS

STK11 RUNX1

SDHA

APC CDH1 HPS1 TP53 MSH2 MEN1 RET CDKN1B GATA2 NF1 NF2 SDHB

Serine/threonine-protein kinase receptor R3 Hamartin Tuberin TNF receptor-associated protein 1

Endoribonuclease Dicer Guanine nucleotide-binding protein G(s) subunit alpha isoforms short Guanine nucleotide-binding protein G(s) subunit alpha isoforms short Hepatocyte growth factor receptor Retinoblastoma-associated protein SWI/SNF-related, matrix-associated, actin-dependent regulator of chromatin subfamily B member Endoglin

Adenomatous polyposis coli protein Cadherin-1 Hermansky–Pudlak syndrome 1 protein Cellular tumor antigen p53 DNA mismatch repair protein Msh2 Menin Proto-oncogene tyrosine-protein kinase receptor Ret Cyclin-dependent kinase inhibitor 1B Endothelial transcription factor GATA-2 Neurofibromin Merlin Succinate dehydrogenase [ubiquinone] iron–sulfur subunit, mitochondrial Succinate dehydrogenase [ubiquinone] flavoprotein subunit, mitochondrial Serine/threonine-protein kinase STK11 Runt-related transcription factor 1

(continued)

12q13.13 9q34.13 16p13.3 16p13.3

9q34.11

7q31 13q14.2 22q11.23

20q13.32

14q32.13 20q13.32

19p13.3 21q22.12

5p15.33

5q22.2 16q22.1 10q24.2 17p13.1 2p21-p16 11q13.1 10q11.21 12p13.1 3q21.3 17q11.2 22q12.2 1p36.13

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M) 51

616276 142900

Coenzyme Q10 deficiency, primary, 7; COQ10D7

Holt–Oram syndrome; HOS

AD

AR

AD AR

125310 607426

AD

615280

AD AD AD AD

AR XL AR

256050 302060 604377

115195 115197 613690 608751

AR

208050

AD AD AD AR AD AD

AD AD AR

106100 109730 609040

115200 613172 601154 604145 192600 615821

Inheritance AD AD AR

OMIM number 181450 193300 278780

Cardiomyopathy, dilated, 1A; CMD1A Cardiomyopathy, dilated, 1DD; CMD1DD Cardiomyopathy, dilated, 1E; CMD1E Cardiomyopathy, dilated, 1G; CMD1G Cardiomyopathy, dilated, 1S; CMD1S Cardiomyopathy, dilated, with woolly hair, keratoderma, and tooth agenesis; DCWHKTA Cardiomyopathy, familial hypertrophic, 2; CMH2 Cardiomyopathy, familial hypertrophic, 4; CMH4 Cardiomyopathy, familial hypertrophic, 7; CMH7 Cardiomyopathy, familial hypertrophic, 8; CMH8 Cardiomyopathy, familial hypertrophic, 8; CMH8 Cerebral arteriopathy, autosomal dominant Coenzyme Q10 deficiency, primary, 1; COQ10D1

Disease Ulnar–mammary syndrome; UMS Von Hippel–Lindau syndrome; VHL Xeroderma pigmentosum, complementation group G; XPG Cardiovascular disorders Angioedema, hereditary, type I; HAE1 Aortic valve disease 1; AOVD1 Arrhythmogenic right ventricular dysplasia, familial, 9; ARVD9 Arterial tortuosity syndrome; ATS Arterial tortuosity syndrome; ATS Atelosteogenesis, type II; AO2 Barth syndrome; BTHS Cardioencephalomyopathy, fatal infantile, due to cytochrome C oxidase deficiency 1 Cardiofaciocutaneous syndrome 4; CFC4

Table 4.1 (continued)

TBX5

COQ4

NOTCH3 COQ2

TNNT2 MYBPC3 TNNI3 MYL3

LMNA RBM20 SCN5A TTN MYH7 DSP

MAP2K2

SLC26A2 TAZ SCO2

SLC2A10

C1NH NOTCH1 PKP2

Gene name/ symbol TBX3 VHL ERCC5

Neurogenic locus notch homolog protein 3 4-Hydroxybenzoate polyprenyltransferase, mitochondrial Ubiquinone biosynthesis protein COQ4 homolog, mitochondrial T-box transcription factor TBX5

Troponin T type 2 (cardiac) Myosin-7 Troponin I, cardiac muscle Myosin light chain 3

Dual specificity mitogen-activated protein kinase kinase 2 Prelamin-A/C RNA binding motif protein 20 Sodium channel protein type 5 subunit alpha Titin Myosin, heavy chain 7, cardiac muscle, beta Desmoplakin

Solute carrier family 2, facilitated glucose transporter member 10 Sulfate transporter Tafazzin Protein SCO2 homolog, mitochondrial

Plasma protease C1 inhibitor Neurogenic locus notch homolog protein 1 Plakophilin-2

Protein name T-box transcription factor TBX3 Von Hippel–Lindau disease tumor suppressor DNA repair protein complementing XP-G cells

12q24.21

9q34.11

19p13.12 4q21.23

1q32.1 11p11.2 19q13.42 3p21.31

1q22 10q25.2 3p22.2 2q31.2 14q11.2 6p24.3

19p13.3

5q32 Xq28 22q13.33

20q13.12

11q12.1 9q34.3 12p11.21

Location 12q24.21 3p25.3 13q33.1

52 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

AR

163950 212350

257300

Noonan syndrome 1; NS1 Sengers syndrome Chromosomal breakage and aneuploidy disorder Aneuploidy by NGS mosaic variegated aneuploidy syndrome 1; MVA1

Cutis laxa, autosomal recessive, type IIB; ARCL2B Cutis laxa, autosomal recessive, type IIIA; ARCL3A Ehlers–Danlos syndrome, classic type Ehlers–Danlos syndrome, type IV, autosomal dominant Ehlers–Danlos syndrome, type VI; EDS6 Ehlers–Danlos syndrome, type VII, autosomal recessive Ehlers–Danlos syndrome, dermatosparaxis type Marfan syndrome; MFS Pseudoxanthoma elasticum; PXE Craniofacial disorders Branchiooculofacial syndrome; BOFS Cranioectodermal dysplasia 2; CED2 Craniofrontonasal syndrome; CFNS Craniosynostosis 2; CRS2

Nijmegen breakage syndrome; NBS Connective tissue disorders Cutis laxa, autosomal dominant 1; ADCL1 Cutis laxa, autosomal recessive, type IA; ARCL1A

AD AR

618447

Long QT syndrome 8; LQT8

AR AR AD AD AR AR

AD AR AD AR XL AD

225400 225410

154700 264800 113620 613610 304110 604757

AD AR

123700 614437 612940 219150 130000 130050

AR

251260

AD

AD

613688

Long QT syndrome 2; LQT2

AD AD

609192 192500

Loeys–Dietz syndrome 1; LDS1 Long QT syndrome 1; LQT1

TFAP2A WDR35 EFNB1 MSX2

FBN1 ABCC6

PLOD1 ADAMTS2

PYCR1 ALDH18A1 COL5A1 COL3A1

ELN FBLN4

NBN

BUB1B

PTPN11 AGK

CACNA1C

KCNH2

TGFBR2 KCNQ1

Transcription factor AP-2 alpha WD repeat-containing protein 35 Ephrin-B1 Homeobox protein MSX-2

Fibrillin 1 Multidrug resistance-associated protein 6

Procollagen-lysine,2-oxoglutarate 5-dioxygenase 1 A disintegrin and metalloproteinase with thrombospondin motifs 2

Elastin EGF-containing fibulin-like extracellular matrix protein 2 Pyrroline-5-carboxylate reductase 1, mitochondrial Delta-1-pyrroline-5-carboxylate synthase Collagen alpha-1(V) chain Collagen alpha-1(III) chain

Mitotic checkpoint serine/threonine-protein kinase BUB1 beta NIBRIN

TGF-Beta receptor type 1 Potassium voltage-gated channel subfamily KQT member 1 Potassium voltage-gated channel subfamily H member 2 Voltage-dependent L-type calcium channel subunit alpha-1C Tyrosine-protein phosphatase non-receptor type 11 Acylglycerol kinase

(continued)

6p24.3 2p24.1 Xq13.1 5q35.2

15q21.1 16p13.11

1p36.22 5q35.3

17q25.3 10q24.1 9q34.3 2q32.2

7q11.23 11q13.1

8q21.3

15q15.1

12q24.13 7q34

12p13.33

7q36.1

9q22.33 11p15.5

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M) 53

Deafness, autosomal dominant 3b; DFNA3B Deafness, autosomal dominant 3b; DFNA3B Deafness, autosomal recessive 3; DFNB3 Deafness, autosomal recessive 8; DFNB8 Deafness, neurosensory, autosomal recessive 1; DFNB1 Pendred syndrome; PDS Usher syndrome, type I; USH1 Usher syndrome, type IF; USHIF Usher syndrome, type IIA; USH2A Usher syndrome, type IIC; USH2C Usher syndrome, type IIC; USH2C Waardenburg syndrome, type 2A; WS2A Warburg Micro syndrome 1; WARBM1 Wolfram syndrome 1; WFS1 Dermatologic disorders Alopecia universalis congenita; ALUNC

Van der Woude syndrome 1; VWS1 Deafness Auriculocondylar syndrome 2; ARCND2

Disease Crouzon syndrome Dentinogenesis imperfecta, Shields type III Holoprosencephaly 2; HPE2 Hydrocephalus due to congenital stenosis of aqueduct of Sylvius; HSAS Hydrocephalus, nonsyndromic, autosomal recessive 2; HYC2 Pfeiffer syndrome Popliteal pterygium syndrome; PPS Saethre–Chotzen syndrome; SCS Treacher Collins syndrome 1; TCS1 Treacher Collins syndrome 2; TCS2

Table 4.1 (continued)

AD AR AR AR AR AR AR AR AR AR AD AR AR AR

612643 600316 601072 220290 274600 276900 602083 276901 605472 605472 193510 600118 222300 203655

AR

614669

AD AD AD AD AD

101600 119500 101400 154500 613717 AD

AR

615219

119300

Inheritance AD AD AD XL

OMIM number 123500 125500 157170 307000

HR

SLC26A4 MYO7A PCDH15 USH2A GPR98 ADGRV1 MITF RAB3GAP1 WFS1

MYO15A TMPRSS3 GJB2

GJB6

PLCB4

IRF6

FGFR1 IRF6 TWIST1 TCOF1 POLR1D

MPDZ

Gene name/ symbol FGFR2 DSPP SIX3 L1CAM

Lysine-specific demethylase hairless

Pendrin Unconventional myosin-VIIa Protocadherin-15 Usherin Adhesion G-protein-coupled receptor V1 G-protein-coupled receptor 98 Microphthalmia-associated transcription factor Rab3 GTPase-activating protein catalytic subunit Wolframin

Unconventional myosin-XV Transmembrane protease serine 3 Gap junction beta-2 protein

1-Phosphatidylinositol 4,5-bisphosphate phosphodiesterase beta-4 Gap junction beta-6 protein

Fibroblast growth factor receptor 1 Interferon regulatory factor 6 Twist-related protein 1 Treacle protein DNA-directed RNA polymerases I and III subunit RPAC2 Interferon regulatory factor 6

Multiple PDZ domain protein

Protein name Fibroblast growth factor receptor 2 Dentin sialophosphoprotein Homeobox protein SIX3 Neural cell adhesion molecule L1 [precursor]

8p21.3

7q22.3 11q13.5 10q21.1 1q41 5q14.3 5q14.3 3p13 2q21.3 4p16.1

17p11.2 21q22.3 13q12.11

13q12.11

20p12.3-p12.2

1q32.2

8p11.23 1q32.2 7p21.1 5q32–q33 13q12.2

9p23

Location 10q26.13 4q22.1 2p21 Xq28

54 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Benign chronic pemphigus; BCPM Cockayne syndrome A; CSA Darier–White disease; DAR Ectodermal dysplasia 10B, hypohidrotic/hair/tooth type, autosomal recessive; ECTD10B Ectodermal dysplasia, hypohidrotic, X-linked; XHED Ectrodactyly, ectodermal dysplasia, and cleft lip/palate syndrome 3; EEC3 epidermolysis bullosa simplex with pyloric atresia; EBSPA Epidermolysis bullosa simplex, Dowling–Meara type; EBSDM Epidermolysis bullosa simplex, Dowling–Meara type; EBSDM Epidermolysis bullosa, junctional, Herlitz type Epidermolysis bullosa, junctional, non-Herlitz type Epidermolytic hyperkeratosis; EHK Geroderma osteodysplasticum; GO Ichthyosis, congenital, autosomal recessive 1; ARCI1 Ichthyosis, cyclic, with epidermolytic hyperkeratosis Ichthyosis, lamellar, 2; LI2 Ichthyosis, spastic quadriplegia, and mental retardation; ISQMR Ichthyosis, X-linked; XLI IFAP syndrome with or without BRESHECK syndrome Mal de Meleda; MDM Palmoplantar keratoderma, Nagashima type; PPKN Pachyonychia congenita 3; PC3 Restrictive dermopathy, lethal Schopf–Schulz–Passarge syndrome; SSPS Endocrinologic disorders Adrenal hyperplasia, congenital, due to 21-hydroxylase deficiency Allan–Herndon–Dudley syndrome; AHDS

AR AR AD AR XL AD AR AD AD AR AR AD AR AD AD AR AR XL XL AR AD AR AR AR XL

169600 216400 124200 224900 305100 604292 612138 131760 131760 226700 226650 113800 231070 242300 607602 601277 614457 308100 308205 615598 615726 275210 224750 201910 300523

SLC16A2

CYP21A2

KRT6A ZMPSTE24 WNT10A

SLURP1

STS MBTPS2

LAMA3 LAMB3 KRT10 GORAB TGM1 KRT1 ABCA12 ELOVL4

KRT14

KRT5

PLEC1

EDA TP63

ATP2C1 ERCC8 ATP2A2 EDAR

Monocarboxylate transporter 8

Steroid 21-hydroxylase

Keratin, type II cytoskeletal 6A CAAX prenyl protease 1 homolog Protein Wnt-10a

Serpin B7

Steryl–sulfatase Membrane-bound transcription factor site-2 protease

Laminin subunit alpha-3 Laminin subunit beta-3 Keratin 10 RAB6-interacting golgin Protein–glutamine gamma-glutamyltransferase K Keratin, type II cytoskeletal 1 ATP-binding cassette subfamily A member 12 Elongation of very long chain fatty acids protein 4

Keratin, type I cytoskeletal 14

Keratin, type II cytoskeletal 5

Plectin

Calcium-transporting ATPase type 2C member 1 DNA excision repair protein ERCC8 Sarcoplasmic/endoplasmic reticulum calcium ATPase 2 Tumor necrosis factor receptor superfamily member EDAR Ectodysplasin-A Tumor protein 63

(continued)

Xq13.2

6p21.33

12q13.13 1p34 2q35

8q24.3

Xp22.31 Xp22.12

18q11.2 1q32.2 17q21 1q24.2 14q12 12q13.13 2q35 6q14.1

17q21.2

12q13.13

8q24.3

Xq13.1 3q28

3q22.1 5q12.1 12q24.11 2q13

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M) 55

Disease Androgen insensitivity syndrome; AIS Blepharophimosis, ptosis, and epicanthus inversus syndrome; BPES Diabetes mellitus, permanent neonatal; PNDM Donohue syndrome Hyperinsulinemic hypoglycemia, familial, 1; HHF1 Hypogonadotropic hypogonadism 1 with or without anosmia; HH1 Hypogonadotropic hypogonadism 1 with or without anosmia; HH1 Hypogonadotropic hypogonadism 2 with or without anosmia; HH2 Hypoparathyroidism–retardation–dysmorphism syndrome; HRDS Johanson–Blizzard syndrome; JBS Pseudovaginal perineoscrotal hypospadias; PPSH Gastrointenstinal disorders Cholestasis, benign recurrent intrahepatic, 2; BRIC2 Cholestasis, progressive familial intrahepatic, 3; PFIC3 Hirschsprung disease, susceptibility to, 1; HSCR1 Infantile liver failure syndrome 1; ILFS1 Pancreatitis, hereditary; PCTT Hematologic disorders and coagulopathy Alpha-thalassemia Amegakaryocytic thrombocytopenia, congenital; CAMT Anemia, nonspherocytic hemolytic, due to G6PD deficiency Antithrombin III deficiency; AT3D Beta-thalassemia Bleeding disorder, platelet-type, 16; BDPLT16 Blood group Kell–Cellano system

Table 4.1 (continued)

AD AR AR XL XL AD AR AR AR AR AR AD AR AD AR AR XL AD AR AD AD

606176 246200 256450 308700 308700 147950 241410 243800 264600 605479 602347 142623 615438 167800 604131 604498 300908 613118 613985 187800 110900

SERPINC1 HBB ITGB3 KEL

G6PD

HBA1 MPL

RET LARS PRSS1

ABCB11 ABCB4

UBR1 SRD5A2

TBCE

FGFR1

KAL1

INS INSR ABCC8 ANOS1

Gene name/ Inheritance symbol XL AR AD FOXL2

OMIM number 313700 110100

Antithrombin III Hemoglobin subunit beta Integrin beta-3 Kell blood group glycoprotein

Glucose-6-phosphate 1-dehydrogenase

Hemoglobin subunit alpha Thrombopoietin receptor

Proto-oncogene tyrosine-protein kinase receptor Ret Leucine–tRNA ligase, cytoplasmic Trypsin-1

Bile salt export pump Phosphatidylcholine translocator ABCB4

E3 ubiquitin-protein ligase UBR1 3-Oxo-5-alpha-steroid 4-dehydrogenase 2

Tubulin-specific chaperone E

Fibroblast growth factor receptor 1

Anosmin-1

Insulin Insulin receptor ATP-binding cassette subfamily C member 8 Anosmin-1 protein

Protein name Androgen receptor Forkhead box protein L2

1q25.1 11p15.4 17q21.32 7q34

Xq28

16p13.3 1p34.2

10q11.21 5q32 7q34

2q31.1 7q21.12

15q15.2 2p23.1

1q42.3

8p11.23

Xp22.31

11p15.5 19p13.2 11p15.1 Xp22.31

Location Xq12 3q22.3

56 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Diamond–Blackfan anemia 1; DBA1 Diamond–Blackfan anemia 2; DBA2 Diamond–Blackfan anemia 3; DBA3 Diamond–Blackfan anemia 5; DBA5 Diamond–Blackfan anemia 9; DBA9 Factor V deficiency Factor VII deficiency Glanzmann thrombasthenia; GT Hemochromatosis, type 1; HFE1 Hemoglobin H disease; HBH Hemophagocytic lymphohistiocytosis, familial, 2; FHL2 Hemophagocytic lymphohistiocytosis, familial, 3; FHL3 Hemophagocytic lymphohistiocytosis, familial, 4; FHL4 Hemophilia A; HEMA Hemophilia B; HEMB Leukocyte adhesion deficiency, type I; LAD Lymphedema, hereditary, III; LMPH3 Lymphedema–distichiasis syndrome Prothrombin deficiency, congenital Pyruvate kinase deficiency of red cells Rhesus blood group, CcEE antigens; RHCE Rhesus blood group, D antigen; RHD Shwachman–Diamond syndrome; SDS Sickle cell anemia Spherocytosis, type 1; SPH1 Spherocytosis, type 2; SPH2 Thrombocythemia 1; THYCT1 Thrombocythemia 1; THYCT1 Thrombocytopenia, X-linked, with or without dyserythropoietic anemia; XLTDA Thrombocytopenia-absent radius syndrome; TAR

AD AD AD AD AD AR AR AR AR AR AR AR AR XL XL AR AR AD AR AD AD AD AR AR AD AD AR XL AR

105650 606129 610629 612528 613308 227400 227500 273800 235200 613978 603553 608898 603552 306700 306900 116920 616843 153400 613679 266200 111700 111680 260400 603903 182900 616649 187950 300367 274000

RBM8A

GATA1

F8 F9 ITGB2 PIEZO1 FOXC2 F2 PKLR RHCE RHD SBDS HBB ANK1 SPTB SH2B3

STX11

UNC13D

RPS19 RPS20 RPS24 RPL35A RPS10 F5 F7 ITGA2B HFE HBA2 PRF1

RNA-binding protein 8A

Erythroid transcription factor

Coagulation factor VIII Coagulation factor IX Integrin beta-2 Piezo-type mechanosensitive ion channel component 1 Forkhead box protein C2 Prothrombin Pyruvate kinase PKLR Blood group Rh(CE) polypeptide Blood group Rh(D) polypeptide Ribosome maturation protein SBDS Hemoglobin beta chain Ankyrin-1 Spectrin beta chain, erythrocytic SH2B adapter protein 3

Syntaxin-11

Protein unc-13 homolog D

40S ribosomal protein S19 40S ribosomal protein S20 40S ribosomal protein S24 60S ribosomal protein L35a 40S ribosomal protein S10 Coagulation factor V Coagulation factor VII Integrin alpha-IIb Hereditary hemochromatosis protein Hemoglobin subunit alpha Perforin-1

(continued)

1q21.1

Xp11.23

Xq28 Xq27.1 21q22.3 16q24.3 16q24.1 11p11.2 1q22 1p36.11 1p36.11 7q11.21 11p15.4 8p11.21 14q23.3 12q24.12

6q24.2

17q25.1

19q13.2 8q12.1 10q22.3 3q29 6p21.31 1q24.2 13q34 17q21.32 6p22.2 16p13.3 10q22.1

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M) 57

Disease Thrombophilia due to protein S deficiency, autosomal dominant; THPH5 Thrombotic thrombocytopenic purpura, congenital; TTP Immunologic disorders Adenosine deaminase deficiency; ADA Agammaglobulinemia, X-linked; XLA Aicardi–Goutieres syndrome 1; AGS1 Autoimmune polyendocrine syndrome, type I, Bare lymphocyte syndrome, type II Familial cold autoinflammatory syndrome 1; FCAS1 Familial Mediterranean fever; FMF Fanconi anemia, complementation group A; FANCA Fanconi anemia, complementation group B; FANCB Fanconi anemia, complementation group C; FANCC Fanconi anemia, complementation group D2; FANCD2 Fanconi anemia, complementation group E; FANCE Fanconi anemia, complementation group F; FANCF Fanconi anemia, complementation group G; FANCG Fanconi anemia, complementation group I; FANCI Fanconi anemia, complementation group J; FANCJ Granulomatous disease, chronic, autosomal recessive, cytochrome B-positive, type II; CDG2 Granulomatous disease, chronic, autosomal recessive, cytochrome B-positive, type I; CDG1 Granulomatous disease, chronic, X-linked; CDGX Griscelli syndrome, type 2; GS2 HLA matching genotyping Hyper-IgE recurrent infection syndrome, autosomal dominant Hyper-IgE recurrent infection syndrome, autosomal recessive immunodeficiency 25; IMD25

Table 4.1 (continued)

AR

AR XL AR AR AR AD AR AR XL AR AR AR AR AR AR AR AR AR XL AR AD AR AR

274150

608958 300755 225750 240300 209920 120100 F249100 227650 300514 227645 227646 600901 603467 614082 609053 609054 233710 233700 306400 607624 147060 243700 610163

CD247

DOCK8

STAT3

CYBB RAB27A

NCF1

FANCE FANCF FANCG FANCI BRIP1 NCF2

ADA BTK TREX1 AIRE RFX5 NLRP3 MEFV FANCA FANCB FANCC FANCD2

ADAMTS13

Gene name/ Inheritance symbol AD PROS1

OMIM number 612336

T-cell surface glycoprotein CD3 zeta chain

Dedicator of cytokinesis protein 8

Signal transducer and activator of transcription 3

Cytochrome b-245 heavy chain Ras-related protein Rab-27A

Neutrophil cytosol factor 1

Fanconi anemia group E protein Fanconi anemia group F protein Fanconi anemia group G protein Fanconi anemia group I protein Fanconi anemia group J protein Neutrophil cytosolic factor 2

Adenosine deaminase Tyrosine-protein kinase BTK Three prime repair exonuclease 1 Autoimmune regulator DNA-binding protein RFX5 NACHT, LRR, and PYD domains-containing protein 3 Pyrin Fanconi anemia group A protein Fanconi anemia group B protein Fanconi anemia group C protein Fanconi anemia group D2 protein

A disintegrin and metalloproteinase with thrombospondin motifs 13

Protein name Vitamin K-dependent protein S

1q24.2

9p24.3

Xp21.1-p11.4 15q21.3 6q21.3 17q21.2

7q11.23

6p21.31 11p14.3 9p13.3 15q26.1 17q23.2 1q25.3

20q13.12 Xq22.1 3p21.31 21q22.3 1q21.3 1q44 16p13.3 16q24.3 Xp22.2 9q22.32 3p25.3

9q34.2

Location 3q11.1

58 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

RAS-associated autoimmune leukoproliferative disorder; RALD Severe combined immunodeficiency Severe combined immunodeficiency, autosomal recessive Severe combined immunodeficiency, X-linked; SCIDX1 Wiskott–Aldrich syndrome; WAS Metabolic disorders 3-Hydroxyisobutyryl-CoA hydrolase deficiency; HIBCHD 3-Methylcrotonyl-CoA carboxylase 2 deficiency; MCC2D 3-Methylglutaconic aciduria with deafness, encephalopathy, and Leigh-like syndrome; MEGDEL Adenine phosphoribosyltransferase deficiency; APRTD Adrenoleukodystrophy; ALD Argininosuccinic aciduria Beta-ureidopropionase deficiency; UPB1D Biotinidase deficiency

Immunodeficiency with hyper-IgM, type 1; HIGM1 Immunodysregulation, polyendocrinopathy, and enteropathy, X-linked; IPEX Incontinentia Pigmenti; IP Lymphoproliferative syndrome, X-linked, 1; XLP1 Neutropenia, severe congenital, 1, autosomal dominant; SCN1 Neutropenia, severe congenital, 3, autosomal recessive; SCN3 Omenn syndrome Periodic fever, familial, autosomal dominant

AR AR XL XL AR AR AR AD XL AR AR AR

300400 301000 250620 210210 614739 614723 300100 207900 613161 253260

AD AD

603554 142680

608971 601457

AR

610738

AD

XL XL AD

308300 308240 202700

614470

XL XL

308230 304790

ABCD1 ASL UPB1 BTD

APRT

SERAC1

MCCC2

HIBCH

WAS

IL2RG

IL7R RAG2

NRAS

RAG1 TNFRSF1A

HAX1

IKBKG SH2D1A ELANE

CD40LG FOXP3

ATP-binding cassette subfamily D member 1 Argininosuccinate lyase Beta-ureidopropionase Biotinidase

Adenine phosphoribosyltransferase

Methylcrotonoyl-CoA carboxylase beta chain, mitochondrial Protein SERAC1

3-Hydroxyisobutyryl-CoA hydrolase, mitochondrial

Wiskott–Aldrich syndrome protein

Cytokine receptor common subunit gamma

Interleukin-7 receptor subunit alpha V(D)J recombination-activating protein 2

V(D)J recombination-activating protein 1 Tumor necrosis factor receptor superfamily member 1A GTPase NRas

HCLS1-associated protein X-1

NF-kappa-B essential modulator SH2 domain-containing protein 1A Neutrophil elastase

CD40 ligand Forkhead box protein P3

(continued)

Xq28 7q11.21 22q11.23 3p25.1

16q24.3

6q25.3

5q13.2

2q32.2

Xp11.23

Xq13.1

5p13.2 11p13

1p13.2

11p12 12p13.31

1q21.3

Xq28 Xq25 19p13.3

Xq26.3 Xp11.23

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M) 59

Congenital disorder of glycosylation, type Ia; CDG1A Congenital disorder of glycosylation, type IIc; CDG2C Congenital disorder of glycosylation, type IIl; CDG2L Congenital disorder of glycosylation, type Ik; CDG1K Congenital disorder of glycosylation, type In; CDG1N Danon disease D-bifunctional protein deficiency Dihydrolipoamide dehydrogenase deficiency; DLDD Encephalopathy, acute, infection-induced, susceptibility to, 3; IIAE3 Fabry disease Fructose intolerance, hereditary Fumarase deficiency; FMRD Galactosemia

Disease Canavan disease Carbamoyl phosphate synthetase I deficiency CPS I deficiency Carnitine deficiency, systemic primary; CDSP Carnitine–acylcarnitine translocase deficiency; CACTD Cerebral creatine deficiency syndrome 1; CCDS1 Ceroid lipofuscinosis, neuronal 2, late infantile; CLN2 Ceroid lipofuscinosis, neuronal, 1; CLN1 Ceroid lipofuscinosis, neuronal, 10; CLN10 Ceroid lipofuscinosis, neuronal, 3; CLN3 Ceroid lipofuscinosis, neuronal, 5; CLN5 Ceroid lipofuscinosis, neuronal, 6; CLN6 Citrullinemia, classic Combined malonic and methylmalonic aciduria; CMAMMA Combined oxidative phosphorylation deficiency 13; COXPD13 Congenital disorder of deglycosylation; CDDG

Table 4.1 (continued)

XL AR AR AR

AR

615273

301500 229600 606812 230400

AR

614932

AR AR AR AR AR AD AR AR AD

XL AR AR AR AR AR AR AR AR

300352 204500 256730 610127 204200 256731 601780 215700 614265

212065 266265 614576 608540 612015 300257 261515 246900 608033

AR AR

212140 212138

GLA ALDOB FH GALT

PMM2 SLC35C1 COG6 ALG1 RFT1 LAMP2 HSD17B4 DLD RANBP2

NGLY1

PNPT1

SLC6A8 TPP1 PPT1 CTSD CLN3 CLN5 CLN6 ASS1 ACSF3

SLC22A5 SLC25A20

Gene name/ Inheritance symbol AR ASPA AR CPS1

OMIM number 271900 237300

Alpha-galactosidase A Fructose-bisphosphate aldolase B Fumarate hydratase, mitochondrial Galactose-1-phosphate uridylyltransferase

Peptide-N(4)-(N-acetyl-beta-glucosaminyl)asparagine amidase Phosphomannomutase 2 GDP-fucose transporter 1 Conserved oligomeric Golgi complex subunit 6 Beta-mannosyltransferase Protein RFT1 homolog Lysosome-associated membrane glycoprotein 2 Peroxisomal multifunctional enzyme type 2 Dihydrolipoyl dehydrogenase, mitochondrial E3 SUMO-protein ligase RanBP2

Sodium- and chloride-dependent creatine transporter 1 Tripeptidyl–peptidase 1 Palmitoyl–protein thioesterase 1 Cathepsin D Battenin Ceroid-lipofuscinosis neuronal protein 5 Ceroid-lipofuscinosis neuronal protein 6 Argininosuccinate synthase Acyl-CoA synthetase family member 3, mitochondrial

Protein name Aspartoacylase Carbamoyl-phosphate synthase [ammonia], mitochondrial Solute carrier family 22 member 5 Mitochondrial carnitine–acylcarnitine carrier protein

Xq22.1 9q31.1 1q43 9p13.3

16p13.2 11p11.2 13q14.11 16p13.3 3p21.1 Xq24 5q23.1 7q31.1 2q13

3p24.2

2p16.1

Xq28 11p15.4 1p34.2 11p15.5 16p12.1 13q22.3 15q23 9q34.11 16q24.3

5q31.1 3p21.31

Location 17p13.2 2q34

60 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

AR AR AR AR XL AD AR AR AR

236250 250940 607014 243500 245200 300322 151660 201710 278000 248600 249900

AR

232700 232800 230500 604397 236200

Glycogen storage disease VI; GSD6 Glycogen storage disease VII; GSD7 Gm1-gangliosidosis, type I Guanine nucleotide-binding protein, alpha-14; GNA14 Homocystinuria due to cystathionine beta-synthase deficiency Homocystinuria due to deficiency of n(5,10)methylenetetrahydrofolate reductase activity Homocystinuria–megaloblastic anemia, CBLG complementation type; HMAG Hurler syndrome Isovaleric acidemia; IVA Krabbe disease Lesch–Nyhan syndrome; LNS Lipodystrophy, familial partial, type 2; FPLD2 Lipoid congenital adrenal hyperplasia; LCAH Lysosomal acid lipase deficiency Maple syrup urine disease; MSUD

Metachromatic leukodystrophy due to saposin B deficiency

AR AR AR AR AR

607839 232200 232220 232400 306000

Glycogen branching enzyme; GBE1 Glycogen storage disease Ia; GSD1A Glycogen storage disease Ib; GSD1B Glycogen storage disease III; GSD3 Glycogen storage disease IXa1; GSD9A1

AR

AR AR AR AR XL

AR AR AR

231670 605899 605899

Glutaric acidemia I Glycine encephalopathy; GCE Glycine encephalopathy; GCE

AR AR AD

256540 230800 606777

Galactosialidosis; GSL Gaucher disease, type I GLUT1 deficiency syndrome 1; GLUT1DS1

PSAP

IDUA IVD GALC HPRT1 LMNA STAR LIPA BCKDHB

MTR

MTHFR

PYGL PFKM GLB1 GNA14 CBS

GBE1 G6PC SLC37A4 AGL PHKA2

GCDH AMT GLDC

CTSA GBA SLC2A1

Alpha-L-iduronidase Isovaleryl-CoA dehydrogenase, mitochondrial Galactocerebrosidase Hypoxanthine-guanine phosphoribosyltransferase Prelamin-A/C Steroidogenic acute regulatory protein, mitochondrial Lysosomal acid lipase/cholesteryl ester hydrolase 2-Oxoisovalerate dehydrogenase subunit beta, mitochondrial Prosaposin

Methionine synthase

Methylenetetrahydrofolate reductase

Lysosomal protective protein Glucosylceramidase Solute carrier family 2, facilitated glucose transporter member 1 Glutaryl-CoA dehydrogenase, mitochondrial Aminomethyltransferase, mitochondrial Glycine dehydrogenase (decarboxylating), mitochondrial 1,4-Alpha-glucan-branching enzyme Glucose-6-phosphatase Glucose-6-phosphate exchanger SLC37A4 Glycogen debranching enzyme Phosphorylase b kinase regulatory subunit alpha, liver isoform Glycogen phosphorylase, liver form ATP-dependent 6-phosphofructokinase, muscle type Beta-galactosidase Guanine nucleotide-binding protein subunit alpha-14 Cystathionine beta-synthase

(continued)

10q22.1

4p16.3 15q15.1 14q31.3 Xq26.2–q26.3 1q22 8p11.23 10q23.31 6q14.1

1q43

1p36.22

14q22.1 12q13.11 3p22.3 9q21.2 21q22.3

3p12.2 17q21.31 11q23.3 1p21.2 Xp22.13

19p13.13 3p21.31 9p24.1

20q13.12 1q22 1p34.2

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M) 61

AR

AR AR AR AR AR XL XL AR AR AR

252500 253200 309900 252900 252920 253000 231680 237310 256550 257200 257220 607625 311250 311601 264470 614882 614883

Mucopolysaccharidosis type VI; MPS6 Mucopolysaccharidosis, type II; MPS2 Mucopolysaccharidosis, type IIIA; MPS3A Mucopolysaccharidosis, type IIIB; MPS3B Mucopolysaccharidosis, type IVA; MPS4A Multiple acyl-CoA dehydrogenase deficiency; MADD

N-Acetylglutamate synthase deficiency; NAGSD Neuraminidase deficiency Niemann–Pick disease, type A Niemann–Pick disease, type C1; NPC1 Niemann–Pick disease, type C2; NPC2 Ornithine transcarbamylase deficiency Pelizaeus–Merzbacher disease; PMD Peroxisomal Acyl-CoA oxidase deficiency Peroxisome biogenesis disorder 10A (Zellweger); PBD10A Peroxisome biogenesis disorder 11A (Zellweger); PBD11A

AR XL AR AR AR AR

AR

252160

AR

251110 AR

AR

251000

252150

AR

277380

PEX13

NAGS NEU1 SMPD1 NPC1 NPC2 OTC PLP1 ACOX1 PEX3

ARSB IDS SGSH NAGLU GALNS ETFA

GNPTAB

MOCS2

MOCS1

MUT

MUT

LMBRD1

Gene name/ Inheritance symbol AR ARSA AR MMACHC

OMIM number 250100 277400

Molybdenum cofactor deficiency, complementation group A; MOCODA Molybdenum cofactor deficiency, complementation group B; MOCODB Mucolipidosis II alpha/beta

Disease Metachromatic leukodystrophy; MLD Methylmalonic aciduria and homocystinuria, cblC type Methylmalonic aciduria and homocystinuria, cblF type; MAHCF Methylmalonic aciduria due to methylmalonyl-COA mutase deficiency Methylmalonic aciduria, cblB type

Table 4.1 (continued)

Peroxisomal membrane protein PEX13

N-Acetylglucosamine-1-phosphotransferase subunits alpha/beta Arylsulfatase B Iduronate 2-sulfatase N-Sulphoglucosamine sulphohydrolase Alpha-N-acetylglucosaminidase N-Acetylgalactosamine-6-sulfatase Electron transfer flavoprotein subunit alpha, mitochondrial N-Acetylglutamate synthase Sialidase-1 Sphingomyelin phosphodiesterase Niemann–Pick C1 protein Epididymal secretory protein E1 Ornithine carbamoyltransferase, mitochondrial Myelin proteolipid protein Peroxisomal acyl-coenzyme A oxidase 1 Peroxisomal biogenesis factor 3

Molybdopterin synthase catalytic subunit

Methylmalonic aciduria (cobalamin deficiency) cblB type Molybdenum cofactor biosynthesis protein 1

Methylmalonyl-CoA mutase, mitochondrial

Protein name Arylsulfatase A Methylmalonic aciduria and homocystinuria type C protein Probable lysosomal cobalamin transporter

2p15

17q21.31 6p21.33 11p15.4 18q11.2 14q24.3 Xp11.4 Xq22.2 17q25.1 6q24.2

5q14.1 Xq28 17q25.3 17q21.2 16q24.3 15q24.2–q24.3

12q23.2

5q11.2

6p21.2

12q24.11

6p12.3

6q13

Location 22q13.33 1p34.1

62 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

AR AR AR AR AR AR AR AR AR AR AR AR AR AR AR AR AR AR AR

214110 614859 614866 261600 263700 170100 606054 606054 610090 268800 271980 272300 272800 276700 277900 611126 203700 614585 256000 256,000 256000 618238

Leigh syndrome; LS

Leigh syndrome; LS Mitochondrial complex I deficiency, nuclear type 16; MC1DN16

AR AR

AR

AR

214100

Peroxisome biogenesis disorder 1A (Zellweger); PBD1A Peroxisome biogenesis disorder 2A (Zellweger); PBD2A Peroxisome biogenesis disorder 3A (Zellweger); PBD3A Peroxisome biogenesis disorder 5A (Zellweger); PBD5A Phenylketonuria; PKU Porphyria, congenital erythropoietic Prolidase deficiency Propionic acidemia Propionic acidemia Pyridoxamine 5-prime-phosphate oxidase deficiency; PNPOD Sandhoff disease Succinic semialdehyde dehydrogenase deficiency; SSADHD Sulfocysteinuria Tay–Sachs disease; TSD Tyrosinemia, type I; TYRSN1 Wilson disease Mitochondrial gene disorders Mitochondrial complex I deficiency due to ACAD9 deficiency Mitochondrial DNA depletion syndrome 4A (Alpers type); MTDPS4A Mitochondrial muscle myopathy Leigh syndrome; LS

SURF1 NDUFAF5

NDUFS8

AR NDUFS1

POLG

ACAD9

SUOX HEXA FAH ATP7B

HEXB ALDH5A1

PAH UROS PEPD PCCA PCCB PNPO

PEX2

PEX12

PEX5

PEX1

12q23.2 10q26.2 19q13.11 13q32.3 3q22.3 17q21.32

8q21.13

17q12

12p13.31

7q21.2

Ferredoxin 1-like protein; FDX1L NADH–ubiquinone oxidoreductase 75 kDa subunit, mitochondrial NADH dehydrogenase [ubiquinone] iron–sulfur protein 8, mitochondrial Surfeit locus protein 1 Arginine-hydroxylase NDUFAF5, mitochondrial

Acyl-CoA dehydrogenase family member 9, mitochondrial DNA polymerase subunit gamma-1

Sulfite oxidase, mitochondrial Beta-hexosaminidase subunit alpha Fumarylacetoacetase Copper-transporting ATPase 2

(continued)

9q34.2 20p12.1

11q13.2

19p13.2 2q33.3

15q26.1

3q21.3

12q13.2 15q23 15q25.1 13q14.3

Beta-hexosaminidase subunit beta 5q13.3 Succinate-semialdehyde dehydrogenase, mitochondrial 6p22.3

Phenylalanine-4-hydroxylase Uroporphyrinogen III synthase 19q13.11 Propionyl-CoA carboxylase alpha chain, mitochondrial Propionyl-CoA carboxylase beta chain, mitochondrial Pyridoxine-5′-phosphate oxidase

Peroxisome biogenesis factor 2

Peroxisome assembly protein 12

Peroxisomal targeting signal 1 receptor

Peroxisome biogenesis factor 1

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M) 63

Alagille syndrome 1; ALGS1 Angelman syndrome; AS Bardet–Biedl syndrome 10; BBS10 Bardet–Biedl syndrome 2; BBS2 Bardet–Biedl syndrome 4; BBS4 Cohen syndrome; COH1 Currarino syndrome DiGeorge syndrome; DGS Donnai–Barrow syndrome Fetal akinesia deformation sequence; FADS Fetal akinesia deformation sequence; FADS fetal akinesia deformation sequence; fads Fraser syndrome Fraser syndrome Harel–Yoon syndrome; HAYOS Lissencephaly, X-linked, 2; LISX2 Lissencephaly, X-linked, 2; LISX2 Marinesco–Sjogren syndrome; MSS Meckel syndrome, type 1; MKS1 Meckel syndrome, type 11; MKS11 Meckel syndrome, type 4; MKS4 Meckel syndrome, type 6; MKS6 Meckel syndrome, type 8; MKS8 Midface hypoplasia, hearing impairment, elliptocytosis, and nephrocalcinosis; MFHIEN Multinucleated neurons, anhydramnios, renal dysplasia, cerebellar hypoplasia

Disease Mitochondrial DNA depletion syndrome 13 Mitochondrial DNA depletion syndrome 2 (myopathic type); MTDPS2 Multiple congenital malformations Aicardi–Goutieres syndrome 2; AGS2 Aicardi–Goutieres syndrome 5; AGS5

Table 4.1 (continued)

AD AD AR AR AR AR AD AD AR AR AR AR AR AR AR XL AR AR AR AR AR AR XL AR

118450 105830 615987 615981 615982 216550 176450 188400 222448 208150 602552 208150 219000 219000 617183 300215 248800 249000 615397 611134 612284 613885 301050 236500

AR AR

610181 612952

CEP55

SIL1 MKS1 TMEM231 CEP290 CC2D2A TCTN2 AMMECR1

JAG1 UBE3A BBS10 BBS2 BBS4 VPS13B MNX1 TBX1 LRP2 RAPSN NUP88 MUSK FRAS1 FREM2 ATAD3A ARX

RNASEH2B SAMHD1

Gene name/ Inheritance symbol AR FBXL4 AR TK2

OMIM number 615471 609560

Centrosomal protein of 55 kDa

Nucleotide exchange factor SIL1 Meckel syndrome type 1 protein Transmembrane protein 231 Centrosomal protein of 290 kDa Coiled-coil and C2 domain-containing protein 2A Tectonic-2 AMME syndrome candidate gene 1 protein

Ribonuclease H2 subunit B Deoxynucleoside triphosphate triphosphohydrolase SAMHD1 Protein jagged-1 Ubiquitin-protein ligase E3A Bardet–Biedl syndrome 10 protein Bardet–Biedl syndrome 2 protein Bardet–Biedl syndrome 4 protein Vacuolar protein sorting-associated protein 13B Motor neuron and pancreas homeobox protein 1 T-box transcription factor TBX1 Low-density lipoprotein receptor-related protein 2 43 kDa receptor-associated protein of the synapse Nuclear pore complex protein Nup88 Muscle, skeletal receptor tyrosine-protein kinase Extracellular matrix protein FRAS1 FRAS1-related extracellular matrix protein 2 ATPase family AAA domain-containing protein 3A Homeobox protein ARX

Thymidine kinase 2, mitochondrial

Protein name

10q23.33

5q31.2 17q22 16q23.1 12q21.32 4p15.32 12q24.31 Xq22.3

20p12.2 15q11.2 12q21.2 16q13 15q24.1 8q22.2 7q36.3 22q11.21 2q31.1 11p11.2 17p13.2 9q31.3 4q21.21 13q13.3 1p36.33 Xp21.3

13q14.3 20q11.23

Location 6q16.1–q16.2 16q21

64 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

AD AD AR AD AR

AR

AR XL AD AR AR AR AR AR AD AD AD AR AR AD

609942 610733 270400 600725 218340

242840

266100 300672 617116 254780 609056 613477 615859 609304 612164 109150 164500 608629 213300 183086 616490 164400 614615 614615 229300 614815

Joubert syndrome 23; JBTS23 Spinocerebellar ataxia 1; SCA1 Joubert syndrome 17; JBTS17 Joubert syndrome 17; JBTS17 Friedreich ataxia 1; FRDA Joubert syndrome 18; JBTS18

AR AD AR AR AR AR

XL

300868

multiple congenital anomalies–hypotonia–seizures syndrome 2; MCAHS2 Noonan syndrome 3; NS3 Noonan syndrome 4; NS4 Smith–Lemli–Opitz syndrome; SLOS Sonic hedgehog; SHH Temtamy syndrome; TEMTYS Temtamy syndrome; TEMTYS Temtamy syndrome; TEMTYS vici syndrome; VICIS Neurologic disorders Epilepsy Epilepsy, pyridoxine-dependent; EPD Epileptic encephalopathy, early infantile, 2; EIEE2 Epilepsy, familial focal, with variable foci 2; FFEVF2 Myoclonic epilepsy of Lafora Salt and pepper developmental regression syndrome; SPDRS Epileptic encephalopathy, early infantile, 5; EIEE5 Epileptic encephalopathy, early infantile, 23; EIEE23 Epileptic encephalopathy, early infantile, 3; EIEE3 epileptic encephalopathy, early infantile, 4; EIEE4 Machado–Joseph disease; MJD Spinocerebellar ataxia 7; SCA7 Joubert syndrome 3; JBTS3 Joubert syndrome 1; JBTS1 Spinocerebellar ataxia 6; SCA6 KIAA0586 ATXN1 C5orf42 CPLANE1 FXN TCTN3

SPTAN1 DOCK7 SLC25A22 STXBP1 ATXN3 ATXN7 AHI1 INPP5E CACNA1A

ALDH7A1 CDKL5 NPRL2 NHLRC1 ST3GAL5

EPG5

KRAS SOS1 DHCR7 SHH C12orf57

PIGA

Spectrin alpha chain, non-erythrocytic 1 Dedicator of cytokinesis protein 7 Mitochondrial glutamate carrier 1 Syntaxin-binding protein 1 Ataxin-3 Ataxin-7 Jouberin 72 kDa inositol polyphosphate 5-phosphatase Voltage-dependent P/Q-type calcium channel subunit alpha-1A Protein TALPID3 Ataxin-1 Ciliogenesis and planar polarity effector 1 Uncharacterized protein C5orf42 Frataxin, mitochondrial Tectonic-3

Alpha-aminoadipic semialdehyde dehydrogenase Cyclin-dependent kinase-like 5 GATOR complex protein NPRL2 E3 ubiquitin-protein ligase NHLRC1 Lactosylceramide alpha-2,3-sialyltransferase

Ectopic P granules protein 5 homolog

Phosphatidylinositol N-acetylglucosaminyltransferase subunit A GTPase Kras Son of sevenless homolog 1 7-Dehydrocholesterol reductase Sonic hedgehog protein Protein C10

(continued)

14q23.1 6p22.3 5p13.2 5p13.2 9q21.11 10q24.1

9q34.11 1p31.3 11p15.5 9q34.11 14q32.12 3p14.1 6q23.3 9q34.3 19p13.13

5q23.2 Xp22.13 3p21.31 6p22.3 2p11.2

18q12.3–q21.1

12p12.1 2p22.1 11q13.4 7q36.3 12p13.31

Xp22.2

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M) 65

Movement disorders Dystonia 1, torsion, autosomal dominant; DYT1 Dystonia 11, myoclonic; DYT11 Dystonia 28, childhood-onset; DYT28 Dystonia 3, torsion, X-linked; DYT3 Neurodegenerative disorders Alzheimer disease 3 Alzheimer disease 4

Mental retardation, autosomal recessive 13; MRT13 Mental retardation, autosomal recessive 38; MRT38 Mental retardation, autosomal recessive 5; MRT5 Mental retardation, X-linked 102; MRX102 Mental retardation, X-linked, associated with fragile site FRAXE Rett syndrome; RTT Migraine Migraine, familial hemiplegic, 1; FHM1

Spinocerebellar ataxia 2; SCA2 Spinocerebellar ataxia 8; SCA8 Intellectual disabilities Dyskinesia, seizures, and intellectual developmental disorder; DYSEIDD Fragile X mental retardation syndrome Hypotonia, infantile, with psychomotor retardation and characteristic facies 3; IHPRF3 Mental retardation Mental retardation, autosomal dominant 35; MRD35

Disease Joubert syndrome 2; JBTS2 Joubert syndrome 21; JBTS21 Joubert syndrome 6; JBTS6 Spinocerebellar ataxia 13; SCA13

Table 4.1 (continued)

AD AD AD XL AD AD

607822 606889

AD

141500

128100 159900 617284 314250

XL

XL AD

NA 616355

312750

XL AR

300624 616900

AR AR AR XL XL

AR

617171

613192 615516 611091 300958 309548

AD AD

Inheritance AR AR AR AD

183090 608768

OMIM number 608091 615636 610688 605259

PSEN1 PSEN2

TOR1A SGCE KMT2B TAF1

CACNA1A

MECP2

TRAPPC9 HERC2 NSUN2 DDX3X AFF2

PPP1R3F PPP2R5D

FMR1 TBCK

DEAF1

ATXN2 ATXN8OS

Gene name/ symbol TMEM216 CSPP1 TMEM67 KCNC3

Presenilin-1 Presenilin-2

Torsin family 1, member A (torsin A) Epsilon-sarcoglycan e-Lysine N-methyltransferase 2B Transcription initiation factor TFIID subunit 1

Voltage-dependent P/Q-type calcium channel subunit alpha-1A

Methyl-CpG-binding protein 2

Protein phosphatase 1 regulatory subunit 3F Serine/threonine-protein phosphatase 2A 56 kDa regulatory subunit delta isoform Trafficking protein particle complex subunit 9 E3 ubiquitin-protein ligase HERC2 tRNA (cytosine(34)-C(5))-methyltransferase ATP-dependent RNA helicase DDX3X AF4/FMR2 family member 2

Synaptic functional regulator FMR1 TBC domain-containing protein kinase-like protein

Deformed epidermal autoregulatory factor 1 homolog

Protein name Transmembrane protein 216 Centrosome and spindle pole-associated protein 1 Meckelin Potassium voltage-gated channel subfamily C member 3 Ataxin-2 Putative protein ATXN8OS

14q24.2 1q42.13

9q34.11 7q21.3 19q13.12 Xq13.1

19p13.13

Xq28

8q24.3 15q13.1 5p15.31 Xp11.4 Xq28

Xp11.23 6p21.1

Xq27.3 4q24

11p15.5

12q24.12 13q21.33

Location 11q12.2 8q13.2 8q22.1 19q13.33

66 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Alzheimer disease; AD Creutzfeldt–Jakob disease; CJD Frontotemporal dementia and/or amyotrophic lateral sclerosis 1; FTDALS1 Frontotemporal lobar degeneration with TDP43 inclusions, GRN-related Gerstmann-Straussler disease; GSD Huntington disease; HD Infantile cerebellar-retinal degeneration; ICRD Leukoencephalopathy with vanishing white matter; VWM Leukoencephalopathy with vanishing white matter; VWM Leukoencephalopathy with vanishing white matter; VWM Megalencephalic leukoencephalopathy with subcortical cysts 1; MLC1 Pontocerebellar hypoplasia, type 1B; PCH1B Pontocerebellar hypoplasia, type 2A; PCH2A Pontocerebellar hypoplasia, type 2D; PCH2D Supranuclear palsy, progressive, 1; PSNP1 Neuronal migration disorders Periventricular nodular heterotopia 1; PVNH1 Polymicrogyria, bilateral frontoparietal; BFPP Polymicrogyria, bilateral frontoparietal; BFPP Neuromuscular disorders Hereditary neuropathies Amyloidosis, hereditary, transthyretin-related Charcot–Marie–Tooth disease, axonal, type 2A2; CMT2A2 Charcot–Marie–Tooth disease, axonal, type 2b; CMT2B Charcot–Marie–Tooth disease, axonal, type 2E; CMT2E

AD AD AD AD AD AD AR AR AR AR AR AR AR AR AD XL AR AR

AD AD AD AD

104300 123400 105550 607485 137440 143100 614559 603896 603896 603896 604004 614678 277470 613811 601104 300049 606854 606854

105210 609260 600882 607684

NEFL

RAB7A

TTR MFN2

FLNA ADGRG1 GPR56

EXOSC3 TSEN54 SEPSECS MAPT

MLC1

EIF2B2

EIF2B5

PRNP HTT ACO2 EIF2B4

PGRN

APP PRNP C9orf72

Neurofilament light polypeptide

Ras-related protein Rab-7a

Transthyretin [Precursor] Mitofusin-2

Filamin-A Adhesion G-protein-coupled receptor G1 Adhesion G-protein-coupled receptor G1

Exosome complex component RRP40 tRNA-splicing endonuclease subunit Sen54 O-Phosphoseryl-tRNA(Sec) selenium transferase Microtubule-associated protein tau

Membrane protein MLC1

Translation initiation factor eIF-2B subunit beta

Translation initiation factor eIF-2B subunit epsilon

Major prion protein Huntingtin Aconitate hydratase, mitochondrial Translation initiation factor eIF-2B subunit delta

Granulins

Amyloid beta A4 protein Major prion protein Guanine nucleotide exchange C9orf72

(continued)

8p21.2

3q21.3

18q12.1 1p36.22

Xq28 16q21 16q21

9p13.2 17q25.1 4p15.2 17q21.31

22q13.33

14q24.3

3q27.1

20p13 4p16.3 22q13.2 2p23.3

17q21.31

21q21.3 20p13 9p21.2

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M) 67

Charcot–Marie–Tooth disease, X-linked, 1; CMTX1 Hereditary motor and sensory neuropathy, type IIC; HMSN2C Hypertrophic neuropathy of Dejerine–Sottas Neuropathy, hereditary sensory and autonomic, type IA; HSAN1A Neuropathy, hereditary sensory and autonomic, type III; HSAN3 Neuropathy, hereditary sensory and autonomic, type VI; HSAN6 Motor neuron diseases Amyotrophic lateral sclerosis 1; ALS1 Amyotrophic lateral sclerosis 4, juvenile; ALS4 Spastic paraplegia 3, autosomal dominant; SPG3A Spastic paraplegia 4, autosomal dominant; SPG4 Spastic paraplegia 49, autosomal recessive; SPG49 Spinal and bulbar muscular atrophy, X-linked 1; SMAX1 Kennedy spinal and bulbar muscular atrophy Spinal muscular atrophy, distal, autosomal recessive, 1; DSMA1 Spinal muscular atrophy, type I; SMA1

Disease Charcot–Marie–Tooth disease, axonal, type 2F; CMT2F Charcot–Marie–Tooth disease, demyelinating, type 1A; CMT1A Charcot–Marie–Tooth disease, demyelinating, type 1B; CMT1B Charcot–Marie–Tooth disease, dominant intermediate C; CMTDIC Charcot–Marie–Tooth disease, type 4C; CMT4C

Table 4.1 (continued)

AR AD AR AR

AD AD AD AD AR XL

AR AR

223900 614653

105400 602433 182600 182601 615031 313200

604320 253300

AR

601596

145900 162400

AD

608323

XL AD

AD

118200

302800 606071

AD

118220

SMN1

IGHMBP2

SOD1 SETX ATL1 SPAST TECPR2 AR

IKBKAP

IKBKAP

PRX SPTLC1

GJB1 TRPV4

SH3TC2

YARS

MPZ

PMP22

Gene name/ Inheritance symbol AD HSPB1

OMIM number 606595

Survival motor neuron protein

DNA-binding protein SMUBP-2

Superoxide dismutase [Cu-Zn] Probable helicase senataxin Atlastin-1 Spastin Tectonin beta-propeller repeat-containing protein 2 Androgen receptor

Dystonin

Elongator complex protein 1

SH3 domain and tetratricopeptide repeat-containing protein 2 Gap junction beta-1 protein Transient receptor potential cation channel subfamily V member 4 Periaxin Serine palmitoyltransferase 1

Tyrosine–tRNA ligase, cytoplasmic

Myelin protein P0

Peripheral myelin protein 22

Protein name Heat shock protein beta-1

5q13.2

11q13.3

21q22.11 9q34.13 14q22.1 2p22.3 14q32.31 Xq12

6p12.1

9q31.3

19q13.2 9q22.31

Xq13.1 12q24.11

5q32

1p35.1

1q23.3

17p12

Location 7q11.23

68 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

XL AR AD XL AR XL AR AR AR AR AR AD AR AR

AR AR XL AD AR AR AR

310300 181350 158900 300376 607855 310200 253600 615356 236670 613153 253800 606072 254090 605809

255320 614399 310400 601419 256030 201450

Acyl-CoA dehydrogenase, very long-chain; ACADVL 609575

Muscular dystrophies Emery–Dreifuss muscular dystrophy 1, X-linked; EDMD1 Emery–Dreifuss muscular dystrophy 2, autosomal dominant; EDMD2 Facioscapulohumeral muscular dystrophy 1; FSHD1 Muscular dystrophy, Becker type; BMD Muscular dystrophy, congenital merosin-deficient, 1A; MDC1A Muscular dystrophy, Duchenne type; DMD Muscular dystrophy, limb-girdle, type 2A; LGMD2A Muscular dystrophy, limb-girdle, type 2S; LGMD2S Muscular dystrophy-dystroglycanopathy (congenital with brain and eye anomalies), type A, 1; MDDGA1 Muscular dystrophy-dystroglycanopathy (congenital with brain and eye anomalies), type A, 5; MDDGA5 Muscular dystrophy-dystroglycanopathy (congenital with brain and eye anomalies), type A, 4; MDDGA4 Rippling muscle disease 2; RMD2 Ullrich congenital muscular dystrophy 1; UCMD1 Myastenia Myasthenic syndrome, congenital, 4A, slow-channel; CMS4A Myopathies congenital Minicore myopathy with external ophthalmoplegia Myopathy, areflexia, respiratory distress, and dysphagia, early-onset; EMARDD Myopathy, centronuclear, X-linked; CNMX Myopathy, myofibrillar, 1; MFM1 Nemaline myopathy 2; NEM2 Myopathies metabolic Acyl-CoA dehydrogenase, medium-chain, deficiency ACADVL

ACADM

MTM1 DES NEB

RYR1 MEGF10

CHRNE

CAV3 Col6A2

FKTN

FKRP

DMD CAPN3 TRAPPC11 POMT1

FRG1 DMD LAMA2

LMNA

EMD

Medium-chain specific acyl-CoA dehydrogenase, mitochondrial Very long-chain specific acyl-CoA dehydrogenase, mitochondrial

Ryanodine receptor 1 Multiple epidermal growth factor-like domains protein 10 Myotubularin Desmin Nebulin

Acetylcholine receptor subunit epsilon

Caveolin-3 Collagen alpha-2(VI) chain

Fukutin

Fukutin-related protein

Dystrophin Calpain-3 Trafficking protein particle complex subunit 11 Protein O-mannosyl-transferase 1

Protein FRG1 Dystrophin Laminin subunit alpha-2

Prelamin-A/C

Emerin

(continued)

17p13.1

1p31.1

Xq28 2q35 2q23.3

19q13.2 5q23.2

17p13.2

3p25.3 21q22.3

9q31.2

19q13.32

Xp21.2-p21.1 15q15.1 4q35.1 9q34.13

4q35 Xp21.2-p21.1 6q22.33

1q22

Xq28

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M) 69

AR AD AD AD AD AR AR XL AD AD AD AD AD AR AR AR AR AR AR AR AR

268200 168300 160800 160900 602668 216900 262300 300500 203200 106210 180500 601777 607541 609218 607313 231300 267750 204100 604393 604537 613826 153700

Gaze palsy, familial horizontal, with progressive scoliosis; HGPPS Glaucoma 3, primary congenital, A; GLC3A Knobloch syndrome 1; KNO1 Leber congenital amaurosis 2; LCA2 Leber congenital amaurosis 4; LCA4 Leber congenital amaurosis 5; LCA5 Leber congenital amaurosis 5; LCA5 Leber congenital amaurosis 6; LCA6

Macular dystrophy, vitelliform, 2; VMD2

AD

Inheritance AR AR AR

Disease Carnitine palmitoyltransferase II deficiency, infantile Glycogen storage disease II; GSD2 Hydroxyacyl-CoA dehydrogenase/3-ketoacyl-CoA thiolase/enoyl-CoA hydratase, alpha subunit; HADHA Myoglobinuria, acute recurrent, autosomal recessive Myotonia Paramyotonia congenita of von Eulenburg; PMC Myotonic disorder Myotonia congenita, autosomal dominant Myotonic dystrophy 1; DM1 Myotonic dystrophy 2; DM2 Ophtalmoligic disorders Achromatopsia 2; ACHM2 Achromatopsia 3; ACHM3 Albinism, ocular, type I; OA1 Albinism, oculocutaneous, type II; OCA2 Aniridia; AN Axenfeld–Rieger syndrome, type 1; RIEG1 Cone-rod dystrophy 6; CORD6 Corneal dystrophy, Avellino type; CDA Foveal hypoplasia 2; FVH2

OMIM number 600649 232300 609016

Table 4.1 (continued)

BEST1

RPGRIP1

CYP1B1 COL18A1 RPE65 AIPL1 LCA5

ROBO3

CNGA3 CNGB3 GPR143 OCA2 PAX6 PITX2 GUCY2D TGFBI SLC38A8

CLCN1 DMPK CNBP

SCN4A

LPIN1

Gene name/ symbol CPT2 GAA HADHA

X-linked retinitis pigmentosa GTPase regulator-­ interacting protein 1 Bestrophin-1

Cytochrome P450 1B1 Collagen alpha-1(XVIII) chain Retinoid isomerohydrolase Aryl-hydrocarbon-interacting protein-like 1 Lebercilin

Cyclic nucleotide-gated cation channel alpha-3 Cyclic nucleotide-gated cation channel beta-3 G-protein-coupled receptor 143 P protein Paired box protein Pax-6 Pituitary homeobox 2 Retinal guanylyl cyclase 1 Transforming growth factor-beta-induced protein ig-h3 Putative sodium-coupled neutral amino acid transporter 8 Roundabout homolog 3

Chloride channel 1, skeletal muscle Myotonin-protein kinase Cellular nucleic acid-binding protein

Sodium channel protein type 4 subunit alpha

Phosphatidate phosphatase LPIN1

Protein name Carnitine O-palmitoyltransferase 2, mitochondrial Lysosomal alpha-glucosidase Trifunctional enzyme subunit alpha, mitochondrial

11q12.3

14q11.2

2p22.2 21q22.3 1p31.3 17p13.2 6q14.1

11q24.2

2q11.2 8q21.3 Xp22.3 15q12–q13 11p13 4q25 17p13.1 5q31.1 16q23.3

7q34 19q13.32 3q21.3

17q23.3

2p25.1

Location 1p32.3 17q25.3 2p23.3

70 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Schizophrenia Rap guanine nucleotide exchange factor 6; RAPGEF6 Renal disorders Alport syndrome, autosomal dominant Alport syndrome, X-linked; ATS Arthrogryposis, renal dysfunction, and cholestasis 1; ARCS1 Bartter syndrome, type 3; BARTS3 Cystinosis, nephropathic; CTNS Gitelman syndrome; GTLMNS Hyperuricemic nephropathy, familial juvenile, 1; HNFJ1 Hypomagnesemia 3, renal; HOMG3 Nephrogenic syndrome of inappropriate antidiuresis Nephrotic syndrome, type 1; NPHS1

Microphthalmia, isolated, with coloboma 3; MCOPCB3 Norrie disease; ND Optic atrophy 1; OPA1 Retinal dystrophy, early-onset, with or without pituitary dysfunction, included Retinitis pigmentosa 13; RP13 Retinitis pigmentosa 2; RP2 Retinitis pigmentosa 25; RP25 Retinitis pigmentosa 3; RP3 Retinitis pigmentosa 4; RP4 Retinoschisis 1, X-linked, juvenile; RS1 Stargardt disease 1; STGD1 Stickler syndrome, type I; STL1 Stickler syndrome, type II; STL2 Overgrowth syndromes Perlman syndrome; PRLMNS Sotos syndrome 1; SOTOS1

AD XL AR AR AR AR AD AR XL AR

607364 219800 263800 162000 248250 300539 256300

AR AD

267000 117550

104200 301050 208085

AD XL AR XL AD XL AR AD AD

600059 312600 602772 300029 613731 312700 248200 108300 604841

AD

XL AD AD

310600 165500 610125

610499

AR

610092

CLDN16 AVPR2 NPHS1

CLCNKB CTNS SLC12A3 UMOD

Col4A3 COL4A5 VPS33B

RAPGEF6

DIS3L2 NSD1

PRPF8 RP2 EYS RPGR RHO RS1 ABCA4 COL2A1 COL11A1

NDP OPA1 OTX2

VSX2

Claudin-16 Vasopressin V2 receptor Nephrin

Chloride channel protein ClC-Kb Cystinosin Solute carrier family 12 member 3 Uromodulin

Collagen alpha-3(IV) chain Collagen, type IV, alpha 5 Vacuolar protein sorting-associated protein 33B

Rap guanine nucleotide exchange factor 6

DIS3-like exonuclease 2 Histone-lysine N-methyltransferase, H3 lysine-36 and H4 lysine-20 specific

Pre-mRNA-processing-splicing factor 8 Protein XRP2 Protein eyes shut homolog X-linked retinitis pigmentosa GTPase regulator Rhodopsin Retinoschisin Retinal-specific ATP-binding cassette transporter Collagen alpha-1(II) chain Collagen alpha-1(XI) chain

Norrin Dynamin-like 120 kDa protein, mitochondrial Homeobox protein OTX2

Visual system homeobox 2

(continued)

3q28 Xq28 19q13.12

1p36.13 17p13.2 16q13 16p12.3

2q36.3 Xq22.3 15q26.1

5q31.1

2q37.1 5q35.3

17p13.3 Xp11.3 6q12 Xp11.4 3q22.1 Xp22.13 1p22.1 12q13.11 1p21.1

Xp11.3 3q29 14q22.3

14q24.3

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M) 71

Disease Nephrotic syndrome, type 2; NPHS2 Nephrotic syndrome, type 5 Polycystic kidney disease 1; PKD1 Polycystic kidney disease 2; PKD2 Polycystic kidney disease, autosomal recessive; ARPKD Renal cysts and diabetes syndrome; RCAD Renal cysts and diabetes syndrome; RCAD Renal tubular acidosis, distal, autosomal recessive; RTADR Renal tubular dysgenesis; RTD Respiratory disorders Alpha-1-antitrypsin deficiency; A1ATD Central hypoventilation syndrome, congenital; CCHS Ciliary dyskinesia, primary, 15; CILD15 Ciliary dyskinesia, primary, 3; CILD3 Cystic fibrosis; CF Surfactant metabolism dysfunction, pulmonary, 2; SMDP2 Surfactant metabolism dysfunction, pulmonary, 3; SMDP3 Skeletal disorders Achondroplasia; ACH Acromesomelic dysplasia, Maroteaux type; AMDM Arthrogryposis, distal, type 2A; DA2A Arthrogryposis, distal, type 2B; DA2B Arthrogryposis, distal, type 2B; DA2B Arthrogryposis, distal, type 3; DA3 Arthrogryposis, distal, type 9; DA9 Brachydactyly, type b1; BDB1 Campomelic dysplasia with autosomal sex reversal Camurati–Engelmann disease; CAEND

Table 4.1 (continued) Inheritance AR AR AD AD AR AD AD AR AR AR AD AR AR AR AD AR

AD AR AD AD AD AD AD AD AD AD

OMIM number 600995 609049 173900 613095 263200 137920 137920 602722 267430 613490 209880 613808 608644 219700 610913 610921

100800 602875 193700 601680 601680 114300 121050 113000 114290 131300 FGFR3 NPR2 MYH3 TNNT3 TNNI2 PIEZO2 FBN2 ROR2 SOX9 TGFB1

ABCA3

SERPINA1 PHOX2B CCDC40 DNAH5 CFTR SFTPC

ACE

HNF1B HNF1B ATP6V0A4

Gene name/ symbol NPHS2 LAMB2 PKD1 PKD2 PKHD1

Fibroblast growth factor receptor 3 Atrial natriuretic peptide receptor 2 Myosin-3 Troponin T, fast skeletal muscle Troponin I, fast skeletal muscle Piezo-type mechanosensitive ion channel component 2 Fibrillin-2 Tyrosine-protein kinase transmembrane receptor ROR2 Transcription factor SOX-9 Transforming growth factor beta-1

ATP-binding cassette subfamily A member 3

Alpha-1-antitrypsin Paired mesoderm homeobox protein 2B Coiled-coil domain-containing protein 40 Dynein heavy chain 5, axonemal Cystic fibrosis transmembrane conductance regulator Pulmonary surfactant-associated protein C

Angiotensin-converting enzyme

Hepatocyte nuclear factor 1-beta Hepatocyte nuclear factor 1-beta V-type proton ATPase 116 kDa subunit a isoform 4

Protein name Podocin Laminin subunit beta-2 Polycystin-1 Polycystin-2 Fibrocystin

4p16.3 9p13.3 17p13.1 11p15.5 11p15.5 18p11.22-p11.21 5q23.3 9q22.31 17q24.3 19q13.2

16p13.3

14q32.13 4p13 17q25.3 5p15.2 7q31.2 8p21.3

17q23.3

17q12 17q12 7q34

Location 1q25.2 3p21.31 16p13.3 4q22.1 6p12.3-p12.2

72 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

XL XL AD AD AD AD AR AR XL AD XL AD AR AR AR AR AR

AD AD AD AR AD XL AD AR

302950 303100 119600 133700 133701 175700 228600 241500 307800 150250 127300 156500 210710 604317 604804 608716 608393

161200 166200 166210 259440 610967 300373 166600 259700

Chondrodysplasia punctata 1, X-linked recessive; CDPX1 Choroideremia; CHM Cleidocranial dysplasia; CCD Exostoses, multiple, type I Exostoses, multiple, type II Greig cephalopolysyndactyly syndrome; GCPS Hyaline fibromatosis syndrome; HFS Hypophosphatasia, infantile Hypophosphatemic rickets, X-linked dominant Larsen syndrome; LRS Leri–Weill dyschondrosteosis; LWD Metaphyseal chondrodysplasia, Schmid type; MCDS Microcephalic osteodysplastic primordial dwarfism, type I; MOPD1 Microcephaly 2, primary, autosomal recessive; MCPH2 Microcephaly 3, primary, autosomal recessive; MCPH3 Microcephaly 5, primary, autosomal recessive; MCPH5 Microcephaly 6, primary, MCPH6 Microcephaly 6, primary, autosomal recessive; MCPH6 Nail–patella syndrome; NPS Osteogenesis imperfecta, type I; OI1 Osteogenesis imperfecta, type II; OI2 Osteogenesis imperfecta, type IX; OI9 Osteogenesis imperfecta, type V; OI5 Osteopathia striata with cranial sclerosis; OSCS Osteopetrosis, autosomal dominant 2; OPTA2 Osteopetrosis, autosomal recessive 1; OPTB1

AR

250250

Cartilage–hair hypoplasia; CHH

LMX1B COL1A1 COL1A2 PPIB IFITM5 AMER1 CLCN7 TCIRG1

CENPJ

ASPM

CDK5RAP2

WDR62

CHM RUNX2 EXT1 EXT2 GLI3 ANTXR2 ALPL PHEX FLNB SHOX COL10A1 RNU4ATAC

ARSE

RMRP

9q33.2

19q13.12

Xq21.2 6p21.1 8q24.11 11p11.2 7p14.1 4q21.21 1p36.12 Xp22.11 3p14.3 Xp22.33;Yp11.3 6q22.1 2q14.2

Xp22.33

9p13.3

LIM homeobox transcription factor 1-beta Collagen alpha-1(I) chain Collagen alpha-2(I) chain Peptidylprolyl isomerase B (cyclophilin B) Interferon-induced transmembrane protein 5 APC membrane recruitment protein 1 H(+)/Cl(−) exchange transporter 7 V-type proton ATPase 116 kDa subunit a isoform 3

Centromere protein J

(continued)

9q33.3 17q21.33 7q21.3 15q22.31 11p15.5 Xq11.2 16p13.3 11q13.2

13q12.12

Abnormal spindle-like microcephaly-associated protein 1q31.3

CDK5 regulatory subunit associated protein 2

WD repeat-containing protein 62

Rab proteins geranylgeranyltransferase component A 1 Runt-related transcription factor 2 Exostosin-1 Exostosin-2 Transcriptional activator GLI3 Anthrax toxin receptor 2 Alkaline phosphatase, tissue-nonspecific isozyme Phosphate-regulating neutral endopeptidase Filamin-B Short stature homeobox protein Collagen alpha-1(X) chain RNA, U4atac small nuclear (U12-dependent splicing)

RNA component of mitochondrial RNA processing endoribonuclease Arylsulfatase E

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M) 73

Disease Pseudoachondroplasia; PSACH Seckel syndrome 1; SCKL1 Short stature, developmental delay, and congenital heart defects; SDDHD Short-rib thoracic dysplasia 10 Short-rib thoracic dysplasia 10 with or without polydactyly; SRTD10 Short-rib thoracic dysplasia 3 with or without polydactyly; SRTD3 Spondyloepimetaphyseal dysplasia, Missouri type Spondyloepiphyseal dysplasia tarda, X-linked; SEDT Symphalangism, proximal; SYM1

Table 4.1 (continued) Inheritance AD AR AR AR

AR AR XL AD

OMIM number 177170 210600 617044 615630

613091 602111 313400 185800 MMP13 TRAPPC2 NOG

DYNC2H1

IFT172

Gene name/ symbol COMP ATR TKT

Collagenase 3 Trafficking protein particle complex subunit 2 Noggin

Cytoplasmic dynein 2 heavy chain 1

Intraflagellar transport protein 172 homolog

Protein name Cartilage oligomeric matrix protein Serine/threonine-protein kinase ATR Transketolase

11q22.3 Xp22.2 17q22

11q22.3

2p23.3

Location 19p13.11 3q23 3p21.1

74 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

4.1 Traditional Indications and Strategies for PGT-M

75

Table 4.2  Clinical outcome of 6204 PGT-M cycles for Mendelian disorders Type of cells tested Polar bodies

# of patients 144

# of cycles 269

# of transfers 202

Polar bodies + blastomere Blastomere

449

747

636

1125

2036

1616

Blastocyst

2040

3152

2176

Total

3758

6204

4630

PGT-M cycles, which is the world’s largest experience in one center. Over a thousand of these PGT-M (1016 cycles) were performed by polar body (PB) analysis, involving the retrieval and testing of more than 10,000 oocytes, with results of sequential PB1 and PB2 testing obtained in 97.6% of these oocytes. This has enabled to preselection for transfer of 1656 embryos originating from these oocytes in 838 (82.5%) of initiated cycles, of which 385 healthy children have resulted. With introduction of blastocyst biopsy, the majority of PGT-M was shifted to embryo biopsy, which has been applied in 5188 cases (3152 by blastocyst and 2036 by blastomere biopsy), resulting in transfer of unaffected embryos in 3792 cycles and yielding 2098 (55.3%) unaffected clinical pregnancies (1365 after blastocyst and 733 after blastomere biopsy) and birth of 2132 disease-free healthy children. Overall, in 6204 PGT-M cycles, performed for 3758 patients at risk, 7061 unaffected embryos were transferred in 4630 cycles (approximately, 1.52 embryos per cycle), resulting in 2447 unaffected pregnancies (52.8% pregnancy rate per transfer) with birth of 2517 apparently healthy children (Table 4.2).

4.1

Traditional Indications and Strategies for PGT-M

4.1.1 Autosomal Recessive Disorders In our experience, more than half of the PGT cycles were performed for autosomal recessive conditions. As mentioned, initially, the most

# of embryos transferred 398 1.97 1258 1.97 2902 1.79 2503 1.15 7061 1.52

Pregnancy 80 39.6% 269 42.3% 733 45.3% 1365 62.7% 2447 52.8%

SAB 18 22.5% 40 14.9% 106 14.5% 105 7.7% 269 11%

# of deliveries 62 77.5% 229 85.1% 627 85.5% 1260 92.3 2178 89%

# of babies 77 308 756 1376 2517

common indications for PGT-M were CFTR and hemoglobin disorders (HBB), performed for an increasing number of mutations. Hemoglobinopathies represent one of the largest groups for which PGT-M was performed (Tables 4.3, 4.4, and 4.5) in our experience. In some communities, such as in Cyprus, Greece, and Turkey, PGT-M is becoming a routine procedure for couples carrying thalassemia mutations who cannot accept prenatal diagnosis and termination of pregnancy [9–11]. Introduced for the first time in 1996 in Cyprus, only 40 PGT cycles for hemoglobin disorders were performed before the year 2000. Hundreds of cycles have been performed since. At present, the proportion of PGT cases for hemoglobin disorders in our overall PGT-M experience is just over 10% (684 of 6204 PGT-M cycles). To improve accuracy of PGT for hemoglobin disorders, a set of polymorphic markers, listed in Fig.  4.1, were used, which makes it realistic to select at least three closely linked informative markers in any case performed to analyze simultaneously with mutation testing. As mentioned, a total of 684 PGT cycles were performed for 410 couples at risk of bearing children with ­hemoglobin (HB) disorders. This included also 23 PGT cycles for alpha-thalassemia and over hundred cycles for sickle cell disease. A total of 188 of these cycles were performed in combination with HLA typing to select unaffected embryos as potential donors for stem cell transplantation (see Chap. 5). Of 684 clinical cycles performed, unaffected embryos for transfer were available in 536 (78.2%), resulting in 222 (41.4%) clinical

HBA1, HBA2

604131 410

14 684

23

Anemias (lack of red blood cells or hemoglobin) and hemolytic anemias (destruction of red blood cells) Anemia, nonspherocytic hemolytic, due to G6PD G6PD 300908 9 11 deficiency Pyruvate kinase deficiency of red cells + HLA PKLR 266200 1 2 Rhesus blood group RHD 111680 9 11 Blood group Kell–Cellano system KEL 110900 14 30 Porphyria, congenital erythropoietic UROS 263700 1 1 Subtotal 34 55 Aplastic anemias Shwachman–Diamond syndrome; Sds SBDS 4 10 10 17 105650 RPS19 Diamond–Blackfan anemia 1; DBA1 606129 RPS20 Diamond–Blackfan anemia 2; DBA2 610629 RPS24 Diamond–Blackfan anemia 3; DBA3 612528 RPL35A Diamond–Blackfan anemia 5; DBA5 613308 RPS10 Diamond–Blackfan anemia 9; DBA9 30 84 227650 Fanconi anemia, complementation group A; FANCA FANCA 227645 Fanconi anemia, complementation group C; FANCC FANCC 227646 FANCD2 Fanconi anemia, complementation group D2; 603467 FANCF FANCD 609053 Fanconi anemia, complementation group F; FANCF FANCI 609054 BRIP1 Fanconi anemia, complementation group I; FANCI Fanconi anemia, complementation group J; FANCJ Subtotal 44 111

Subtotal

Hemoglobin–alpha locus 1; HBA1

12 1 16 29 1 59 4 20

72

96

1 9 17 1 37 3 14

46

63

978 1.79

31

21

2 8

0 6 4 1 17

6

244 44.6%

4

4

0 0

0 0 0 0 0

0

41 16.8%

41 17.5% 0

940 1.79 38

234 44.5% 10

SAB

# of embryos transferred Pregnancy

9

546

21

# of # of # of Disease Gene OMIM patients cycles transfers Hemoglobinopathies (congenital abnormality of the hemoglobin molecule or of the rate of hemoglobin synthesis) Hemoglobin–beta locus; HBB HBB 141900 396 661 525

Table 4.3  Hematologic disorder and coagulopathy

27

17

2 8

0 6 4 1 17

6

203 83.3%

193 82.5% 10

33

23

2 8

0 7 5 1 19

6

240

10

230

Delivery Birth

76 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Coagulopathies (disorders of bleeding and coagulation) Bleeding disorder, platelet-type, 16; BDPLT16 ITGB3 Amegakaryocytic thrombocytopenia, congenital; MPL CAMT Prothrombin deficiency, congenital F2 Factor V deficiency F5 Factor VII deficiency F7 Hemophilia A F8 Hemophilia B F9 Thrombasthenia of Glanzmann ITGA2B Thrombotic Thrombocytopenic purpura, congenital; ADAMTS13 TTP Wiskott–Aldrich syndrome; WAS WAS Thrombocythemia 1; THCYT1 SH2B3 Thrombocytopenia-absent radius syndrome; TAR RBM8A Thrombophilia due to protein S deficiency, autosomal PROS1 dominant; THPH5 Subtotal Total 15 2 6 1 146 973

6 1 4 1 89 577

301000 187950 274000 612336

2 1 1 103 9 2 2

2 1 1 62 7 1 1

613679 227400 227500 306700 306900 273800 274150

1 1

1 1

187800 604498

132 757

11 2 6 0

3 2 1 95 7 2 2

0 1

190 1285 1.69

18 2 6 0

3 2 1 138 11 4 4

0 1

76 358 47.3%

9 2 4 0

2 1 1 48 6 1 1

0 1

11 56 15.6%

1 1 0 0

0 0 1 7 0 1 0

0 0

65 302 84.4%

8 1 4 0

2 1 0 41 6 0 1

0 1

68 350

8 1 4 0

2 1 0 44 6 0 1

0 1

4.1 Traditional Indications and Strategies for PGT-M 77

78

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Table 4.4  List of thalassemia mutation for which PGT was performed in our experience

Legacy name Beta thalassemia Transcriptional mutations nt -90 C>T nt -88 C>T nt -87 C>G nt -42 C>G nt -32 C>T nt -31 A>C nt -30 T>A nt -29 A>G nt -28 A>G CAP +1 (A->C) silent nt 1 A>C Pro5Ala Pro5Ser Glu6Lys Glu6Val Leu11Pro Arg12Ser Trp15Stop Lys18Stop Glu26Lys Ala27Ser Gln39Stop Glu121Ala Glu121Val Poly A (A->G) Splicing mutations IVS-I-1 (G->T) IVS-I-5 (G->C) IVS-I-6 (T->C) IVS-I-110 (G->A) IVS-I-116 (T->G) IVS-II-1 (G->A) IVS-II-654 (C->T) IVS-II-745 (C->G) IVS-II-848 (C->A) Insertions/deletions Codons 7/8 (+G) Codons 27/28 (+C) 45 kb deletion; the Filipino deletion beta0 Codon 5 (-CT) Codon 6 (-A); Codon 8 (-AA) Codons 36/37 (-T) Codons 41/42 (-TTCT) Codon 44 (-C) Codon 76 (-C) 619 bp deletion beta0 Hb Lepore Hollandia Sicilian (deltabeta)0-Thal

HGVS nomenclature

HBB:c.-140C>T HBB:c.-138C>T HBB:c.-137C>G HBB:c.-92C>G HBB:c.-82C>T HBB:c.-81A>C HBB:c.-80T>A HBB:c.-79A>G HBB:c.-78A>G HBB:c.-50A>C HBB:c.16C>G HBB:c.16C>T HBB:c.19G>A HBB:c.20A>T HBB:c.44T>C HBB:c.93G>C HBB:c.47G>A HBB:c.52A>T HBB:c.79G>A HBB:c.82G>T HBB:c.118C>T HBB:c.365A>C HBB:c.365A>T c.∗113A-G HBB:c.92+1G>T HBB:c.92+5G>C HBB:c.92+6T>C HBB:c.93-21G>A HBB:c.93-15T>G HBB:c.315+1G>A HBB:c.316-197C>T HBB:c.316-106C>G HBB:c.316-3C>A HBB:c.24_25insG HBB:c.84_85insC NG_000007.3:g.66258_184734 del118477 HBB:c.17_18delCT HBB:c.20delA HBB:c.25_26delAA HBB:c.112delT HBB:c.126_129delCTTT HBB:c.135delC HBB:c.230delC NG_000007.3:g.71609_72227del619 NG_000007.3:g.63290_70702del NG_000007.3:g.64336_77738 del13403

4.1 Traditional Indications and Strategies for PGT-M Table 4.4 (continued)

79

Legacy name Chinese Ggamma(Agammadeltabeta)0-Thal Alpha-thalassemia (MC); a deletion of at least 46 kb involving both alpha genes and zeta gene alpha-Thal-1 (THAI); a deletion of 34–38 kb involving the alpha1, alpha2, and zeta genes alpha-Thal-1 (FIL); a deletion of 30–34 kb involving the alpha1, alpha2, and zeta genes alpha-Thal-1 (MED-I); deletion of ~17.5 kb including both alpha-globin genes alpha-Thal-1 (SEA); deletion of ~20 kb including both alpha-globin genes alpha-Thal-1

HGVS nomenclature NG_000007.3:g.48795_127698 del78904 NG_000006.1:g.164_43364del43201

NG_000006.1:g.10664_44164 del33501 NG_000006.1:g.11684_43534 del31851 NG_000006.1:g.24664_41064 del16401 NG_000006.1:g.26264_45564 del19301

Table 4.5  Clinical outcome PGT for hemoglobinopathies Mutation (HGVS # of nomenclature/legacy name) patients c.-140C>T (nt -90 C>T) 1 c.-138C>T (nt -88 C>T) 1 -81A>C (nt -31 A>C) 1 c.-79A>G (nt -29 A>G 2 c.-78A>G (nt -28 A>G) 1 c.-50A>C (CAP +1) 2 HBC c.19G>A (Glu6Lys) 7 HbS c.20A>T (Glu6Val) 148 c.47G>A (Trp15Stop) 1 c.52A>T (Lys18Stop) 1 HBE c.79G>A (Glu26Lys) 5 c.92G>C (Arg30Thr) 1 c.118C>T (Gln39Stop) 26 c.208G>A Gly69Ser 1 c.365A>T (Glu121Val) 1 2 c.∗113A-G (Poly A) c.92+1G>T (IVS-I-1) 11 c.92+5G>C (IVS-I-5) 24 c.92+6T>C (IVS-I-6) 18 c.93-21G>A (IVS1-110) 75 c.93-15T>G (IVS1-116) 1 c.315+1G>A (IVS-II-1) 5 c.316-197C>T (IVS-II-654) 3 c.316-106C>G (IVS-II-745) 4 c.316-3C>A (IVS-II-848) 1 c.17_18delCT (Codon 5-CT) 2 c.20delA (Codon 6-A) 2 c.25_26delAA (Codon 8-AA) 10 c.84_85insC (Codons 27/28 7 +C) c.112delT (Codons 36/37-T) 1

# of cycles 1 2 2 3 1 3 9 227 1 1 9 12 39 1 2 2 17 37 31 140 1 14 4 10 1 3 4 19 8 1

# of transfers 1 2 2 1 1 2 7 183 2 1 7 8 27 1 2 2 13 25 25 115 1 9 4 7 2 3 4 16 7 0

# of embryos transferred 3 4 4 2 2 2 11 322 1 1 13 13 43 2 4 2 23 36 49 234 3 15 8 12 2 6 6 27 10 0

Pregnancy SAB 0 0 0 0 1 0 1 0 0 0 0 0 4 1 101 16 1 0 1 0 5 3 0 0 17 1 1 1 0 0 1 0 4 0 13 3 12 3 35 8 0 0 2 0 2 0 1 1 1 0 1 0 3 0 7 0 5 0 0

0

Delivery Birth 0 0 0 0 1 1 1 1 0 2 0 0 3 3 85 97 1 1 1 1 2 2 0 0 16 18 0 0 0 0 1 1 4 6 10 10 9 14 27 34 0 0 2 3 2 3 0 0 1 1 1 1 3 3 7 7 5 5 0

0 (continued)

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

80

Table 4.5 (continued) Mutation (HGVS nomenclature/legacy name) c.126_129delCTTT (Codons 41/42-CTTT) c.135delC (Codon 44-C) IVS1-25bp DELETION The Filipino deletion beta0 (45 kb deletion) 619 bp deletion beta0 Chinese Ggamma(Agammadelt abeta) 0-Thal (30 kb) 13.4 kb deletion HBB Exon 1–3 deletion Sicilian (delta beta)0-Thal Hb Lepore Subtotal beta thalassemia Alpha-thalassemia Total

# of patients 8

# of cycles 12

# of transfers 6

# of embryos transferred 9

Pregnancy SAB 2 1

Delivery Birth 1 1

1 1 1

1 1 1

1 1 0

2 2 0

1 1 0

0 0 0

1 1 0

2 1 0

11 1

25 2

22 2

35 4

5 0

1 0

4 0

6 0

1 3 3 1 396 14 410

5 3 5 1 661 23 684

5 2 5 1 525 21 546

11 2 12 3 940 38 978 1.79

2 1 2 1 234 10 244 44.6%

1 1 0 1 1 1 0 1 41 193 0 10 41 203 16.8% 83.3%

1 1 2 2 230 10 240

−87 −42 −31 −29 −28 (C-G) (C-G) (A-G) (A-G) (A-G)

a

IVS I-110 IVSII-745 IVS I-6 Cd 41/42 619 bp DEL IVSII-645 IVS I-5 Cd 39 IVSII-1 R30T IVS I-1

Cd 8/9 Cd 8 E6V CAP

HBE1

(TA)n

HBG2

(CA)n

HBG1

HBD

EXON 1

EXON 2

IVS 1

Sfa NI

IVS 2

Bam HI Ava II

Hinf I

EXON 3

Hinf I

D11S1338 (CA)n

D11S1241 (CA)n

D11S1323 (CA)n

D11S1997 (GATA)n

(TG)n(CG)n

Barn HI Hpa I Hinf I (ATTTT)n

b

Rsa I Hinf I (AT)xTy

Polymorphic markers List of mutations: RNA Processing mutations:

Transcriptional mutations:

IVSI−1 (G−A) IVSI−1 (G−T) IVSI−5 (G−C)

−87 (C−G) −42 (C−G) −31 (A−G)

IVSI−5 (G−T) IVSI−5 (G−A) IVSI−6 (T−C) IVSI−110 (G−A) IVS2−1 (G−A) IVS2−654 (C−T) IVS2−745 (C−G)

−29 (A−G) −28 (A−G) E 6V (Sickle cell anemia) R30T Deletion: 619 bp

Fig. 4.1  Mutations in beta-globin gene for which PGT was performed and polymorphic markers used in multiplex PCR analysis. Map of human beta-globin gene,

Nonfunctional mRNA: Codon 8 (−AA) Codon 39 (C−T) Codon 41/42 (−CTTT)

Cap site: +1 (A−C) +2 (T−C)

showing sites and location of mutations (a), and linked polymorphic markers used for avoiding misdiagnosis (b). List of mutations is also presented in the lower panel

4.1 Traditional Indications and Strategies for PGT-M

p­ regnancies and the birth of 221 healthy children. Because the majority of thalassemia cases, as mentioned, were done for Eastern Mediterranean patients, approximately, over a half of them were performed for IVSI-110 mutation, which is the most common thalassemia mutation in the Mediterranean region (Table 4.5). While PGT-M cycles were mainly performed for heterozygous carriers, eight cycles were done for couples with homozygous or compound heterozygous male or female patients at 50% risk of bearing an affected offspring. In these couples, PGT involved testing for either three different mutations or, in the majority of cases, two different mutations or the same maternal and paternal mutation. Analysis of these mutations was done either simultaneously or in sequence by testing the maternal mutation in PB1 and PB2 and the paternal one by embryo biopsy (see details in corresponding section on PGT for affected homozygote or compound heterozygous patients). While the majority of PGT-M cases were performed for beta-globin gene mutations, as mentioned, 23 cycles were performed for α-thalassemia mutations, which resulted in preselection and transfer of 38 unaffected embryos in 10 cycles, yielding 8 clinical pregnancies and birth of 10 unaffected children, confirmed to be free from hydrops fetalis. To avoid misdiagnosis, the haplotype analysis for five polymorphic markers was performed, with confirmatory testing on the non-transferred embryos showing a correct diagnosis. An example of PGT for α-thalassemia is presented in Fig.  4.2. As seen from this case, both parents were carriers of this mutation, with the father having also hemoglobin H disease. The couple had one previous pregnancy resulting in spontaneous abortion, caused by hydrops fetalis. To avoid misdiagnosis the haplotype analysis with at least five polymorphic markers was performed. Of 15 embryos tested, 8 were affected, with the remaining 7 carrying 1 copy of the deleted α-globin gene, of which 2 (embryos #7 and #15) were transferred, with the mutant embryos confirmed to be affected, showing the reliability of the approach.

81

Cystic Fibrosis (CFTR)  has been one of the two major PGT-M indications from the very beginning of PGT-M application [12–15], with 724 cycles in our experience (Table  4.6), with testing for more than 2 dozen different mutations in the CFTR gene (see below). PGT for CFTR mutations is usually performed simultaneously with at least three strongly linked polymorphic markers, which may realistically be selected from a set of available markers listed in Fig. 4.3. In close to 90% of our cases of PGT for CFTR (in 638 of 724 cycles), unaffected embryos were selected for transfer, resulting in clinical pregnancies and birth of 360 unaffected children. Because of the high prevalence of CFTR mutations, there has been a need to test simultaneously for two or three CFTR mutations in the same reaction, as presented in Fig. 4.4. As can be seen from the pedigree, PGT-M for this couple seems to be the only choice, as the father is a compound heterozygote for DF508 and R117H and the mother is a carrier of the D1507 mutation in the CFTR gene. To avoid testing for three different mutations in the same reaction from the embryo biopsy, taking into consideration a high risk for ADO for each of the three alleles that may lead to a potential misdiagnosis, PB1 and PB2 testing was also performed to ensure the accuracy of the preselection of the mutation-free oocytes. As can be seen from Fig. 4.4, testing for DI507 maternal mutation, simultaneously with four closely linked markers, allowed the identification of three mutation-free oocytes from the six oocytes available for testing. The two embryos resulting from oocytes #1 and #9 had acceptable development potential and were transferred, yielding a singleton pregnancy and the birth of a healthy boy confirmed to be an unaffected carrier of the paternal mutation. In increasing number of cases, PGT for CFTR is performed simultaneously not only for two or three CFTR mutations, such as in PGT for CF patients, but also for a few other conditions in the same couples, as will be described below. Although the majority of autosomal recessive disorders are much rarer than those presented above (listed in Table  4.1), overall, they have

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

82

a

D16S521 (CA)n

REPEAT (A3T)n(N26)26(AC)n

D16S3399 (CA)n

D16S3024 (CA)n

5′HVR

D16S423 (CA)n

3′HVR ψζ1

ζ2

ψα2

b α3+α4 = NORMAL 230BP

ψα1

α2

α6

α3+C = DEL 124BP α7+α8 = NORMAL 160BP α5+α6 = NORMAL 316BP

C C1

α8

α7

α4

α3

θ

α1

α5

α5

α1

c

D16S3134 (CA)n

45Kb SEA (-- /--)

Markers order: D16s 521 D16s3399 HBA D16S3024 D16S3134 D16S423

143 157

143 143

172 174

172 170

140 140

146 142

-- αα -Hb H

147 150 153 142

-- / αα

150 147 144 140

Haplotype is based on PB1, PB2 and Blastomere results

Haplotype is based on Sperm and Blastomere result SAB PGT

d Embryo # 1

2

3

4

6

7

8

9

11

13

14

15

16

17

18

143 143

143

143 143

143 143

143 143

143 143

143

143

157 143

143

143

143 143

143

157 143

143

172 170

172

172 170

172 170

172 170

172 170

172

172

174 172

172

172

172 170

172

174 170

172

--

--

--

--

--

--

α- --

--

--

--

α- αα

--

--

αα

--

αα

αα

αα

αα

--

αα

140 142

146

140 142

140 142

140 142

140 142

146

146

140 146

146

146

140 142

146

140 142

146

147 147

150

147 147

147 147

147 147

147 147

150

150

150 150

150

150

147 147

150

150 147

150

153 140

144

153 140

153 140

153 140

153 140

144

144

142 144

144

144

153 140

144

142 140

144

/ --

/ --

α-/--

/ --

Predicted Genotype

/ -ET

Fig. 4.2  PGT for alpha thalassemia. (a) Map of human alpha-globin gene, showing the position of 45 Kb SEA deletion and polymorphic markers used in multiplex PCR analysis; (b) size of fragments; (c) family pedigree showing both parents carrying SEA deletion, the father also having HbH disease; parental haplotypes are also shown,

become an important and established PGT-M indications in genetic practices. Practical implication of PGT for these rare recessive disorders is demonstrated below on the example of PGT for severe neurodegenerative conditions such as spinal muscular atrophy (SMA) and hereditary ­sensory autonomic neuropathy, also known as familial dysautonomia (FD).

/ --

/ --

/ --

ET

with paternal haplotypes obtained from sperm testing, and maternal haplotypes obtained from PB1 and PB2 analysis; (d) results of testing of 15 embryos showing the presence of six heterozygous embryos for deletion, of which two were transferred back to the patients

Familial Dysautonomia (FD)  is associated with the mutation affecting the donor splice site of intron 20 of the IKBKAP gene, assigned to chromosome 9q31 [16, 17]. It is the most common congenital sensory neuropathy, present in 1/3600 live births in Ashkenazi Jews. Characteristic features include absence of fungiform papillae on the tongue, absence of flare

219700 483 608644 2 613490 9 613808 1 610921 1 610913

CFTR DNAH5 SERPINA1 CCDC40 ABCA3 SFTPC

Cystic fibrosis; CF Total Ciliary dyskinesia, primary, 3; CILD3 Alpha-1-antitrypsin deficiency; A1ATD Ciliary dyskinesia, primary, 15; CILD15 Surfactant metabolism dysfunction, pulmonary, 3; SMDP3 Surfactant metabolism dysfunction, pulmonary, 2; SMDP2 Total 3 748

497

724 2 16 1 2

347

# of cycles 377

1

219700 232

CFTR

Cystic fibrosis; CF + PGT-A

# of OMIM patients 219700 251

Gene CFTR

Disease Cystic fibrosis; CF

Table 4.6  PGT-M for pulmonary disorders

661

3

638 2 14 1 3

285

# of transfers 353

1078 1.6

3

# of embryos transferred 635 (1.8) 415 (1.4) 1050 2 18 1 4

356 54%

1

Pregnancy 181 51% 163 57% 344 1 8 1 1

1

# of deliveries 154 85% 153 93.9% 307 1 7 1 1

38 318 10.7% 89.3%

0

SAB 27 15% 10 6.1% 37 0 1 0 0

371

1

360 1 7 1 1

168

# of babies 192

4.1 Traditional Indications and Strategies for PGT-M 83

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

84

a

R75XH C–T

EXON 3

b

R117H G–A

A120T G–A

EXON 4

I148T T–C

A349V C–T

621+1 G–T

Intron 4

1259 Ins A

EXON 7

IVS 1

IVS 6a

(CA)n

(GATT)n

Delta I 507

Delta F 508

1717-1 G–A

R553X C–T

EXON 14a

(CA)n (CA)n Poly T 1540 A/G tract Mnl I

Fig. 4.3  Mutations in CFTR gene, for which PGT was performed and polymorphic markers used in multiplex PCR analysis. Map of CFTR gene, showing sites and location of mutations (a), and polymorphic markers used

G551D G–A

R560T G–C

EXON 11

EXON 10

IVS 8

G550X G–T

Intron 10

EXON 10

EXON 8

G542X G–T

2694 T/G Ava II

IVS 17b

R1162X C–T

W1282X G–A

N1303K C–G

EXON 19

EXON 20

EXON 21

IVS 18

(TA)n (CA)n TUB 18 C/A Hinf I

IVS 20

TUB 20 A/G Pvu II

for avoiding misdiagnosis (b). (a) Upper level – mutations in CFTR gene. (b) Bottom level – polymorphic markers in CFTR gene

Family Pedigree

N 98 +

1.2

1.1 DF508 / R117H

N /DI507

Markers order: Exon 10(dI507) Intron 17 Intron 18 Intron 20

dI507 106 + -

PGT 2.1 PGT by Sequential Polar Body analysis

N 98 + Oocyte #

N 98 +

dI507 106 + -

dI507 106 + -

dI507 106 + -

N 98 +

1

2

4

7

8

9

N

N

I507

I507

I507

N

ET

ET

Fig. 4.4  PGT for a couple with three different mutations in CFTR gene. (Top) The mother (1.2) is a carrier of the delta I507 mutation in CFTR gene. The father (1.1) is affected with CF and had delta F508 and R117H mutations in CFTR gene. (Bottom) PGT was performed by sequential PB1 and PB2 analysis. The mother is informa-

tive for three inside CFTR gene polymorphic markers (introns 17, 18, and 20). Multiplex heminested PCR of PB1 and PB2 revealed three normal (#1, #2, and #9) and three affected oocytes (#4, #7, and #8). Embryos #1 and #9 were transferred and a healthy boy was born

after injection of intradermal histamine, decreased or absent deep tendon reflexes, and absence of overflow of emotional tears. This devastating and debilitating disorder is further characterized by the poor development and pro-

gressive degeneration of the sensory and autonomous nervous system, gastrointestinal and respiratory dysfunctions, vomiting crisis, excessive sweating, and postural hypotension. Despite a remarkable variability of the disease pheno-

4.1 Traditional Indications and Strategies for PGT-M

type within and between f­amilies, expected to reflect different mutations producing inactivation of the gene, a single major mutation predominates [18]. A single T  →  C change at the base pair 6 of the splice donor site, responsible for 99% of cases of FD, results in the skipping of exon 20 (74 bp) from the IKBKAP mRNA only in the brain, with varying level of the gene expression in other tissues. This may explain the severe progressive degeneration of the ­sensory and autonomous nervous system, leading to continued neuronal depletion with age and early death. The product of the IKBKAP gene is a part of a multiprotein complex, hypothesized to play a role in general transcriptional regulation; thus, the complete inactivation of the gene might cause a lethal phenotype at any stage of embryonic development [18]. Variability of the disease phenotype may be explained by the presence of a partially functional gene product in some tissues, including the brain, given even a small

85

amount of the encoded protein expressed at critical developmental stages permitting sufficient neuronal survival. In addition, a very rare minor FD missense mutation has been described (G → C change at base pair 17 in exon 19 of the gene) to be associated with a mild phenotype in patients with heterozygous status for the major mutation [18]. Despite progress in understanding of the nature and pathogenesis of the disease, FD is still fatal, with no effective management available at the present time, making PGT-M a useful option for those at-risk couples that cannot accept prenatal diagnosis and termination of pregnancy as an option for avoiding FD in their offspring. One of such couples presented for PGT-M due to a previous child with FD.  Both parents were of Ashkenazi Jewish descent and, as seen from the pedigree (Fig.  4.5), are carriers of the gene for FD based on marker analysis available for all members of the extended family [19].

Family Pedigree

Markers order: Major mutation (intron 20) D9S1677 D9S58

N M 135 135 151 116

M M 140 135 116 116

Fig. 4.5  Family pedigree of the couple undergoing PGT for familial dysautonomia (FD). The father is a carrier of mutation of the donor splice site of intron 20 of the IKBKAP gene, which is linked to 135  bp repeat of D9S1677, and 116 bp repeat of D9S58, while the normal allele is linked to 135 and 151 bp repeats of the same polymorphic markers respectively. The mother is also a carrier of the same mutation, linked to 140 and 116 bp repeats,

N M 126 140 99 116

PGT

PGT

N N 135 126 151 99

N N 135 126 151 99

PGT M N 135 126 116 99

the normal gene being linked to 126 and 99  bp repeats respectively. As seen from this panel, the paternal and maternal sisters and maternal brother are also carriers of the mutation, which was inherited from the paternal father and maternal mother, respectively. Reproductive outcomes of this couple, including one previous affected child with FD and unaffected triplets born following PGD, are shown at the bottom

86

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

PGT-M cycle was performed using a sequential PB1 and PB2 biopsy and multiplex nested PCR analysis, involving mutation testing simultaneously with different linked markers. The mutation analysis involved detection of T to C change in the donor splice site of intron 20, based on HhaI restriction digestion that does not cut the normal allele, thus creating two fragments of 60 and 94 bp in the mutant allele (Fig. 4.6). As described above, there may be three genetic possibilities for the PB1 genotype from a heterozygous mother. If no crossover occurs, PB1 will be homozygous (either normal or mutant), but in the event of a crossover, PB1 will

be heterozygous. If crossover does not occur and the PB1 is homozygous for the mutant gene, the oocyte must contain two copies of the normal gene; thus, any embryo resulting from this oocyte can be transferred. This was not the case in any of the oocytes shown in Fig.  4.6. If the PB1 is homozygous for the normal gene, the maternal contribution to the embryo must be the mutant gene, which was also not the case as seen from Fig.  4.6. In both of these occasions, the extruded PB2 will have identical genotype to oocyte (opposite to genotype of PB1). In the event of crossover, PB1 is heterozygous, and the analysis of PB2 is required to predict which maternal alleles have been extruded with PB2

Donor splice site T C

D9S1677 (CA)n

a

D9S58 (CA)n

Intron 20

Hha I GCG’C

b

60

Restriction map:

94

Mutant Normal

154

Polar body

Polar body

1 2 1 2 1 2 1 2 1 2 1 2U

1

2 1 2 1 2 1 2 1 2 P Ma U

154 Normal

c

94 Mutant L

Oocyte # 1 3 N N ET ET

60 Mutant

L

6 N ET

7 N

8 M

9 N

Fig. 4.6  Preimplantation genetic testing for major mutation in IKBKAP gene, causing FD. (a) Position of major splice donor mutation T– C in IKBKAP gene and linked markers. (b) Restriction map. Major mutation creates restriction site for Hha I enzyme. (c) PB analysis of normal and mutant sequences of IKBKAP gene. Of 11

10 M

11 N

12 N

13 M

15 M

oocytes tested, seven were mutation-free based on heterozygous PB1 and affected PB2, of which oocytes #1, #3, and #6 were transferred resulting in unaffected triplets. L 100 bp ladder, N normal, M mutant, ET embryo transfer, U undigested PCR product, Ma maternal, P paternal

4.1 Traditional Indications and Strategies for PGT-M

and which are left in the maternal pronucleus following fertilization. Accordingly, if the normal gene is extruded with PB2 (e.g., PB2 is hemizygous normal), the resulting maternal contribution to the embryos is the mutant gene, or, in reverse if the mutant gene is extruded with PB2 (e.g., PB2 is hemizygous mutant), the resulting maternal contribution to the embryos is the normal gene. It is furthermore possible that even the oocytes predicted as mutant may form unaffected heterozygous embryos, following fertilization by a mutation-free sperm. Therefore, with insufficient number of mutation-­ free oocytes, preselected by PB1 and PB2 sequential analysis, testing of the resulting embryos may allow the identification of heterozygous unaffected carrier embryos for transfer. Preselection of mutation-free oocytes was performed based on the simultaneous mutation detection and linked marker analysis, involving two strongly linked markers D9S58 and D9S1677, which were shown not to be involved in recombination in the analysis of 435 FD chromosomes [19]. Therefore, prior to PGT-M, a single-­sperm testing was performed to identify the paternal haplotypes, which were as follows: the mutant allele was linked to 116  bp, and the normal to 151  bp repeat of the D9S58 marker, while the D9S1677 marker was not informative. The maternal haplotypes were established based on PB analysis as follows: the mutant allele was linked to 116 bp repeat of the D9S58 marker, and 140 bp repeat of the D9S1677 marker, while the normal allele was linked to 140 bp repeat of the D9S58 marker and 126 bp repeat of the D9S1677 marker (Fig.  4.6). Overall, 15 oocytes were ­available for testing, of which 11 were with the information for both PB1 and PB2. Of these 11 oocytes, 4 were predicted to be mutant based on the heterozygous PB1 and hemizygous normal (mutation-free) PB2 (oocytes #8, #10, #13, and #15), while the remaining 7 oocytes were free of the mutant gene, as evidenced by the heterozygous PB1 and hemizygous mutant PB2. These results were in agreement with both markers, except for oocytes #7 and #11, in which ADO of

87

D9S1677 allele linked to the mutant gene was observed. Three embryos resulting from the above seven oocytes (embryos #1, #3, #6; Fig. 4.6), having mutation-free status confirmed by both markers, and reaching the blastocyst stage, were transferred back to the patient. This yielded a triplet pregnancy leading to birth of three unaffected children, two homozygous normal and one heterozygous carrier. These results and further similar cases performed by the present time demonstrate diagnostic accuracy of PGT for FD by sequential PB1and PB2 analysis. Follow-up analysis of the embryos resulting from either mutant or normal (mutation-­ free) oocytes was in agreement with the sequential PB1 and PB2 analysis in all the embryos tested. Although ideally at least three linked markers are considered necessary to completely exclude the risk for misdiagnosis due to ADO, use of the two linked markers or even one in the present study proved reliable because it was possible to follow both normal and mutant alleles through the first and second meiotic divisions. Presence of both mutant and normal alleles in PB1 following meiosis I, as well as detection of the mutant allele extruded with PB2 following meiosis II, left no doubt of the mutation-free status of the maternal pronucleus, even if no linked markers were available for testing. However, testing of a sufficient number of linked markers would be absolutely essential to exclude the possible ADO of the normal allele if PB1 had appeared to be homozygous mutant; failure of detecting ADO could lead to the opposite interpretation for PB2. With undetected ADO in PB1, the presence of a normal allele in PB2 would erroneously suggest the normal (mutation-free) status of the resulting oocyte, which is actually mutant. The presented experience of PGT for FD demonstrates the clinical relevance of PGT in those couples who cannot accept prenatal diagnosis and termination of pregnancy [20]. Because of the high prevalence of FD in Ashkenazi Jews, with carrier frequency 1 in 32, this approach may have practical implications; at-risk couples

88

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

require information about the availability of PGT-M. The presented PGT design for FD may be applied without extensive preparatory work in different couples, due to a single major mutation being involved and existence of a sufficient number of informative linked markers from which to choose. As shown also by the above example of PGT for FD, it may be predicted that PGT may in the future be applied for gene expression ­abnormalities that might be limited to a particular tissue or particular stage of embryonic development. This may also allow preselecting the embryos with best potential to establish a viable pregnancy, based on the progress on the understanding of stage-specific gene expression. Spinal Muscular Atrophy (SMA)  is another relatively frequent indication and presents a real complexity for PGT-M [21, 22]. This severe autosomal recessive disorder has a newborn prevalence of 1/10,000 and a carrier frequency of 1 in 50 individuals. All types of SMA are caused by mutations in the survival motor neuron (SMN) gene locus mapped on chromosome 5q11.2–13 [16]. The SMN gene consists of two highly homologous copies, SMN1 and SMN2, of which homozygous loss of functional survival motor neuron 1 (SMN1) alleles results in SMA but homozygous absence of SMN2 gives no clinical phenotype. Since 95% of SMA patients lack both copies of SMN1 in exon 7, PGT for SMA is based on the absence of SMN1 homozygous deletions. Doing so is complex because sequence similarity between the SMN1 and SMN2 genes requires simultaneous linkage analysis involving STR or SNP markers. To test for paternal SMN1 and SMN2 deletion patterns on each chromosome by embryo biopsy, single-sperm testing is required to establish the linkage between normal and deleted alleles with a multi-copy markers on the promoter region of both gene copies. PGT-M is the most practical option chosen by couples at risk, presented in Fig. 4.7, which shows PGT for a couple with both parents carrying the deletion in the SMN1 gene. Of six embryos tested for deletion and linked markers, one embryo was mutant, four were predicted to be heterozygous for the deletion in SMN1 gene, and one (embryo 23)

has only a copy of the normal gene, and was also free of aneuploidy. This embryo was transferred, resulting in the birth of an unaffected child.

4.1.2 Autosomal Dominant Disorders Autosomal dominant conditions are important candidates for PGT-M, as couples have a 50% risk of producing an affected child. PGT for autosomal dominant disorders represents under one-­ fifth of our experience, which was extremely accurate and effective in detection and transfer of mutation-free embryos. Autosomal dominant conditions for which PGT was performed are presented in Table 4.1. Examples of two conditions  – early-onset primary torsion dystonia (PTD) and Charcot–Marie–Tooth (CMT) disease – are described below. Primary Torsion Dystonia (PTD)  Is caused in most cases by a 3 bp deletion of the DYT1 gene, located on chromosome 9q34 [16, 17, 23–25]. PTD is the most severe and common form of hereditary movement disorders, present in 1/15,000 live births and characterized by sustained twisting, contractures that begin in an arm or leg between 4 and 44 years, spreading to other limbs within about 5 years. Although phenotypic expression of the disease is similar in all ethnic populations, the highest prevalence is reported among Ashkenazi Jews. Despite low penetrance (30–40%), the disease phenotype varies greatly among families. In contrast to other neurodegenerative disorders above, PTD does not show any distinct neuropathology. A 3 bp deletion in the coding sequence of the DYT1 gene is believed to result in loss of a pair of glutamic acid residues in a conserved region of an ATP-binding protein torsin A that has resemblance to the heat shock proteins and may lead to imbalance of neuronal transmission in the basal ganglia implicated in dystonia. Low levels of dopaminergic metabolites in the cerebrospinal fluid of these patients show no response to dopa, probably caused by a defect in release rather than failure of synthesis of dopamine. The remarkable phenotypic variability of the disease may be explained by the interaction of

4.1 Traditional Indications and Strategies for PGT-M

a

89

Family Pedigree DEL / N

DEL / N

PGT

b

PGT

N/N

DEL / DEL

Embryo #

D5S 2046

D5S 435

D5S 1414

EXON 7

EXON 8

D5S 610

D5S 351

D5S 1491

Predicted Genotype

NGS

1

128/134

159/154

202/200

N / DEL

N / DEL

101/103

129/123

146/150

CARRIER

45, XX, -10

15

128/134

159/154

202/200

N / DEL

N / DEL

101/103

129/123

146/150

CARRIER

47, XY, +16

16

126/132

145/154

204/202

DEL / N

DEL / N

112/93

143/125

141/146

CARRIER

46, XY

17

128/134

159/154

202/200

N / DEL

N / DEL

101/103

129/123

146/150

CARRIER

46,XX

23

128/132

159/154

202

N

N

101/93

129/125

146

NORMAL

46, XY

24

126/134

145/154

204/200

DEL

DEL

112/103

143/123

141/150

AFFECTED

46, XY

SMA 440

126/128

145/159

204/202

DEL / N

DEL / N

112/101

143/129

141/146

PARTNER DEL/N

SMA 441

132/134

154/154

202/200

N / DEL

N / DEL

93/103

125/123

146/150

PATIENT N/DEL

Embryo #23

46, XY

Embryo #1

45, XX, -10

Embryo #15 47, XY, +16

Fig. 4.7  Combined testing PGT-M and PGT-A for spinal muscular atrophy (SMA) caused by deletion mutation in SMN1 gene. Couple’s first baby was affected with SMA and died. Both parents are carriers for the deletion in SMN1 gene. On a pedigree (upper middle panel) red bar corresponds to chromosome 5 harboring a deletion, and a white bar represents a normal sequence. Whole genome amplification (WGA) products were used to amplify exon 7 and exon 8 of SMN1 and SMN2 genes and linked polymorphic markers surrounding SMN1/SMN2 on both sides. The outcome of the first cycle is presented in table (middle panel). Left column shoes tested embryo numbers and parents’ DNA, followed by columns with informative markers, embryo genotype prediction, and aneuploidy test results by NGS.

Six embryos were developed to blastocyst, and trophectoderm biopsy was performed on Day 5. One embryo was predicted normal (#23), one embryo was predicted to be affected (#24), and four embryos were carriers (#1, 15, 16, and 17). Three of these embryos inherited maternal deletion (#1, #15, and #17) and one inherited a deletion from father (#16). Of six embryos tested for aneuploidy, four were euploid (#16, #17, #23, and #24), of which the first three were recommended for frozen transfer. Among aneuploidy embryos, #1 had monosomy, and #10 and #15 had trisomy 16 (bottom panel). Embryo #23, unaffected and free of aneuploidy, was transferred, resulting in birth of a healthy boy

the 3 bp deletion with modifying genes, such as polymorphic variations in torsin by mutations in the associated proteins, as well as by interaction with environmental factors such as trauma, high body temperature, or exposure to toxic agents. Although understanding these relationships may allow elucidating neuronal mechanisms underlying loss of movement control, there is no effective treatment. This makes PGT-M a useful option for those at-risk couples who cannot accept prenatal diagnosis and termination of pregnancy as an option for avoiding PTD in their offspring.

A couple to be described presented for PGT with the affected paternal partner carrying the DYT1 3  bp deletion (GAG) inherited from his mother who had no other children (Fig. 4.8a, b). PGT-M cycle was performed using embryo biopsy, tested by multiplex nested PCR analysis, involving testing the DYT1 mutation simultaneously with a set of linked polymorphic markers. The mutation analysis for the GAG deletion in the coding sequence of DYT1 was based on fragment-­ size analysis using capillary electrophoresis or BSeRI restriction digestion. The latter

90

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

creates three fragments of 161, 24, and 8 bp in the normal allele but only two fragments of 185 and 8 bp in the mutant gene. Three closely linked markers  – D9S62, D9S63, and ASS (intron 14) – were shown not to be involved in recombination with the DYT1 gene [23, 25–27] and were used in the multiplex nested PCR system. To identify the paternal haplotype, single-sperm testing was performed prior to PGT. This showed linkage of the mutant allele to markers of lengths 121, 157, and 134 bp; the normal paternal allele was linked to 123, 155, and 130  bp repeat of markers D9S62, D9S63, and

ASS.  The maternal haplotypes were 123/123, 140/155, and 126/136  bp repeats of D9S62, D9S63, and ASS, respectively. Embryos derived from oocytes free of DYT1 3 bp deletion, based on the above polymorphic markers, were preselected for transfer, whereas those predicted to be mutant or with insufficient marker information were subjected to confirmatory analysis using genomic DNA from these embryos to evaluate the accuracy of single-cell-­ based PGT-M.  Eight embryos were available for testing; two were free of the 3 bp deletion, all three markers not only excluding a possible ADO of the mutation [28], but con-

Fig. 4.8 (a) Pedigree of a couple whose PGT for DYT1 resulted in the birth of a mutation-free baby. (Upper panel) Patient’s parents, showing that he inherited 3  bp deletion of DYT1 gene from his father. (Middle panel) The father is a carrier of a 3 bp deletion of DYT1 gene, which is linked to 121, 157, and 134 bp repeats of D9S62, D9S63, and ASS markers, respectively, while the normal allele is linked to 128, 140, and 124 bp repeats of the same polymorphic markers, respectively. The mother is normal, with one normal DYT1 allele linked to 123, 150, and 128 and the other to 123, 155, and 124 bp repeats of D9S62, D9S63, and ASS markers, respectively. As seen from the upper panel the mutation was inherited from the paternal mother, with no other family members available in the pedigree. (Lower panel) Reproductive outcomes following PGT, showing the 3 bp deletion-free baby, which is in agreement with polymorphic markers, also suggesting the presence of both paternal and maternal normal genes. This embryo originates from the transfer of embryo #6, as shown in (b). (b) Preimplantation genetic testing for GAG deletion of DYT1 gene, resulting in the birth of mutationfree baby. Capillary electrophoregrams of fluorescently labeled PCR products of linked markers D9S62 (first from the left), D9S63 (third from the left), and ASS (first from the right), scored by Genotyper TM. The data of genotyping of only three embryos are shown as examples, including two transferred normal (embryos #6 and #8) and one affected (embryo #10) embryo. Paternally derived 128,

140, and 124 dinucleotides indicative of the DYT1 mutation are evident in blastomeres of embryos #6 and #8, together with the presence of maternal normal alleles. (Second panel from the left) The location (top) of the mutation in DYT1, restriction map for Dse RI digestion (second panel from the top), creating three fragments of 161, 24, and 8 bp in the normal allele, in contrast to only two fragments of 185 and 8  bp in the mutant gene. However, because this required a long incubation and high amount of enzymes, fluorescent genotyping was also performed (bottom of this panel). (Middle section of the same panel) The polyacrylamide gel electrophoregram of Dse RI-digested PCR products of eight blastomeres from one of the cycles of PGT, paternal DNA from sperm (P), and maternal (normal) DNA. Hdx – the extra fragment in the heterozygous mutant embryos as a result of heteroduplex formation. (Bottom section of this panel) Capillary electrophoregrams of fluorescently labeled PCR products of some of the above blastomeres, including two normal (embryos #6 and #8) and one affected (embryo #10). Paternally derived GAG deletion shown by an arrow is evident in embryo #10, which is absent in embryos #6 and #8, in agreement with the linked marker analysis (see relevant panels on the left and right). These embryos inherited the paternal normal chromosome, but may be distinguished from each other by the inheritance of different maternal chromosomes, allowing the identification of the origin of the resulting mutation-free baby (a)

4.1 Traditional Indications and Strategies for PGT-M

a

91

Family Pedigree

Markers order: D9S62 DYT1 D9S63 ASS

123 N 149 128

121 128 Del N 157 140 134 124

Paternal haplotype is based on single sperm typing, confirmed by Blastomere analysis

PGT 128 N 140 124

b

123 N 155 124

Maternal haplotype is based on Blastomere results

123 N 149 128

PGT for DYT resulting in birth of an unaffected child Del GAG

(CA)n

D9S62

DYT 1

(CA)n

D9S63

ASS Intron 14

3000

3000

2000

2000

Restriction map:

1000

185

128

161

Embryo #8 (Normal)

2000 1500 1000 500

1000

8 24 8

Mutant Normal

140

124 128

150

Embryo #8 (Normal)

Embryo #8 (Normal) 4000 3000 2000 1000

3000 2000 1000

123

Embryo #6 (Normal)

Embryo #6 (Normal)

Embryo #6 (Normal)

123

(CA)n

140

128

124

155

Embryo #10 (Mutant)

Embryo #10 (Mutant) Embryo #10 (Mutant)

4000 3000 2000 1000

Hdx Mutant Normal

3000 2000 1000

155 157

121 123 Embryo #

3 6 7 8 10 11 12 13 M F ET ET

4000 3000 2000 1000

900 600 300

124

134

92

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

firming the presence of both maternal and paternal normal alleles. These two embryos were transferred to the patient resulting in the birth of a mutation-free boy. These results represented the first experience of PGT for TDY1, demonstrating the clinical usefulness of PGT in those couples that cannot accept prenatal diagnosis that leads to termination of pregnancy. Because a single unique 3 bp deletion is involved in more than 70% cases of early-onset PTD, the PGT design for testing this DYT1 mutation may be applied without extensive preparatory work in other couples, taking also into consideration the limited number of founder mutations [25]. As seen from the sperm haplotype analysis of our patients with GAG deletion of the DYT1 gene, the same haplotypes usually surround the DYT1 gene. Availability of a sufficient number of highly variable and closely linked markers allows testing for the mutation simultaneously with at least three markers to exclude misdiagnosis due to ADO. To ensure reliable preselection of mutation-free embryos for transfer, PGT should include detection of both paternal and maternal normal alleles in addition to the exclusion of GAG deletion, which may be masked by ADO.  This also allows identification of an individual embryo that is implanted among a cohort of more than one embryos transferred [29]. Although prenatal genetic diagnosis for PTD is also available, PGT-M may seem a more attractive option. That approximately 70% of the offspring will not develop the disease in the obligate carriers of the mutation makes the decision of what to do in the case of a mutation carrier especially difficult for parents. Charcot–Marie–Tooth (CMT) Disease  Represents a clinically and genetically heterogeneous group of hereditary peripheral neuropathies (Table  4.7), affecting approximately 1  in 2500  in the United States. Prenatal diagnosis is available, but termination of pregnancy is not acceptable for many couples. Thus, PGT was an attractive option for CMT, initially reported for five couples with CMT1A. This is the most frequent autosomal dominant type of CMT, caused by 1.5  Mb tandem duplication that includes a dosage-sensitive gene for peripheral myelin protein 22 (PMP22) on chromosome 17p11.2–12 [30]. PCR design for PGT of this condition

involved the application of 13 highly polymorphic microsatellite markers, located within the duplicated area and closely linked to the PMP22 gene, some showing three alleles in patients with CMT1A duplication. PGT was performed for CMT type 1A and 1B, with both paternally and maternally derived mutations. In those with paternally derived duplication, single-sperm analysis was performed to determine normal and mutant haplotypes. In the PGT cycles with maternal mutation, both PB1 and PB2 and embryo biopsy were analyzed, using markers D17S1357, D17S2229, D17S2226, D17S2225, D17S839, D17S2224, D17S2221, D17S2220, D17S291, D17S2219, D17S2218, D17S2217, and D17S2216. These were amplified in a multiplex heminested PCR system, followed by fragment analysis on an ABI 3100 analyzer. As a result the embryos free of PMP 22 duplication were transferred, yielding unaffected pregnancies and the birth of unaffected children. Overall, we performed 50 cycles for CMT1A, resulting in the birth of 25 healthy children free of PMP 22 duplication. Among rare indication was autosomal dominant CMT-type 2E, for which PGT was performed for one patient. This was a maternally derived mutation in the light polypeptide neurofilament protein gene (NEFL), which was tested by PB1 and PB2 analysis, followed by embryo biopsy to detect the presence of the mutation P8R, simultaneously with microsatellite marker D8S137. This resulted in the birth of a child with a normal NEFL gene. PGT has also been performed for six patients with X-linked form of CMT, caused by mutations in connexin-32 (Cx32) gene, by testing for the presence of the V95M mutation in the Cx32 gene, simultaneously with STRs (DXS453, DXS8052, DXS8030, DXS559, DXS441). This resulted in the transfer of mutation-free embryos and in the birth of six unaffected babies. An example of PGT for paternally derived CMT1A is presented in Fig. 4.9, showing the outcome of testing of ten embryos using ten linked markers. Five unaffected embryos were identified, two of which were transferred resulting in the birth of a healthy child, again demonstrating the reliability and accuracy of PGT designs applied to PGT for CMT.

Disease Muscular dystrophy Emery–Dreifuss muscular dystrophy 1, X-linked; EDMD1 Emery–Dreifuss muscular dystrophy 2, autosomal dominant; EDMD2 Facioscapulohumeral muscular dystrophy 1; FSHD1 Muscular dystrophy, Becker type; BMD, DMD Muscular dystrophy, congenital merosin-deficient, 1A; MDC1A Muscular dystrophy, limb-girdle, type 2A; LGMD2A Muscular dystrophy, limb-girdle, type 2S; LGMD2S Muscular dystrophy-dystroglycanopathy (congenital with brain and eye anomalies), type A, 4; MDDGA4 Muscular dystrophy-dystroglycanopathy (congenital with brain and eye anomalies), type a, 5; MDDGA5 Ullrich congenital muscular dystrophy 1; UCMD1 Subtotal Myopathy congenital Myopathy, areflexia, respiratory distress, and dysphagia, early-onset; EMARDD Myopathy, centronuclear, X-linked; CNMX Myopathy, myofibrillar, 1; MFM1 Nemaline myopathy 2; NEM2 Subtotal Myopathy metabolic Carnitine palmitoyltransferase II deficiency, infantile Glycogen storage disease II; GSD2 Hydroxyacyl-CoA dehydrogenase/3-ketoacyl-CoA thiolase/enoyl-CoA hydratase, alpha subunit; HADHA Myoglobinuria, acute recurrent, autosomal recessive Leigh syndrome; LS Subtotal Myotonic dystrophy and other myotonic disorders Myotonic dystrophy 1; DM1 Myotonic dystrophy 2; DM2

Table 4.7  PGT for neuromuscular disorders

4 17

3 7 25 67 7 1 1 2 1 1 114 1 5 1 6 13 4 7 4 1 1 17 94 1

310300 181350 158900 300376 607855

254090

614399 310400 601419 256030

600649 232300 609016 268200 256000

160900 602668

FRG1 DMD LAMA2 CAPN3 253600 TRAPPC11 615356 FKTN 253800 613153

EMD LNMA

FKRP Col6A2 10 genes MEGF10 MTM1 DES NEB 4 genes CPT2 GAA HADHA LPIN1 SURF1 5 genes DMPK CNBP

147 2

1 1 26

8 9 7

6 1 6 14

1

1 203

3

1 2 2

54 112 8

# of cycles

# of OMIM patients

Gene

107 2

1 2 16

5 4 4

4 1 5 11

1

0 149

3

0 2 2

42 74 7

3 16

# of transfers

188 4

1 3 26

5 9 8

7 1 8 17

1

0 255

3

0 2 3

71 132 13

6 25

# of embryos transferred

55 2

1 0 10

2 3 4

4 1 4 10

1

0 95

1

0 2 2

22 49 6

3 10

10 0

0 0 1

0 0 1

0 0 0 0

0

0 12

0

0 0 0

2 8 1

0 1

Pregnancy SAB

45 2

1 0 9

2 3 3

4 1 4 10

1

0 83

1

0 2 2

20 41 5

3 9

(continued)

50 2

1 0 10

2 3 4

5 1 4 11

1

0 97

1

0 2 2

24 48 6

4 10

Delivery Birth

4.1 Traditional Indications and Strategies for PGT-M 93

Disease Myotonia congenita, autosomal dominant Paramyotonia congenita of von Eulenburg; PMC Subtotal Motor neuron disease Amyotrophic lateral sclerosis 1; ALS1 Amyotrophic lateral sclerosis 4, juvenile; ALS4 Spastic paraplegia 3, autosomal dominant; SPG3A Spastic paraplegia 4, autosomal dominant; SPG4 Spinal and bulbar muscular atrophy, X-linked 1; SMAX1 Spinal muscular atrophy, distal, autosomal recessive, 1; DSMA1 Spinal muscular atrophy, type I; SMA1 Subtotal Neuropathy hereditary Amyloidosis, hereditary, transthyretin-related Charcot–Marie–Tooth disease, xonal, type 2A2; CMT2A2 Charcot–Marie–Tooth disease, axonal, type 2B; CMT2B Charcot–Marie–Tooth disease, axonal, type 2E; CMT2E Charcot–Marie–Tooth disease, axonal, type 2F; CMT2F Charcot–Marie–Tooth disease, demyelinating, type 1A; CMT1A Charcot–Marie–Tooth disease, demyelinating, type 1B; CMT1B Charcot–Marie–Tooth disease, X-linked, 1; CMTX1 Hereditary motor and sensory neuropathy, type IIC; HMSN2C Neuropathy, hereditary sensory and autonomic, type III; HSAN3 Neuropathy, hereditary sensory and autonomic, type VI; HSAN6 Subtotal Total

Table 4.7 (continued)

102 116 3 2 1 1 1 27 3 6 1 13 1

253300

105210 609260 600882 607684 606595 118220 118200 302800 606071 223900 614653

SMN1 7 genes TTR MFN2 RAB7A NEFL HSPB1 PMP22 MPZ GJB1 TRPV4 IKBKAP DST

59 418

2 1 1 5 3 2

105400 602433 182600 182601 313200 604320

SOD1 SETX ATL1 SPAST AR IGHMBP2

11 genes

# of OMIM patients 160800 1 168300 3 99

Gene CLCN1 SCN4A 4 genes

107 676

2

20

9 1

6

7 6 1 4 1 50

151 173

2 1 1 10 5 3

# of cycles 1 3 153

79 511

2

16

9 2

2

5 3 1 4 1 34

124 142

2 1 1 8 4 2

# of transfers 1 4 114

118 813 (1.6)

3

27

16 2

5

6 4 2 7 1 45

201 229

3 2 1 12 6 4

# of embryos transferred 2 4 198

47 311 (60.8%)

3

9

5 1

0

3 2 1 1 1 21

75 88

2 1 1 6 2 1

Pregnancy 1 3 61

2

9

5 1

0

2 2 1 1 1 18

67 77

1 1 1 5 1 1

2

11

6 1

0

2 2 2 1 1 25

80 93

1 1 1 6 2 2

Delivery Birth 1 1 2 2 50 55

5 42 53 40 271 319 (12.8%) (87.2%)

1

0

0 0

0

1 0 0 0 0 3

8 11

1 0 0 1 1 0

SAB 0 1 11

94 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

D17S2216

D17S2217

D17S921

D17S2220

D17S2221

D17S839

D17S2224

D17S2225

95

PMP 22

Proximal CEN

D17S2226

D17S1357

4.1 Traditional Indications and Strategies for PGT-M

CMT1A-Rep

Distal TEL

CMT1A-Rep 0.5Mb

1Mb

1. 5 M b

17p12

Markers order: D17S1357 D17S2226 D17S2225 D17S2224 D17S839 D17S2221 D17S2220 D17S921 D17S2217 d17S2216

123 136 158 196 124 107 351 124 113 121

123/111 140/136 162/164 196/204 126/142 111/111 312/328 116/116 111/118 113/119

N/N

N/Dup PGT

111 128 160 193 142 109 328 122 116 105

125 124 162 193 126 113 316 120 113 105

N/N Embryo #

2

123/111 140/136 162/164 196/204 126/142 111/111 312/328 116/116 111/118 113/119

125 124 162 193 126 113 316 120 113 105

AFFECTED DUPLICATION

3

123 136 158 196 124 107 351 124 113 121

125 124 162 193 126 113 316 120 113 105

NORMAL

4

111 128 160 193 142 109 328 122 116 105 AFFECTED DELETION

6

123 136 158 196 124 107 351 124 113 121

125 124 162 193 126 113 316 120 113 105

7

123 136 158 196 124 107 351 124 113 121

125 124 162 193 126 113 316 120 113 105

NORMAL

NORMAL

ET

ET

Fig. 4.9  PGT for CMT1A resulted in the birth of a healthy baby. (Upper panel) Schematic representation of 1.5  Mb tandem duplication including dosage-sensitive gene for peripheral myelin protein 22 (PMP22) and linked markers, showing the complexity of the testing. (Middle panel) Family pedigree, showing the affected father with duplication and the results of haplotype analysis established through sperm testing in the father and blastomere

Given the mutation rate for dominant conditions is much higher than for recessive and X-linked disorders, the strategy for PGT-M cycles for dominant disorders caused by de novo mutations will be presented in a special section below.

8

123 136 158 196 124 107 351 124 113 121

125 124 162 193 126 113 316 120 113 105

NORMAL

9

123 136 158 196 124 107 351 124 113 121

10

125 124 162 193 126 113 316 120 113 105

NORMAL

111 128 160 193 142 109 328 122 116 105 AFFECTED DELETION

11

123/111 140/136 162/164 196/204 126/142 111/111 312/328 116/116 111/118 113/119

111 128 160 193 142 109 328 122 116 105

AFFECTED DUPLICATION

12

123 136 158 196 124 107 351 124 113 121

125 124 162 193 126 113 316 120 113 105

NORMAL

analysis in the mother. (Lower panel) Results of multiplex PCR analysis for ten markers, which identified five duplication-free unaffected embryos and five affected ones, including two with duplication, one recombinant containing partial duplication, and two with deletion. Two of the unaffected embryos were transferred resulting in the birth of an unaffected child

the 50% of male embryos that were not affected. Couples at risk choose between prenatal diagnosis at the downside of potential pregnancy termination and PGT by gender determination at the cost of discarding 50% of male embryos that were normal. However, because X-linked disorders are maternally derived an attractive option is preselection of mutation-free oocytes through 4.1.3 X-Linked Disorders testing for specific causative mutations in PB1 and PB2. This allows avoiding any further testing X-linked conditions were the most straightfor- of the resulting embryos, which may be transward indication from the very beginning of PGT-­ ferred irrespective of gender or any contribution M. Initially this was done by gender determination from the male partner. not only because sequencing information was not The first PGT for the X-linked disorders was always available but because the approach was performed 30 years ago by gender determination technically straightforward: identifying female [1]. This was initially done by single-cell PCR embryos despite the obvious pitfall of discarding analysis, while later performed by the FISH tech-

96

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

nique in interphase cells. Although accurate, the necessity to discard the 50% of unaffected male embryos was not accepted by many patients. Thus, there has been a need for specific genetic tests allowing the identification of the unaffected healthy male embryos for transfer as well. In order to preselect the X-linked mutation-free embryos for transfer, sequential PB1 and PB2 analysis was applied as an alternative to gender determination. One of such cases, involving PGT for ornithine transcarbamylase (OTC) deficiency, was first reported 20 years ago, showing the feasibility of the approach [31]. Couples for whom PGT-M was applied have also included those with previous male offspring affected by Duchenne muscular dystrophy (DMD), Charcot–Marie–Tooth disease X1 (CMTX1), fragile X syndrome (FMR1), hemophilia A (F8) and B (F9), hyperimmunoglobulin syndrome (HIGM1), Wiskott–Aldrich syndrome (WAS), X-linked hydrocephalus (L1CAM), Hunter syndrome (IDS), choroideremia (CHM), myotubular myotonic dystrophy (MTMD), Norrie disease (NDP), X-linked adrenoleukodystrophy (ALD), ornithine transcarbamylase (OTC) deficiency, Pelizaeus–Merzbacher disease (PMLD), incontinentia pigmenti (IP), and Alport disease (ATS), to mention only a few. Examples of PGT for these conditions will be presented in the relevant sections according to classification used in Table 4.1. For example, one of X-linked disorders, Alport syndrome, causes severe kidney disorder (list of renal disorders for which PGT was performed is, presented in Table 4.8). Of 16 PGT-M cycles for X-linked Alport syndrome, 22 unaffected embryos were transferred in 15 cycles, resulting in 11 pregnancies and birth of 9 healthy children free of Alport syndrome. As mentioned, PGT for X-linked conditions may be reliably performed without embryo biopsy, based on sequential PB1 and PB2 testing, involving mutation and/or linked marker analysis. Results of PGT-M in this approach are available on Day 1 (approximately, 30–32  h after oocyte retrieval). Embryos resulting from oocytes predicted to be free from maternal mutation were transferred back to the patients, while those predicted to be affected were further tested for con-

firmation of diagnosis when available. In our earlier experience, an average of 7.7 oocytes were obtained with PB1 results per cycle, heterozygous in 66% and homozygous normal or mutant in 34%. PB2 results were available in 79% of these oocytes (6.1 oocytes per cycle on an average). In the remaining oocytes, genotype prediction was not possible, so additional testing by embryo biopsy was applied. ADO if undetected remains to be the major problem in avoiding misdiagnosis. This was observed in one PGT case for FMR1. ADOs were detected either by the use of the linked markers or by sequential PB1 and PB2 analysis. For example, in the case of PGT for X-linked hydrocephalus, both PB1 and PB2 in one of the oocytes had identical homozygous mutant genotype, suggesting that PB1 was apparently heterozygous, with the normal allele not detected due to ADO. However, at least two embryos in each cycle for this condition were available for transfer, originating from the mutation-free oocytes preselected based on the heterozygous status of PB1 and homozygous mutant PB2, which not only predicted the mutation-free status of the resulting embryos but also reliably excluded ADO. Fragile X Syndrome (FMR1)  Is the largest group of X-linked disorders for which PGT has been offered. It is tested indirectly using linkage analysis given no test is available for direct analysis of the expanded allele in a single cell [32]. As the dynamic mutation causing this condition cannot reliably be amplified, preselection of the mutation-free oocytes must be inferred from the presence of the linked markers for the normal allele. Only a few of these cycles were performed by embryo biopsy alone, with the majority performed either by PB1 and PB2 only or PB1 and PB2 analysis followed by embryo biopsy, if necessary. Only one misdiagnosis was observed, involving undetected ADO. Three embryos were transferred, two deriving from oocytes with heterozygous PB1 and mutant PB2 and one from the oocyte with presumably homozygous mutant PB1. In fact, the latter turned out to be heterozygous because of an undetected ADO of both markers linked to the normal gene (Fig.  4.10).

Disease Alport syndrome, autosomal dominant Alport syndrome, X-linked Arthrogryposis, renal dysfunction Bartter syndrome, type 3 Cystinosis, nephropathic Gitelman syndrome Hyperuricemic nephropathy, familial juvenile, 1 Nephrogenic syndrome of inappropriate antidiuresis Nephrotic syndrome, type 1; NPHS1 Nephrotic syndrome, type 2 Nephrotic syndrome, type 5 Polycystic kidney disease 1 Polycystic kidney disease 2 Polycystic kidney disease, autosomal recessive Renal tubular acidosis, distal Renal tubular dysgenesis Total

Table 4.8  PGT-M for renal disorders # of OMIM patients 607364 1 301050 8 208085 1 CLCNKB 1 219800 1 263800 1 162000 1 1 1 1 1 48 2 16 1 1 86

300539 256300 600995 609049 173900 613095 263200

Gene Col4A3 COL4A5 VPS33B CLCNKB CTNS SLC12A3 UMOD

AVPR2

NPHS1 NPHS2 LAMB2 PKD1 PKD2 PKHD1

ATP6V0A4 602722 ACE 267430

1 2 152

3 1 2 84 2 29

3

# of cycles 4 16 1 1 1 1 1

2 2 124

3 1 2 64 2 25

3

# of transfers 0 15 0 2 1 1 1

2 2 191 1.5

7 1 4 98 3 41

6

# of embryos transferred 0 22 0 2 1 1 1

1 1 71 57.2%

1 1 2 35 2 15

1

Pregnancy 0 11 0 1 0 0 0

0 0 1 32 2 14

1

# of deliveries 0 9 0 1 0 0 0

0 1 0 1 9 62 12.6% 87.4%

1 1 1 3 0 1

0

SAB 0 2 0 0 0 0 0

1 1 70

0 1 1 36 3 16

1

# of babies 0 9 0 1 0 0 0

4.1 Traditional Indications and Strategies for PGT-M 97

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

98

Oocyte

Cell type

DXs1193

DXs548

Predicted Genotype

1

PB1

154/156

245/247

AFFECTED

PB2

156

247

PB1

154/156

245/247

PB2

156

247

PB1

154/156

245/247

3 5 6 9 10

PB2

154

245

PB1

245/247

245/247

PB2

154

247

PB1

154*

245*

PB2

156

247

PB1

245/247

245/247

PB2

156

247

154

245

Cord blood

Family Pedigree 247 N 156

AFFECTED NORMAL NORMAL NORMAL* AFFECTED

245 247 Exp N 1.2 154 145

1.1

1 2.1 247 N 154

2.2 245 247 Exp N 154 156

3.1 PGT

3.2 PGT

245 Exp 154

247 247 N N 154 156

2.3

2.4 2.5 245 247 247 Exp N N 154 156 145

2.6 247 N 145

2.7 247 N 145

3.3 245 Exp 154

AFFECTED

Fig. 4.10  PGT for FRM1 with misdiagnosis due to undetected ADO in the first polar body. Family pedigree (right) and PGD results of haplotype analysis for CGG expansion in FMR1 gene using two linked markers (left). The mother is a carrier of an expanded allele linked to 154 and 245 markers. Her sister also has an expanded allele inherited from their mother. One of the sisters who presented for PGT is shown by arrow. The other sister has an affected son, who carries only an expanded allele (linked to 154 and 245 markers). On the left of the pedigree, an affected child carrying only an expanded allele (linked to 154 and 245 markers) was born following PGT, due to the pre-

dicted 5% risk of misdiagnosis, resulting from undetected ADO of the normal allele in PB1 of the corresponding oocyte (shown by ∗ in the table on the left), from which the transferred embryo was derived. As can be seen from the table, misdiagnosis originates from the undetected ADO of both markers linked to the expanded allele in PB1 of oocyte #9, which was erroneously considered homozygous mutant, while it was actually heterozygous; following the extrusion of PB2 with the normal allele, the maternal contribution to the resulting oocytes should have been considered mutant

As only two of four available markers were informative for linkage analysis in this cycle, absence of a direct test for the expanded allele necessitated that the chance of ADO of the normal allele in this case be predicted to be over 5%, based on our previous observations described in detail above.

PB2, but with the linked marker information available to confirm the diagnosis. Despite transferring only one or two embryos resulting from mutationfree oocytes in each of these cycles, each of them resulted in unaffected pregnancies and birth of healthy children. An increasing number of patients presenting for PGT are of advanced reproductive age. Thus, combined aneuploidy testing is useful. Avoidance of the birth of children with chromosomal disorders improves PGT clinical outcome, as demonstrated in the example of PGT for PMLD.

Ornithine Transcarbamylase (OTC) Deficiency  Was the first X-linked disorder for which the PB approach was utilized [31]. In this report, five PGT cycles were performed for four couples at risk for producing offspring with OTC; unaffected embryos for transfer were predicted in every cycle (1.6 embryos per cycles), resulting in four clinical pregnancies and the birth of four healthy children free of mutation, confirmed by prenatal genetic diagnosis. Preselection of mutation-­free oocytes was based on ­heterozygous PB1 and mutant PB2, except only a few with the homozygous mutant PB1 and normal

Pelizaeus–Merzbacher Disease (PMLD)  Is an X-linked recessive demyelinating disorder of the central nervous system that leads to deterioration of coordination, motor ability, and intellectual function. Varied expressivity is common. PMLD is caused by mutation of the proteolipid protein 1

4.1 Traditional Indications and Strategies for PGT-M

(PLP1) gene, located on the long arm of the X (Xq22) chromosome. This gene consists of seven exons 15 kbp in length, encoding two major proteins PLP1 and DM20 that result from alternative splicing. Both provide an important structural component of compact myelin [16, 33]. PMD manifests in infancy or early childhood, with a span of continuum of neurological impairments from nystagmus, delayed motor and cognitive impairment, to severe spasticity and ataxia with childhood mortality from respiratory complication. Mutations in PLP1 may result in misfolding of coded proteins, failing to progress through the intracellular processes. This results in the accumulation of misfolding proteins in the endoplasmic reticulum, triggering apoptosis and adversely affecting oligodendrocyte survival and myelination. Several non-disease-causing polymorphisms have been described, and at least 100 different mutations in the PLP1 gene are known to be associated with PMD. These include duplications and the most frequent small insertions, deletions, and single-base substitutions leading to missense, nonsense, or splicing mutations. Because no specific treatment for PMD is available, prenatal diagnosis is offered as an option for couples at risk to avoid the birth of the affected child [34]. Because of the wide individual phenotypic variability even within families, the decision about termination of the affected pregnancy may be extremely difficult even for those couples who accept prenatal diagnosis. Fig. 4.11 presents PGT for PMD in the couple with a previous affected child. The phenotypically normal 35-year-old mother was a carrier of the T257C (L86P) mutation in exon 3 of the PLP1 gene, representing T to C substitution in nucleotide position 257 and resulting in lysine to proline substitution in 86 position of amino acid sequence of PLP1 protein. The couple had two previous pregnancies, one resulting in spontaneous abortion and the other terminated because the fetus was diagnosed with PMD at chorionic villus sampling (CVS). The PGT-M cycle was performed using both PB and embryo biopsy analysis. PB1 and PB2 were removed sequentially following maturation and fertilization of the oocytes and tested by the multiplex PCR

99

analysis, involving the mutation testing simultaneously with two closely linked polymorphic markers, PLP-intragenic microsatellite PLP5′ (CA)n and the nearest flanking extragenic microsatellite DXS8020. Mutation analysis involved detection of T to C change in nucleotide position 257 of exon 3, based on Msp I restriction digestion that does not cut the normal allele but creates two fragments of 48 and 22  bp in the mutant allele (Fig. 4.11). Because the oocytes predicted as mutant may further form unaffected heterozygous female embryos following fertilization by a sperm carrying the X chromosome, testing for paternal normal gene in the resulting embryos was also performed. Maternal haplotypes (Fig. 4.11) were established based on PB analysis: the mutant allele was linked to 168  bp repeat of DXS8020 and 99  bp repeat of PLP5′ (CA)n marker; the normal allele was linked to 172  bp repeat of DXS8020 and 118  bp repeat of PLP5′ (CA)n marker. Paternal haplotypes were established based on family blood sample DNA analysis: the normal paternal allele was linked to 168 bp repeat of DXS8020 and 116 bp repeat of PLP5′ (CA)n marker. Of 16 oocytes available for testing in 1 PGT cycle, 9 were mutation-free based on heterozygous PB1 and affected PB2 in 2 of them (embryos #7 and #14) and mutant PB1 and normal PB2 in the remaining 7 oocytes. Six embryos were affected, based on heterozygous PB1 and normal PB2 in three of them (embryos #5, #6, and #13) and normal PB1 and mutant PB2 in the remaining three (embryos #1, #8, and #10) (Fig. 4.11). All were in accordance with the two tightly linked polymorphic markers DXS 8020 and 5′ (CA)n, as shown in three normal oocytes (oocytes #2, #3, and #11). In all but one (oocyte #9), both PB1 and PB2 mutation analysis demonstrated the normal genotype, whereas marker analysis revealed both normal and mutant alleles in PB1, suggesting allele dropout (ADO) of the mutant allele. Of 12 embryos available for analysis, 3 were affected (embryos #5, #9, and #13), 5 chromosomally abnormal, including one with uniparental disomy for chromosome 16 (embryo #8) and one with monosomy 16 in the affected embryo

100

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Fig. 4.11 PGT for PMD with aneuploidy testing. (a) Pedigree showing that the mother (1.2) is a carrier of L86P mutation in exon 3 of the PLP1 gene, which is linked to 99  bp repeat of 5′ PLP1 (CA)n, and 168  bp repeat of DXS8020, while the normal allele is linked to 118 and 172  bp repeats of the same polymorphic markers, respectively. The father (1.1) is unaffected, carrying the normal PLP1 allele, linked to 116 and 168 bp repeats, respectively. (Lower panel) Reproductive outcomes of this couple, including one previous affected child with PMD (2.1), one spontaneous abortion (2.2), and one prenatal diagnosis, resulting in identification of the affected fetus and termination of pregnancy (2.3). SAB spontaneous abortion, TAB termination of pregnancy following chorionic villus sampling (CVS). (b) Sequential PB1 and PB2 analysis of 16 oocytes, of which nine were predicted to be free of L86P mutation in exon 3 of the PLP1 gene, and seven affected. (c) Blastomere analysis for causative gene (upper panel) and

aneuploidy (lower panel), showing three mutant embryos (embryos #5, #9, and #13) and nine unaffected, including three heterozygous female embryos. The remaining four embryos were inconclusive, including one with failed amplification. Aneuploidy testing revealed two embryos with trisomy 22 (embryos #4 and #6), one with monosomy 22 (embryos #1), one with monosomy 16 (embryo #9), and one with uniparental disomy 16 (embryo #8). Three unaffected embryos (embryos #2, #3, and #11) were transferred, resulting in the birth of healthy twins. ∗Not available for testing (the IVF was performed in a different institution in another state, and it is assumed that biopsy material was not sent for testing, because embryos did not develop further. For the same reason, the fate of the other two unaffected embryos (embryos #12 and #15) is also unknown, although it is presumed that they were frozen). FA failed amplification, M mutant, Mo maternal genotype, F paternal genotype, C affected child’s genotype, ND normal (control) DNA

(embryo # 9), and 5 unaffected. Three of these embryos, free of the mutant gene and free also of aneuploidy (embryos #2, #3, and #12), were transferred, resulting in a twin pregnancy and the birth of two unaffected children confirmed to be euploid and free of T257C mutation in the PLP1 gene. This was the first PGT case performed for PMD, demonstrating the realistic option avail-

able for couples at risk for producing offspring free of the most severe type of PMD caused by the T257C mutation. PGT-M may be useful also for milder but more frequent types of PMD caused by duplications; however, the varied expressivity may present difficulties in making decision on interruption of affected pregnancies in invasive prenatal diagnosis.

4.2 Affected Homozygous or Compound Heterozygous Conditions

One incidental finding in this case was uniparental disomy of chromosome 16, detected with PCR-based aneuploidy testing. Because both chromosomes 16 were of maternal origin, the extra chromosome 16 should have been derived from maternal meiosis I, followed by subsequent trisomy rescue at the cleavage stage with loss of the paternal chromosome 16. This embryo was unfortunately not tested for presence of the trisomic cell line. Detection of uniparental disomies may in the future help in avoiding their being transferred. This would contribute to avoidance of some proportion of imprinting disorders described in association with assisted reproductive technologies [35] (see Chap. 6). With progress in obtaining sequence information for underlying X-linked disorders, PGT for an increasing number of X-linked disorders is being performed by specific genetic testing, thus avoiding discarding the 50% of male embryos that are unaffected. On the other hand, testing for X-linked genetic disorders may be entirely limited to oocytes, making needless any further manipulation and testing of the resulting embryos, which may be transferred irrespective of gender or any contribution from the father. PGT using specific diagnosis for X-linked disorders naturally can also be tested using embryo biopsy [36].

4.2

 ffected Homozygous or A Compound Heterozygous Conditions

PGT is of special value for those couples in which one partner is homozygous or compound heterozygous-affected; only 50% chance exists for having an unaffected child. This was first applied for a couple with compound heterozygous male partner affected by phenylketonuria (PKU) and female partner carrying the third PKU mutation [37]. Phenylketonuria  One in every 10,000 infants in the United States is born with PKU, an inherited metabolic disorder that causes mental retardation if untreated. When infants are fed a strict phenylalanine-free diet from birth, they have normal

101

development and a normal life span. However, after childhood dietary relaxation is possible, but during pregnancy female patients must avoid phenylalanine-restricted diet to prevent potential detrimental effects on the fetus. Because of considerable progress in screening and treating newborns for PKU with dietary modification treatment, PKU has not been a frequent indication for invasive prenatal diagnosis. With introduction of PGT-M, couples are increasingly interested in avoiding birth to a child who will need a lifelong dietary treatment. PGT may be particularly useful for couples in whom one partner is homozygous or compound heterozygous-­ affected because there is a 50% chance of producing a child affected with PKU. The first such case of PGT for PKU was performed in a couple having a compound heterozygous-­ affected partner presenting for PGT in connection with their first offspring with PKU [37]. Their producing another affected child was 50%. The 31-year-old mother was a carrier of the R408W mutation in exon 12 of phenylalanine hydroxylase (PAH), whereas the affected father was compound heterozygous for R408 and Y414C mutations in the same exon. Because the male partner was an affected compound heterozygote, the PGT strategy was based on the preselection of the maternal mutation-free oocytes using sequential PB1 and PB2 DNA analysis, using heminested multiplex PCR.  Restriction digestion of the PCR product for mutation was performed with Sty1 enzyme (Promega), followed by acrylamide gel electrophoresis. To detect possible ADO for the mutant or normal allele, two linked markers were amplified simultaneously with the PAH gene, one representing an STR in intron 3 and the other a restriction fragment length polymorphism (RFLP) XmnI in intron 8. To detect potential contamination with extraneous DNA, an additional non-linked VNTR was also amplified. Embryos resulting from oocytes predicted to contain only normal maternal alleles were transferred back to the patient, resulting in pregnancy followed up by prenatal diagnosis to confirm the results of PGT-­ M.  Accordingly, those embryos resulting from oocytes predicted to contain the mutant maternal

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

102

gene were also analyzed further to investigate the accuracy of the results of the sequential PB1 and PB2 analysis. Of 15 oocytes available for testing, results for both PB1 and PB2 were obtained in 11 to allow a reliable diagnosis. Six oocytes with both PB1 and PB2 data were predicted normal, based on the het-

erozygous PB1 containing both the normal and mutant maternal alleles and the mutant PB2 suggesting that no mutant allele was left in the resulting oocyte (Fig.  4.12). Another oocyte was predicted normal based on the presence of the mutant allele in PB1 and the normal allele in PB2. These results were in agreement with the linked

Maternal Haplotype

Paternal Haplotype 3 rep

R408W

3 rep

PB1 PB2

- Site

PB2

VNTR

PB1

intron12

PB1 PB2

XmnI

PB1 PB2

PB1 PB2

PB1 PB2

240 bp

Y414C

+Site

STR

3 rep

R408W PB2

PB1 PB2 PB1 PB2

- Site

PB1 PB2

240 bp

240 bp

VNTR

intron12

XmnI

PB1 PB2

STR

6 rep

N

+ Site

PB1

232 bp

NORMAL ADO

R408W (StyI) intron12

MUTANT

MUTANT

- site + site

RFLP (Xmn I) intron8

+ site 6 Rep 3 Rep

VNTR

STR intron3 Oocyte #

ADO

1

2

3

4

5

7

Fig. 4.12  PGT for R408W mutation in phenylalanine hydroxylase gene in a couple with the male partner homozygous affected. (Upper panel) Schematic representation of maternal (left) and paternal (right) haplotypes. Heterozygous mother has R408W mutation linked to short tandem repeats (STRs) in intron 3, variable number of tandem repeats (VNTRs) close to 3′ of gene (3 rep), and restriction fragment-length polymorphism (RFLP) in intron 8 (−site). Homozygous affected father has two different mutations, one similar to maternal (R408W) mutation, and the other, Y414C mutation, with its own linkage pattern. (Bottom panel) Genotyping oocytes by sequential analysis of PB1and PB2 for R408W mutation and informative linked markers  – STR, VNTR, and RFLP.  All series include PB1 followed by PB2  in the lane to its immediate right, corresponding to 11 oocytes studied (oocytes are numbered at the bottom). As R408W mutation creates restriction site for StyI enzyme, oocytes #2, #3, #4, #7, #9, and #15 were predicted normal based on

ADO

9

11

12

13

240bp 232bp

15

heterozygous PB1 and homozygous PB2. Oocyte #5 was also predicted normal, but based on homozygous mutant PB1 and normal PB2, which was in agreement with marker analysis, excluding the possibility for ADO in the corresponding PB1. ADO of mutant allele is evident from identical genotype of both PB1 and PB2  in oocyte #11 (confirmed by all three markers), suggesting affected status of this oocyte. Other three affected oocytes were predicted based on heterozygous PB1 and normal PB2 (oocytes #1 and #12), and homozygous normal PB1 and mutant PB2 (oocyte #13). ADO was also detected in oocytes #7 and #15 (identical genotype of PB1 and PB2 for intron 3 STR). These are not in conflict with unaffected genotype of resulting embryos, which were transferred together with two other unaffected embryos (#2 and #4), resulting in twin pregnancy and the birth of two healthy children, following confirmation of PGT by prenatal diagnosis

4.2 Affected Homozygous or Compound Heterozygous Conditions

103

Table 4.9  PGT-M for beta and alpha-thalassemia Number of mutations tested per couple Shared same mutation in the family Two different mutations Three mutations (affected family member and carrier) Subtotal Beta thalassemia Alpha-thalassemia Total

# of patients 236

# of cycles 375

# of transfers 303

# of embryos transferred 550

Pregnancy SAB 148 27

Delivery Birth 121 142

148 12

271 15

210 12

371 19

80 6

12 2

68 4

396

661

525

14 410

23 684

21 546

940 1.79 38 978 1.79

234 44.5% 10 244 44.6%

41 17.5% 0 41 16.8%

193 82.5% 10 203 83.3%

marker analysis, showing that the homozygous status of PB1 was not due to ADO of the normal allele. Three oocytes were predicted affected, the diagnosis of two of which were based on the heterozygous status of PB1 and the normal PB2; diagnosis of the third was based on the homozygous normal status of PB1 and the mutant PB2. ADO of the mutant allele was observed in one oocyte, evidenced by the identical genotype of PB1 and PB2. This was confirmed by presence of the linked polymorphic markers, suggesting that the resulting oocytes were mutant. Marker analysis also confirmed the mutant status of these three oocytes. ADO was observed in the STR analysis of two oocytes, evidenced by the identical patterns in PB1 and PB2. Three of four embryos resulting from these mutant oocytes were followed up by mutation and marker analysis, confirming presence of the mutant allele in all three. Four of seven embryos resulting from the mutation-free oocytes were available for transfer. A clinical twin pregnancy was established, both fetuses confirmed by chorionic villus sampling (CVS) to be unaffected. Healthy unaffected twins were born, with subsequent normal mental development. The application of trophectoderm biopsy would be the choice for couples with PKU maternal partner, which may involve testing for three mutations at the same reaction, representing a particular challenge. Thalassemia  In thalassemia PGT usually involves two heterozygous partners. However, with progress in the treatment, PGT-M is requested by affected and well-treated patients

83 5 230 10 240

who wish to reproduce (Table  4.9). Life expectancy has been significantly improved for thalassemia, which may be treated radically by stem cell transplantation. Our strategy in such cases depends on whether the affected partner is male or female, because testing may be entirely based on oocytes if the affected partner is male or, in contrast, embryo testing when the female partner is affected. However, if a male partner is affected with thalassemia, male factor infertility is usually involved; thus, male partners often need treatment prior to PGT-M and undergo testicular biopsy. This was the case in one of our couples having an affected male partner. A sufficient number of oocytes with homozygous mutant or heterozygous PB1 may be available, but no sperm for fertilization; thus, matured oocytes were frozen for possible future use after treatment. Still we observed a case in which the male affected partner had two different mutations (IVSI-110 and IVSII-745) and the female partner heterozygous carrier of one of the same mutation (IVSI-110) [11]. Of 25 embryos tested, 5 failed to amplify, 11 were homozygous mutant, and 9 were heterozygous normal. PGT resulted in the birth of an unaffected child. In other cycles in which the female partner was affected, embryo biopsy was the only option, allowing preselection of an unaffected embryo for transfer. However, in affected females a limited number of oocytes may be available after stimulation regiments, making PGT in some cases quite challenging. Cystic Fibrosis  We currently performed PGT for 108 different CFTR mutations (Table  4.10),

104

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Table 4.10  List of CF mutations for which PGT was performed 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50

Legacy name -94G>T 125G>C P5L G27X 306insA E60X K68E R74W R75Q R75X G85E 394delTT 406- 1G>A D110H P111L R117C R117H A120T I148T M152V L159S 621+1G>T R170H G178R N189S 711+ 1G>T P205S L206W E217G 852del22 R258G R560T 1078delT L320F 1154insTC A349V Q359K; T360K Y301C T338I R347P R347H L375F A455E 153 1C>T Q493R 1609delCA ΔI507 ΔF508 1677delTA

cDNA name c.-226G>T c.-8G>C c.14C>T c.79G>T c.174_175insA c.178G>T c.202A>G c.220C>T c.224G>A c.223C>T c.254G>A c.262_263delTT c.274-1G>A c.328G>C c.332C>T c.349C>T c.350G>A c.358G>A c.443T>C c.454A>G c.476T>C c.489+1G>T c.509G>A c.532G>A c.566A>G c.579+1G>T c.613C>T c.617T>G c.650A>G c.720_741 c.772A>G c.890G>A c.948delT c.960A>T c.1022_1023insTC c.1046C>T c.1075C>T 1079C>A c.902A>G c.1013C>T c.1040G>C c.1040G>A c.1125A>C c.1364C>A c.1399C>T c.1478A>G c.1477_1478delCA c.1519_1521delATC c.1521_1523delCTT c.1545_1546delTA

Protein name

p.Pro5Leu p.Gly27X p.Arg59LysfsX10 p.Glu60X p.Lys68Glu p.Arg74Trp p.Arg75Gln p.Arg75Opa p.Gly85Glu p.Leu88IlefsX22 p.Asp110His p.Pro111Leu p.Arg117Cys p.Arg117His p.Ala120Thr p.Ile148Thr p.Met152Val p.Leu159Ser p.Arg170His p.Gly178Arg p.Asn189Ser p.Pro205Ser p.Leu206Trp p.Glu217Gly p.Gly241GlufsX13 p.Arg258Gly p.Arg297Gln p.Phe316LeufsX12 p.Leu320Phe p.Phe342HisfsX28 p.Ala349Val p.Gln359Lys Thr360Lys p.Tyr301Cys p.Thr338Ile p.Arg347Pro p.Arg347His p.Leu375Phe p.Ala455Glu p.Leu467Phe p.Gln493Arg p.Gln493ValfsX10 p.Ile507del p.Phe508del p.Tyr515X

Location Promoter Promoter Exon 1 Exon 2 Exon 3 Exon 3 Exon 3 Exon 3 Exon 3 Exon 3 Exon 3 Exon 3 Intron 3 Exon 4 Exon 4 Exon 4 Exon 4 Exon 4 Exon 4 Exon 4 Exon 4 Intron 4 Exon 5 Exon 5 Exon 5 Intron 5 Exon 6 Exon 6 Exon 6 Exon 6 Exon 7 Exon 8 Exon 8 Exon 8 Exon 8 Exon 8 Exon 8 Exon 8 Exon 8 Exon 8 Exon 8 Exon 9 Exon 10 Exon 11 Exon 11 Exon 11 Exon 11 Exon 11 Exon 11

4.2 Affected Homozygous or Compound Heterozygous Conditions

105

Table 4.10 (continued) 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88 89 90 91 92 93 94 95 96 97 98 99 100 101

Legacy name V520F 1717-1G>A 1717- 2A>G G542X S549N G551S G551D R553X I556V R560T G576A E585X 1898+1G>A G622D 1898+ 3A>G 2184delA R668C 2183AA>G 2184insA N703S S737F P750L R785X 2622+1G>A E831X D836Y 2752-26A>G 2789+5G>A Q890X S945L 3007delG L967S D979A 3120G>A L997F 3199del6 I1027T Y1014C M1028R F1052V R1066C G1069R Y1092X E1104X A1136T I1139V D1152H R1158X R1162X 3659delC S1235R

cDNA name c.1558G>T c.1585-1G>A c.1585-2A>G c.1624G>T c.1646G>A c.1651G>A c.1652G>A c.1657C>T c.1666A>G c.1679G>C c.1727G>C c.1753G>T c.1766+1G>A c.1865G>A c.1766+3A>G c.2052delA c.2002C>T c.2051_2052delAAinsG c.2052_2053insA c.2108A>G c.2210C>T c.2249C>T c.2353C>T c.2490+1G>A c.2491G>T c.2506G>T c.2620-26A>G c.2657+5G>A c.2668C>T c.2834C>T c.2875delG c.2900T>C c.2936A>C c.2988G>A c.2991G>C c.3067_3072delATAGTG c.3080T>C c.3041A>G c.3083T>G c.3154T>G c.3196C>T c.3205G>A c.3276C>A c.3310G>T c.3406G>A c.3415A>G c.3454G>C c.3472C>T c.3484C>T c.3528delC c.3705T>G

Protein name p.Val520Phe

p.Gly542X p.Ser549Asn p.Gly551Ser p.Gly551Asp p.Arg553X p.Ile556Val p.Arg560Thr p.Gly576Ala p.Glu585X p.Gly622Asp p.Lys684AsnfsX38 p.Arg668Cys p.Lys684SerfsX38 p.Gln685ThrfsX4 p.Asn703Ser p.Ser737Phe p.Pro750Leu p.Arg785X p.Glu831X p.Asp836Tyr

p.Gln890X p.Ser945Leu p.Ala959HisfsX9 p.Leu967Ser p.Asp979Ala p.Leu997Phe p.Ile1023_Val1024del p.Ile1027Thr p.Tyr1014Cys p.Met1028Arg p.Phe1052Val p.Arg1066Cys p.Gly1069Arg p.Tyr1092X p.Glu1104X p.Ala1136Thr p.Ile1139Val p.Asp1152His p.Arg1158X p.Arg1162X p.Lys1177SerfsX15 p.Ser1235Arg

Location Exon 11 Intron 11 Intron 11 Exon 12 Exon 12 Exon 12 Exon 12 Exon 12 Exon 12 Exon 12 Exon 13 Exon 13 Intron 13 Exon 14 Intron 13 Exon 14 Exon 14 Exon 14 Exon 14 Exon 14 Exon 14 Exon 14 Exon 14 Intron 14 Exon 15 Exon 15 Intron 15 Intron 16 Exon 17 Exon 17 Exon 17 Exon 17 Exon 18 Exon 18 Exon 19 Exon 19 Exon 19 Exon 19 Exon 19 Exon 20 Exon 20 Exon 20 Exon 20 Exon 20 Exon 21 Exon 21 Exon 21 Exon 22 Exon 22 Exon 22 Exon 22 (continued)

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

106

Table 4.10 (continued) Legacy name 3849+10kbC>T 3849+40A>G 3905insT W1282X A1285V N1303K Q1352H(G>C)

102 103 104 105 106 107 108

a

Family Pedigree

MARKERS ORDER D7S486 D7S522 D7S633 D7S677 INTRON 1 INTRON 6 INTRON 8-1 CFTR D7S847

cDNA name c.3717+12191C>T c.3717+40A>G c.3773_3774insT c.3846G>A c.3854C>T c.3909C>G c.4056G>C

117 137 150 135 208 104 124 R117h 161

124 131 150 131 212 100 139 F508 161

1.1

Protein name

Location Intron 22 Intron 22 Exon 23 Exon 23 Exon 23 Exon 24 Exon 25

p.Leu1258PhefsX7 p.Trp1282X p.Ala1285Val p.Asn1303Lys p.Gln1352His

117 135 150 131 212 100 126 F508 161

1.2

2.1

2.2

PGT

PGT

R117H/N

R117H/N

122 131 148 125 217 104 124 N 149

b Sequential Polar Body Analysis 1

Oocyte #

PB1

PB2

117 135 150 131 212 100 126 F508 161

122 131 148 125 217 104 124 N 149 122 131 148 125 217 104 124 N 149

2

117 135 150 131 212 100 126 F508 161

3

122 131 148 125 217 104 124 N 149 122 131 148 125 217 104 124 N 149

122 131 148 125 217 104 124 N 149 117 135 150 131 212 100 126 F508 161

4

117 135 150 131 212 100 126 F508 161

122 131 148 125 217 104 124 N 149

117 135 150 131 212 100 126 F508 161

5

122 131 148 125 217 104 124 N 149 117 135 150 131 212 100 126 F508 161

9

117 135 150 131 212 100 126 F508 161 122 131 148 125 217 104 124 N 149

10

117 135 150 131 212 100 126 F508 161 122 131 148 125 217 104 124 N 149

11

117 135 150 131 212 100 126 F508 161

122 131 148 125 217 104 124 N 149 122 131 148 125 217 104 124 N 149

Predicted Genotype: AFFECTED

AFFECTED

NORMAL*

ET

NORMAL

AFFECTED*

NORMAL*

NORMAL*

AFFECTED

ET

Fig. 4.13  PGT for compound heterozygous CF male patient, with three different mutations in the CFTR gene performed, in which the paternal partner was affected carrying both delta F508 and R117H mutations in the CFTR gene. To avoid testing for two different mutations simultaneously in the same blastomere, taking into consideration up to 20% risk of ADO for each of the alleles tested, potentially leading to a misdiagnosis, PGT was was based on a sequential PB1 and PB2 analysis, to identify the

mutation-free oocytes. As a result of testing for Delta F508 maternal mutation, performed simultaneously with multiple closely linked markers, four mutation free oocytes were detected from the eight oocytes available for testing. Two of these embryos resulting from oocytes #3 and #4 with acceptable development potential were transferred, yielding a twin pregnancy and birth of two healthy baby girls, confirmed to be unaffected carriers of the paternal mutation

with requests also from affected CFTR patients, with both female and male partners being affected. As shown in Fig. 4.13, this may involve testing for two or even three mutations simultaneously. PGT seems to be the only choice in this case, as the male partner is compound heterozygous for Delta F508 and R117H and the female

partner a carrier of the Delta F508 mutation in CFTR gene. To avoid testing for two different mutations simultaneously in the same embryo biopsy sample, sequential PB1 and PB2 analysis was applied first to identify the mutation-free oocytes. As result of testing for Delta F508 maternal mutation, performed simultaneously

4.3 Concomitant PGT for Two or More Single-Gene Disorders

with multiple closely linked markers, four mutation-­free oocytes were detected from eight oocytes available for testing. Embryos resulting from oocytes #3 and #4 were transferred, yielding a twin pregnancy that resulted in the birth of two healthy baby girls confirmed to be unaffected carriers of the paternal mutation. Thus, PGT is of special value for couples with homozygous and compound heterozygous-affected partners, given only 50% chance of having an unaffected child and making invasive prenatal genetic diagnosis a daunting option. With progress in treatment of some genetic disorders, PGT-M will have an increasing impact on the decision of the affected and well-treated patients to reproduce. For example, life expectancy has been significantly improved for both CF and thalassemia, both representing the most frequent indications in the PGT-M practice. In each of these disorders, strategy depends on whether the affected partner is male or female. Testing may be restricted to oocyte testing if a male partner is affected. By contrast, embryo testing will be required if the female partner is affected. If the female partner is affected, as shown in Fig.  4.14, testing for the two different CFTR mutations is involved. In this case the female partner was a compound heterozygote (R117H and G542X) and the male partner a carrier of R117H mutation; PGT-M had to be based on embryo biopsy samples. To be able to test simultaneously for the required number of the linked markers, paternal and maternal haplotypes were first established, paternal using single-sperm PCR and the maternal by PB1 analysis. The couple had two previous pregnancies, the first resulting in spontaneous abortion of twins and the second terminated following prenatal diagnosis of affected fetus with CFTR.  Multiplex PCR performed for the two mutations in CFTR gene in this case was combined with age-related aneuploidy testing because of the mother’s advanced reproductive age. Multiplex heminested PCR analysis was performed on 14 embryos to allow simultaneous detection of the paternal and maternal CFTR haplotypes and non-syntenic short tandem repeats (STRs) located on chromosomes 13, 16, 18, 21, 22, and XY.  Six embryos were predicted to be

107

unaffected CFTR mutation carriers, based on the presence of the normal paternal CFTR gene. In addition to avoiding the transfer of affected compound heterozygous embryos, three aneuploid embryos were identified and excluded from transfer and freezing. Two unaffected embryos (embryos #6 and #7) were transferred, resulting in an unaffected pregnancy and birth of a healthy baby girl confirmed to be a carrier of maternal G542X mutation. DNA analysis of the baby revealed a genetic profile identical to embryo #7, evidencing the usefulness and ­accuracy of combined mutation, linkage, and aneuploidy analysis in patients of advanced reproductive age. Overall, of 44 PGT-M cycles performed for 25 CF-affected patients, including 7 female and 18 male partners, 72 unaffected embryos were identified for transfer in 36 cycles, resulting in 17 clinical pregnancies and births of 14 unaffected children with no misdiagnosis.

4.3

 oncomitant PGT for Two or C More Single-Gene Disorders

The higher proportion of some PGT-M performed for some indications may reflect their higher prevalence, such as PGT for thalassemia or CFTR. Thus, it is not unexpected that these couples presenting for PGT may incidentally be also at risk for some other conditions; thus, more complex PGT-M is required for two or more conditions using the same biopsy sample. We performed a total of 19 PGT-M cycles for CFTR in 12 couples also at risk for producing offspring with other genetic conditions, including Darier disease, facioscapulohumeral muscular dystrophy (FSHD), Aicardi–Goutieres syndrome, hereditary hemochromatosis, and Robertsonian translocations (Table  4.11). In these cycles, PGT-M resulted in preselection of 25 unaffected embryos for transfer, yielding 14 clinical pregnancies and birth of 12 healthy children free of both CFTR and each of the above conditions [38]. One of the above cases is presented in Fig.  4.15, demonstrating the results of PGT for Delta F508 mutation combined with testing for a Darier disease de novo mutation in ATP2A2.

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

108

a

Family Pedigree

b

Markers order: CFTR Intron 1 Intron 6 Intron 8-1 Intron 8-2 Mutation

109 101 139 176 N

115 105 124 172 1717

PGT 2

3

115 115 105 101 124 139 172 174 1717 G542X

Embryo #

Predicted Genotype Predicted Karyotype

1717/ G542X

13,13; 16,16; 18,18; 21,21; 22,22 XX

4

5

109 115 101 101 139 139 176 174 N G542X

115 111 105 105 124 124 172 172 1717 R117H

115 115 105 101 124 139 172 174 1717 G542X

N/ G542X

1717/ R117H

1717/ R117H

13,13; 13,13; 13,13,13; 16,16; 16,16; 16,16; 18,18; 18,18,18; 18,18; 21,21; 0 ,21; 21,21; 22,22 22,22 22,22 XY XY XXY

1.1

1.2

N / R117H

R117H / G542X

2.1 2.2

2.3

SAB 10 weeks

TAB 23 weeks

6

7

8

109 115 109 115 115 115 101 101 101 101 105 101 139 139 139 139 124 139 176 174 176 174 172 174 N G542X N G542X 1717 G542X

N/ G542X

13,13; 16,16; 18,18; 21,21; 22,22 XY

ET

111 115 105 101 124 139 172 174 R117H G542X

N/ G542X

13,13; 16,16; 18,18; 21,21; 22,22 XX

1717/ G542X

13,13; 16,16; 18,18; 21,21; 22,22 XY

2.4 PGT

9

10

109 115 101 101 139 139 176 174 N G542X

11

115 115 115 111 109 111 105 101 105 105 101 105 124 139 124 124 139 124 172 174 172 172 176 172 1717 G542X 1717 R117H N R117H

1717/ G542X

13,13; 16,16; 18,18; 21,21; 22,22 XX

1717/ R117H

13,13; 16,16; 18,18; 21,21; 22,22 XY

13

14

115 111 115 115 105 105 105 101 124 124 124 139 172 172 172 174 1717 R117H 1717 G542X

N/ R117H

1717/ R117H

13,13; 16,16; 18,18; 21,21; 22,22 XX

13,13; 16,16; 18,18; 21,21; 22,22 XY

1717/ G542X

13,13; 16,16; 18,18; 21,21; 22,22 XX

15

17

109 111 109 111 101 105 101 105 139 124 139 124 176 172 176 172 N R117H N R117H

N/ R117H

13,13; 16,16; 18,18; 21,21; 22,22 XX

N/ R117H

13,13; 16,16; 18,18; 21,21; 22,22 XX

ET

Fig. 4.14  PGT for a couple with female affected double heterozygous partner with two different CFTR mutations combined with aneuploidy testing. (a) Family pedigree of a couple with three different mutations in CFTR gene. The mother is affected with cystic fibrosis (CF) and had two different mutations (R117H and G542X). The father is a carrier of 1717 mutation in CFTR gene. Paternal haplotype was established by multiplex heminested singlesperm PCR.  Maternal linkage was based on DNA amplification of polar bodies. Marker order of the mutations and polymorphic markers in CFTR gene are shown on the upper left. The first pregnancy with twins resulted in spontaneous abortion and the second pregnancy in an affected fetus which was terminated; PGT was performed for three different mutations in CFTR gene, resulting in an unaffected pregnancy and the birth of a healthy girl predicted and confirmed to be the carrier of maternal mutation G542X. (b) Outcome of PGT cycle for three

mutations in CFTR gene combined with age-related aneuploidy testing. Multiplex heminested PCR performed on blastomeres from 14 embryos allowing simultaneous detection of the paternal and maternal CFTR haplotypes and non-syntenic short tandem repeats (STRs) located on chromosomes 13, 16, 18, 21, 22, and XY.  Six embryos (#3, #6, #7, #11, #15, and #17) were predicted to be carriers based on the presence of the maternal mutation and normal paternal CFTR gene. Monosomy 21 was found in the blastomere from embryo #3, which was excluded from the transfer and freezing. Trisomy 18 and XXY was observed in the blastomere from embryo #4, and trisomy 13 was detected in the blastomere from embryo #5. Embryos #6 and #7 were transferred, resulting in pregnancy and the birth of a healthy girl. DNA analysis of the newborn baby revealed a genetic profile identical to that of embryo #7

PGT-M was based on sequential PB1 and PB2 testing for both mutations, followed by biopsy of the embryos resulting from mutant oocytes to detect carrier embryos. Of 15 oocytes tested by sequential PB1 and PB2, only 2 (oocytes #9 and #14) were free of both mutations; embryos resulting from these oocytes were transferred, resulting in a singleton pregnancy and birth of a healthy baby girl free of both diseases. Sequential blasto-

mere analysis of the four embryos resulting from the oocytes with DF508 mutation and without ATP2A2 deletion allowed preselection of one embryo (embryo #2) – a carrier of DF508 mutation, frozen for future use by the couple. Sequential PB1 and PB2 analysis, followed by embryo biopsy, was also applied for combined PGT for Delta F508 mutation and FSHD, as well as for Delta F508 and hereditary hemochromato-

CF + other disorders Aicardi–Goutieres syndrome 5; AGS5 Spinal muscular atrophy, type I; SMA1 Facioscapulohumeral muscular dystrophy 1; FSHD1 Hemochromatosis, type 1; HFE1 Darier–White disease; DAR Fragile X Familial Mediterranean fever; FMF Huntington disease; HD Phenylketonuria; PKU Duplication 22q11 Translocation Total

OMIM 612952 253300 158900 235200 124200 300624 249100 143100 261600

Gene SAMHD1 SMN1 FRG1 HFE ATP2A2 FMR1 MEFV HTT PAH

1 1 1 1 1 1 1 2 12

# of patient 1 1 1 1 1 1 1 1 1 2 2 19

# of cycle 2 3 4

Table 4.11  Combined PGT-M for cystic fibrosis (CFTR gene) and additional genetic disorders

1 1 1 1 1 0 1 2 16

# of transfers 2 2 4 1 1 1 2 3 0 2 3 25 (1.47)

# of embryos transferred 3 3 6 0 1 1 1 2 0 2 1 14 73%

Pregnancy 2 2 2

0 0 0 0 1 0 1 0 4 28

SAB 1 1 0

0 1 1 1 1 0 1 1 10 72%

0 1 1 1 1 0 1 2 12

Delivery Birth 1 1 1 2 2 2

4.3 Concomitant PGT for Two or More Single-Gene Disorders 109

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

110

Family Pedigree MARKERS ORDER FOR CFTR INTRON1 INTRON6 INTRON 8-1 INTRON 8-2 DF508 INTRON 17b

225 124 122 N

115 100 126 174 DF508 98

205 122 124 N

ATP2A2

117 104 124 172 N 106

209 134 139 N

CFTR

117 115 104 100 124 139 174 174 N DF508 98 98

225 124 122 DEL

ATP2A2

MARKERS ORDER FOR DARIER DISEASE: D12S1583 D12S1328 D12S1645 ATP2A2- 4bp DEL

CFTR

PGT NORMAL DARIER; NORMAL CF

Sequential Polar Body Ananlysis Oocyte #1

2

3

4

5

6

7

8

9

12

13

PB1 - FA

14

16

17

19

Predicted Genotype:

CFTR

ATP2A2

115 100 139 DF508 98

117 104 124 N 98

117 104 124 N 98

115 100 139 DF508 98

115 100 139 DF508 98

115 100 139 DF508 98

117 104 124 N 98

115 100 139 DF508 98

117 104 124 N 98

115 100 139 DF508 98

209 134 139 N

225 124 122 N

225 124 122 DEL

209 134 139 N

225 124 122 DEL

225 124 122 DEL

225 124 122 DEL

209 134 139 N

209 134 139 N

225 124 122 DEL

1

2

3

4

5

6

7

8

9

115 100 139 DF508 98

209 134 139 N

225 124 122 DEL

ET

ET

Blstomeres Analysis EMBRYO#

PB1 - FA

117 104 124 N 98

12

13

17

115 100 139 DF508 98

115 100 139 DF508 98

225 124 122 DEL

225 124 122 DEL

19

Predicted Genotype:

CFTR 115

115 100 100 139 126 174 174 DF508 DF508 98 98 225

ATP2A2 124 122 N

209 134 139 N

AFFECTED CF NORMAL DARRIER

115 100 126 174 DF508 98 225 124 122 N

117 104 124 174 N 98 209 134 139 N

CARRIER CF NORMAL DARRIER

115 115 100 100 139 126 174 174 DF508 DF508 98 98 225 124 122 N

209 134 139 N

AFFECTED CF NORMAL DARRIER

115 115 100 100 139 126 174 174 DF508 DF508 98 98 225 124 122 N

115 115 100 100 126 139 174 174 DF508 DF508 98 98 225 124 122 N

209 134 139 N

AFFECTED CF NORMAL DARRIER

209 134 139 N

AFFECTED CF NORMAL DARRIER

FROZEN

Fig. 4.15  PGT for Delta F508 mutation combined with testing for Darrier disease de novo mutation in ATP2A2. PGT was based on sequential PB1 and PB2 testing for both mutations, followed by blastomere biopsy of the embryos containing the CFTR mutation for possible detection of unaffected carrier embryos. Of 15 oocytes tested by sequential PB1 and PB2, only two (oocytes #9 and #14) appeared to be free of both mutations, so the

embryos resulting from these oocytes were transferred, resulting in a singleton pregnancy and birth of a healthy baby girl, free of both diseases. A sequential blastomere analysis of the four embryos resulting from the oocytes with DF508 mutation and without ATP2A2 deletion allowed preselecting one embryo (embryo #2) – a carrier of DF508 mutation, frozen for future use by the couple

sis (C282Y mutation in HFE gene). PGT-M in the latter case allowed preselecting two of five embryos; one without C282Y mutation was a carrier of Delta F508; one resulted from Delta F508 mutation-free oocyte. Concomitant PGT for Delta F508 and FSHD in the former case resulted in detecting two of eight embryos free from FSHD deletion for transfer, yielding a singleton pregnancy and birth of unaffected carrier of Delta F508 mutation, free of FSHD. Two PGT-M cycles were performed for CFTR and Aicardi– Goutieres syndrome using trophectoderm biopsy. This resulted in an unaffected pregnancy, free from Delta F508 and Aicardi–Goutieres syndrome. A PGT cycle for two CFTR mutations

combined with testing for spinal muscular atrophy (SMA) is presented in Fig. 4.16. Concomitant PGT for two different genetic disorders has been described in a childless Ashkenazi Jew couple at risk for producing offspring with Tay–Sachs (TS) and Gaucher disease (GD); each parent carried different mutations in both hexosaminidase A (HEX A) and B glucocerebrosidase (GBD) genes [39]. Six embryos were analyzed: one wild type for both TS and GD, three wild types for GD and a carrier of TS, and two compound heterozygotes for TS.  Two of the four transferable embryos that developed into blastocysts were transferred, resulting in a singleton pregnancy

4.3 Concomitant PGT for Two or More Single-Gene Disorders

111

Family Pedigree MARKERS ORDER: D5S2019 D5S435 D5S1414 D5S1556 SMN1 D5S610 D5S351 D5S1491 D5S2122

124 135 151 131 212 ∆F508 187 139

162 145 202 129 DEL 112 127 151 118

122 131 148 125 214 N 182 164

CFTR GENE

37y.o.

137 156 204 127 N 112 127 151 122

SMN1 GENE

143 137 158 160 190 208 127 121/135 N DEL 105 103 125 123 151 146 120 122

SMN1 GENE

124 135 148 125 214 Y1424 187 160

119 137 153 133 222 N703S 136 160

CFTR GENE

MARKERS ORDER: D7S486 D7S522 D7S633 D7S677 INTRON1 ∆F508 N703S INTRON17b Y1424 D7S847

PGT

PGT SINGLE SPERM HAPLOTYPING

SEQUENTIAL POPAR BODY ANALYSIS Cystic fibrosis & SMA

Normal for SMA ∆F508 & N703S CF*

Carrier for SMA ∆F508 & Y1424Y variant CF

1

OOCYTES

3

137 124 160 135 208 148 121/135 125 N 214 103 Y1424 123 187 146 160 122

Normal SMN1 Y1424Y variant CF BLASTOMERE ANALYSIS FOR Mutations in CFTR, SMN1 genes and aCGH

162 145 202 129 DEL 112 127 151 118

124 124 135 135 151 148 131 125 212 214 ∆F508 Y1424 187 187 160 139

46, XY

ET

143 158 190 127 DEL 105 125 151 120

119 137 153 133 222 N703S 136 160

Monosomy 5 Affected N703S

1

EMBRYO #

137 160 208 121/135 N 103 123 146 122

0

5

0

124 119 135 137 151 153 131 133 212 222 ∆F508 N703S 136 187 160 139

45,XY(-5)

124 135 148 125 214 Y1424 187 160

Affected SMN1 Y1424Y variant CF

3

162 145 202 129 DEL 112 127 151 118

6

5

162 145 202 129 DEL 112 127 151 118

143 124 158 135 190 151 127 131 DEL 212 105 ∆F508 125 187 151 139 120

46,XX

7

143 158 190 127 DEL 105 125 151 120

137 119 160 137 208 153 121/135 133 N 222 103 N703S 123 136 146 160 122

Normal SMN1 Affected N703S

Affected SMN1 Y1424Y variant CF

6

124 135 148 125 214 Y1424 187 160

137 137 124 119 160 156 135 137 208 204 151 153 127 121/135 131 133 N N 212 222 103 ∆F508 N703S 112 123 127 136 187 146 151 160 139 122 122

46,XX

ET*

124 135 148 125 214 Y1424 187 160

7

137 156 204 127 N 112 127 151 122

143 158 190 127 DEL 105 125 151 120

124 124 135 135 151 148 131 125 212 214 ∆F508 Y1424 187 187 160 139

46XY,(+10,-16)

Fig. 4.16  Concomitant PGT for SMA, Delta F508, and 24-chromosome aneuploidy testing. Both partners are carriers of SMN1 gene, while father is also a carrier of DF508 and mother also the carrier of N703 S CFTR mutation. Sequential PB1 and PB2 testing was performed to detect the oocytes free of CFTR and SMN1 mutations, with further testing of the resulting embryos for 24-chromosome aneuploidy. Of five oocytes available for testing, only one (oocyte #1) appeared to be free of both SMN1 and N703S mutations, while others were either with CFTR or SMN1 mutations. Testing of the resulting embryos confirmed unaffected status of the embryo result-

ing from oocyte #1, which was also 24-chromosome aneuploidy free. The embryos resulting from the oocytes # 2 and #7, although unaffected carriers of CFTR and SMA mutations, appeared to be with monosomy 5 (embryo # 2), and trisomy 10, and monosomy 16 (embryo # 7). The embryo resulting from oocyte #5 was affected for SMA and carrier of DF508 mutation in CFTR. Finally, the embryos resulting from the oocyte #6 appeared to be unaffected carrier of DF508 mutation and free of aneuploidy. Two embryos (embryos #1 and #6) were transferred, resulting in a twin pregnancy and birth of two unaffected carriers of SMA and CFTR

and birth of a healthy child free from both conditions. Concomitant PGD was also performed for Charcot–Marie–Tooth (CMT) and Fabry diseases in a couple in which both partners carried the Fabry mutation; the male partner had also CMT disease. Testing was performed for Fabry disease by sequential PB1 and PB2 analysis followed by embryo biopsy and testing for CMT.  This allowed identification and transfer of an unaffected embryo resulting in a triplet pregnancy and birth of three healthy children that appeared to be monozygotic triplets.

Although the mechanism for formation of monozygotic triplets in this case is not understood, data nonetheless showed that concomitant PGT for more than one condition is feasible and may be performed using the combination of different biopsy techniques, allowing accurate detection of both conditions. A similar approach has been used for concomitant PGT cycles for BRCA1 and SMA and for BRCA2 and MEN1. Testing for both mutations in each of these cycles allowed identification of two unaffected embryos for transfer in each case, to be described below.

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

112

Of special interest is a combined simultaneous PGT for five different conditions [40]. This consanguineous couple presented for PGT-M to avoid the risk of producing another affected child homozygous for four different autosomal recessive disorders identified in their previous offspring. The disorders were (1) early infantile epileptic encephalopathy 5 (EIEE5), caused by SPTAN1 mutation; (2) xeroderma pigmentosum, complementation group C (XPG), caused by ERCC5 mutation; (3) merosin-deficient congenital muscular dystrophy 1A (MDC1A), caused by LAMA2 mutation; and (4) phenylketonuria (PKU) caused by PAH mutation. Parents requested aneuploidy testing as well. Thus, the

PGT strategy was to combine PGT-A and PGT-­ M, involving mutation and linked marker testing by multiplex nested PCR to avoid the undetected ADO of each of the genes tested. Overall, 12 of 16 embryos reaching the blastocyst stage were tested together with PGT-A; ten were affected, including six affected by one mutation and four by two mutations (Fig. 4.17). The remaining two embryos were each carriers for all four mutations; one had trisomy 13, but the other was euploid. Transfer of this embryo resulted in a healthy unaffected baby. Although the cumulative risk for producing an offspring affected by all four conditions was only 0.4% (1/256), the couple still had been unfortunate enough to pro-

Family pedigree CHANCES TO FIND EMBRYO FREE FROM PKU ¾ FREE FROM EIEE5 ¾ FREE FROM XPG FREE FROM MDCA1

¾ ¾

FREE FROM ALL CONDITIONS: ¾ X ¾ X ¾ X ¾ = 81/256 (31.6%) AFFECTED FOR ALL 4 CONDITIONS: ¼X¼ X¼ X¼= 1/256

(0.4%)

DOB3/1988

PAH SPTAN1 ERCC5 LAMA2 IVS1+3 R2142C R69X R2352C

PAH SPTAN1 ERCC5 LAMA2 IVS1+3 R2142C R69X R2352C

PGT PAH SPTAN ERCC5 LAMA2 IVS1+3 N R69X R2352C

AFFECTED FOR 4 CONDITIONS

1

3

4

5

6

CARRIER

AFFECTED

CARRIER

7

8

9

10

AFFECTED

CARRIER

NORMAL

AFFECTED

NORMAL

CARRIER

NORMAL

CARRIER

CARRIER

11

12

CARRIER

AFFECTED

AFFECTED

CARRIER

NORMAL

AFFECTED AFFECTED

16

PAH12q22-24.2 AFFECTED

CARRIER

CARRIER

SPTAN1 9q34.11 CARRIER

NORMAL

AFFECTED NORMAL

CARRIER

CARRIER

CARRIER

CARRIER

CARRIER

CARRIER

CARRIER

AFFECTED

AFFECTED

CARRIER

CARRIER

CARRIER

CARRIER

NORMAL

AFFECTED

CARRIER

46,XY

46,XY

NORMAL

ERCC5 13q33.1 CARRIER

LAMA2 6q22-23

NGS

0

45, XX,-22

46,XY

46,XX

46,XX

47,XX+13

48,XX+21,+22

AFFECTED

47XY,+22

-6 44,XX-2,-6

NORMAL

46,XX

NORMAL

47,XY+13

FET

Fig. 4.17  PGT for four conditions together with PGTA.  A total of 12 of 16 embryos reaching the blastocyst stage were tested together with NGS for PGT-A, of which ten were affected, including six affected by one mutation and four by two mutations. Only two embryos were unaf-

fected carriers of all four gene mutations (embryos #3 and #6), of which one (embryo 6) was with trisomy 13. Thus, only a single embryo euploid and carrier of all four gene mutations (embryo #3) was transferred, resulting in birth of a healthy unaffected baby

4.4 De Novo Mutations

duce such a child in a natural conception cycle. Accordingly, the chance for detecting an unaffected embryo was 31.6%, and taking into consideration the additional 50% risk for aneuploidy, only 15.8% chance may be expected to identify unaffected embryo for transfer. Still one of 12 embryos tested was both euploid and a normal carrier for all the mutations, resulting in a healthy, unaffected child. This is the world’s first PGT for five different genetic conditions (EIEE5, XPG, MDC1A, PKU, and aneuploidy) in a single test, resulting in a transfer of euploid embryo free of all the conditions tested, demonstrating feasibility and accuracy of simultaneous combined PGT for multiple genetic conditions. The above cases demonstrate feasibility and advantages of analyzing a large number of markers in a single multiplex reaction allowing the analysis of multiple diseases in cases where couples are carriers of mutations in several genes. It is particularly common in the Ashkenazi Jewish population, in which there are a large number of prevalent autosomal recessive diseases. Similar circumstances arise in populations of high consanguinity.

4.4

De Novo Mutations

With the increasing number of different genetic disorders for which PGT-M is being applied each year, PGT is 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. Performing PGT-M is facilitated by knowledge of sequence information for Mendelian diseases; however, PGT-M may also be performed when the exact mutation is not known, again through the application of the linkage analysis [41–43]. With expanding use of polymorphic markers, linkage analysis may allow PGT for any genetic disease irrespective of the availability or not of the mutation-specific sequence. This approach is more universal, mak-

113

ing it possible to track the inheritance of the mutation without actual testing or knowing the gene itself. The approaches described above cannot, however, be applied in cases of de novo mutations (DNM) in parent(s) or affected children, when neither origin nor relevant haplotypes can be determined in single cells biopsied from embryos or oocytes. On the other hand, with the improved awareness of PGT, increasing numbers of couples request PGT for a genetic disease first diagnosed in one of their parents or for an autosomal dominant disorder manifested in their own affected child. Thus, PGT strategies for genetic conditions arising by de novo mutations are presented below. Our experience is the world’s largest series, including  526 PGT-M cycles for 283 couples with 82 different DNMs. The majority of PGT cycles were performed for dominant mutations (480 cycles for 257 patients), with only 14 for autosomal recessive and 19 for X-linked DNM (Table 4.12) [44]. First performed in 2000, numbers of annual cases of PGT for DNM have steadily increased, reaching over 150 cases performed only in the last 2 years (Fig. 4.18). It is of interest that despite the expected predominance of dominant DNM of a paternal origin, reflecting an increasing proportion of older paternal partners, DNM of both paternal and maternal origin was observed (Tables 4.12 and 4.13). To perform DNM of paternal origin, the DNM was first investigated using paternal DNA from blood and total sperm, followed by single-sperm typing to determine the proportion of sperm with DNM and relevant normal and mutant haplotypes. Maternal DNM presents a greater challenge, frequently requiring polar body analysis to determine maternal haplotypes. Another major challenge arises from gonadal mosaicism. In addition, up to 10% of the PGT-M tested DNM (35 cases) were first detected in affected children, with no evidence of detectable mutation in either parent despite the finding the corresponding mutant haplotype associated with a normal parental haplotype.

Conditions Autosomal-dominant Achondroplasia Alzheimer disease 3 Angelman syndrome Angioedema, hereditary, type I Aniridia Arthrogryposis Arthrogryposis, distal, type 2B Axenfeld–Rieger syndrome, Basal cell nevus syndrome; BCNS (Gorlin) Benign chronic pemphigus Blepharophimosis Brachydactyly, type B1 Branchiooculofacial syndrome Breast–ovarian cancer Cardiomyopathy, dilated Cardiomyopathy, familial hypertrophic, 4; CMH4 Carney complex (mosaic) Charcot–Marie–Tooth disease, CMT1A Charcot–Marie–Tooth disease, CMT2A2 Cleidocranial dysplasia Colorectal cancer, hereditary nonpolyposis, type 2; HNPCC2 Colorectal cancer, hereditary nonpolyposis, type 4; HNPCC4 Craniofrontonasal syndrome 1 5 1

2 1

SCN4A PMP22

MFN2

RUNX2 MLH1

PMS2

EFNB1

1 2

1 1

1 2 1 1 1 4

4 1

ATP2C1 FOXL2 ROR2 TFAP2A BRCA1,2 MYH7 MYBPC3

FGFR3 PSEN1 UBE3A SERPING1 PAX6 FBN2 TNNI3 PITX2 PTCH1

Gene

1

1 1

3

1 3

1

1 1

1

2

1

2

Origin of de novo mutation Detected in Paternal Maternal child

Table 4.12  Different de novo mutations for which PGT was performed

1

1

1 3

1

1 8

1 1 1 1 3 1 2

6 1 2 1 3 1 1 3 5

# of patients with PGT-M

1

2

3 10

4

10

1 1 3 1 7 3 3

9 1 2 1 6 2 2 13 6

# of cycles

1

1

2 7

2

12

1 1 3 0 7 2 4

10 1 2 1 5 2 1 12 6

# of transfers

1

0

4 15

3

14

2 1 4 0 8 2 5

14 1 3 1 5 2 2 14 9

# of embryos transferred

1

0

1 4

1

7

0 0 2 0 3 1 3

7 1 1 1 3 2 0 5 3

1

0

1

2

Pregnancy SAB

1

0

1 4

1

7

0 0 2 0 3 1 2

5 1 1 1 3 2 0 4 3

Birth

114 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Dystonia 1, torsion Emery–Dreifuss muscular dystrophy 2 Exostoses, multiple, types I and II Facioscapulohumeral muscular dystrophy 1 Familial adenomatous polyposis Familial cold autoinflammatory syndrome Holt–Oram syndrome Hyperkeratosis Larsen syndrome Li–Fraumeni Syndrome 1 Loeys–Dietz syndrome 1 Long QT syndrome 8; LQT8 Lynch syndrome I Marfan syndrome Mental retardation, autosomal dominant 35; MRD35 Multiple endocrine neoplasia Multiple endocrine neoplasia, type IV Myotonia congenita Myotonic dystrophy 1 Nail–patella syndrome Neurofibromatosis, types I and II Neutropenia, severe congenital Noonan syndrome 1 Optic atrophy 1

Crouzon syndrome Cutis laxa Darier–White disease Diamond–Blackfan anemia all types

MEN 1,2 CDKN1B CLCN1 DMPK LMX1B NF1,2 ELANE PTPN1 OPA1 2 2

3 1 1 3 1 13

1 12

1

22

1

1

2 1 1 1 6

1 3

1 4 1 1 4 1 36 3 3 2

1 1 1 4 1 1 2 18 1

1 1 1 2

11 1

6

5 1

APC NLRP3

TBX5 KRT1 FLNB TP53 TGFBR2 CACNA1C MSH2 FBN1 PPP2R5D

6 3

2

4 3

1 1

EXT1,2 FRG1

9

6 1 1 10 5 2

4

5 2

2 1

FGFR2 ELN ATP2A2 RPS10,19, 20 TOR1A LMNA

15 1 1 7 1 64 4 5 2

1 3 2 1 4 38 1

4

22 1

10 9

11 5

11 4 1 18

10 1 2 5 1 53 6 5 2

1 3 2 1 5 45 2

5

20 1

10 6

11 4

8 2 1 14

16 1 4 8 1 84 5 5 4

1 3 4 1 5 69 2

4

30 1

17 10

17 9

13 3 2 20

5 1 1 2 1 32 3 3 0

1 2 2 1 3 25 1

2

10 1

8 3

6 2

6 2 1 8

1 2 2 1 2 21 1

2

9 1

7 3

4 2

5 2 1 7

4 1 1 2 1 3 29 1 2 1 2 0 (continued)

1

1 4

1

1

2

1

1

4.4 De Novo Mutations 115

Conditions Osteogenesis imperfecta all types Periventricular nodular heterotopia Xl Peutz–Jeghers syndrome Pfeiffer syndrome Polycystic kidney disease 1 and 2 Popliteal pterygium syndrome Rap guanine nucleotide exchange factor 6 Renal cysts and diabetes syndrome mosaic (4–19) Retinoblastoma Rett syndrome Sotos syndrome 1 Spinocerebellar ataxia 6 Telangiectasia, hereditary hemorrhagic Telangiectasia, hereditary hemorrhagic Treacher Collins syndrome 1 Treacher Collins syndrome 2 Tuberous sclerosis 1 and 2 Ulnar–mammary syndrome Von Hippel–Lindau syndrome Subtotal Autosomal recessive Hypophosphatasia AR Ataxia–telangiectasia Cystic fibrosis Fanconi anemia, FANCA Fanconi anemia, FANCI

Table 4.12 (continued)

ALPL1 ATM CFTR FANCA FANCI

TCOF1 POLR1D TSC1,2 TBX3 VHL

ENG

RB1 MECP2 NSD1 CACNA1A ACVRL1

HNF1B

Gene Col1A1,2 FLNA STK11 FGFR1 PKD1,2 IRF6 RAPGEF6

1 1 1 2 1

5 134

6

2

1 2 1

1

101

2 1 6 1

1

4

5

22

1 2

1

Origin of de novo mutation Detected in Paternal Maternal child 12 8 1 1 2 2 5 2 1 1

1 1 1 2 1

4 1 12 1 5 257

1

10 2 2 2 1

1

# of patients with PGT-M 21 1 2 2 7 1 1

2 3 1 5 2

4 1 21 3 9 480

1

21 3 3 3 3

# of cycles 42 3 5 2 16 1 2

2 2 1 4 2

4 1 22 3 9 442

0

21 3 2 2 5

# of transfers 32 2 4 2 17 1 3

2 3 2 6 3

7 0 34 4 10 642

0

33 3 2 4 6

# of embryos transferred 50 3 5 4 24 1 3

2 2 1 2 0

4 0 9 1 5 245

0

7 2 2 1 2

1

32

2

1 1

Pregnancy SAB 17 2 2 1 3 1 2 7 1 0 3 2

2 1 1 2 0

4 0 7 1 5 213

0

6 1 2 1 2

Birth 15 1 2 2 6 0 1

116 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Spinal muscular atrophy, type I; SMA1 Subtotal X-linked condition Epileptic encephalopathy Granulomatous disease, chronic Muscular dystrophy Ectodermal dysplasia Charcot–Marie–Tooth disease, Incontinentia pigmenti Hypophosphatemic rickets Wiskott–Aldrich syndrome Subtotal Total

CDKL5 CYBB DMD EDA GJB1 IKBKG PHEX WAS

SMN1

2 143

1

1

4 105

3

1

1 13 35

3

1 3 5 1 1 6 1 1 19 283

7

7 1 3 5

1

1

1 7 6 1 1 13 1 2 32 526

14

1

1 7 5 1 2 9 0 2 27 481

12

1

2 9 8 1 2 14 0 3 39 699 1.45

18

2

1 3 4 1 1 5 0 2 17 270 56.1%

8

1 7

1 1 4 1 1 5 0 1 1 3 14 36 234 13.3% 86.7% 2

1

1

4.4 De Novo Mutations 117

118

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Fig. 4.18 Steady increase of PGT cycles for de novo mutations after first description in 2000

160 140 120 100 80 60 40 20 0

2000–2003 2004–2006 2007–2009 2010–2012 2013–2015 2016–2018 284 patients

PGT strategies required for these families differ depending on the origin of DNM. An extensive DNA analysis of the parents and affected children is required prior to PGT, with mutation verification, polymorphic marker evaluation, whole- and single-sperm testing for paternal haplotypes, and PB analysis in order to establish the normal and mutant maternal haplotypes. Without this PGT for DNM cannot be performed (Fig. 4.19). When DNM is detected first in children, the mutation is first verified in their whole blood DNA.  Performing PGT-M for DNM required extensive preparatory DNA work before performing the actual PGT. Additional tests include paternal DNA from blood, total and single sperm, and potentially performing sequential PB1 and PB2, followed by blastocyst analysis. Overall, PGT for DNM in our experience has resulted in preselection and transfer of 699 DNM-free embryos in 481 cycles (average of 1.45 embryos per transfer), yielding 270 (56.1%) unaffected pregnancies and birth of 234 healthy children confirmed to be free of the DNM tested (Tables 4.12 and 4.13). An example of PGT strategy and design for DNM of dominant inheritance is presented below for a couple in which a NF2 splicing mutation (c114 + 2T-C) was detected in the hus-

535 cycles

82 genes

band who had no previous family history of the disease (Fig.  4.20). DNA analysis in paternal blood confirmed the presence of NF2 splicing mutation (c114 + 2T-C), while testing of single sperms showed gonadal mosaicism, represented by three types of sperms corresponding to three different haplotypes. Only 30% of sperm were represented by the mutant haplotype, 36% were normal characterized by a normal haplotype, and 34% contained a normal allele within the mutant haplotype. Strategies differ depending on the type of DNM inheritance, but the most commonly applied approach involves single-sperm typing (Fig. 4.19). This has been performed in approximately 50% of cases: we test at least 15 single sperms per patient and as many as 50 per patient to exclude gonadal mosaicism. Even if DNM is not identified, single-sperm typing may identify a “benign” mutant haplotype, characterized by a mutant haplotype without DNM.  It is an important requirement to identify the relevant linked markers in both parents, even though only one is a DNM carrier. Although not always possible, PGT by the PB approach to detect or differentiate maternal normal from maternal mutant haplotypes is always the method of choice, performed in one-third of our maternal DNM cases.

119

4.4 De Novo Mutations Table 4.13  Summary OF PGT-M outcome for de novo mutations # of Type of inheritance patients Autosomal Dominant 257 (67) Autosomal Recessive (6) 7 X-linked (8) 19 Total 283

a

# of cycles 480

# of transfers 442

# of embryos 642

# of pregnancies 245

SAB 32

14 32 526

12 27 481

18 39 699 1.45

8 17 270 56.1%

1 7 3 14 36 234 13.3% 86.7%

b

Paternal

Maternal

MUTATION CONFIRMATION AND

MUTATION CONFIRMATION :

POLYMORPHIC MARKERS EVALUATION ON DNA ROM BLOOD

ON DNA FROM BLOOD

c

Delivery 213

Detected in child

MUTATION DETECTION AND POLYMORPHIC MARKERS EVALUATION ON DNA FROM BLOOD

ON DNA FROM TOTAL SPERM

ON DNA FROM TOTAL SPERM

MUTATION TESTING ON SINGLE LIMPHOCYTES

SINGLE SPERM HAPLOTYPNG (15-50) 120 115 135 N 190 201 155

128 112 143 M 184 210 155

198 125 167 N 117 132 176

SINGLE SPERM HAPLOTYPNG

PGD DESIGN

(198 (125 (167 (N (115 (132 (176

198 129 169 N 117 138 178

201) 127 ) 165) M) 112) 136) 176)

MUTATION TESTING ON SINGLE LIMPHOCYTES

PGT

PGT

Polar Body analysis (to detect gonadal mosaicism)

Normal

Blastocyst confirmation

Polar Body analysis is highly recommended 120 115 135 N 190 201 155

128 112 143 M 184 210 155

PGT

122 117 141 N 192 203 155

126 112 145 N 192 205 155

Normal

Affected

Blastocyst testing is possible

Fig. 4.19  PGT strategies for DNM of different origin. (a) Case workout for DNM detected in male partner: (1) Pedigree in two generations. (2) Mutation verification in DNA extracted from blood and total sperm. (3) Amplification of partner’s single sperm to establish normal and affected haplotypes required for PGT cycle preparation. (4) Amplification of patient’s DNA to identify the most informative markers for PGT. (5) PGT by embryo biopsy for combined mutation and linkage analysis. (b) Case workout 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 analyzed on a single sperm for more accurate embryo genotype predic-

tion. (3) PGT by PB1 and PB2 analysis to identify DNMfree oocytes and establish maternal haplotypes, followed by blastomere or blastocyst analysis to confirm the diagnosis. (c) Case workout 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 on DNA extracted from parents’ whole blood and total sperm. (3) Mutation evaluation on single lymphocytes and single-sperm testing to rule out paternal gonadal mosaicism. (4) PGT by polar body analysis to detect potential maternal gonadal mosaicism. (5) Blastomere or blastocyst analysis to confirm the absence of the mutation

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

120

a

Family Pedigree 92 118 8 114 14 2 204 138 N 129 9 12 24 124

102 11 112 10 107 2 212 4 142 M 127 12 128

a

104 120 114 200 134 N 129 122

102 112 107 2 212 142 N* 127 128

b

c

36% 29%

34%

PGT

PGT

Markers order D22S1144 D22S1163 D22S535 D22S689 D22S1150 NF2 gene IVSI (GT)n IVS10(GT)n

102 126 114 216 153 N 127 122

PGT

NORMAL

b

PGT

CYCLE #1 Embryo # 4

92 118 114 204 138 N 129 124

Predicted Genotype:

102 126 114 216 153 N 127 122

5

92 118 114 204 138 N 129 124

N

104 120 114 200 134 N 129 122

102 112 107 212 142 N* 127 128

N ET

c

6

102 126 114 216 153 N 127 122

N* TE-N*

7

92 118 114 204 138 N 129 124

8

102 126 114 216 153 N 127 122

92 118 107 212 142 N* 127 128

N ET

102 126 114 216 153 N 127 122

N* TE-N*

PGT CYCLE #2 2

102 112 107 212 142 N* 127 128

102 126 114 216 153 N 127 122

N* TE-N*

3

92 118 114 204 138 N 129 124

102 126 114 216 153 N 127 122

N ET

4

92 118 114 204 138 N 129 124

5

104 120 114 200 134 N 129 122

N ET

102 112 107 212 142 N* 127 128

6

104 120 114 200 134 N 129 122

N*

22,0

TE-N*

Fig. 4.20  PGT for DNM in NF2 gene (c114+ 2 T-C splicing mutation) of paternal origin. (a) Family pedigree showing that DNM in NF2 gene was first detected in the father. Single-sperm analysis via multiplex heminested PCR revealed gonadal mosaicism with three different haplotypes: a– normal; b– mutant containing c.114  +  2  T-C allele; and c– mutant without c.114 + 2 T-C allele in NF2. Maternal linkage was based on DNA amplification of blastomeres in PGT cycle. Mutation and marker order are printed in upper left corner. (b) Outcome of the first PGT 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 dropout (ADO) of the dominant mutation in single blastomere. Trophectoderm (TE) biopsy

92 118 114 204 138 N 129 124

7

102 112 107 212 142 N* 127 128

102 126 114 216 153 N 127 122

N*

9

92 118 114 204 138 N 129 124

N

104 120 114 200 134 N 129 122

11

102 112 107 212 142 N* 127 128

104 120 114 200 134 N 129 122

N*

12

92 118 114 204 138 N 129 124

102 126 114 216 153 N 127 122

N

13

102 112 107 212 142 M 127 128

102 126 114 216 153 N 127 122

M

TE-N*

from these embryos confirmed the presence of the normal sequence of NF2 gene. Embryos #5 and #7 were transferred, resulting in an unaffected pregnancy and the birth of a healthy boy and girl confirmed by postnatal testing. (c) Outcome of the second PGT cycle. Combined mutation and linkage analysis by multiplex heminested PCR was performed on blastomeres from ten 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 of them (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 (embryos #3 and #4) were transferred, resulting in clinical pregnancy and the delivery of a healthy girl confirmed by postnatal analysis-

4.4 De Novo Mutations

Thus, PGT for DNM was based on detecting and, hence, avoiding the transfer of embryos with mutant haplotype with or without a mutant gene; thus, only embryos with normal haplotypes of paternal and maternal origin were transferred. The rationale is that ADO could have occurred and been undetected if the haplotype alone were the basis of diagnosis. Figure 4.20 shows the five tested embryos from the PGT cycle, all unaffected despite finding of the mutant haplotype in two (embryos #6 and #8). Yet both embryos lacked the mutant gene. The remaining three embryos had normal paternal and maternal haplotypes, of which two (embryos #5 and #7) were transferred and resulted in a twin pregnancy and birth of two unaffected children. In the second PGT cycle for this couple, ten embryos were examined, but only one contained the actual mutant haplotype. Three showed the mutant haplotype but without the mutant gene; the remaining six were with normal haplotypes. Two of these embryos (embryos #3 and #4) were transferred, resulting in a singleton pregnancy and birth of an unaffected child. An example of dominant DNM of maternal origin is presented in Fig. 4.21, in which gonadal mosaicism was also detected. DNM in the NF1 gene (intron 17–38 deletion) first presented in the affected child and appeared to be originated from the mother, who had three cell populations represented by three different haplotypes: normal haplotype, mutant haplotype with intron 17–38 deletion, and mutant haplotype without deletion. PGT was based on preselection and transfer of embryos with either normal maternal haplotype or mutant maternal haplotype lacking intron 17–38 deletion. Of 11 embryos examined, 2 of the 6 embryos with mutant maternal haplotypes were affected (embryos #6 and #8); the other 4 (embryos #1, #2, #10, and #12) were unaffected because their mutant haplotype did not contain the mutant intron 17–38 deletion allele. Of the remaining five embryos, two were monosomic for the maternal chromosome encoded for NF1 (embryos #7 and #11); three

121

(embryos #3–5) 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. As presented in Fig. 4.22, DNM for FANC I was first detected in a child who was compound heterozygous for the C750G/E837X mutations. Testing both parents for the presence of these mutations showed that the mother was a carrier of the E837X mutation and was characterized by two relevant haplotypes; the C750G mutation allele was not found in either paternal blood or whole sperm. Yet normal and mutant haplotypes were alternatively found in single sperm. Because the couple also requested HLA typing for possible stem cell transplantation required for the affected sibling, embryos were also tested for HLA haplotypes. Of the six embryos tested, only one embryo (embryo #2) inherited both maternal and paternal normal haplotypes, whereas two others had both paternal and maternal mutant haplotypes (embryos #3 and #6) but were unaffected heterozygous carriers because the paternal mutant haplotype lacked the paternal FCAN750G mutation. The remaining three embryos had normal maternal and mutant paternal haplotypes, without the mutant FANC allele involved. Thus, none of the embryos were actually affected. Two heterozygous carriers appeared to be also an HLA match to the affected child (embryos #3 and #6) and were transferred. Figure 4.23 presents an example of PGT-M performed for X-linked DNM, namely, chronic granulomatosis caused by DNM IVS 9 + 5G-A in the CYBB gene. DNM in this case was first detected in an affected child, who also required HLA-matched stem cell transplantation. DNA analysis in maternal blood failed to detect the mutant gene, although both normal and mutant haplotypes were present despite the latter lacking the actual mutant gene. PGT was performed by sequential PB1 and PB2 analysis in nine oocytes, showing that all were normal; however,

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

122

a

b

Family Pedigree 172 199 150 122 157 121 148 145 122 148 240

176 203 157 114 163 128 140 154 146 143 248

a

b

176 203 157 114 163 128 140 154 146 143 248

157 207 154 114 159 126 140 152 124 136 252

172 203 150 DEL DEL DEL DEL DEL 122 143 260

172 203 150 118 159 119 144 143 122 143 260

Markers order: D17S1873 D17S1294 D17S1863 NF1-IVS27-1 NF1-IVS27-2 NF1-IVS27-3 NF1-IVS36 NF1-IVS38 D17S1166 D17S1849 D17S798

172 203 150 DEL DEL DEL DEL DEL 122 143 260

c d e

b e

PGT

1

172 199 150 122 157 121 148 145 122 148 240

172 203 150 118 160 119 144 143 122 143 260

N*

2

172 199 150 122 157 121 148 145 122 148 240

172 203 150 118 160 119 144 143 122 143 260

3

172 199 150 122 157 121 148 145 122 148 240

157 207 154 114 159 126 140 152 124 136 252

N*

N

ET

ET

4

176 203 157 114 163 128 140 154 146 143 248

157 207 154 114 159 126 140 152 124 136 252

N

5

176 203 157 114 163 128 140 154 146 143 248

157 207 154 114 159 126 140 152 124 136 252

N

6

176 203 157 114 163 128 140 154 146 143 248

172 203 150 DEL DEL DEL DEL DEL 122 143 260

DEL

7

172 199 150 122 157 121 148 145 122 148 240

0

17,0

8

176 203 157 114 163 128 140 154 146 143 248

172 203 150 DEL DEL DEL DEL DEL 122 143 260

DEL

10

172 199 150 122 157 121 148 145 122 148 240

N*

172 203 150 118 159 119 144 143 122 143 260

11

172 199 150 122 157 121 148 145 122 148 240

17,0

0

12

176 203 157 114 163 128 140 154 146 143 248

172 203 150 118 159 119 144 143 122 143 260

N*

Fig. 4.21  PGT for DNM in NF1 gene (intron 27–38 deletion) of maternal origin. (a) Family pedigree of a couple with an 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. Position and order of the markers and deletion in NF1 gene are shown

on the upper left. (b) Outcome of PGT cycle, performed by multiplex heminested PCR on blastomeres from 11 embryos. Three embryos (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 (embryos #1, #2, #10, and #12) inherited the “benign” mutant maternal haplotype (d) and were also predicted normal (N∗) and suitable for embryo transfer (ET). Of the remaining four embryos, embryos #7 and #11 were predicted to have monosomy of chromosome 17, based on 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 (embryos #2 and #3) were transferred and resulted in a biochemical pregnancy

four (oocytes #2, #3, #7, and #11) contained the maternal mutant haplotype without the mutant gene. Testing of the embryos resulting from each of these oocytes confirmed the PB haplotype analysis, showing lack of any with the mutant allele. All IVS 9 + 5 G-A embryos were

found to be aneuploidy-free, of which four (embryos #7, #8, #9, and #11) appeared to be also the exact HLA match to the affected sibling. Two of these embryos (embryos #8 and #9) were transferred, resulting in a clinical pregnancy that spontaneously aborted in the first tri-

4.4 De Novo Mutations

a

123

142 117 145 172 N 130 167 106

Family Pedigree

HLA

133 130 147 162 N 123 165 130

HLA markers order D6S1629 D6S150 Ring D6S244LH D6S273 TNF-A D6S62 MIC A RF MOG a

142 142 128 117 150 145 172 172 C750G E837X 132 130 162 167 127 106

FANCI 1

PGT

FANCI

FANCI HLA

FANCI

DNA FROM TOTAL SPERM

b

Markers and mutations order: D15S655 D15S1046 D15S979 D15S1045 C750G E837X D15S202 D15S116 D15S127

133 142 130 128 150 150 162 172 N E837X 128 132 154 162 107 127

142 117 145 172 N 130 167 106

2

133 130 150 162 N 128 154 107

133 130 147 162 N 123 165 130

133 130 150 162 N 128 154 107

HLA

3

4

142 142 117 128 145 150 172 172 N E837X 130 132 167 162 106 127

142 117 145 172 N 130 167 106

133 130 150 162 N 128 154 107

5

142 117 145 172 N 130 167 106

133 130 150 162 N 128 154 107

6

142 142 117 128 145 150 172 172 N E837X 130 132 167 162 106 127

HLA

ET Normal*, Non-match

Normal, Non-match

ET

Normal*, Normal*, Non-match Non-match

Carrier* Match

Carrier* Match

Fig. 4.22  PGT for autosomal-recessive DNM detected first in an affected child who was compound heterozygous for C750G E837X mutations in the FANC I gene, combined with 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 sperm. Mutation and marker orders are printed in the upper right corner. (b) PGT cycle combined with HLA testing. (Upper portion) Multiplex heminested and fully nested amplification performed on blastomeres from six embryos

did not reveal the paternal mutation. Four embryos (embryos #1, #2, #4, and #5) were predicted normal (N) 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. (Lower portion) HLA marker analysis demonstrated the presence of two HLA-matched embryos (#3 and #6), which were transferred, but no pregnancy was achieved. N∗ – shows benign paternal haplotype a similar to the mutant haplotype of the affected child

mester. Of special interest also is another case in which two de novo maternal deletions occurred in DNM exon 8–9 and 9q34.3 (Fig. 4.24). The experience presented demonstrates not only complexity but also practical utility of

s­ trategies used for PGT in DNM. Initially, PGT could not be offered in the absence of family history and other affected family members who could allow testing for the origin of mutation and trace the inheritance of the mutant and nor-

124

a

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M) 157 N 112 Y 199 141

Family Pedigree HLA

164 N 110 199 143

CYBB

CYBB 160 IVS9+5G-A Y 114 189 141

CYBB

b

c

PGT by Polar Body

PGT by Blastomere

OOCYTES

2

3

CYBB

N*

N*

EMBRYOS

CYBB

2

4

N

3

8

N*

N

8

7

157 160 N N* 112 114 199 189 141 141

6, 6; 13,13; 16,16; 18,18; 21,21; 22,22;

160 N* 114 189 141

Y

164 N 110 199 143 Y

6, 6; 13,13; 16,16; 18,18; 21,21; 22,22;

6, 6; 13,13 16,16 18,18 21,21 22,22

6, 6; 13,13 16,16 18,18 21,21 22,22

HLA

HLA markers order: D6S1618 D6S1568 RING 1 D6S2447 LH 9N2 D6S265 D6S510

Markers and mutation order: DXS8090 CYBB DXS8014 DXS8025 DXS8018

HLA

7

4

157 160 N N* 112 114 199 189 141 141

160 N* 114 189 141

164 N 110 199 143

9

157 N 112 199 141

10

11

12

N

N

N*

N

9

10

11

157 N 112 199 141

164 N 110 199 143 Y

164 N 110 199 143

6, 6; 13,13; 16,16; 18,18; 21,21; 22,22;

6, 6; 13,13; 16,16; 18,18; 21,21; 22,22;

157 160 N N* 112 114 199 189 141 141

12

164 N 110 199 143

157 N 112 199 141

HLA

XX

XY

XY

XX

Normal, Non-match male Normal* Normal*, Non-match Non-match female male

Normal* Match female

6, 6; 13,13; 16,16; 18,18; 21,21; 22,22;

6, 6; 13,13; 16,16; 18,18; 21,21; 22,22;

XX XY XX XX Normal Normal, Normal, Match Match Non-match female male female ET

ET

6, 6; 13,13; 16,16; 18,18; 21,21; 22,22;

XX Normal, Non-match

Normal* Match female

Fig. 4.23  PGT for chronic granulomatous disease, determined by X-linked DNM IVS9+ 5G-A, combined with HLA genotyping and aneuploidy testing. (a) Family pedigree showing the mutation and closely linked to CYBB gene markers, and HLA-matched haplotypes depicted as white and non-matched colored in gray. (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) Embruyo biopsy analysis, including multiplex heminested PCR for combined mutation analysis (1c), HLA g (2c), and karyotyping (3c) for six chromosomes by PCR on blastomeres. Two of four embryos (embryos #8 and #9), predicted to be HLA-matched and free of mutation and aneuploidy, were transferred resulting in a singleton pregnancy and the birth of an unaffected child. ET embryo

mal alleles in oocytes and embryos. PGT-M is now possible as a result of  tracking parental origin of DNM in sperm  or  oocyte and constructing of relevant haplotypes. Highly accurate preselection of oocytes or embryos free from DNM can be provided [44].

Gonadal mosaicism carriers has an implication for genetic counseling of dominant disorders, such as NF1, TSC1, TSC2, lethal osteogenesis imperfecta, familial adenomatous polyposis, retinoblastoma, and X-linked dominant trait incontinentia pigmenti [45–50]. Although germinal

4.4 De Novo Mutations

125

PGT-M for TWO DE NOVO MATERNAL DELETIONS: DMD – exon 8-9 and 9q34.3 (Kleefstra syndrome) and Xp22.31 duplication

a

Family Pedigree Xp22.31 DMD

Xp22.31 DMD

Y 9q34.3

2.1

DOB 1978 2.2

Xp22.31

Xp22.31

DMD

DMD DE NOVO

b

PGT

EMBRYOS

Xp22.31

3

DMD

Y

Y 9q34.3 DEL DE NOVO

DMD Xp22.31 9q34.3

9q34.3

2.3

PGT-M + PGT-A

NORMAL PGT-M

5

Y NGS

9q34.3 46, XY NORMAL FET

9q34.3 47, XX, +15

Fig. 4.24  PGT for two de novo mutation detected in patient’s son: exon 8–9 deletion in DMD gene and 9p34.3 deletion causing a Kleefstra syndrome, characterized by intellectual disability, childhood hypotonia, severe expressive speech delay, and a distinctive facial appearance with a spectrum of additional clinical features. Patient also has

an Xp22 duplication, detected in her healthy daughter and affected son. PGT for three conditions were performed by linkage analysis combined with PGT-A. Out of two tested embryos one normal euploid male was recommended for transfer and healthy baby boy was delivered

mosaicism is believed to be common, its presence is usually difficult to detect. Clinically, all oocytes and embryos tested from such cases in our experience appeared to be unaffected, irrespective of the origin of DNM.  However, PGT is still justified, because possibility exists for a low-level mosaicism in parents’ gonads. In cases of DNM detected first in children, we found no mutation in either parent; however, presence of mutant haplotypes without the mutant gene was evident, suggesting that a proportion of germ cells with the mutation remain undetected. Thus, PGT-M is indeed indicated in such cases to exclude the possibility of the 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. A close proportion of DNM are of paternal (45%) or maternal (55%) origin, thus requiring testing for the presence of DNM in both parents. On the other hand, all cases of DNM of recessive inheritance were of paternal origin, although the number of cases (a total of 14) is not sufficient for definitive conclusions. In conclusion, the data presented in this section show that despite its complexity, PGT for DNM is highly accurate. No misdiagnoses were observed in PGT-M of 526 cycles, i.e., 100% accuracy of the applied technique of PGT-­ M. PGT for DNM has thus become an important

126

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

addition to the practice of PGT-M, making it applicable to any couple at risk for producing offspring with genetic disease, despite the traditional requirement of family data. Application of the above strategy will become even wider with the utilization of whole-genome sequencing (WGS), which will identify an increasing number of carriers of de novo dominant mutations. This will especially be expected in older fathers. Increasing numbers of new mutations may also be due to environmental hazards related to the technological developments in the modern society.

4.5

Late-Onset Common Disorders with Genetic Predisposition

Diseases with genetic predisposition – as opposed to complete penetrance if the mutation is transmitted  – have not always been considered an indication for prenatal diagnosis. The reasoning was that this could lead to pregnancy termination that was not considered well justified on the basis of genetic predisposition alone. That is, the tested fetus or embryo was not certain to become affected. On the other hand, choosing the embryos free of genetic predisposition for transfer would obviate the need for considering pregnancy termination, as only potentially normal pregnancies are established. PGT for such conditions gained acceptability on ethical grounds because only a limited number of the embryos available following hyperstimulation are selected for transfer. The number of PGT requests due to inherited predisposition for common late-onset conditions has been increasing. We shall focus on three groups of high clinical importance: (1) cancer, (2) neurodegenerative diseases, and (3) cardiac diseases.

4.5.1 Cancer Cancers are the largest group of conditions with genetic predisposition for which PGT-M is performed. The list of cancers we have performed

PGT is shown in Table  4.14, currently the world’s largest experience. PGT was performed in 874 PGT cycles for 484 couples at risk. This involves 45 different inherited cancers including BRCA 1 and 2, Li–Fraumeni disease; Fanconi anemia (FA); familial adenomatous polyposis (FAP); familial colorectal cancer; hereditary nonpolyposis coli (HNPCC) (type 1 and 2); Von Hippel–Lindau (VHL) syndrome; familial posterior fossa brain tumor (hSNF5); retinoblastoma (RB); neurofibromatosis 1 and 2 (NF1 and NF2); nevoid basal cell carcinoma (NBCCS) or Gorlin syndrome; tuberous sclerosis (TSC type 1 and type 2); ataxia telangiectasia; xeroderma pigmentosum, complimentary group G; exostosis multiple EXT1 and EXT2; dyskeratosis congenita AD2/AD3/DKCX; gastric cancer; paragangliomas 5 (PGL5); Peutz–Jegher syndrome; multiple endocrine neoplasia (MEN 1/2A/4); and pleuropulmonary blastoma, to mention only a few. The example of PGT for pleuropulmonary blastoma is shown in Fig.  4.25. Overall, our PGT program resulted in preselection of 966 predisposition-­ free embryos in 634 transfer cycles (1.52 embryos on an average), yielding 387 clinical pregnancies (61.0%) with birth of 407 healthy children apparently free from predisposition to those cancers (Table 4.14) (Some of these cases were reported previously [51– 53]). Except for breast cancer, most disorders are relatively rare autosomal dominant conditions, with prevalence of 1  in 5000  in the American populations for FAP, 1 in 15,000 for RB, and 1 in 36,000 for VHL, and even rarer for remaining cancers. The first PGT for inherited predisposition was performed in 1999 for a couple carrying p53 tumor suppressor gene mutations [53]. This tumor suppressor is known to determine a strong predisposition to many cancers, a major factor for genetic instability [54, 55]. Since then, the spectrum of indications for PGT has been extended to genetic predisposition for common diseases never before considered appropriate for prenatal diagnosis. The attraction of PGT for preselection of the mutation-free embryos and establishment of an unaffected pregnancy over-

Disease Albinism, oculocutaneous, type IA; OCA1A Albinism, oculocutaneous, type II; OCA2 Albinism, oculocutaneous, type III; OCA3 Subtotal (OCA1, OCA2, OCA3) Ataxia–telangiectasia; AT Basal cell nevus syndrome; BCNS (Gorlin) Birt–Hogg–Dube syndrome; BHD Breast–ovarian cancer, familial, susceptibility to, 1; BROVCA1 Breast–ovarian cancer, familial, susceptibility to, 2; BROVCA2 Subtotal (BRCA1+ BRCA2) Breast and colorectal cancer, susceptibility Colorectal cancer, hereditary nonpolyposis, type 1; HNPCC1 (Lynch syndrome I) Colorectal cancer, hereditary nonpolyposis, type 2; HNPCC2 Colorectal cancer, hereditary nonpolyposis, type 4; HNPCC4 Colorectal cancer, hereditary nonpolyposis, type 5; HNPCC5 Subtotal (HNPCC 1,2,4 and 5) Dyskeratosis congenita, autosomal dominant 3, DKCA3 Dyskeratosis congenita, autosomal dominant, 2, DKCA2 Dyskeratosis congenita, autosomal recessive, 5, DKCB5 Dyskeratosis congenita, X-linked; DKCX Subtotal (DKCA2, DKCA3, and DKCB5) Epidermolysis bullosa dystrophica, autosomal dominant; DDEB

Table 4.14  Overall experience of PGT for cancers Gene TYR OCA2 TYRP1 ATM PTCH1 FLCN BRCA1 BRCA2

CHEK2 MSH2 MLH1 PMS2 MSH6

TINF2 TERT RTEL1 DKC1 COL7A1

MIM 203100 203200 203290 208900 109400 135150 604370 612555

604373 120435 609310 614337 614350

613990 613989 615190 305000 131750

1 4 8

1

1 6 9

1

3

52 1

27 1 1

11

2

5

1

18

284 3 21

155 1 11 10

125

1 4 8

1

1

37 1

9

1

15

199 3 12

85

# of # of cycles transfers 9 8 4 3 1 0 14 11 10 6 7 6 2 1 159 114

65

# of patients 4 2 1 7 6 6 1 90

2 5 10

1

1

52 1

11

1

25

280 3 15

117

# embryos transferred 14 6 0 20 10 10 1 163

1 3 4

1

0

22 1

5

1

9

131 1 7

55

Pregnancy 4 2 0 6 5 4 1 76

0 0 0

0

0

1 0

0

0

0

9 0 1

4

SAB 0 0 0 0 1 0 0 5

1 3 4

1

0

21 1

5

1

9

122 1 6

51

1 3 4

1

0

22 1

5

1

10

134 1 6

55

Birth 4 2 0 8 4 4 1 79

(continued)

Delivery 4 2 0 6 4 4 1 71

4.5 Late-Onset Common Disorders with Genetic Predisposition 127

Disease Epidermolysis bullosa, junctional, Herlitz type Epidermolysis bullosa, junctional, non-Herlitz type Subtotal (Herlitz type, non-Herlitz type) Exostoses, multiple Familial adenomatous polyposis 1; FAP1 Fanconi anemia, complementation groups A, C, D, E, D, F, J Gastric cancer, hereditary diffuse; HDGC Hereditary leiomyomatosis and renal cell cancer; HLRCC Hermansky–Pudlak syndrome 1; HPS1 Li–Fraumeni syndrome 1; LFS1 Multiple endocrine neoplasia, type I; MEN1 Multiple endocrine neoplasia, type IIA; MEN2A Multiple endocrine neoplasia, type IV; MEN4 Subtotal (MEN1, MEN2A, MEN4) Myelodysplastic syndrome; MDS Neurofibromatosis, type I; NF1 Neurofibromatosis, type II; NF2 Subtotal (NF1, NF2) Nijmegen breakage syndrome; NBS Pancreatic cancer, susceptibility to, 3 Paraganglioma and gastric stromal sarcoma Paragangliomas 5; PGL5 Peutz–Jeghers syndrome; PJS Platelet disorder, familial, with associated myeloid malignancy; FPDMM Pleuropulmonary blastoma; PPB Renal cell carcinoma, papillary, 1; RCCP1 Retinoblastoma; RB1 Rhabdoid tumor predisposition syndrome 1; RTPS1

Table 4.14 (continued) Gene LAMA3 LAMB3

EXT1, EXT2 APC FANCA,C, D,E,F,G,J CDH1 FH HPS1 TP53 MEN1 RET CDKN1B GATA2 NF1 NF2 NBN PALB2 SDHB SDHA STK11 RUNX1 DICER1 MET RB1 SMARCB1

MIM 226700 226650

133700 175100 227650 137215 150800 203300 151623 131100 171400 610755 614286 162200 101000 251260 613348 606864 614165 175200 601399 601200 605074 180200 609322

1 1 17 1

1 16 9 6 1 16 1 51 7 58 1 1 1 1 4 1

1 2

18 19 22 29

# of patients 4 6

1 1 31 1

4 22 22 11 3 34 1 93 10 103 1 2 3 1 9 1

1 2

25 29 42 83

1 2 26 1

3 17 14 11 1 26 1 79 9 88 2 1 2 1 6 1

1 2

20 22 31 45

# of # of cycles transfers 9 8 7 4

1 2 51 1

6 24 22 16 1 39 1 121 17 138 2 2 2 1 9 1

2 2

31 38 51 71

# embryos transferred 14 7

1 1 14 1

2 13 7 8 1 16 1 46 7 53 1 1 1 1 4 1

1 1

12 15 14 20

Pregnancy 6 2

0 0 1 1

0 2 3 0 0 3 0 5 0 5 0 0 0 0 0 0

0 0

0 2 2 4

SAB 0 0

1 1 13 0

2 11 4 8 1 13 1 41 7 48 1 1 1 1 4 1

1 1

12 13 12 16

Delivery 6 2

1 1 16 0

3 11 6 10 1 17 1 46 9 55 1 1 1 1 4

2 1

16 16 13 22

Birth 9 3

128 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Telangiectasia, hereditary hemorrhagic, of Rendu, Osler, and Weber; HHT Telangiectasia, hereditary hemorrhagic, type 2; HHT2 Tuberous sclerosis 1; TSC1 Tuberous sclerosis 2; TSC2 Subtotal (TSC1, TSC2) Von Hippel–Lindau syndrome; VHL Xeroderma pigmentosum, complementation group G; XPG Total 21 8 29 19 1 484

TSC1 TSC2 VHL ERCC5 56 GENES

191100 613254 193300 278780

4

ACVRL1

600376

4

ENG

187300

874

30 14 44 25 2

8

11

634

25 8 33 16 1

4

6

966 (1.52)

49 12 61 25 1

6

7

387 (61%)

16 4 20 10 1

2

3

34 (8.8%)

2 0 2 0 0

0

1

353 (91.2%)

14 4 18 10 1

2

2

407

23 4 27 11 1

2

2

4.5 Late-Onset Common Disorders with Genetic Predisposition 129

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

130

a

Family pedigree

Markers order: D14S81 137 145 D14S1434 121 117 DICER1 c.1234delT N D14S62 124 131 D14S987 180 161 DICER1

31 YO

150 117 N 127 180

145 117 N 120 180

DICER1

PGT

145 145 117 117 c.1234delT N 131 120 161 180

b

137 121 N 124 180

NOT TESTED

150 117 N 127 180

PGT

Embryo #

1

2

145 150 117 117 c.1234delT N 131 127 161 180

145 150 117 117 c.1234delT N 131 127 161 180

MUTANT

10 TOTAL 5 NORMAL 4 EUPLOID

MUTANT

3

137 121 N 124 180

4

150 117 N 127 180

NORMAL EUPLOID

137 121 N 124 180

145 117 N 120 180

NORMAL EUPLOID

5

6

145 150 117 117 c.1234delT N 131 127 161 180

145 150 117 117 c.1234delT N 131 127 161 180

MUTANT

MUTANT

7

137 121 N 124 180

8

150 117 N 127 180

NORMAL EUPLOID

137 121 N 124 180

9

150 117 N 127 180

NORMAL EUPLOID

137 121 N 124 180

10

150 117 N 127 180

NORMAL ANEUPLOID

145 145 117 117 c.1234delT N 131 120 161 180

MUTANT

FET

Fig. 4.25  Combined PGT-M and PGT-A for pleuropulmonary blastoma caused by c.1234 deletion “T” in DICER1 gene. (a) Family pedigree. Partner and first baby are carriers of c.1234 deletion “T” in DICER1 gene. Mutation and linked markers are colored in red, while normal sequence and linked informative markers are depicted in black. A 27-year-old father was diagnosed with papillary thyroid cancer, and his first daughter developed pleuropulmonary blastoma at the age 2. Cancer

spread to her spine, brain, and bones. Second child was not tested, but was healthy. (b) Couple requested combined PGT-M and PGT-A.  Ten embryos developed to blastocyst stage and were analyzed. Five of them were predicted to be normal and the other five had a mutation. One of five mutation-free embryos tested for aneuploidy by NGS was aneuploid (embryo # 9). Embryo #3 which was mutation free and euploid was transferred resulting in birth of a healthy baby

weigh testing and the termination of a clinically affected pregnancy. In fact, at-risk couples have had the unfortunate experience of repeated prenatal diagnoses and termination of affected pregnancies. PGT is a better option to have unaffected offspring of their own, despite having to undergo IVF.  PGT-M is particularly attractive for couples at risk for late-onset disorders with the genetic predisposition, the fact that explains the steady increase of the number of PGT for cancer predisposition after our first description of PGT for cancer in 1999 (Fig. 4.26). Over 100 annual PGT-M cases have been performed during the last 2 years, and cur-

rently PGT for cancer constitutes 13.3% of all PGT-M cases. Neurofibromatosis (NF) was the second most frequent cancers for which PGT was performed in our experience (see Table 4.14). NF is a common autosomal dominant neurological disorder, of which there are at least two distinct major forms. NF type I (NF1) is more common (1:4000) and characterized by fibromatous skin tumors with café au lait spots, known also as Von Recklinghausen disease. NF type II (NF2) is less common (1:100,000) and characterized by bilateral acoustic neuromas, meningiomas, schwannomas, and neurofibromas [16]. The

4.5 Late-Onset Common Disorders with Genetic Predisposition Fig. 4.26 Steady increase of PGT cycles for cancer predisposition after first description in 1999

131

140 120 100 80 60 40 20

429 patients

NF1 gene is located on chromosome 17ql 1.2, whereas the NF2 gene is mapped on chromosome 22q12.2. Alterations in the sequence of these genes affect the tumor suppressor function of their gene products (neurofibromin and merlin, respectively), leading to a strong predisposition to malignancies. Different mutations in these genes have been described, resulting in a variety of clinical manifestations. Approximately half of these mutations are sporadic [56, 57], the rest representing germline mutations, which may be detected before the establishment of pregnancy to ensure unaffected pregnancy and the birth of a healthy child without an inherited predisposition to malignancy. Although PGT for inherited cancer predisposition is still controversial [58, 59], the possibility of establishing mutation-free pregnancies makes PGT an attractive option for neurofibromatosis, as preselection of predisposition-free embryos avoids the risk for clinical pregnancy termination. PGT for NF1/2 is presented in Figs. 4.20 and 4.21, demonstrating the practical relevance. We performed PGT for NF1 and NF2 either by PB sampling or embryo biopsy, depending on the parental origin of NF1 and

792cycles

2018

2017

2016

2015

2014

2013

2012

2011

2010

2009

2008

2007

2006

2005

2004

2003

2002

2001

2000

1999

0

46 genes

NF2 mutations, using multiplex PCR system with the application of available linked markers described previously [60–65]. We reported the first series of PGT cases for NF in 2002 [66]. At present, we performed 103 cycles for 58 at-risk couples, resulting in transfer of 138 embryos free from predisposition to NF1 or NF2  in 88 cycles, yielding 53 clinical pregnancies and birth of 55 NF predisposition-­ free children (Table 4.14).

4.5.1.1 Breast Cancer Breast cancer is presently the commonest indication for PGT-M in conditions having genetic predisposition [52, 67–71] (Table 4.14). Overall, we have performed 284 PGT cycles for 155 patients at risk for breast cancer caused by BRCA1 and BRCA2 mutations. This resulted in identification and transfer of 280 embryos free from mutations predisposing to breast cancer in 199 cycles, yielding 131 pregnancies and birth of 134 children without risk of developing breast cancer due to BRCA1 and BRCA2 (list of BRCA1 and BRCA2 mutations for which PGT has been ­performed is listed in Tables 4.15 and 4.16). Of the 38 different BRCA1 mutations, 187delAG was the most frequent originating comparably

132

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Table 4.15  List of BRSA1 mutation for which PGT was performed BRCA1 mutations 187 del AG 2813 ins A 3100 del GT 3977 del 4bp 5382 ins C 5385 ins C c.5154 c.5256 del G or 52 66? c5407-25T>A Exon 17del Exon 8-13 DEL IVS163 2del 3835 IVS22 (510 bp del) K679X Q1313X 3005del E6-8DEL R1699W 5360delA 2813ins A 3977 del4 IVS17+1 G>A 3100del GT c.5077_5079delGCT E 6-8 del 3005del C61G c3756del4 c.4065-4068del R1835X dup ex13 c2679del4 V1736A W1837R R1692H K135x R1752x Ivs23-1 a>g C2216-17 del aa c.3756 del 4bp 5266 dup c Y1703x C44f E20 del R1835x 4184 del 4 3481-2491? E577x c.4689 c>g c.5241 del E 1-2 del R1699w Total (38)

Maternal 33 3 0 1 4 2 1 2 1 1 1 1 1 1 1 1 1 1 1 1 1 1 0 1 1 1 2 1 1 1 0 1 1 1 1 1 1 1 1

Paternal 27 0 1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 1 0 0 0 1 0 0 0 1 0 0 0 0

1 1 1 1 1 1 1 1 1 1 1 1 1 87

33

Total 60 3 1 1 4 2 1 2 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 3 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 120

Table 4.16  List of BRSA2 mutation for which PGT was performed BRCA2 mutations Maternal Paternal Total 1417 ins 4 bp 2 0 2 2041 del A 0 1 1 2041 dup A 0 1 1 2776 del C 1 0 1 2942 ins 4 bp 1 1 2 3036-4 bp del 3 1 4 3398 del AAAG 1 0 1 3773 del TT 1 0 1 3782 del CA 1 0 1 4355 del 4 1 0 1 4682 del T 0 1 1 5578 del AA 1 0 1 5681 dup A 1 0 1 5722-23 del CT 1 0 1 5849 del 4 0 1 1 6174 Del T (c.5946 del T) 27 14 41 6486-89 del4 1 0 1 6644-47 del 4 1 0 1 886 del GT 1 1 2 955del CA 2 0 2 9686 del G 1 0 1 c.1189-90 ins4 TTAG 1 0 1 c.1813dup A 0 1 1 c.4075 del GT 0 1 1 c.4359 ins 6 2 0 2 c.5849 del 4 0 1 1 c.5851 del 4 AGTT 1 0 1 c.8673_74 del AA 0 1 1 c.9097 dup 1 0 1 c.9117 G>A 1 0 1 C7409 dp T 1 0 1 Exon 20 dup 1 0 1 Exon 1-2 del 1 0 1 Exon 8 del 1 0 1 G2748d 0 1 1 IVS13-2A>G 1 0 1 IVS17del3ins2 1 0 1 Q583X 1 0 1 S1955X 1 0 1 Y1655x 1 0 1 Total (40) 62 26 88

from the paternal and maternal partners, although, overall, maternally derived BRCA1 mutations were almost three times more frequent. Similar ratios were observed for BRCA2 mutations, applicable for both the major BRCA2 mutation and other rare ones. The most frequent BRCA2 mutation was 6174 Del T, present in almost half

4.5 Late-Onset Common Disorders with Genetic Predisposition

of the cases (41 of 88 cases performed). Of these 41 BRCA2 mutations, 27 were maternally derived, while, overall, 62 of 88 cases of BRCA2 were maternally derived. Thus, a total of only 59 of 175 (1 in 3) of the tested BRCA1/2 mutation were paternally derived. Interestingly, increasing number of cycles performed for BRCA1/2 was at concomitant risk for other genetic condition(s). In these cases PGT for both conditions was performed, as summarized in Table 4.17. Overall 37 PGT cycles were performed for BRCA1/2 mutations (24 cycles for BRCA1 and 12 for BRCA2), combined with PGT for other conditions, resulting in the birth of 15 healthy children unaffected for that second condition and free from predisposition to breast cancer (10 free of BRCA1 and 5 free of BRCA2 mutations). Of special interest were three cycles combined with PGT for additional two different conditions, each resulting in the birth of unaffected, predisposition-free children (see Table 4.17). Examples are shown for BRCA1 and SMA, or BRCA2 and MEN1, and are presented in Fig. 4.27, enabling preselection and transfer of unaffected embryos for SMA and MEN1, also free from genetic predisposition to breast and ovarian cancer. Another case of special interest is PGT-M in a couple with paternal partner carrying BRCA2 and maternal partner carrying BRCA1 mutations (Fig. 4.28). Of six tested blastocysts, two were carriers of BRCA1 and one BRCA2 mutations. Of the three remaining embryos free from BRCA1 and BRCA2 mutations, two had chromosomal aneuploidy, and the one remaining euploid embryo was a non-carrier of BRCA1 and BRCA2 mutations. This was transferred, resulting in a breast cancer predisposition-free child. Also, increasing numbers of patients with inherited predisposition to breast cancer have shown interest in nondisclosure PGT, previously used for Huntington disease. Either direct mutation or indirect linkage-based analysis can be applied. Indirect nondisclosure PGT for BRCA1 was performed for a couple, in which the breast cancer maternal partner had undergone oophorectomy and hysterectomy; her younger sister

133

was utilized as an egg donor. This resulted in transfer of cancer predisposition-free embryo, originating from younger sister’s donor oocyte, with birth of healthy child free of predisposition to breast cancer.

4.5.1.2 Other Cancers As seen from Table 4.14, another frequent cancer indication for PGT is familial adenomatous polyposis (FAP), which we have performed in 42 cycles. Patients with FAP usually present with colorectal cancer in early adult life, secondary to the extensive adenomatous polyps in their colon. Etiology is a mutation in the adenomatous polyposis coli (APC) gene located on chromosome 5 (5q21–q22). Mutations in APC gene cause premature truncation of the APC protein through single amino acid substitutions or frameshifts. The most common mutation is a 5  bp deletion resulting in a frameshift mutation at codon 1309. APC mutations lead to a premalignant disease with one or more polyps eventually progressing through dysplasia to malignancy with a median age at diagnosis of 40 years. Because mutations in the APC gene are almost always fully penetrant, albeit with striking variation in expression, pre-symptomatic diagnosis and treatment of carriers cannot exclude the progression of polyps to malignancy. PGT is an attractive approach for couples carrying APC mutations [72]. In our 42 cycles, 51 embryos free of APC mutation were detected for transfer in 31 cycles, resulting in 14 clinical pregnancy and birth of 13 of children without predisposition to FAP. A total of 25 PGT cycles were performed for Von Hippel–Lindau (VHL) syndrome, which is a cancer syndrome with age-related penetrance. The disorder is characterized by hemangioblastomas of the brain, spinal cord, and retina; bilateral renal cysts and renal carcinoma; pheochromocytoma; and pancreatic cysts. Depending on the combination of these clinical features, four ­different clinical phenotypes have been described. The gene responsible for VHL syndrome consists of three exons, located on chromosome 3 (3p26-­ p25). Specific VHL gene mutations correlate with a given clinical phenotype. The wild-type gene product is a tumor suppressor protein, expressed

Cystic fibrosis; Darier–White disease Cystic fibrosis; familial Mediterranean fever

All breast cancer type+ second condition Cystic fibrosis; 22q11.23 duplication

BRCA2+ second condition Breast cancer; beta-thalassemia

Conditions Breast–ovarian cancer, familial, susceptibility to, 1; breast–ovarian cancer, familial, susceptibility to, 2 Breast–ovarian cancer, familial, susceptibility to, 1; biotinidase deficiency Breast–ovarian cancer, familial, susceptibility to, 1; dystonia 1, torsion, autosomal dominant Breast–ovarian cancer, familial, susceptibility to, 1; fragile X mental retardation syndrome Breast–ovarian cancer, familial, susceptibility to, 1; mucopolysaccharidosis, type IIIA Breast–ovarian cancer, familial, susceptibility to, 1; spinal muscular atrophy, type I Breast–ovarian cancer, familial, susceptibility to, 1; arthrogryposis, renal dysfunction, and cholestasis 1 BRCA1+ second condition Breast–ovarian cancer, familial, susceptibility to 2; breast and colorectal cancer, susceptibility Breast–ovarian cancer, familial, susceptibility to 2; Tay–Sachs disease Breast–ovarian cancer, familial, susceptibility to 2; multiple endocrine neoplasia, type I Breast–ovarian cancer, familial, susceptibility to 2; Alzheimer disease 4 Breast–ovarian cancer, familial, susceptibility to 2; translocation

Gene BRCA1 BRCA2 BRCA1 BTD BRCA1 TOR1A BRCA1 FMR1 BRCA1 SGSH BRCA1 SMN1 BRCA1 VPS33B Subtotal BRCA2 CHEK2 BRCA2 HEXA BRCA2 MEN1 BRCA2 PSEN2 BRCA2 TL Subtotal PALB2 HBB Total CFTR 22q11.23DUP CFTR ATP2A2 CFTR MEFV

Table 4.17  Overall experience of combined PGT-M for more than one condition

2 2 1 4 24 3 1 6 1 1

1 1 1 1 13 1 1 2 1 1

1 1

20 1

1 1

37 2

12 1

1

1

6 1

1

# of cycles 13

1

# of patients 7

1 1

25 2

8 0

1

1

3

1

17 2

3

1

1

1

1

0

# of transfers 10

1 2

28 3

9 0

0

2

5

0

19 2

4

2

1

1

1

0

# of embryos transferred 10

1 1

15 2

4 0

0

1

2

0

11 1

2

1

0

1

1

0

0 0

2 1

0 0

0

0

0

0

2 0

1

0

0

0

0

0

1 1

13 1

4 0

0

1

2

0

9 1

1

1

0

1

1

0

1 1

15 1

5 0

0

1

3

0

10 1

1

2

0

1

1

0

Pregnancy SAB Delivery Birth 6 1 5 5

134 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Fragile X mental retardation syndrome; Leber congenital amaurosis 5 Fragile X mental retardation syndrome; translocation

Fragile X mental retardation syndrome; Canavan disease

Hemophilia A; Huntington disease

Cystic fibrosis; fragile X mental retardation syndrome Cystic fibrosis; facioscapulohumeral muscular dystrophy 1 Cystic fibrosis; hemochromatosis, type 1 Cystic fibrosis; Huntington disease nondisclosure Cystic fibrosis; phenylketonuria Cystic fibrosis; Aicardi–Goutieres syndrome 5 Cystic fibrosis; spinal muscular atrophy, type I Cystic fibrosis; translocation CFTR + second condition Renal tubular dysgenesis; surfactant metabolism dysfunction, pulmonary, 2 Familial adenomatous polyposis 1; achromatopsia 3 Familial adenomatous polyposis 1; deafness, neurosensory, autosomal recessive 1 Biotinidase deficiency; Smith–Lemli–Opitz syndrome Muscular dystrophy, limb-girdle, type 2A; choreoacanthocytosis Microcephaly 6, primary; mental retardation, autosomal recessive 5 Congenital disorder of glycosylation, type IIL; gaze palsy, familial horizontal, with progressive scoliosis Joubert syndrome 21; Perlman syndrome Smith–Lemli–Opitz syndrome; translocation Muscular dystrophy, Duchenne type; Glanzmann thrombasthenia Muscular dystrophy, Duchenne type; neuropathy, hereditary sensory and autonomic, type III Prothrombin deficiency, congenital; factor V deficiency 2 1 2 1 2 1 2 1 2

1 1 1 1 1 1 1 1 2

BTD DHCR7 CAPN3 GNA14 CENPJ NSUN2 COG6 ROBO3 CSPP1 DIS3L2 DHCR7 TL DMD ITGA2B DMD IKBKAP F2 F5 F8 HTT FMR1 ASPA FMR1 LCA5 FMR1 TL

2 1

1

1 1

1

1

1 1

1 1

APC CNGB3 APC GJB2

1

1 3 2 1 1 2 3 2 19 2

1 1 1 1 1 1 1 2 12 1

CFTR FMR1 CFTR FRG1 CFTR HFE CFTR HTT ND CFTR PAH CFTR SAMHD1 CFTR SMN1 CFTR TL Subtotal ACE SFTPC

0

2

1

1

2

2 0 2 0

0 0 2 1

2 1

1 3 1 2 0 2 2 1 16 2

0

2

1

1

2

2 0 2 0

0 0 2 1

2 1

1 5 1 3 0 3 3 3 25 3

0

1

1

1

2

1 0 2 0

0 0 2 0

1 0

1 2 0 2 0 2 2 1 14 1

0

0

0

1

0

0 0 2 0

0 0 1 0

0 0

0 0 0 1 0 1 1 0 4 0

0

1

1

0

2

1 0 0 0

0 0 1 0

1 0

1 2 0 1 0 1 1 1 10 1

(continued)

0

1

1

0

2

1 0 0 0

0 0 1 0

1 0

1 2 0 1 0 1 2 2 12 1

4.5 Late-Onset Common Disorders with Genetic Predisposition 135

Phenylketonuria; propionic acidemia Albinism, oculocutaneous, type IA; albinism, oculocutaneous, type II Polycystic kidney disease 1; Niemann–Pick disease, type C1 Polycystic kidney disease 1; polycystic kidney disease 2 Charcot–Marie–Tooth disease, demyelinating, type 1A; Fabry disease Charcot–Marie–Tooth disease, demyelinating, type 1A; Charcot– Marie–Tooth disease, demyelinating, type 1B; Charcot–Marie–Tooth disease, demyelinating, type 1A; neurofibromatosis, type I Charcot–Marie–Tooth disease, demyelinating, type 1A; Warburg Micro syndrome 1 Charcot–Marie–Tooth disease, demyelinating, type 1A; rhabdoid tumor predisposition syndrome 1 Alpha-1-antitrypsin deficiency translocation Spinal muscular atrophy, type I; anemia, nonspherocytic hemolytic

Familial Mediterranean fever; Noonan syndrome 1 Familial Mediterranean fever; tuberous sclerosis 1 Charcot–Marie–Tooth disease, axonal, Type 2A2; hypertrophic neuropathy of Dejerine–Sottas Usher syndrome, type I; Usher syndrome, type IIA Nemaline myopathy 2; fetal akinesia deformation sequence Nemaline myopathy 2; glycogen storage disease II

Infantile liver failure syndrome 1; translocation

Blood group Kell–Cellano system; rhesus blood group, D antigen

Conditions Fraser syndrome 1; Fraser syndrome 2 Tay–Sachs disease; translocation

Table 4.17 (continued)

1 1 1 1 4 1 1 1 2

1 1 1 1 1 1 1 1 1

PKD1 NPC1 PKD1 PKD2 PMP22 GLA PMP22 MPZ PMP22 NF1 PMP22 RAB3GAP1 PMP22 SMARCB1 SERPINA1 TL SMN1 G6PD

1 1 3 2 1 1

1 1 1

1

4

1 1 1

1

2

2 1

# of cycles 1 3

1 1

# of patients 1 1

Gene FRAS1 FREM2 HEXA TL KEL RHD LARS TL MEFV PTPN11 MEFV TSC1 MFN2 PRX MYO7A USH2A NEB MUSK NEB GAA PAH PCCA TYR OCA2

1 2

1

1

1

1

1 1 1

2 1

1 1 2

1 1 3

1

0

# of transfers 1 3

2 2

1

0

1

1

2 2 2

2 1

1 1 2

1 1 3

1

0

# of embryos transferred 1 3

1 1

0

0

1

1

1 1 1

1 1

1 1 2

1 1 1

1

0

0 0

0

0

0

0

0 0 0

0 0

0 0 1

0 0 0

0

0

1 1

0

0

1

1

1 1 1

1 1

1 1 1

1 1 1

1

0

1 1

0

0

1

1

1 1 3

1 1

1 1 1

1 1 1

1

0

Pregnancy SAB Delivery Birth 1 0 1 1 3 0 3 3

136 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

All combined conditions

Spinal muscular atrophy, type I; epidermolysis bullosa, junctional Ataxia–telangiectasia; deafness, neurosensory, autosomal recessive 1; muscular dystrophy, congenital merosin-deficient, 1A Metaphyseal chondrodysplasia, Schmid type; immunodysregulation, polyendocrinopathy, and enteropathy; cardiofaciocutaneous syndrome 4 Muscular dystrophy, Duchenne type; 9q34deletion; Xp22 triplication Epileptic encephalopathy, early infantile, 5; xeroderma pigmentosum, complementation group G; muscular dystrophy, congenital merosin-deficient, 1A; phenylketonuria Subtotal Total

DMD 9q34DEL Xp22 TRIPL SPTAN1 ERCC5 LAMA2 PAH

SMN1 LAMB3 ATM GJB2 LAMA2 COL10A1 FOXP3 MAP2K2 1 1

1 1

64 120

2

1

45 77

1 1

1 1

49 90

1

1

1

1 0

52 105 1.2

1

1

1

0 0

36 65 72%

1

1

1

0 0

1

1

1

0 0

5 31 11 54 17% 83%

0

0

0

0 0

33 60

1

1

1

0 0

4.5 Late-Onset Common Disorders with Genetic Predisposition 137

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

138

a

b

Family Pedigree MARKERS FOR SMA D5S2046 D5S2019 D5S435 D5S1414 AG1 SMN1 exon& SMN1 exon8 D5S610 D5S351 D5S1491 D5S2122

SMN1

BRCA1 SMN1

Family Pedigree

MARKERS FOR BRCA1 D17S800 D17S1787 D17S1793 BRCA1 Intron 8 Intron 41 D17S951 D171861 D17S934 D17S950

MARKERS FOR BRCA2

MARKERS FOR MEN1

D13S1229 D13S260 3036 del 4bp D13S171 D13S267 D13S220

D11S2006 D11S4191 D11S4076 IVS 7-del GT D11S1889 D11S4155

BRCA1

N N

PGT

DEL N

DEL N

DEL N 187 AG

PGT

AFFECTED SMA

1

2

Predicted genotype: BRCA2

?

N

N*

DEL DEL

MEN1

N

?

N

DEL DEL DEL

OOCYTE #

PGT Embryo #

1

2

3

5

SMN1

6

EMBRYO # 1

AFFECTED

CARRIERS

BRCA1

Predicted genotype: BRCA2 MEN1

NORMAL ET

3

4

5

6

N

7

8

9

DEL

DEL DEL

DEL

DEL

N

10

11

N

N

DEL

12

DEL

?

DEL

2

3

DEL

N

N

N

N

N

N

N

13,13; 16,16; 18,18; 21,21; 22,22; XX

13,13; 16,16; 18,18; 21,21; 22,22; XX

13,13; 16,16; 18,18; 21,21; 22,22; X XY

13,0;

ET

MEN1 IVS7-GT del

BRCA2 c.3036 –del 4 bp

PGT

ANEUPLOIDY 16,16; TESTING 18, 0; 21,21; 22,22; XX

ET

11

ET

Fig. 4.27  Concomitant PGT for breast cancer, BRCA1 and BRSA2, and SMA and MEN1. (a) PGT for BRCA1 and SMA in the same couple. (Upper panel) Pedigree showing that the patient and her mother are carriers of BRCA1 mutation (Del 187 AG; shown in red) – linked markers are listed on the right. The patient is also a carrier of SMN1 mutation (deletion, shown in green), inherited from her father. The male partner (the father) is also an unaffected carrier of the same deletion in SMN gene (shown in blue). The couple had one previous pregnancy resulting in the birth of an affected child with SMA who died (linked polymorphic markers for SMN1 mutation are listed on the left). (Lower panel) Five embryos were tested for both SMN1 and BRCA1 in the same reaction, showing that embryos #1 and #2 contained deletion in SMNA from both parents (blue and green), and the remaining three embryos were carriers of either maternal (embryos #4 and #5) or paternal (embryo #3) deletion. Two of these embryos (embryos #1 and #5) were also carriers of BRCA1 mutation, so three embryos (embryos #2, #3, and #4) were predicted to be free of BRCA 1 mutation and unaffected by SMA. Two of these embryos (embryos #3 and #4) were transferred, resulting in the birth of an unaffected child, shown in pedigree as PGD

(ET embryo transfer). (b) Combined PGT for BRCA2, MEN1, and aneuploidy. (Upper panel) Pedigree showing that the patient and her father are carriers of BRCA2 (c.3036-del4 bp, shown in red) and MEN1 (IVS7-GT del, shown in blue) mutations. Polymorphic markers for testing of BRCA2 mutation are shown on the left, and for MEN1 deletion on the right. (Middle panel1) Twelve oocytes were tested by sequential PB1 and PB2 analysis simultaneously for both mutations, which detected only one oocyte (oocyte #3) to be free of both mutations (in addition, oocyte #2 had insufficient marker information to confirm a normal allele for MEN1), so the resulting four embryos (embryos #1, #2, #3, and #11) were further tested by blastomere biopsy, presented in the middle panel2. (Middle panel2) Blastomere analysis of embryos #1, #2, #3, and #11, showing that all but one (embryo #1) are free of both mutations. (Lower panel) Two of these embryos were chromosomally abnormal (embryos #1 monosomic for chromosomes 13 and 18; and embryo #11 with extra chromosome X), while the other two (embryos #2 and #3) were euploid. These two embryos were transferred, resulting in an unaffected singleton pregnancy and the birth of a healthy child free of both BRCA2 and MEN1 mutations

in most cells and carrying a variety of functions including transcriptional and posttranscriptional regulation. More than 300 germline mutations have been identified in families with VHL syndrome, consisting of partial or complete gene

deletions and frameshift, nonsense, m ­ issense, and splice site mutations most commonly affecting codon 167. Mutations in the VHL gene either prevent its expression completely or lead to the expression of an abnormal protein. Only 20% of

4.5 Late-Onset Common Disorders with Genetic Predisposition

139

a Family pedigree 170 138 122 N 175 134

174 138 124 N 175 136

BRCA1

154 152 135 137 102 114 886delGT N 97 112 175 179 189 200

34 yo

NGS

c

154 135 104 N 100 179 194

154 133 108 N 110 183 200

BRCA2

PGT Normal BRCA1 & BRCA2

1

Embryo #

BRCA 2

172 134 132 N 175 129

BRCA1

BRCA2

b PGT

BRCA 1

170 136 127 C61G 173 131

2

3

4

5

6

170 138 122 N 175 134

172 134 132 N 175 129

170 138 122 N 175 134

170 136 127 C61G 173 131

170 138 122 N 175 134

170 136 127 C61G 173 131

174 138 124 N 175 136

172 134 132 N 175 129

170 138 122 N 175 134

154 135 102 N 97 175 189

154 135 104 N 100 179 194

154 135 102 N 97 175 189

154 135 104 N 100 179 194

154 135 102 N 97 175 189

154 135 104 N 100 179 194

154 135 102 N 97 175 189

154 135 104 N 100 179 194

152 137 114 886delGT 112 179 200

47, XX, +2 Embryo # 1

46, XX 47, XX, +2

46, XY

Embryo # 4

47, XX, +16 47, XX, +16

172 134 132 N 175 129

154 133 108 N 110 183 200

46, XX Embryo # 6

174 138 124 N 175 136

172 134 132 N 175 129

154 135 102 N 97 175 189

154 135 104 N 100 179 194

46, XX FET 46, XX

Fig. 4.28  Combined PGT-M for BRCA1, BRCA2, and PGT-A by NGS. (a) Three generations of family members, harboring BRCA1 mutation (depicted in red) in patient’s family and BRCA2 mutation (depicted in blue) in partner’s family. (b) Tropectoderm sample analysis for BRCA1, BRCA2, and NGS for chromosome copy numbers. Out of six tested embryos two had maternal mutation and one paternal and three embryos were predicted to be

free from both mutations. (c) NGS-based testing of embryos detected trisomy 2 in embryo #1 and trisomy 16 in embryo #4, while embryo #6 was free from aneuploidy and transferred (at the time of the testing, patient was 34 years old). Out of these three embryos only one was euploid (embryo #6) and transferred. A Baby free of both mutations was delivered

cases of VHL are sporadic, the remaining 80% familial. PGT is thus an attractive option for couples carrying these mutations, avoiding inheritance by their children of these tumor suppressor mutations. In our experience of 25 PGT cycles for VHL, 25 genetic predisposition-free embryos were preselected for transfer in 16 cycles, resulting in the birth of 11 children free from predisposition to VHL. With accelerating progress in understanding the molecular basis of cancers and sequencing of the genes, inherited cancer predispositions are becoming the major emerging PGT indication. Most of them are dominant and may be also secondary to germline mutations [73]. As mentioned, these conditions account already for 13.3% of all PGT-M cases, despite still remaining controversy, because these diseases may present beyond early

childhood and may not be expressed in 100% of the cases. Despite extensive discussions of the ethical and legal issues involved in PGT for lateonset disorders with genetic predisposition, an increasing number of patients clearly regard the procedure not only as their preferable option but the only possible reason for forgoing the pregnancy. Establishing a pregnancy free of mutation from the onset allows avoiding a difficult decision to terminate a pregnancy at high risk if the fetus carries a mutant gene. Genetic counseling and oncologic services should inform patients at risk for these cancers that having children with genetic predisposition to cancers can be avoided through PGT.  Options exist for couples who might have chosen to remain childless because of their concern to avoid prenatal diagnosis and possible pregnancy termination.

140

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

4.5.2 Neurodegenerative Diseases Neurologic diseases, including neurodegenerative conditions, represent another large group of late-onset common conditions with a strong inherited predisposition, for which PGT has become an attractive option. The list of these conditions for which we are offering PGT is presented in Table 4.18. The majority of cases (210 cycles) were performed for Huntington disease, but PGT for some rarer conditions were also involved, such as 55 cycles for hereditary ataxias, including spinocerebellar ataxia 1, 2, 3, 6, 7, and 8 (ATXN1, ATXN2, ATXN3, ATXN7, ATXN8), 6 for Gerstmann–Sträussler Schinker disease (PRNR), and 5 for Alzheimer disease (PSENT1, PSENT2, APP), to mention only a few. PGT for the most of these condition listed in Table  4.18 resulted in clinical pregnancies and birth of offspring free of predisposition to these diseases.

4.5.2.1 Huntington Disease Huntington disease is a rare hereditary neurodegenerative disorder, with the prevalence of 1 in 10,000 in the United States caused by an expansion of a repeating CAG triplet series in the huntingtin gene on chromosome 4. This dynamic mutation results in a protein with an abnormally long polyglutamine sequence, representing one of a larger family of polyglutamine repeat disorders, all of which are neurodegenerative diseases. Huntington disease is inherited in an autosomal dominant fashion; thus, each child of an affected parent has a strong genetic predisposition, with 50% chance of developing the disease. As no cure is available, PGT provides a realistic approach for parents at risk to avoid the disease in their offspring. Huntington disease is unusual in that a germline mutation is found in almost all families. Penetrance is almost 100% of adjusted for age and number of CAG repeats. Almost a half of patients referring for PGT of HD request nondisclosure of genotype in PGT, which has actually first been developed particularly for couples at risk for HD. Transfer of only disease-free embryos is possible while the prospective parents do not learn their own status. Parents receive no information about the num-

ber of oocytes obtained after hormonal stimulation, nor information on the number of embryos formed, and nor the number of embryos available for transfer. The rationale underscoring absolute nondisclosure is that stating no embryos for transfer, or that all embryos were mutant. An alternative nondisclosure PGT method is exclusion testing. Embryos inheriting a grandparental allele would be excluded from transfer notwithstanding that only half of these known embryos could contain the affected allele. As described above, nondisclosure PGT-M is no longer restricted to HD and also has been requested in PGT for other conditions, including breast cancer, Machado–Joseph disease, spinocerebellar ataxia type 6, CMT1A, early-onset Alzheimer disease, Gerstmann–Sträussler– Scheinker disease, myotonic dystrophy, multiple endocrine neoplasia type IV (MEN4), myofibrillar myopathy 1, frontotemporal dementia, and lateral amyotrophic sclerosis (Table  4.19). Our experience of 210 PGT cycles for HD resulted in transfer of 263 genetic predisposition-free embryos, yielding 104 clinical pregnancies and birth of 101 children with no predisposition to develop the disease in their life span (Table 4.18). As mentioned, more than half of PGT patients preferred not knowing the actual diagnosis of their own genotypes, thus choosing nondisclosure PGT-­ M.  This situation arises when the patient is aware of HD in one of his or her parents, but is reluctant to know his or her own genotype, while still does not wish to transmit a potential mutation to offspring. Thus, PGT-M is performed to preselect only unaffected embryos for transfer. No results are disclosed to the patient, but transfer can avoid any with the mutant gene. This minimizes the concern whether the patient inherited or not the abnormal genotype for which they are at 50% risk. As alluded before, if only two of ten embryos were said to be transferable, presence of the mutation might be suspected. At present nondisclosure has become more practical with the introduction of concomitant PGT-A.  A potential pitfall is that the same procedure must be repeated in all subsequent cycles, even if the (undisclosed) patient had been shown not to carry the mutant gene in a previous

Disease Ataxia hereditary Friedreich ataxia 1; FRDA Joubert syndrome 1; JBTS1 Joubert syndrome 2; JBTS2 Joubert syndrome 3; JBTS3 Joubert syndrome 6; JBTS6 Joubert syndrome 17; JBTS17 Joubert syndrome 21; JBTS21 Joubert syndrome 23; JBTS23 Machado–Joseph disease; MJD Spinocerebellar ataxia 1; SCA1 Spinocerebellar ataxia 2; SCA2 Spinocerebellar ataxia 6; SCA6 Spinocerebellar ataxia 7; SCA7 Spinocerebellar ataxia 8; SCA8 Subtotal Epilepsy Epileptic encephalopathy, early infantile, 2; EIEE2 Epileptic encephalopathy, early infantile, 23; EIEE23 Epileptic encephalopathy, early infantile, 3; EIEE3 Epileptic encephalopathy, early infantile, 5; EIEE5 Subtotal Intellectual disability Dyskinesia, seizures, and intellectual developmental disorder; DYSEIDD Fragile X mental retardation syndrome Mental retardation, autosomal recessive 38; MRT38 Rett syndrome; RTT Subtotal Migraine, familial hemiplegic, 1; FHM1 Movement disorder Dystonia 1, torsion, autosomal dominant; DYT1 1 610 2 5 618 6 37

1 1 1 1 4 1

300672 615859 609304 613477

617171 300624 315 615516 1 312750 3 320 141500 3 128100

CDKL5 DOCK7 SLC25A22 SPTAN1 4 genes DEAF1 FMR1 HERC2 MECP2 4 genes CACNA1A TOR1A

17

6 1 1 1 3 1 3 1 7 7 14 6 3 1 55

2 1 1 1 3 1 2 1 5 4 6 3 2 1 33

229300 213300 608091 608629 610688 614615 615636 616490 109150 164400 183090 183086 164500 608768

FXN INPP5E TMEM216 AHI1 TMEM67 CPLANE1 CSPP1 KIAA0586 ATXN3 ATXN1 ATXN2 CACNA1A ATXN7 ATXN8OS 14 genes 1 1 1 2 5

# of cycles

Gene

# of OMIM patients

39

441 2 3 447 3

1

1 2 0 2 5

4 2 1 0 3 2 3 2 6 6 14 3 3 1 50

# of transfers

Table 4.18  PGT for neurologic disorders with genetic predisposition, including neurodegenerative conditions

64

652 3 3 658 5

0

1 2 0 2 5

7 2 1 0 5 2 5 2 8 8 27 5 7 1 80

# of embryos transferred

18

232 1 2 235 2

0

1 2 0 1 4

2 1 1 0 2 1 2 1 6 4 6 2 2 1 31

3

25 0 1 26 0

0

0 1 0 0 1

0 0 0 0 0 0 0 0 1 0 0 0 1 0 2

Pregnancy SAB

15

207 1 1 209 2

0

1 1 0 1 3

2 1 1 0 2 1 2 1 5 4 6 2 1 1 29

(continued)

17

228 1 1 230 3

0

1 1 0 1 3

2 1 1 0 2 1 2 1 6 4 8 3 1 1 33

Delivery Birth

4.5 Late-Onset Common Disorders with Genetic Predisposition 141

Disease Dystonia 3, torsion, X-linked; DYT3 Dystonia 11, myoclonic; DYT11 Dystonia 28, childhood-onset; DYT28 Subtotal Neurodegenerative disorders Alzheimer disease 3 Alzheimer disease 4 Alzheimer disease; AD Creutzfeldt–Jakob disease; CJD Gerstmann–Sträussler disease; GSD Huntington disease; HD Infantile cerebellar–retinal degeneration; ICRD Leukoencephalopathy with vanishing white matter; VWM Pontocerebellar hypoplasia, type 1B; PCH1B Supranuclear palsy, progressive, 1; PSNP1 Subtotal Total

Table 4.18 (continued)

3 1 5 6 3 210 1 1 1 3 234 960

607822 2 606889 1 104300 2 123400 3 137440 2 143100 142 614559 1 603896 1 614678 601104

PSEN1 PSEN2 APP PRNP PRNP HTT ACO2 EIF2B2 EXOSC3 MAPT 10 genes

1 2 157 537

# of cycles 1 3 1 42

# of OMIM patients 314250 1 159900 1 617284 1 20

Gene TAF1 SGCE KMT2B 4 genes

1 3 190 738

2 2 3 6 5 166 1 1

# of transfers 2 1 1 43

1 5 294 1110 (1.5)

3 3 5 7 5 263 1 1

# of embryos transferred 2 1 1 68

1 1 120 412 (55.8%)

2 1 2 4 3 104 1 1

Pregnancy 1 1 0 20

0 0 13 45 (11%)

0 0 0 0 0 11 1 1

SAB 0 0 0 3

1 1 107 367 (89%)

2 1 2 4 3 93 0 0

1 1 118 406

3 1 3 4 4 101 0 0

Delivery Birth 1 1 1 1 0 0 17 19

142 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

4.5 Late-Onset Common Disorders with Genetic Predisposition

143

Table 4.19  Conditions for which nondisclosure PGT-M was performed Condition ALS SCA3 (Machado–Joseph disease) Spinocerebellar ataxia type 6 BRCA Cerebral arteriopathy (CADASIL) CMT1A Early Alzheimer Gerstmann–Sträussler–Scheinker disease Myotonic dystrophy Multiple endocrine neoplasia, type IV; MEN4 Myofibrillar myopathy 1 Frontotemporal dementia Huntington disease Total

Gene SOD1 ATXN3 CACNA1A BRCA1 NOTCH3 PMP22 PSEN1 PRNP DMPK CDKN1B DES PGRN HTT

cycle. In our experience 64 (45.1%) couples still had direct HTT testing. An example of PGT with direct mutation analysis for Huntington disease is presented in Fig. 4.29.

4.5.2.2 Alzheimer Disease The first PGT for Alzheimer disease was reported in 2002 [74]. Alzheimer disease occurs in 1–2% of the US population, usually at the age 65–80 years. Familial tendencies exist, with concordance in approximately 50% in MZ twins and 20% in DZ twins or sibs. No single gene is causative. There are at least three rarer autosomal dominant genes conferring familial predisposition to a pre-senile form of dementia (age T), leading to an amino acid change from Arg to Trp in position 335 of the proteins lamin A and lamin B in the cardiac muscle. This mutation was originally detected by MspI digestion, which generates two fragments of 90 and 95 bp in the PCR product of the normal LMNA allele, leaving the mutant one uncut. Four polymorphic markers were also tested simultaneously with the mutation analysis  – D1S2714, D1S82777, D1S2624, and D1S506  – to avoid misdiagnosis due to preferential amplification or allele dropout (ADO) of the mutation tested. The second largest group of cardiac disorders for which PGT was performed was familial hypertrophic cardiomyopathy (CMH), including patients with CMH1, CMH2, CMH4,

MYH7 (AD) 3 TNNT2 (AD) 1 MYBPC3 (AD) 14 TNNI3 (AD) 1 MYL3 (AD) 1 TAZ (XLR) 1

COQ4 (AR) 1 EMD (XLR) 3 TBX5 (AD) 5 TGFBR2 (AD) 2 KCNQ1 (AD) 4 KCNH2 (AD) 3 CACNA1C 1 (AD) 601419 DES (AD) 1 163950 PTPN11 (AD) 6 23 genes 67

616276 310300 142900 609192 192500 613688 618447

192600 115195 115197 613690 608751 302060

1

615280 MAP2K2 (AD) 115200 LMNA (AD) 613172 RBM20 (AD) 601154 SCN5A (AD) 604145 TTN (AD) 615821 DSP (AD) 7 1 1 1 2

2

604377 SCO2 (AR)

# of OMIM Gene patients 609575 ACADVL (AR) 5

AD autosomal dominant, AR autosomal recessive, XLR X-linked recessive

Myopathy, myofibrillar, 1; MFM1 Noonan syndrome 1; NS1 Total

Cardiomyopathy, dilated, 1A; CMD1A Cardiomyopathy, dilated, 1DD; CMD1DD Cardiomyopathy, dilated, 1E; CMD1E Cardiomyopathy, dilated, 1G; CMD1G Cardiomyopathy, dilated, with woolly hair, keratoderma, and tooth agenesis; DCWHKTA Cardiomyopathy, familial hypertrophic, 1; CMH1 Cardiomyopathy, familial hypertrophic, 2; CMH2 Cardiomyopathy, familial hypertrophic, 4; CMH4 Cardiomyopathy, familial hypertrophic, 7; CMH7 Cardiomyopathy, familial hypertrophic, 8; CMH8 Cardioskeletal myopathy with neutropenia and abnormal mitochondria Coenzyme Q10 deficiency, primary, 7; COQ10D7 Emery–Dreifuss muscular dystrophy 1, X-linked; EDMD1 Holt–Oram SYNDROME; HOS Loeys–Dietz syndrome 1; LDS1 Long QT syndrome 1; LQT1 Long QT syndrome 2; LQT2 Long QT syndrome 8; LQT8

Disease Acyl-CoA dehydrogenase, very long-chain, deficiency of; ACADVLD Cardioencephalomyopathy, fatal infantile, due to cytochrome C oxidase deficiency 1; CEMCOX1 Cardiofaciocutaneous syndrome 4; CFC4

Table 4.20  PGT for Cardiac Diseases with Genetic Predisposition

2 8 109

1 4 8 5 6 3 1

6 1 22 1 2 1

17 2 2 1 3

2

5

# of cycles 6

2 5 89

1 4 7 4 3 1 1

4 1 16 1 1 1

15 3 2 1 2

1

5

# of transfers 8

3 7 123 (1.38)

1 6 8 6 3 1 1

4 2 23 1 1 1

24 3 2 1 2

1

10

# of embryos transferred 12

1 4 55 (61.7%)

1 3 3 2 3 1 1

2 1 11 0 0 1

9 2 1 1 2

1

3

0 1 5 (9%)

0 0 0 1 0 0 0

0 0 2 0 0 0

0 0 0 0 1

0

0

1 3 50 (91%)

1 3 3 1 3 1 1

2 1 9 0 0 1

9 2 1 1 1

1

3

1 4 54

1 3 4 1 3 1 1

2 2 9 0 0 1

10 2 1 1 1

1

3

Pregnancy SAB Delivery Birth 2 0 2 2

150 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

4.5 Late-Onset Common Disorders with Genetic Predisposition

151

a Family pedigree

Markers: D1S2714 D1S2777 LMNA-R335T D1S2624 D1S506

143 150 N 126 89

140 156 R335T 132 95

PGT 143 150 N 126 89

b

140 152 N 128 95

143 152 N 128 97

143 152 N 128 97

PGT Embryo # 1

143 150 N 126 89

143 152 N 128 97

N ET

2

4

143 140 150 152 N N 126 128 89 95

143 143 150 152 N N 126 128 89 97

N

N

FR

5

143143 150152 N N 126128 89 97

6

143143 150152 N N 126128 89 97

N

N

FR

FR

7

143143 150152 N N 126128 89 97

N

8

143 143 150 152 N N 126 128 89 97

N ET

9

140 143 156 152 R335T N 132 128 95 97

M

10

143 140 150 152 N N 126 128 89 95

N

11

FA

12

140 143 156 152 R335T N 132 128 95 97

M

Fig. 4.34 PGT for dilated cardiomyopathy (CMD), determined by dominant mutation in LMNA gene. (a) Family pedigree of a couple with affected husband carrying R335T mutation in LMNA gene. Paternal linked polymorphic markers are shown on the left, and maternal on the right, and the order of the markers and mutation in LMNA gene are shown on the upper left. (b) Blastomere

results revealed two embryos carrying R335T mutation in LMNA gene (embryos #9 and #12), while the remaining nine were free of R335T mutation. Two of these embryos (#1 and #8) were transferred, resulting in a singleton pregnancy and birth of a healthy child without the predisposing gene to CMD (as indicated in the family pedigree by PGT). ET embryo transfer, FR frozen embryos

CMH7, and CMH8, determined by mutation in MYH7, TNNT2, MYBPC3, TNNI3, and MYL3 genes, respectively. Neither of these couples had previous progeny but had a family history existed for premature or sudden death. PGT for CMH4 is presented in Fig. 4.35. PGT was performed in one of the families having the frameshift mutation D1076 fs in the MYBPC3 gene. This was detected by RSAI and BsaHI digestion, the former cutting the mutant gene into two fragments of 72 and 60 bp and the second cutting the normal allele into two fragments of the same size. In addition, five polymorphic markers were also used to exclude the possibility of ADO, including D11S1978, D11S1344,

D11S4117, D11S1350, and D11S4147. Of seven embryos tested, three (embryos #7, #9, and #10) were carriers of the frameshift mutation D1078fr in the MYBPC3 gene, three were unaffected, and one did not amplify. Two of the normal embryos were transferred following freezing, resulting in an unaffected pregnancy and birth of predisposition-free child. PGT for CMH7 is shown in Fig.  4.36, performed for a family with a A157V mutation in the TNNI3 gene. The A157V mutation in the TNNI3 gene was detected by the use of the two enzymes, HaeII, cutting the normal, and BspMI, cutting the mutant gene into two fragments. Of 11 tested embryos, 10 were amplified, of which

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

152

Family pedigree a

MARKERS: D11S4174 D11S1344 MYBPC3-D1076fs D11S4117 D11S1350 D11S1978

144 172 N 116 194 186

148 174 N 114 187 167

PGT

148 174 N 114 187 167

b

3

EMBRYO #

148 174 N 114 187 167

137 159 N 116 191 186

NORMAL

144 164 D1-76fs 112 179 182

137 159 N 116 191 186

137 159 N 116 191 186

PGT 5

FA

7

144 144 172 164 N D1-76fs 116 112 194 179 186 182

AFFECTED

FET 1

Fig. 4.35 PGT for hypertrophic cardiomyopathy (CMH4). (a) Family pedigree of a couple with affected mother carrying frameshift mutation D1076 fs in MYBPC3 gene. Paternal linked polymorphic markers are shown on the left, and maternal on the right, and the order of the markers and frameshift mutation in MYBPC3 gene are shown on the upper left. (b) Embryo biopsy results

8

148 174 N 114 187 167

137 159 N 116 191 186

NORMAL

9

144 144 172 164 N D1-76fs 116 112 194 179 186 182

AFFECTED

10

144 144 172 164 N D1-76fs 116 112 194 179 186 182

AFFECTED

12

148 174 N 114 187 167

137 159 N 116 191 186

NORMAL

FET 2

revealed three embryos (embryos #7, #9, and #10) carrying the frameshift mutation D1078fr in MYBPC3 gene, four unaffected, and one did not amplify. Two of the normal embryos were transferred (embryos #3 and #8), following the freezing (frozen embryo transfer (FET)), resulting in an unaffected pregnancy (as indicated in the family pedigree by PGT)

4.5 Late-Onset Common Disorders with Genetic Predisposition

153

Family pedigree MARKERS: D19S902 D19S867 D19S904 D19S246 TNNI 3-A157V D19S206 D19S571

a

149 117 119 219 A157V 126 123

145 126 127 183 N 136 104

134 132 127 206 N 121 100

PGT 145 126 127 183 N 136 104

134 126 119 222 N 121 127

134 126 119 222 N 121 127

b PGT 1

149 117 119 219 A157V 126 123

134 126 119 222 N 121 127

2

149 117 119 219 A157V 126 123

3

134 132 127 206 N 121 100

145 126 127 183 A157V 126 123

4

134 132 127 206 N 121 100

AFFECTED AFFECTED AFFECTED

145 126 127 183 N 136 104

134 126 119 222 N 121 127

5

145 126 127 183 N 136 104

134 132 127 206 N 121 100

NORMAL NORMAL

ET*

6

145 117 119 219 A157V 126 123

7

134 132 127 206 N 121 100

AFFECTED

FR*

149 117 119 219 A157V 126 123

8

134 126 119 222 N 121 127

AFFECTED

FA

9

149 117 119 219 A157V 126 123

10

134 132 127 206 N 121 100

AFFECTED

149 117 119 219 A157V 126 123

134 126 119 222 N 121 127

AFFECTED

11

145 126 127 183 N 136 104

134 132 127 206 N 121 100

NORMAL

FR*

* Normal for 24 chromosome by microarray Fig. 4.36  PGT for CMH7 (a) Family pedigree of a couple with affected father carrying A157V mutation in TNNI3 gene. Paternal linked polymorphic markers are shown on the left, and maternal on the right, and the order of the markers and mutation in A157V mutation in TNNI3 gene are shown on the upper left. (b) Embryo biopsy results revealed three mutation-free embryos, based on the

testing of the mutation and six polymorphic markers (embryos #4, #5, and #11), seven mutant ones, and one did not amplify. Unaffected embryos were tested for 24-chromosome aneuploidy at the blastocyst stage, of which one (embryo #4) was euploid and was transferred in the subsequent cycle. FR frozen, FET frozen embryo transfer, FA failed amplification

3 (embryos #4, #5, and #11) were unaffected, based on the testing of the mutation and 6 polymorphic markers. Because these embryos were also tested for 24-chromosome aneuploidy by array CGH analysis at the blastocyst stage, embryos were frozen. One (embryo #3) was also aneuploidy-free and was transferred in the subsequent cycle. PGT cycle for cardioencephalomyopathy was performed in a couple having a previously

affected child with left ventricular hypertrophic cardiomyopathy, whose first symptoms were manifested at 1.5 months as severe respiratory attack (Fig.  4.37). Maternal mutation E140K of the SCO2 gene was detected by HindIII and BsrBI digestion, the first cutting the mutant allele and the second cutting the normal allele. Paternal mutation R262del (CA) was tested by sequencing, resulting in detection of a 139 bp fragment in the normal allele

154

a

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Family pedigree

MARKERS:

D22S1153 D22S1160 D22S1161 D22S922 SCO2 SNP-NlaII

150 162 157 138 R262delCA +

158 133 155 138 N -

148 150 133 162 155 157 138 136 E140K N + -

PGT 150 150 162 162 157 157 138 136 R262delCA E140K + + AFFECTED, RECOMBINANT

b

PGT

EMBRYO #

1

2

3

4

158 133 155 138 N -

150 162 157 136 N -

NORMAL

5

6

7

150 148 150 148 158 150 150 150 158 150 158 150 150 150 162 133 162 133 133 162 162 162 133 162 133 162 162 162 157 155 157 155 155 157 157 157 155 157 155 157 157 157 138 138 138 138 138 136 138 136 138 136 138 136 138 136 R262 E140K R262 E140K N N R262 N N N N N R262 N delCA delCA delCA delCA + + + + + + AFFECTED AFFECTED NORMAL CARRIER NORMAL NORMAL CARRIER

ET

8

158 133 155 138 N -

9

150 162 157 136 N -

NORMAL

150 162 157 138 R262 delCA + MONOSOMY22

0

ET

Fig. 4.37 PGT for cardioencephalomyopathy. (a) Family pedigree of a couple with a previous affected child, who was double heterozygous for E140K and R262del (CA) in the SCO2 gene. Paternal polymorphic markers are shown on the left, and maternal on the right, with the order of the markers and mutation shown on the upper left. (b) Embryo biopsy results revealed two embryos (embryo #1 and #2) homozy-

gous affected, two (embryos #4 and #7) carriers of the paternal mutation, four mutation-free embryos (embryos #3, #5, #6, and #8), and one monosomic for chromosome 22, based on the testing of the mutation and six polymorphic markers. Two mutation-free embryos (embryos#3 and #5) were transferred, resulting in a singleton pregnancy and the birth of an unaffected child. ET embryo transfer

and 137 bp fragment in the mutant allele. Five polymorphic markers – D22S1153, D22S1160, D22S1161, D22S922, and SNP NlaIII  – were also tested simultaneously to avoid misdiagnosis due to ADO. Figure 4.37 shows that of nine embryos tested, two embryos (embryos #1 and #2) were homozygous-­affected, two (embryos #4 and #7) were carriers of the mutant gene, one (embryo #9) monosomic for chromosome 22, and four (embryos #3, #5, #6, and #8) free of the mutation. Two of these embryos (embryos #3 and #5) were transferred, resulting in a singleton pregnancy and birth of a healthy child free from cardio-encephalopathy.

4.5.3.1 Conclusion Results presented show that PGT is a realistic option for couples at risk for producing offspring with cardiac disease, as characterized by inherited predisposition. Inheritance of such susceptibility factors places the individual at risk for a serious cardiac disease, clinically manifested from as early as the first year of life as in cardio-­encephalopathy or later in life with the only clinical realization of premature or sudden death, as in CMD and CMH. Among the conditions in the family history of the couples at risk that may indicate a possible need of PGT may be a heart attack and sudden death at young age and family members with pacemakers or internal cardiac defibrillators,

4.6 Other Genetic Conditions with Important Health-­Related Implications

arrhythmia, and cardiac surgery. The likelihood that offspring of these patients will develop the same heart disease will depend on the mode of inheritance, but penetrance is difficult to predict because many inherited cardiac conditions are difficult to diagnose and will manifest at different ages. Disease may also be induced by certain medications or activities such as excessive exercise that may lead to cardiac arrest or sudden death. All these justly request for PGT-M. In some cases a common, apparently “milder” disease susceptibility gene may contribute to premature death, major disability, or hardship in a family. Personal experience may alter a family’s perception of severity of the condition and is pivotal toward a decision to undertake PGT-­M. Couples already going through IVF for infertility may have questions about the implications of genetic susceptibility for offspring, the option to test embryos, and the appropriateness of using PGT-M for susceptibility to inherited cardiac disease. Because symptoms of inherited cardiac disease may be easily overlooked, findings in the family history may alone provide the reason to test for presence of predisposing gene mutations or the need for PGT.  This may be a lifesaving procedure for individuals at risk. With future advances in identification of genes predisposing to inherited cardiac disease, PGT might appear as a useful tool for couples at risk for producing offspring with inherited cardiac diseases that have high probability of premature or sudden death during their life span.

4.6

 ther Genetic Conditions O with Important Health-­ Related Implications

This heterogeneous group of conditions may be of relevance depending on high prevalence (e.g., rhesus disease (RhD)), whereas others may be of broader importance (e.g., congenital malformations, dynamic mutations). Several other approaches could be applied to assist couples at risk, but PGT is one of the most attractive options for avoiding the risk and ensuring the birth of an unaffected child.

155

4.6.1 Blood Group Incompatibility Mother-Fetus Kell incompatibility represents one of the important causes of blood group incompatibility, similar to rhesus disease (RhD). Although pregnancies at risk for this blood group incompatibility may be detected by prenatal diagnosis and treated by an intrauterine transfusion, complications for the fetus exist and cannot be completely excluded even after the procedure. Pregnancy termination in such cases may also be unacceptable, as the antibodies to K1, for example, are developed only in 5% of persons obtaining incompatible blood. On the other hand, some at-risk couples have had such unfortunate experience in hemolytic disease of the newborn (HDN), resulting in neonatal death, that they regard PGT-M as their only option for another pregnancy. This makes PGT-M attractive for patients at risk for alloimmunization, even if conditions have rarely been an indication for prenatal diagnosis in ongoing pregnancies. The K1 system is one of the major antigenic systems in human red blood cells, comparable in importance to RhD. It may cause maternofetal incompatibility leading to a severe hemolytic disease of the newborn (HDN) in sensitized mothers. The K1 allele is present in 9% of the populations, in contrast to its highly prevalent allelic variant K2. The locus is on chromosome 7 (7q33), consisting of 19 exons. In K1 a C to T base substitution in exon 6 in K1 differs from K2 antigen and leads to a threonine to methionine change at amino acid residue 193, preventing N-glycosylation [82, 83]. C to T base substitution also creates a BsmI restriction enzyme site, providing a reliable DNA test for diagnosis of KEL genotype. In pregnancy in which the fetus is K1 and the mother K2, antibodies to K1 may develop, leading to maternofetal incompatibility causing severe HDN. As mentioned, although prenatal diagnosis is available for identification of pregnancies at risk for HDN, this alone does not prevent potential complications for fetus, stillbirth, or neonatal death. Thus, PGT is an attractive option for preventing both Kell and rhesus hemolytic diseases. We present here PGT for a couple, having paternal K1/K2 genotype with K1 heterozygous

156

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

for C to T base substitution in exon 6. In this couple, a 36-year-old mother had a previous dizygotic twin pregnancy, resulting in death of one of twins carrying the K1 allele due to HDN (Figs. 4.38 and 4.39). As a result of PGT, two K1 allele-free embryos were transferred, yielding two healthy twins with K2/K2 genotype. In the other couple, a 37-year-old mother had three previous pregnancies. The first resulted in the birth of a healthy boy carrying the K1 allele. The second resulted in the birth of a normal K2/ K2 boy and the third in a premature delivery of a 32-week female carrying the K1 allele who died the next day with the clinical features of a severe HDN. To establish paternal haplotypes for PGT, single-­sperm analysis was performed to be able to undertake linked marker analysis in addition to KEL genotyping. Short tandem repeats (STRs) associated with the cystic fibrosis (CFTR) gene were used, given this locus is close to K1 and K2 alleles with extremely rare recombination rates [84]. This analysis showed the presence of infor-

mative linked markers CFTR intron 1 D7S550, CFTR intron 6, and CFTR intron 8. The K1 allele was linked to the 158 (D7S550), 7 (CFTR intron 6), and 124 (CFTR intron 8) repeats. Thus, in PGT-M cycle, multiplex nested PCR analysis was performed, using the BsmI restriction site simultaneously with these linked polymorphic markers, the D7S550, CFTR intron 6, and CFTR intron 8. Outside and inside primer sequences and primer melting temperatures for DNA analysis were reported elsewhere [85–87]. Overall, 41 PGT cycles were performed for 23 couples, including 11 cycles for rhesus and 30 for Kell incompatibility, resulting in transfer of 45 compatible embryos in 26 transfers, with 12 healthy children born. Although the predicted genotypes were confirmed overall, six cases of recombination between K alleles and two linked markers (CFTR intron 6 and CFTR intron 8) were observed, suggesting limited value of these two markers on their own. However, no ­recombination was observed between the gene and intron 1 CFTR, verifying absence of the KI

Family pedigree

Markers order: 1. KELL (Bsm I) 2. Intron 1 CFTR

I

K1

K2

K2

K2

118

116

118

112

II

PGT

K2

K2

116

118

K1

Fig. 4.38  PGT for Kell genotype: family pedigree. (I) The father (left) has K1/K2genotype, K1 allele linked to 118 bp repeats, and K2 allele to 116 bp repeats of intron 1 of CFTR polymorphic marker, while the mother (right) has K2/K2 genotype, one allele linked to 118 bp repeats, and the other to 112 bp repeats of intron 1 of CFTR poly-

K2

PGT

K2

K2

K2

K2

116

112

116

118

morphic marker. (II) Reproductive outcomes of this couple, including previous twin pregnancy resulting in the death of one of the twins near birth due to HDN.  Two healthy twins with K2/K2 genotype resulting from PGT were born confirmed also by linked polymorphic markers

4.6 Other Genetic Conditions with Important Health-­Related Implications

KEL1

157

Thr 193 Met ( C-T ) S

a

Exon 6 R

KEL2

Restriction map : 156 bp

b

K2 ( Thr ) K1 ( Met )

100 bp

56 bp

BsmI ( 5’GAATGCT 3’) ET

ET L 1

3 4

5

6 7 8

9 10 11 12 13 Un

L

14 16 17 18 F M

c 156 bp K2 100 bp K1

K2

K2 /K1 K2 /K1

K2 /K1 K2 /K1 K2

K2 /K1

K2 K2 K2 /K1

K2 /K1 K2 K2 /K1 K2

K2 K2 /K1

K2 K2 /K1

56 bp K1

Fig. 4.39  PGT for Kell genotype. (a) Schematic diagram showing C  >  T substitution in exon 6 of KEL gene on chromosome 7: black arrows demonstrate the positions of nested primers. (b) Restriction map for BsmI digestion, showing the gain of BsmI site by the K1 allele (lower line). (c) Polyacrylamide gel electrophoresis of the BsmI digested PCR products of 16 blastomeres from the first PGT couple, demonstrating K1 allele-free genotype in

embryos #1, #4, #7, #9, #10, #11, and #17, from which embryos #1 and #9 were transferred, resulting in a twin pregnancy and the birth of healthy K1 allele-free children. The remaining nine embryos have K1/K2 genotype. L standard, F paternal DNA amplified from sperm, M maternal normal amplified DNA, Un undigested PCR product, K1/K2 affected blastomere, K2/K2 normal blastomere

allele and detecting both maternal and paternal K2 alleles in the embryo, thus improving considerably the reliability of PGT-­M.  It is of importance that preselection of the K1-free embryos was based not only on the absence of the K1 allele, which may be also explained by ADO, but also on the absence of the linked intron 1 CFTR (118 repeats) marker and presence of both the paternally and maternally derived K2 alleles, as evidenced by the presence of polymorphic markers linked to the paternal and maternal K2 alleles. In other words, absence of the K1 allele together with the presence of both paternal and maternal K2 alleles allowed reliably preselecting the KI allele-free embryos for transfer.

Rhesus disease has been an indication for PGT in a number of cases but initially did not result in a clinical pregnancy [88]. However, later PGT-M attempts were successful, yielding a clinical pregnancy and the birth of a healthy girl confirmed to be blood type Rh-negative [89]. This was done for a couple having an Rh-negative mother and an Rh-positive father with two children, one of whom was affected by HDN characterized by hyperbilirubinemia, neonatal jaundice, and hemolytic anemia. Because of the RhD alloimmunization, the couple was presented with the dilemma of whether to attempt a further pregnancy, given the tendency of rhesus-incompatible pregnancy in sensitized women worsening in

158

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

subsequent pregnancies. Using embryo biopsy and direct PCR amplification with analysis by capillary electrophoresis of fluorescently labeled amplicons, RhD-negative embryos were preselected for transfer. As seen from Table  4.3, we have currently performed 30 cycles for rhesus disease, with the transfer of RhD-negative embryos in 17 of 30 cycles, resulting in the birth of five Rh-compatible children. As mentioned, both Kell and Rh disease are frequent, with 15% prevalence for RhD and 9% for KEL antigen. Alloimmunization may lead to HDN in the offspring of some of at-risk couples. Therefore, PGT-M may be a practically useful option for couples in avoiding establishment of RhD or K1 pregnancies in the sensitized mothers [90].

4.6.2 Congenital Malformations Congenital malformations are highly heterogeneous conditions for which applicability of PGT depends on the gene involved. Prevalence of these conditions depends on definition but has cited as high as 29.3/1000 live births. Usually cases are sporadic, and their reduction has come mainly from population-based preventive measures such as folic acid fortification of major foodstuffs (Chap. 1). However, reduction by this approach will not have sufficient impact on the prevention of the well-established inherited forms, for which PGT is clearly the preferable option. The Human Genome Project has enabled a number of inherited forms, which can be avoided through PGT. Our experience of PGT for multiple congenital malformations is summarized in Table  4.21, with PGT for craniofacial disorders shown separately in Table 4.22. Overall, 84 PGT cycles were performed for multiple malformation, resulting in 46 (55.4%) unaffected pregnancies and birth of 45 healthy children. Of special interest in this experience are Currarino syndrome and sonic hedgehog (SHH) described below. We performed 16 PGT cycles for Crouzon syndrome and 52 for craniofacial disorders, described below in the example of sonic hedgehog (SHH) mutation.

Sonic Hedgehog (SHH)  Mutations are an example for which PGT is an attractive option. The SHH gene is a human homolog of the Drosophila gene encoding for inductive signals involved in patterning the early embryo, known to be (functionally) highly conserved in many species. SHH mutations may cause failure of cerebral hemispheres to separate into distinct left and right halves, leading to holoprosencephaly (HPE), one of the most common developmental anomalies of the forebrain and midface [91]. The gene is mapped to chromosome 7 (7q36), previously designated as the locus for the gene involved in holoprosencephaly (HPE3) [92]. SHH protein is an intercellular signaling molecule, synthesized as a precursor undergoing autocatalytic internal cleavage into both a highly conserved domain (SHH-N) with signaling activity and a more divergent domain (SHH-C), which in addition to precursor processing acts as an intramolecular cholesterol transferase crucial for proper patterning activity in animal development. Although the effect of the above nonsense mutation on SHH function is unknown, the resulting protein may not fulfill its expected signaling function in early morphogenesis [16, 92]. Although the majority of HPE are sporadic, familial cases are not rare, as clear autosomal dominant inheritance shows. Intra-familial clinical variability exists in HPE: alobar HPE and cyclopia to cleft lip and palate, microcephaly, ocular hypertelorism, and even normal phenotype. This suggests interaction of the SHH gene with other genes during craniofacial development, as well as possible involvement of environmental factors, which may explain the almost one-third of the carriers of SHH mutations who are clinically normal. Therefore, even in familial cases, the detection of SHH mutations in prenatal diagnosis might confront the decision for pregnancy termination, making PGT a more attractive option for couples at risk for producing progeny with HPE. A couple undergoing PGT for HPE is presented in Figs. 4.40 and 4.41. This couple had two children with clinical signs of HPE [93]. A female with severe HPE and cleft lip and

Disease Aicardi–Goutieres syndrome 2; AGS2 Aicardi–Goutieres syndrome 5; AGS5 Alagille syndrome 1; ALGS1 Angelman syndrome; AS Bardet–Biedl syndrome 2; BBS2 Bardet–Biedl syndrome 4; BBS4 Bardet–Biedl syndrome 10; BBS10 Cohen syndrome; COH1 Currarino syndrome DiGeorge syndrome; DGS Donnai–Barrow syndrome Fetal akinesia deformation sequence; FADS1 Fetal akinesia deformation sequence; FADS2 Fetal akinesia deformation sequence; FADS4 Fraser syndrome 1; FRASRS1 Harel–Yoon syndrome; HAYOS Lissencephaly, X-linked, 2; LISX2 Marinesco–Sjogren syndrome; MSS Meckel syndrome, type 1; MKS1 Meckel syndrome, type 4; MKS4 Meckel syndrome, type 6; MKS6 Meckel syndrome, type 8; MKS8 Midface hypoplasia, hearing impairment, elliptocytosis, and nephrocalcinosis; MFHIEN Multinucleated neurons, anhydramnios, renal dysplasia, cerebellar hypoplasia Smith–Lemli–Opitz syndrome; SLOS Sonic hedgehog; SHH Temtamy syndrome; TEMTYS Total

Table 4.21  PGT-M for multiple malformation

4 1 1 0 1 3 2 1 3 2 3 15

6

2 1 1 1 1 3 1 1 3 1 3 13

8 1 32 2 1 84

1

236500 270400 19 600725 1 218340 1 53

CEP55 DHCR7 SHH C12orf57

23 2 2 83

2

# of transfers 1 0 1 2 8

# of cycles 1 1 1 2 4

# of OMIM patients 610181 1 612952 1 118450 1 105830 2 615981 3 615982 615987 216550 2 176450 1 188400 1 222448 1 208150 1 618388 2 602552 1 219000 1 617183 1 300215 1 248800 1 249000 8 611134 612284 613885 301050 2

Gene RNASEH2B SAMHD1 118450 UBE3A BBS2 BBS4 BBS10 VPS13B MNX1 TBX1 LRP2 MUSK RAPSN NUP88 FRAS1 ATAD3A ARX SIL1 MKS1 CEP290 CC2D2A TCTN2 AMMECR1

32 3 2 109 1.3

2

9

4 1 1 0 1 5 2 1 3 3 5 22

# of embryos transferred 1 0 1 3 8

15 1 0 46 55.4%

1

3

3 2 1 0 1 3 1 1 1 0 2 6

Pregnancy 0 0 1 1 3

1

2

2 1 1 0 1 2 1 0 0 0 1 6

# of deliveries 0 0 1 1 2

0 15 0 1 0 0 8 38 17.4% 82.6%

0

1

1 1 0 0 0 1 0 1 1 0 1 0

SAB 0 0 0 0 1

17 1 0 45

1

2

3 1 1 0 1 2 1 0 1 0 1 8

# of babies 0 0 1 2 2

4.6 Other Genetic Conditions with Important Health-­Related Implications 159

Disease Branchiooculofacial syndrome; BOFS Cranioectodermal dysplasia 2; CED2 Craniofrontonasal syndrome; CFNS Crouzon syndrome Holoprosencephaly 2; HPE2 Hydrocephalus due to congenital stenosis of aqueduct of Sylvius; HSAS Pfeiffer syndrome Popliteal pterygium syndrome; PPS Treacher Collins syndrome 1; TCS1 Treacher Collins syndrome 2; TCS2 Total

Table 4.22  PGT-M FOR craniofacial disorders OMIM TFAP2A WDR35 EFNB1 FGFR2 SIX3 L1CAM FGFR1 IRF6 TCOF1 POLR1D

Gene 113620 613610 304110 123500 157170 307000 101600 119500 154500 613717

2 5 6 1 38

# of patients 1 1 1 9 1 11 2 5 8 1 52

# of cycles 1 1 1 16 1 16 2 5 8 1 50

# of transfers 1 1 1 14 1 16 4 6 14 1 88 1.76

# of embryos transferred 2 1 1 23 2 34 2 3 7 0 29 58%

Pregnancy 0 1 0 9 0 7

0 0 0 0 3 10.3%

SAB 0 0 0 1 0 2

2 3 7 0 26 89.7%

# of deliveries 0 1 0 8 0 5

2 3 9 0 29

# of babies 0 1 0 8 0 6

160 4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

4.6 Other Genetic Conditions with Important Health-­Related Implications

161

Family pedigree SHH 1

SHH 2

Markers order: N 152

1. SHH 2. D7S550

N 138

M 156

N 158

PGT SHH 3 N 158

PERINATAL DEATH

M 156

N 152

N 158

Fig. 4.40  PGT for sonic hedgehog (SHH) mutation: family pedigree. (Upper panel) The father has a gonadal mosaicism for SHH mutation, which is linked to 156 bp dinucleotide C-A repeat allele of D7S550 polymorphic marker, while the mother is normal, with one normal allele linked to 158  bp repeat and the other to 138  bp repeat alleles. (Lower panel) Reproductive outcomes of this couple, including three previous pregnancies, one resulting in the birth of an affected child with holoprosen-

cephaly, carrying the mutant gene (lower left), one in perinatal death, also carrying the mutant gene (lower middle(circle)), and one in a spontaneously aborted fetus with Turner syndrome, free from SHH mutation (lower middle(triangle)). The lower right (PGT) shows the outcome of PGT resulting in an unaffected clinical pregnancy and the birth of a healthy child, following confirmation of the mutation-free status by amniocentesis

palate died shortly after birth. Chromosomal analysis performed using peripheral blood lymphocytes of both this child and parents was normal, but DNA analysis in the child’s autopsy material demonstrated a SHH nonsense mutation, a GAG  >  TAG sequence change leading to premature termination of the protein at position 256 (Glu256  →  stop) [91] (Fig. 4.41). The same mutation was found in their 5-year-­ old son, who was born after a full-term normal pregnancy weighing 6 lb, with a birth length of 18 1/3 inches. This child had less severe facial dysmorphic features but did show microcephaly, Rathke’s pouch cyst, a single central incisor, cho-

anal stenosis (treated surgically after birth with dilatation), and clinodactyly of the fifth fingers and incurved fourth toes bilaterally. Growth was slow in the first 2 years, but thereafter maintained a reasonably progress and normal social and cognitive development. The couple had another pregnancy that ended in spontaneous abortion due to Turner syndrome (45, X), but showed no inheritance of the SHH mutation. The mutation was not found in either parent’s genomic DNA, although paternity testing showed that the father was the true biological father of both affected children. This suggested a de novo gonadal mutation in one of the parents, as indeed identified by a single-sperm genotyping.

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

162

a

e

Glu256 Stop

SHH Exon 3 b

D7s550

Blastomere from embryo # 2

213 bp

Normal

Paternal affected Maternal

Affected

133 bp

80 bp Xba I

c

L

2 4 5

(CA)n

Blastomere from embryo # 9 Paternal affected

8 9 10 16 17 19 F Mo S Un

Maternal

ADO

Normal

Blastomere from embryo # 10 Paternal affected

Affected Maternal

ADO

ADO

A N N N A

d

L

2

ADO

A

N A

Affected A A N A

Blastomere from embryo # 17 Maternal

9 10 17 19 S Un

Normal Affected

Paternal affected

Whole embryo #17 Maternal

Paternal affected

ADO Affected A

A A A A A

Fig. 4.41 PGT for sonic hedgehog (SHH) mutation involving confirmation of the presence of both maternal and paternal normal genes in preselected mutation-free embryo. (a) Schematic diagram of the mutation and D7S550 linked marker on chromosome 7. Black arrows demonstrate the positions of heminested primers. (b) Restriction map for XbaI digestion, showing the gain of XbaI site by the mutant allele (lower line). (c) Polyacrylamide gel electrophoresis of the XbaI digested PCR products of nine blastomeres from PGT cycle. (d) Follow-up DNA analysis of genomic DNA from five embryos predicted to be affected by blastomere testing. (e) Capillary electrophoregrams of fluorescently labeled PCR product of tightly linked marker D7S550. Paternally derived 156  bp dinucleotide C-A repeat linked to SHH

mutation are shown by arrow (noted as “paternal affected”) in blastomeres of embryos #2, #9, #10, and #17 and the genomic DNA of the whole embryo #10, in which ADO of mutant gene was seen in the follow-up study (see panel b, embryo #10). Maternally derived 158 bp dinucleotide C-A repeats of D7S550 polymorphic marker are shown by arrow (noted as “maternal”) in blastomeres of embryos #2, #9, and #10, while the other maternally derived 138  bp repeats of D7S550 are shown in blastomeres of embryo #17. L 100  bp standard, ADO allele dropout, F paternal DNA amplified from sperm, Mo maternal normal amplified DNA, amplified DNA from affected baby, Un undigested PCR product, A affected blastomere

4.6 Other Genetic Conditions with Important Health-­Related Implications

Two PGT-M cycles were performed by embryo biopsy and multiplex nested PCR analysis, involving specific mutation testing simultaneously with linked marker analysis. Of the 15 embryos obtained in the first cycle, 12 were available for embryo biopsy. Four failed to amplify, leaving eight with available data for mutation analysis. Seven of these eight embryos appeared to contain the mutant allele; thus, only one embryo was mutation-free and transferred, yielding no clinical pregnancy. A second PGT-M cycle was performed 1 year later, producing 19 embryos, of which 10 were biopsied; 9 of the 10 amplified were tested for the SHH gene and the marker to identify mutation-free embryos for transfer (Fig. 4.41). Prior to PGT cycles, a single-sperm testing had been performed, identifying mosaicism for SHH mutation. As the mutation was shown to lead to the gain of an XbaI restriction site [91], the normal allele was identified on the basis of an undigested PCR product; the mutant allele was represented by two fragments (Fig. 4.41). To avoid misdiagnosis in mutation analysis due to ADO, a closely linked microsatellite DNA marker D7S550 was tested in the same reaction as the internal control. Primers used in the first- and second-round PCR for mutation and linked marker analysis and reaction conditions were described elsewhere [93]. Haplotype analysis showed that the mutant allele was linked to the 156 bp dinucleotide CA repeat, whereas the normal gene was linked to 152 bp repeat allele in 7q36 (Figs. 4.40 and 4.41). Other linked markers have been described [92], but not informative in this couple. As seen from Fig.  4.41, four ADOs were observed in the mutation analysis: ADO of the mutant allele in embryos #2, #9, and #17 and ADO of the normal allele in embryo #10. This was based on marker analysis showing that in all four cases, embryos were heterozygous. That is, embryos (#2, #9, and #17) could have been misdiagnosed as normal without linked marker analysis. Embryo #19 also contained the mutant gene. Remaining

163

four embryos were free of the mutant gene, as confirmed by marker analysis showing that all these embryos contained two normal alleles. The paternal allele was linked to the 152 repeat; one normal maternal allele was either linked to 138 repeat (embryos #4 and #5) or to other normal maternal allele linked to 158  bp repeat (embryos #8 and #16). Two of these embryos (embryos #4 and #5) were transferred, resulting in a singleton pregnancy and the birth of a healthy child following confirmation of the mutation-free status by amniocentesis. The other two mutation-free embryos (embryos #8 and #16) were frozen. The presented data demonstrate diagnostic accuracy using multiplex PCR-based embryo analysis despite availability of only one linked marker (not three as concurrently recommended). Preselection of mutation-free embryos has been based not only on the presence of the paternally derived normal allele, but also on the presence of a second normal allele linked to a maternal-­ linked marker. That is, absence of the mutant gene together with presence of two normal alleles, identified by different linked makers, allowed identification of embryos as either normal or carrying the SHH mutation. Currarino Triad  Is another example of the usefulness of PGT for avoiding congenital malformations [94, 95]. Application of prenatal diagnosis is less attractive because multiple factors may modify clinical manifestations and confound prediction of an individuals’ phenotype. Obviously, PGT is a more attractive choice in these circumstances. Currarino triad, also known as Currarino syndrome (CS), is a severe autosomal dominant disorder caused by a homeobox gene HLXB9 mutation. Phenotype involves partial sacral agenesis, presacral mass, and anorectal malformations, which are one of the commonest digestive anomalies requiring neonatal surgery [16, 17]. The causative homeobox gene is located on chromosome 7q36, linked to microsatel-

164

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

lite DNA markers D7S559 and D7S2423 [96]. Abnormalities observed in CS are caused by disturbances in early embryonic development of the human tail bud, leading to aberrant positioning defects involving the neural tube, notochord, somites, and hindgut. More than two dozen different HLXB9 intragenic mutations and microdeletions have been detected in patients with CS, including frameshift and nonsense mutations and missense mutations in the homeodomain resulting in non-conservative substitutions of a highly conserved amino acid [16, 97–99]. Although carrier screening and prenatal diagnosis of CS are currently available, the decision about termination of a clinical pregnancy is controversial because approximately half of the carriers of the mutation are asymptomatic [99]. An abnormal genotype does not necessarily indicate phenotype abnormalities and thus confounding decision on termination of clinical pregnancy. Absence of robust genotype–phenotype correlation and high variability of phenotype in the carriers  – severe anomalies to minor sacral abnormalities undetectable without X-ray to even a completely asymptomatic carrier status – make PGT an important alternative for the at-risk couples wishing to avoid CS in their liveborn offspring. Figure 4.42a describes a couple presented for PGT-M because a previous child was diagnosed to have CS.  The child was born with imperforate anus, an anterior meningocele, and characteristic sickle-shaped hemisacrum revealed by X-ray of the sacrum region (Fig.  4.42a; IV 1). The father also had an anal stricture (Fig. 4.42; III 3) requiring anal dilatation. A sacral defect was detected by X-ray, involving central anomaly from S2 downward. One of his two sisters (Fig. 4.42a; III 1) was born with imperforate anus, anterior meningocele rectovaginal fistula, and vesicoureteric reflux resulting in the need of renal transplantation; sacral X-ray showed the similar to the father the central defect with absence of the distal one-third of the sacrum. The other sister (Fig. 4.42a; III 2) was clinically asymptomatic, but sacral X-ray revealed absence of a coccyx; MRI scan disclosed an anterior meningocele. The mother of the father of the index case had an undeveloped coccyx and

urinary tract bilateral ureteropelvic junction obstruction (Fig. 4.42a; II 4). Of this grandmother’s three siblings, one of her two brothers had anal stricture (Fig.  4.42a; II 1), but the asymptomatic sister (Fig. 4.42a; II 2) also appeared to be a carrier because her grandson had an imperforate anus. The father inherited the mutation from his grandmother (Fig. 4.42a; I 2), who was probably the first affected member of the family, having constipation but a normal sacral X-ray. DNA analysis in this family demonstrated a homeobox HLXB9 mutation due to frameshift insertion of a cytosine into a stretch of six cytosines at positions 125–130 in exon 1, leading to the introduction of premature codon termination [96]. The requested PGT cycle was performed using embryo biopsy and multiplex nested PCR analysis, to test for the specific mutation simultaneously with linked markers. The PCR product was identified by fragment length analysis using capillary electrophoresis. Prior to initiating PGT cycles, single-sperm testing had been performed to establish paternal haplotypes. To identify ADO in the mutation analysis, four closely linked microsatellite dinucleotide DNA markers, D7S559, D7S550, D7S637, and D7S594, were tested in the same reaction with the HLXB9 gene [96]. The mutant allele was linked to markers of 119, 155, 124, and 162 bp repeats; the normal allele was linked to 123, 159, 116, and 168 bp repeats of D7S559, D7S550, D7S637, and D7S594 markers, respectively. Testing for these markers in the affected child and the mother showed that the parents shared markers of the same size for two of the four markers linked to the normal allele, making these markers of limited value for preselection of mutation-free embryos for transfer. Overall, 17 embryos were available for testing in a single PGT cycle; 3 failed to amplify either alleles or polymorphic markers, suggesting lack of a nucleus in these biopsy samples; 2 others showed amplification of only polymorphic markers, with no signal detected for the HLXB9 gene. Of the total of 12 embryos with results for the mutation and linked marker analysis, 11 showed conclusive results; 5 were pre-

4.6 Other Genetic Conditions with Important Health-­Related Implications

a

165

Family pedigree

I

1

II

1

2

2

3

4

III 1

2

3 162 M 119 155 124

4 Markers order: D7S594 HLXb9 D7S559 D7S550 D7S637

168 170 N N 117 123 155 155 116 122

170 N 123 158 116

IV

PGT

b

1

2

162 168 M N 119 117 155 155 124 116

170 170 N N 123 123 158 155 116 122

Insertion C 125-130

(CA)n

HLXB 9

D7S594

Exon 1

(CA)n

(CA)n

(CA)n

D7S559

D7S550

D7S637

Embryo # 4 ( Normal –transferred ) Hex 168 170

Fam

Fam 105

117

Fam

Hex 155

123

159

116

Embryo # 16 ( Normal – transferred ) Fam Hex 168 170

Fam

Fam

Hex 155

123

105

159

116

122

Embryo # 5 ( Affected ) Fam

Fam

Hex 162

170

105 106

119

123

Fig. 4.42 (a) PGT for Currarino syndrome: Family pedigree. I and II Patient’s parents and grandparents with signs of symptoms of the disease in his mother and grandfather. Affected father (III, 3) has two affected sisters; homeobox gene HLXB 9 mutation was inherited from their mother (II, 4), who also had two affected siblings (brother II, 1; and sister II. 2). Reproductive outcome is shown in lower panel, with the previous affected child (IV. 1) and the healthy baby boy born after PGT (IV. 2). (b) PGT for homeobox gene HLXB9 mutation in exon 1 causing Currarino syndrome. Upper panel shows the location of the mutation in HLXB9 gene and linked

Fam

Hex 155 /155

122 124

markers on chromosome 7. The other three panels show capillary electrophoregrams of fluorescently labeled PCR products of HLXB9 alleles and each of the four linked markers. Paternally derived mutant allele is shown by arrow in embryos #5 (lower panel), in agreement with paternally derived markers (CA repeats) linked to the mutant gene. The mutant allele is absent in embryos # 4 and #16 (middle panels), also in agreement with all four markers. These embryos have been transferred back to the patient resulting in the birth of mutation-free baby. According to the marker analysis the baby originated from the transfer of embryo #16

166

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

dicted to contain the mutant allele in agreement with the presence of repeat markers. One of these embryos contained only paternal alleles, due to absence of maternal chromosome 7. The remaining six embryos were predicted to be free of mutation, but only in three (shown in Fig. 4.42b) could ADO of the mutant allele be excluded based on linked marker analysis. Two embryos were transferred, yielding a singleton pregnancy and birth of a mutation-free child, followed by confirmation of diagnosis by amniocentesis. In the other three embryos predicted to contain the normal allele, ADO of the mutant gene could not be excluded because of the parents’ shared same size polymorphic markers linked to the normal allele. For example, one embryo was informative only for a single linked marker, D7S637 (116/122  bp). This might suggest the presence of both paternal and maternal normal alleles, assuming both of these alleles were not of maternal origin. Alternatively, uniparental disomy 7 of maternal origin could exist, albeit not supported by presence of other linked markers. In the other two embryos, ADO of the mutant allele could not be excluded, despite these embryos being heterozygous for the two linked markers, D7S637 (116/122) and D7S550 (159/155). Alleles detected (116 and 159 bp) and potentially linked to the paternal normal gene may have also been derived from the mother, who shares the same size of linked marker linked to the normal allele. In one embryo there was failure of amplification of HLXB9 alleles and might have been predicted to be mutant based on presence of three of the four polymorphic markers linked to the mutant gene. However, this was not in agreement with the presence of 170  bp repeat of the D7S594 marker, linked to the normal paternal allele, which may probably have been due to recombination of the paternal alleles. Unfortunately, parental sharing of two of the four markers linked to the normal allele left only three of six potentially normal embryos for transfer because of the given inability to exclude completely the risk for misdiagnosis due to ADO of the mutant paternal allele. This case was the first PGT performed for a homeobox-containing gene mutation, demon-

strating clinical application of PGT for this group of genes, which are involved in transcriptional regulation during early embryonic development. Because of the high prevalence of congenital anomalies determined by the mutations causing the formation and positioning defects, this approach for PGT has practical implications also for at-risk couples with other groups of familial dysmorphologies. PGT is an attractive new option for many couples wishing to avoid the risk of having children with a wide range of formation and positioning abnormalities at different stages of embryogenesis. Data on the stage-­ specific expression of homeobox genes observed in CS are also of relevance for the development of PGT for these transcriptionally relevant anomalies to offer different options for the couples at risk for producing progeny with inherited predisposition to congenital malformations. Crouzon Syndrome (CFD1)  Is another example of usefulness of PGT technology for couples at risk for offspring with congenital malformations (Fig. 4.43). CFD1 is an autosomal dominant craniofacial dysostosis, caused by mutations in the fibroblast growth factor receptor 2 (FGFR2) gene on chromosome 10q [16, 100]. Crouzon syndrome is quite rare, with birth prevalence of 15–16 per million [101]. However, FGFR2 causes a number of disorders and in aggregate contributes a large portion of de novo paternal age-related mutations. The first reported PGT case for Crouzon syndrome was applied by testing of G/A base pair substitution at codon 568  in a couple with one affected child, who had died aged 18  months during corrective surgery. Of two PGT cycles, one resulted in the birth of an unaffected child [94]. As shown in Table 4.22, we performed 16 PGT cycles for 9 couples at risk of producing offspring with Crouzon syndrome, resulting in transfer of 23 unaffected embryos in 14 cycles, yielding 9 unaffected clinical pregnancies and birth of 8 healthy children. In one of these couples, the mother was a carrier of the C3422Y mutation in exon B of the FGFR2 gene, and had an affected child with Crouzon syndrome (Fig.  4.43). Two cycles were performed, using

4.6 Other Genetic Conditions with Important Health-­Related Implications

Family pedigree

Markers order: FGFR2 D10S190

1.1 N N 126 118

1.2 N M 120 118

PGT 2.1 N M 118 118

2.2 N N 126 120

Fig. 4.43  PGT for Crouson syndrome: family pedigree. The mother (1.2) is the carrier of a mutation in FGFR2 gene and had a previous child with Crouson syndrome (2.1). PGT resulted in the birth of a healthy unaffected boy (2.2)

PB1 and PB2 testing and restriction digestion with HpyCH4 V.  The restriction endonuclease resulted in the normal allele having 74 and 20 bp fragments while leaving the mutant allele intact (94 bp). Two informative linked markers, D10S190 and Msp I polymorphism, were amplified simultaneously with the causative gene for testing PB1 and PB2. This allowed preselection of mutation-free oocytes. Of nine oocytes tested in the first cycle, five appeared to be mutation-free, but the transfer did not result in clinical pregnancy. In the other cycle, 22 oocytes were available for testing; 9 were mutation-free (Fig.  4.44). Transfer of two of these embryos (embryos #7 and #11) resulted in a singleton pregnancy. Birth of an unaffected child was confirmed to be free of the causative gene.

4.6.2.1 Embryonic Lineage and Differentiation With improvements in embryo culture media, ovulation stimulation, and embryo transfer, livebirth rates in ART will surely become higher than the 50–60% currently expected. The impediment reflects in part our inability to exclude mutations

167

in single (protein coding) genes that are responsible for errors in embryogenesis. To appreciate our incomplete knowledge, recall that congenital anomalies in liveborns are more often caused by perturbation of single genes (1%) or complex factors (1%) than chromosomal abnormalities (0.6%). All three causes are assumed to play roles in embryonic cell lineage and differentiation. A well-known example involves embryonic gene OCT 4, required for progression beyond the four-­ cell embryo. As mutant genes deleterious for perturbations of cell lineage and differentiation increasingly become enumerated, they can be interrogated through PGT-M or cell-free DNA platforms. It would be helpful to exclude embryos having such mutations from transfer. This strategy is analogous to use in prenatal genetic diagnosis targeted organ system platforms, whole-exome sequencing (WES), or whole-­genome sequencing (WGS) in chorionic villi or amniotic fluid cells. The current difficulty with the above approach is a lack of data on human embryo genes. For only one-third of the 21,000 protein-coding genes function is known, while the function of the remaining two-thirds is still unclear, although doubtless many play roles in embryogenesis and if perturbed would have deleterious effects despite being euploid. Once identified, it should be facile to test and exclude these mutations from transfer. In fact, evidence exists that euploid miscarriages show many of the same aberrant features characteristic of aneuploid embryos. Using array CGH and embryoscopy in a cohort of early miscarriages, it was reported that the first-trimester euploid miscarriages were usually characterized by growth retardation and developmental abnormalities. Only one-quarter were morphologically normal and euploid. WES and WGS detect additional mutations responsible for miscarriages. These genes can be sequenced or aggregated into panels interrogated to exclude embryos for transfer.

4.6.3 Dynamic Mutations Dynamic mutations represent another important group of inherited conditions caused by trinucleotide repeat expansion, already mentioned for

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

168

Cys 342 Tyr G>A

1F

a

(CA)n 0.6cM

FGFR 2

D10S190

Exon B 4R

e

2R

Normal allele 5’. . . ATACGTGCTT….. 3’ Mutant allele 5’. . . ATACGTACTT ….. 3’ 3’ end of 4R primer 3’ ….GTA CC…. 5’ HpyCH4 V restriction site TG’CA

b

Oocyte # 7 ( NORMAL )

PB1 118 120

PB2

74 bp

c

20 bp

Oocyte # 11( NORMAL )

Mutant allele

94 bp

PB1 PB2

PB1 PB2

PB1 PB2

PB1 PB2

PB1 PB2

L PB1 PB2

PB1 PB2

PB1 PB2

PB1 PB2

PB1 PB2

PB1 PB2

PB1 L PB1 PB2

d

118

Normal allele

118 120

PB2 118

Ado

Mutant allele

Oocyte # 1 ( AFFECTED )

Normal allele Oocyte # 1

3

4

5

6

7

ET

8

11

12

ET

13

14

15

PB1 118 120

PB2 120

Fig. 4.44  PGT for mutation in FGFR2 gene causing Crouson syndrome. (a) Map of human FGFR2 gene, showing sites and location of C342Y mutation and position of linked D10s190 dinucleotide STR.  Horizontal arrows show primer sets for heminested PCR. (b) Primer design and (c) restriction map for normal and abnormal alleles. (d) Mutation analysis of 12 oocytes by sequential first (PB1) and second (PB2) polar bodies. Allele dropout

(ADO) of the mutant allele (∗) was detected both by sequential PB1 and PB2 study (identical genotype of both) and linked marker analysis (presence of 118 bp and 120 bp bands). STR profile for PB1 and PB2 from oocytes #1, #7, and #11 confirmed the results of mutation analysis (e), suggesting that the latter two are normal and may be transferred, which resulted in birth of unaffecetd child. L size standard, bp base pair, ET embryo transfer

fragile X syndrome; these are currently among the most frequent indications for PGT-M, along with X-linked disorders, breast cancer, CFTR, and hemoglobin disorders. PGT for a dynamic mutation was first introduced for the couples at risk for producing offspring with myotonic dystrophy (DM) [102]. As was also showm above in PGT for FXRM1, PGT for this group of diseases presents a significant complexity [103, 104]. Other dynamic mutations for which PGT was applied include the prototypic disorder Huntington disease (HD), spinal and bulbar muscular atrophy (SBMA), and spino-­ cerebellar ataxia (SCA) type 2, 3, 6, and 7.

For this group of conditions, identification of normal alleles is mainly based on testing for a sufficient number of linked markers. The reason is that amplification of the mutant region is hazardous given the nature of dynamic repeats. To avoid misdiagnosis, at least three closely linked informative markers should be present in order to confirm inheritance of the normal allele from the affected parent. In practice, presence of normal alleles from both parents should be confirmed by analysis of the maternal and paternal haplotypes, as demonstrated by the example of PGT for DM and Machado–Joseph disease (SCA3), presented in Figs. 4.45 and 4.46.

4.6 Other Genetic Conditions with Important Health-­Related Implications Family pedigree

Markers order: D19S559 ApoC (CA)n DMPK

169

167 128 214(N)

171 153 205(N)

159 151 Exp

167 128 232(N)

PGT

PGT

167 128 214(N)

Predicted Genotype:

167 128 232(N)

Embryo #1 Normal ET

167 128 214(N)

167 128 232(N)

Embryo #2 Affected Trisomy 19

159 151 Exp

171 153 205(N)

167 128 232(N)

Embryo #8 Normal

171 153 205(N)

159 151 Exp

Embryo #9 Affected

171 153 205(N)

167 128 232(N)

Embryo #11 Normal ET

171 153 205(N)

159 151 Exp

Embryo #15 Affected

Fig. 4.45  PGT for myotonic dystrophy combined with chromosome 19 aneuploidy testing. (Top) Maternal haplotype based on PB1 and PB2 multiplex DNA amplification of the normal allele of DMPK gene (19q13.2–13.3) and linked markers (right). Paternal haplotype based on blastomere analysis (open and darker). As the expansion of (CTG) repeat in DMPK gene is not detectable at singlecell level (red bar in maternal haplotype corresponds to the affected allele), linked polymorphic markers are used for PGT; grey bar represents the normal allele and tightly linked markers. (Bottom) Six embryos tested for the presence of the normal number of (CTG) repeat in DMPK gene using polymorphic marker pattern. Embryos #1, #8, and #11 were predicted normal based on the presence of

the normal maternal chromosome (haplotype) (grey bar) and one paternal chromosome 19. Embryos #2, #9, and #15 were affected evidenced by the presence of the maternal affected haplotype (red bar). Embryo #2 contains two sets of maternal and one set of paternal polymorphic markers, suggesting trisomy 19 of maternal origin. Therefore, amplification of only (CTG) repeat in DMPK gene would have revealed the normal status based on the presence of only normal maternal and paternal alleles, which would have led to misdiagnosis. The follow-up FISH analysis confirmed trisomy 19, as predicted, and also incidental trisomy 13. DMPK dystrophic myotonia protein kinase gene, ET embryo transfer

In Fig.  4.45, an affected mother with DM (DMPK) had an expanded allele linked to 159 and 151 repeat markers. Of six embryos tested using three closely linked markers, three were affected (embryos #2, #9, and #15), as evidenced by presence of mutant maternal allele and one set of paternally derived markers. Embryo #2 was trisomic for chromosome 19, in which the gene for DMPK is located (19q13.2–q13.30). The remaining three embryos (embryos #1, #8, and #11) were normal; all three markers were in agreement with presence of the maternal normal allele together with presence of one of the paternal alleles. Two of these embryos (embryos #1 and #11) were transferred, resulting in an unaffected clinical pregnancy. PGT for another dynamic mutation, Machado– Joseph disease, is presented in Fig.  4.46. Machado–Joseph disease is a neurodegenerative

disorder characterized by perturbation of the gene cerebellar ataxia 3 (SCA3). Phenotype consists of pyramidal and extrapyramidal signs, peripheral nerve palsy, external ophthalmoplegia, facial and lingual fasciculation, and bulging. The mother in this family was a carrier of the expanded allele, inherited from her mother. Closely linked were 166, 137, 109, and 126  bp repeat markers for D14S617, D14S1015, D14S1016, and D14S1050, respectively. PGT was performed by sequential PB1 and PB2 analysis in seven oocytes, five of which appeared to be affected with only two without expansion, which were transferred back to patients, yielding an unaffected twin pregnancy and the birth of healthy children, as confirmed by amniotic fluid analysis. Endocrinal Disorders  The majority of these disorders are of complex nature, such as diabetes;

170

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Fig. 4.46  PGT for Machado–Joseph disease (SCA3) resulting in birth of healthy twins. (a) Family pedigree with haplotype analysis, showing that the mother (2.2) inherited the expanded allele (red bar) from her mother (1.2). PGT resulted in the birth of two unaffected twins (3.1 and 3.2), following amniocentesis. (b) Seven oocytes

were tested for the presence of the expanded allele by sequential PB1 and PB2 analysis, showing that only two oocytes (oocyte #1 and #4) were free of the expansion. Both embryos resulting from these mutation-free oocytes were transferred yielding the birth of unaffected children

however with identification of the major gene associated with such complex conditions, PGT may be considered as an option. In our experience 37 couples requested PGT for 10 different endocrinal disorders, listed in Table 4.23. As can be seen from the Table 4.23, more than a half of the cycles were performed for adrenal hyperplasia, which resulted in the birth of 24 unaffected children. However, of special interest are 11 cycles which were done for hyperinsulinemia (HHF1), determined by mutation in ABBC8 gene. PGT for this condition is demonstrated in Fig. 4.47, which resulted in birth of an unaffected child in each cycle performed for this couple. Overall, 86 unaffected embryos were selected for transfer in 51 PGT cycles for endocrinal disor-

ders, resulting in the birth of 30 healthy children, free from mutant genes responsible for the above conditions. The groups of conditions for which increasing number of PGT is being performed are also a genetically determined deafness and blindness, with the majority of cycles for deafness done for neurosensory conditions, caused by autosomal dominant mutation in GJB2 gene (Table 4.24). It is of interest that up to 80% of these cases were presented through expanded carrier screening (Fig.  4.48), which is introduced in the areas of high prevalence of inherited forms of deafness, such as in China. Blindness is another group of considerable number of inherited conditions, also requiring PGT in the majority of them

Disease Adrenal hyperplasia, congenital, due to 21-hydroxylase deficiency Allan–Herndon–Dudley syndrome; AHDS Androgen insensitivity syndrome; AIS Blepharophimosis, ptosis, epicanthus inversus; with ovarian failure Diabetes insipidus, nephrogenic, X-linked Diabetes mellitus, permanent neonatal; PNDM Hyperinsulinemic hypoglycemia, familial, 1; HHF1 Hypogonadotropic hypogonadism 1 with or without anosmia; HH1 Hypoparathyroidism–retardation–dysmorphism syndrome; HRDS Pseudovaginal perineoscrotal hypospadias; PPSH Total

Table 4.23  PGT-M for endocrine disorders

2 5 7

1 3 3 1 1 2 1 1 2 38

SLC16A2 300523 AR 313700 FOXL2 110100 304800 606176 256450 308700 241410 264600

AVPR2 INS ABCC8 KAL1 TBCE SRD5A2 (10)

2 67

1

1

3 1 11

# of cycles 34

# of Gene OMIM patients CYP21A2 201910 23

2 54

1

2

3 1 8

2 4 5

# of transfers 26

4 92 1.67

2

2

6 1 19

3 6 7

# of embryos transferred 42

1 28 52.94%

0

0

1 1 3

1 2 3

0

0

1 1 2

1 1 3

1 2 26 7.4% 92.6%

0

0

0 0 1

0 1 0

# of Pregnancy SAB deliveries 16 0 16

1 31

0

0

1 1 2

1 2 3

# of babies 20

4.6 Other Genetic Conditions with Important Health-­Related Implications 171

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

172

Family pedigree Markers order: D11S4099 D11S4060 ABCC8 D11S902 D11S4138 D11S1310 D11S4096

N/F1388 del

N/c.3989-9G>A

1.1

2.1

Affected

1.2

2.2

Normal

PGT 1

2.3

Normal

PGT 2

Cycle 1 – 2016 (Sequential testing) Embryo #

PGT-M ABCC8 gene

1

Affected NA

NGS

2

Carrier 46, XX

3

4

6

8

Affected

Normal

Affected

Carrier

NA

46, XY

NA

44, XX, -16, -18

Cycle 2 – 2019 (Sequential testing) Embryo # 1

9

Carrier Complex

PGT-M ABCC8 gene

Normal

NGS

46, XY

2

4

Carrier

Carrier

46,XX,del(6)(q16.1 ,q27)

46, XY

Embryo # 9

44, XX, -16, -18

Normal 46, XY

6

8

10

11

Carrier

Carrier

Carrier

Affected

46, XX

46, XY

NA

46, XX

FET 2

FET 1

Embryo # 4

5

Embryo # 2

46,XX,del(6)(q16.1, q27)

Embryo # 2

46, XY

Fig. 4.47  PGT for Familial Hyperinsulinism (c.3989-9 G>A and deletion F1388 in ABCCB gene), combined with PGT-A by NGS. Couple had an affected boy with deletion F1388 in ABCCB gene. PGT was performed by blastocyst biopsy with results in the first cycle for 7 embryos, 5 of which were affected, one carrier and one

normal (embryos #4). This embryo was transferred and resulted in birth of mutation free boy. In the next cycle, 8 embryos reach the blastocyct, of which 1 was affected, 5 carriers and 2 normal. One of these two embryos were transferred and resulted in birth of an unaffected child

(Table  4.25). Both of these groups are determined by gene mutations with different modes of inheritance, with the most severe ones being autosomal recessive, representing an obvious indication for PGT-M. Figure 4.49 demonstrates the case PGT for 2 monozygotic twins with blindness determined by de novo 32 base pair deletion in PITX2 gene causing Axenfeld–Rieger syndrome. The first PGT cycle was performed by sequential polar body analysis to establish maternal haplotypes. Out of five oocytes, three were predicted to be normal and two oocytes were affected. One of normal embryos was transferred on Day 5, and pregnancy resulted in spontaneous abortion. The other normal embryo reaching blastocyst was frozen, and a healthy baby boy was delivered after the frozen transfer of this embryo.

The second PGT cycle was performed by blastocyst biopsy and analysis for 32  bp deletion, together with PGT-A by NGS. One of four embryos tested was affected, another was unaffected but aneuploid, and two were available for transfer. The first frozen transfer resulted in SAB, while the second resulted in the birth of a healthy baby girl. In a PGT for the twin sister, three PGT cycles were performed for the PITX2 gene mutation also combined with PGT-A.  No transferrable embryos were available in the first two cycles, while the third one resulted in an unaffected baby boy delivery. Finally, inherited forms of renal disorders are becoming important indications for PGT, although they are usually components of a different syndrome, as can be illustrated by PGT for X-linked Alport disease (Fig. 4.50).

Waardenburg syndrome, type 2A; WS2A Wolfram syndrome 1; WFS1 Total

Disease Auriculocondylar syndrome 2; ARCND2 Deafness, autosomal dominant 3b; DFNA3B Deafness, neurosensory, autosomal recessive 1; DFNB1 Usher syndrome, type I; USH1 Usher syndrome, type IF; USH1F Usher syndrome, type IIA; USH2A Usher syndrome, type IIC; USH2C

Table 4.24  PGT-M for deafness

276900 602083 276901 605472

MYO7A PCDH15 USH2A ADGRV1 GPR98 MITF WFS1 (10) 193510 222300

220290 52

GJB2

2 1 63

1 1 3 1

# of OMIM Patients 614669 1 612643 1

Gene PLCB4 GJB6

6 2 103

3 4 6 2

75

# of cycles 3 2

5 1 84

1 4 4 2

62

# of transfers 3 2

5 1 109 1.3

1 6 5 3

80

# of embryos transferred 5 3

3 1 45 53.5%

0 4 2 1

31

0 2 1 1

29

# of deliveries 1 1

0 3 0 1 6 39 13.3%

0 2 1 0

2

Pregnancy SAB 2 1 1 0

3 1 42

0 2 1 1

32

# of babies 1 1

4.6 Other Genetic Conditions with Important Health-­Related Implications 173

174

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

Fig. 4.48 Increased Number PGT Cycles for non-syndromic hearing loss, determined by GJB2 gene through expanded carrier screening

35

32

30

27

25

22

20

23 20

21

22 19

16 15 10 5

5 0

2005–2015

2016

Total requests for GJB2PGT-M

4.6.4 Variants of Unknown Significance (VUS) With recent technological progress, DNA sequencing is becoming robust and replicable. Ideally, sequencing results are unambiguous, either normal or abnormal for a given gene. However, variants unexpectedly are present, which is not always clear. The term “variants of uncertain significance” (VUS) is then applied. For example, a single nucleotide change may alter transcription of one amino acid to that of another; the change may or may not be deletions. It could just be a newly discovered synonymous missense mutation (SMM) present in other phenotypically normal individuals. By contrast, it is tempting if the variant is found in a phenotypically abnormal individual to conclude that such a rare variant is causative of a given disorder. This is especially true if it inadvertently detected during carrier screening or in the context of another family member being tested. Determining the clinical consequence of a VUS is important. Deletion or duplication of a 1,000,000 or more base pair sequence (1 mb) would be predicted to be pathogenic, while a frameshift generates an incorrect message. To determine clinical significance of a VUS, data on the significance of a given DNA variant in the general population is sought. Unfortunately, little data may exist.

2017

2018

2019

Referred after carrier screening

A VUS is not correlated with parental age, but may be familial. For this reason parental studies may be indicated if a VUS is detected in a fetus or a child under study. The likelihood of transmission of a VUS to offspring or embryo is 50%. Phenotype may not, however, necessarily be predictable even if transmitted. If other family members are heterozygous for the same VUS, this may be helpful in deducing the phenotype. Phenotype prediction may also be helped by computer-based protein prediction models, which judge if the projected 3D altered gene product (protein) is likely or is not likely to be damaging. The long-term strategy is that additional data will allow a VUS to be reclassified. DNA variants are stratified into five categories: benign, probably benign, VUS, probably pathogenic, and pathogenic. About 80% of VUS become reclassified as probably benign or benign. Yet there is no universally recommended management policy after recognition of a VUS. A typical scenario might involve a couple in which one partner has a pathogenic mutation for an autosomal recessive disorder, whereas the other partner has a VUS.  A decision may be needed on whether to undergo PGT for the disorder potentially at risk. We exercise a relatively liberal policy for justification for PGT and would not exclude the option of PGT or an invasive prenatal genetic diagnostic procedure if

Disease Achromatopsia 2; ACHM2 Achromatopsia 3; ACHM3 Albinism, ocular, type I; OA1 Albinism, oculocutaneous, type II; OCA2 Aniridia Axenfeld–Rieger syndrome, type 1 Cone-rod dystrophy 6; CORD6 Glaucoma 3, primary congenital, A; GLC3A Knobloch syndrome 1; KNO1 Leber congenital amaurosis 10 Leber congenital amaurosis 2 Leber congenital amaurosis 5; LCA5 Macular dystrophy, vitelliform, 2; VMD2 Microphthalmia, isolated, with coloboma 3 Norrie disease; ND Optic atrophy 1; OPA1 Retinal dystrophy, early-onset Retinitis pigmentosa 2 Retinitis pigmentosa 3 Retinitis pigmentosa 4 Retinoschisis 1, X-linked, juvenile Stargardt disease 1; STGD1 Stickler syndrome, type I; STL1 Stickler syndrome, type II; STL2 Total 4 3 1 1 1 1 1 1 1 2 5 3 1 1 5 3 1 4 4 2 52

267750 611134 204100 604537 153700 610092 310600 165500 610125 312600 300029 613731 312700 248200 108300 604841

COL18A1 CEP290 RPE65 LCA5 BEST1

VSX2

NDP OPA1 OTX2 RP2 RPGR RHO RS1 ABCA4 COL2A1 COL11A1

# of OMIM patients 216900 1 262300 3 300500 1 203200 2

PAX6 106210 PITX2 180500 GUCY2D 604537 CYP1B1 231300

Gene CNGA3 CNGB3 GPR143 OCA2

Table 4.25  PGT-M for ophtalmologic disorders

8 5 1 1 6 5 2 10 4 7 99

2

1 1 1 1 1

7 13 1 1

# of cycles 1 4 12 4

6 5 0 1 5 1 1 5 2 6 71

2

1 2 0 1 1

5 12 1 1

# of transfers 1 4 5 3

13 9 0 1 7 1 2 6 4 13 105 1.48

2

0 2 0 1 1

6 14 1 1

# of embryos transferred 1 5 9 6

2 0 0 1 3 1 1 2 1 1 35 49.3%

1

0 1 0 1 1

4 5 0 1

Pregnancy 1 2 4 2

0 0 0 0 0 1 1 0 1 0 6 17.1%

0

0 0 0

0

0 1 0 1

SAB 0 0 1 0

2 0 0 1 3 0 0 2 0 1 29 82.9%

1

0 1 0 1 1

4 4 0 0

# of deliveries 1 2 3 2

3 0 0 1 3 0 0 2 1 1 32

1

0 1 0 1 1

4 4 0 1

# of babies 1 2 3 2

4.6 Other Genetic Conditions with Important Health-­Related Implications 175

4  Strategies and Indications for Preimplantation Genetic Testing for Monogenic Disorders (PGT-M)

176

PGT-M FOR DE NOVO MATERNAL 32 base pair DELETION IN PITX6 GENE (AXENFELD –RIEGER SYNDROME)

Family pedigree 32 bp Deletion De novo

DOB 1984

FET3 FET2

PGT1 - 2013 1

2

3

182 170 DEL 132 146 108

182 170 DEL 132 146 108

M

M

154 165 N 139 150 121

4

5

154 165 N 139 150 121

154 165 N 139 150 121

N

N

N FET 1 SAB

FET4 SAB

FET 2 PGT1

2

4

6

M 46,XX

M

N

46,XY

45, XX,-15

1

N 47,+16, XX

N 46,XY FET 3 SAB

5

N 46,XX FET 4 PGT2

N

10

M 46,XX

M 46, XX

2

46,XX 4

8

10

N

M

44, XX,-3, -16 46, XY

3

5

PGT2 2018

PGT2 - 2017 3

7

PGT1 2017

M

N

45, XY,-22

46, XX

N 47, XY, +21

FET2

FET1 3

5

N

M

6

7

9

PGT3 2019

45,XY, -20 46, XX

M 47, XY, +21

N

N

46,XY FET3

46,XX

Fig. 4.49  PGT for affected monozygotic twins with congenital blindness caused by de novo 32 base pairs deletion in PITX2 gene. The first PGT-M cycle was performed in 2013 by sequential polar body analysis to establish the maternal haplotypes. Of the five oocytes tested, three were predicted to be normal and two oocytes were affected. One normal embryo was transferred on Day 5 and pregnancy resulted in SAB.  The other embryo developed to blastocyst, and the frozen transfer of this embryo resulted in birth of unaffected baby boy. In 2017, the patient had another PGT-M cycle, performed by blastocyst biopsy

and analysis for 32  bp deletion in PITX2 gene together with NGS-based PGT-A. One of the four embryos tested appeared to be with 32 bp deletion, one aneuploidy, while two unaffected euploid embryos were transferred in two sequential frozen cycles, resulting in an unaffected birth of healthy baby girl in the second cycle, as the first one resulted in SAB. Three PGT-M cycles were performed for the other twin, also combined with PGT-A. Of the three PGT cycles performed, only the third one resulted in a healthy baby boy delivery

there is plausible clinical significance. That said, genetic counseling should not convey the same level of certainty communicated if both partners were carriers for a pathogenic mutation, e.g., ΔF508 cystic fibrosis. Offspring are still at risk if only one partner had a pathogenic variant and the other a VUS, but not to the same extent as if both partners had known pathogenic variants. In other circumstances, reasoning may differ. For example if a severe inborn error of metabolism existed in an offspring homozygous for a VUS transmitted in a consanguineous union, the VUS could logically seem as deleteri-

ous and, hence, an explanation for the disorder. Yet, even this VUS could still be present only coincidental. The molecular explanation could tie elsewhere (e.g., regulatory region). In the USA, the American College of Medical Genetics and Genomics recommends laboratories disclose VUS, a policy providing justification for PGT or an invasive procedure [105]. In Europe, laboratories usually do not disclose the presence of a VUS. In our experience, a total of 44 VUS cases presented, of which 36 were accepted for PGT-M and 8 declined, due to inadequate genotype/phenotype correlation in the context of

4.6 Other Genetic Conditions with Important Health-­Related Implications a FAMILY

177

b STRATEGY FOR MATERNAL DE NOVO EXON

PEDIGREE

DELETIONS

E1

E2

E14 15 16 17 18 19

E1

E2

E 14-19

E 20

E 52

NORMAL SEQUENCE Col4A5 133 172 117 126 106 N 197 160 148

PGT Normal

135 172 117 137 118 N 197 160 148

c

131 170 115 132 106 DEL 195 158 146

MUTANT SEQUENCE Col4A5

E 20

E 52

DELETED

PGT -M +PGT-A for DE NOVO EXON 14– 19 DELETION IN Col4A5 gene (ALPORT SYNDROME) 2

Markers order: DXS1191 DXS8048 DXS8097 DXS1120 DXS571 Col4A5 IVS (CA)n IVS30(CCCT)n DXS1210

13 5 17 2 11 7 13 7 11 8 N 19 7 16 0 FET 14 8

Y

NORMAL NGS

46, XY

3

Y

5

13 3 17 2 11 7 12 6 10 6 N 19 7 16 0 14 8

13 1 17 0 11 5 13 2 10 6 DE L 19 5 15 8 14 6

DEL

13 5 17 2 11 7 13 7 11 8 N 19 7 16 0 14 8

NORMAL

46, XY

47, XX, +16

6

13 3 17 2 11 7 12 6 10 6 N 19 7 16 0 14 8

13 5 17 2 11 7 13 7 11 8 N 19 7 16 0 14 8

NORMAL 46, XX

8

Y

13 1 17 0 11 5 13 2 10 6 DE L 19 5 15 8 14 6

DEL 46, XY

10

13 3 17 2 11 7 12 6 10 6 N 19 7 16 0 14 8

13 1 17 0 11 5 13 2 10 6 DE L 19 5 15 8 14 6

CARRIER 45, XX, -22

Fig. 4.50  Combined PGT-M and PGT-A testing for X-linked de novo mutation in Col4A5 gene causing Alport symdrome. (a) Family pedigree. Patient was clinically diagnosed with Alport syndrome. Genetic testing of family members for deletion/duplication in Col4A5 gene showed that patient was positive for 10.09  kb deletion involving exon 14–19 of Col4A5 gene. Both her parents were negative. Based on this results, de novo deletion was confirmed in patient. Sequence of linked markers and Col4A5 gene located in the left corner. (b) Because the exact breakpoint of the deletion was not known, primers were designed to amplify sequences in all deleted exons in order to distinguish normal male embryos and avoiding the presence of deleted exons that may consistently fail to

amplify. (c) Combined PGT-M and PGT-A testing results in six embryos biopsied on Day 5 and analyzed for the presence and absence of exons 14–19. Of three male embryos one demonstrated the presence of all tested exons (embryo #2) and two were affected (embryo #3 and #8), based on the complete absence of exon 14–19. These allowed for linkage prediction, and normal (colored in green) and affected (colored in red) haplotypes were established. Based on linkage analysis three female embryos were predicted to be either normal (embryos #5 and #6) or carriers at risk (embryo #10). Overall, two normal euploid embryos were recommended for transfer, and the transferred embryo #2 resulted in birth of a healthy baby boy

family history. For cases that were accepted for PGT-M, a letter of support and recommendation were required from the referring provider as well as documentation of prior genetic counseling regarding the molecular finding. Our data show the importance of clinical history and additional family member testing in assessing

feasibility of PGT-M. In cases of dominant inheritance, VUS may segregate in affected and unaffected family members, supporting the decision for PGT-M, while in the others, when VUS is identified in children and one of their parents, PGT-M is not required, because the patient and parents’ phenotype did not correlate

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adequately with the expected phenotype of the gene identified. However, in the majority of cases VUSs require additional interpretation and correlation with family and clinical history to determine if PGT-M is an appropriate option.

4.7

Contemporary Selection of Molecular Technologies

Presented in this chapter is our experience, which is the world’s largest PGT-M series comprising 6204 PGT-M cycles performed for 3758 at risk couples. The number of conditions for which PGT performed is expanding, with the present number as high as 581; (Table  4.1). As can be seen from Table 4.2, outcome of PGT-M is even more favorable than in a routine IVF, resulting in over 2447 pregnancies (52.8% pregnancy rate), despite transfer of only one or two embryos per cycle on the average. This doubtless reflects in part the already proven fertility in couples unexpectedly at risk for monogenic disorders. It is of note that unaffected embryos suitable for transfer were available in 4630 of 6204 cycles (74.6%). Only in 1574 cycles (25.4%) were either no unaffected embryos available or embryos predicted to be normal failing to develop for transfer. Overall, 2517 children were born following PGT-M, with high accuracy and reliability of PGT-M close to 99%. This provides strong evidence that PGT-M is a safe and well-­established clinical procedure with great impact on the improvement in ART. Its application not only allows primary prevention of genetic disorders but provides an option for couples at risk who wish to avoid the risk of producing offspring with expanding number of common late-onset conditions with genetic predisposition. PGT has also opened the possibility of improving access to HLA-matched embryos needed for stem cell transplantation treatment for congenital and acquired disorders. Given our 30-year experience in this field, this chapter has been illustrated with pioneering examples, often the first or unique PGT cases. These involved utilization of all the available PGT approaches, earlier using polar body or blastomere analysis, and more recently trophectoderm samples from blastocysts.

Similarly, PGT technologies have been improved, such as from FISH to 24-chromosome analysis by NGS.  While linkage analysis using polymorphic markers remains routine, introduction of vitrification allows obviating the need for testing fresh embryos, so the reader could always be able to relate with the reference labs to ensure performing PGT with the use of the contemporary methodologies.

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References 78. Murrel J, Hake AM, Quaid KA, Farlow MR, Ghetti B.  Early-onset Alzheimer disease caused by a new mutation (V717L) in the amiloid precursor proteen gene. Arch Neurol. 2000;57:885–7. 79. Tupler R, Rogaeva E, Vaula G, et al. A highly informative microsatellite repeat polymorphism in intron 1 of the human amyloid precursor protein (APP) gene. Hum Mol Genet. 1993;2:620–1. 80. He J, McDermont DA, Song Y, Gilbert F, Kligman I, Basson C.  Preimplantation genetic diagnosis of human congenital heart disease and Holt-Oram syndrome. Am J Med Genet. 2004;126A:93–8. 81. Papadopoulou LC, Sue CM, Davidson MM, et  al. Fatal infantile cardioencephalomyopathy with COX deficiency and mutations in SCO2, a COX assembly gene. Nat Genet. 1999;23:333–7. 82. Lee S, Zambas E, Wu X, Reid M, Zelinsky T, Redman C. Molecular basis of the Kell (K1) phenotype. Blood. 1995;85:912–6. 83. Lee S, Zambas E, Green ED, Redman C. Organization of the gene for encoding the human kell group protein. Blood. 1995;85:1364–70. 84. Purohit KR, Weber JL, Ward LJ, Keats JB. The kell blood group locus is close to the cyctic fibrosis locus on chromosome 7. Hum Genet. 1992;89:457–8. 85. Reid ME, Rios M, Powell VI, Charles-Pierre D, Malavade V. DNA from blood samples can be used to genotype patients who have recently received a transfusion. Transfusion. 2000;40:48–53. 86. Zielenski J, Rozmahel R, Bozon D, Kerem BS, Grzelczak Z, Riordan J, Rommens J, Tsui L-H.  Genomic DNA sequence of the cystic fibrosis transmembrane conductance regulator (CFTR) gene. Genomics. 1991;10:214–28. 87. Chehab FF, Johnson J, Louie E, Goossens M, Kawasaki E, Elrich H. A dimorphic 4-bp repeat in the cystic fibrosis gene is in absolute linkage disequilibrium with the delta F508 mutation: implications for prenatal diagnosis and mutation origin. Am J Hum Genet. 1991;48:223–6. 88. Van Den Veyver IB, Chong SS, Cota J, et al. Fetus-­ placenta-­newborn: single cell analysis of the RhD blood type for use in preimplantation diagnosis in the prevention of severe haemolitic disease of the newborn. Am J Obstet Gynecol. 1995;172:533–40. 89. Secho SKM, Burton G, Leigh D, Marshall JT, Pearsson JW, Morris JM. The role of PGD in the management of severe alloimmunization: first unaffected pregnancy. Case report. Hum Reprod. 2005;20:697–701. 90. Verlinsky Y, Rechitsky S, Seckin O, Masciangelo C, Ayers J, Kuliev A. Preimplantation diagnosis for Kell genotype. Fertil Steril. 2003;80:1047–51. 91. Nanni L, Ming JE, Bocian M, et  al. The mutational spectrum of the sonic hedgehoc gene in holoprosencephaly: SHH mutations cause a significant proportion of autosomal dominant holoprosencephaly. Hum Mol Genet. 1999;8:2479–88. 92. Muenke M, Gurrieri F, Bay C, et  al. Linkage of a human brain malformation, familial holoprosenceph-

181 aly, to chromosome 7 and evidence for genetic heterogeneity. Proc Natl Acad Sci U S A. 1994;91:8102–6. 93. Verlinsky Y, Rechitsky S, Verlinsky O, et  al. Preimplantation diagnosis for sonic hedgehog mutation causing familial holoprosencephaly. N Engl J Med. 2003;348:1449–54. 94. Abou-Sleiman PM, Apessos A, Harper JC, Serhal P, Delhanty JDA.  Pregnancy following preimplantation genetic diagnosis for Crouson syndrome. Mol Hum Reprod. 2002;8:304–9. 95. Verlinsky Y, Rechitsky S, Schoolcraft W, Kuliev A.  Preimplantation diagnosis for homeobox Gene HLXB9 mutation causing currarino syndrome. Am J Med Genet A. 2005;134A:103–4. 96. Ross AJ, Rui-Perez V, Wang Y, Hagan DM, Scherer SW, Lynch SA, Lindssy S, Custard E, Belloni E, Wilson DI, Wadey R, Goodman F, Orstavic KH, Monclair T, Robson S, Reardon W, Burn J, Scambler P, Strachan T.  A homeobox gene, HLXB9, is the major locus for dominantly inherited sacral agenesis. Nat Genet. 1998;20:358–61. 97. Belloni E, Martucciello G, Verderio D, Ponti E, Seri M, Jasonni V, Torre M, Ferrari M, Tsui L-C, Scherer SW, et  al. Involvement of the HLXB9 homeobox gene in Currarino syndrome. Am J Hum Genet. 2000;66:312–9. 98. Hagan DM, Ross AJ, Strachan T, Lynch SA, Ruiz-­ Perez V, Wang YM, Scambler P, et  al. Mutation analysis and embryonic expression of the HLXB9 Currarino syndrome gene. Am J Hum Genet. 2000;66:1504–15. 99. Kochling J, Karbasiyan M, Reis A.  Spectrum of mutations and genotype-phenotype analysis in Currarino syndrome. Eur J Hum Genet. 2001;9: 599–605. 100. Reardon W, Winter RM, Rutland P, et al. Mutations in the fibroblast growth factor gene cause Crouzon syndrome. Nat Genet. 1994;8:98–103. 101. Cohen MM, Kreiborg S.  Birth prevalence studies of the Crouzon syndrome: comparison of direct and indirect methods. Clin Genet. 1992;41:12–5. 102. Sermon K, Lissens W, Joris H, et al. Clinical application of preimplantation diagnosis for myotonic dystrophy. Prenat Diagn. 1997;17:925–32. 103. Sermon K, Seneca S, Vanderfaeillie A, et  al. Preimplantation diagnosis for fragile X syndrome based on the detection of the non-expanded paternal and maternal CGG. Prenat Diagn. 1999;19:1223–30. 104. Verlinsky Y, Rechitsky S, Verlinsky O, Strom C, Kuliev A. Polar body based preimplantation diagnosis for X-linked genetic disorders. Reprod Biomed Online. 2002;4:38–42. 105. Morales A, Hershberger RE.  Variants of uncertain significance: should we revisit how they are evaluated and disclosed? Circ Genom Precis Med. 2018;11:e002169. https://doi.org/10.1161/ CIRCGEN.118.002169.

5

Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)

Preimplantation HLA matching (PGT-HLA) per se has never been a standard indication for prenatal diagnosis practice because the decision on clinical pregnancy termination simply on the basis of a given HLA genotype cannot alone be justified. However, PGT-HLA is acceptable with the current single embryo transfer (SET) policy in the ART practices as only a single embryo has to be preselected anyway, which in this case will represent unaffected embryo with a perfect match for affected siblings in need of a HLA-compatible transplant. The world’s first case of preimplantation HLA typing was introduced in combination with PGT for Fanconi anemia (FA) with the objective establishing an unaffected pregnancy yielding a potential donor progeny who could provide bone marrow for stem cell transplantation into an affected sibling [1, 2]. This case opened a new chapter in reproductive medicine, demonstrating feasibility of PGT- HLA as part of PGT.  Prospect was opened for application to other inherited conditions, which require an HLA-compatible donor for bone marrow transplantation. A realistic option was provided for couples not only desiring to avoid the birth of an affected child but providing an HLA match for their older affected sibling. It was shown that PGT-HLA in single or small number of cells biopsied from preimplantation embryo is accurate. It was also shown applicable as primary indication for cases not requiring mutation testing (e.g., childhood leukemia) but awaiting an © Springer Nature Switzerland AG 2020 A. Kuliev et al., Practical Preimplantation Genetic Testing, https://doi.org/10.1007/978-3-030-43157-0_5

HLA-compatible donor. This new indication made PGT a genuine and unique alternative to conventional prenatal diagnosis, providing patients with important prospects not only to avoid an inherited risk without facing termination of pregnancy but also to establish a pregnancy with particular genetic parameters to benefit the affected member of the family. Our experience of PGT-HLA is presented in Table 5.1, summarizing the results of 485 PGT-­ HLA cycles performed for 239 patients. A total of 424 HLA-matched embryos were identified for transfer (1.46 HLA-matched embryos per transfer on the average) in 291 of 485 (68.6%) cycles, resulting in 125 (43.0%) clinical pregnancies and birth of 117 healthy HLA-matched children, representing a stem cell donor for their affected siblings. Among conditions requiring HLA-compatible stem cell transplantation, hemoglobinopathies are one of the most prevalent, representing the commonest autosomal recessive disease in the Mediterranean region, Middle East, and Southeast Asia, with the heterozygous frequencies as high as 14% in Greece and Cyprus.

5.1

Thalassemia

Beta-thalassemia affects the production of beta-­ globin chains resulting in severe anemia that makes patients transfusion-dependent starting 183

Disease HLA genotyping HLA + adenosine deaminase deficiency; ADA HLA + adrenoleukodystrophy; ALD HLA + cardiomyopathy, familial hypertrophic, 4; CMH4 HLA + granulomatous disease, chronic, autosomal recessive; CDG1 HLA + Diamond–Blackfan anemia 1; DBA1 Diamond–Blackfan Anemia 2; DBA2 Diamond–Blackfan anemia 3; DBA3 Diamond–Blackfan anemia 5; DBA5 Diamond–Blackfan anemia 9; DBA9 HLA + glanzmann thrombasthenia; GT Muscular dystrophy, Duchenne type; DMD HLA + myotonic dystrophy 1; DM1 HLA + ectodermal dysplasia and immunodeficiency 1; EDAID1 HLA + epidermolysis bullosa dystrophica, autosomal dominant; DDEB HLA + Fanconi anemia, complementation group A; FANCA HLA + Fanconi anemia, complementation group C; FANCC HLA + Fanconi anemia, complementation group D2; FANCD2 HLA + Fanconi anemia, complementation group F; FANCF HLA + Fanconi anemia, complementation group G; FANCG HLA + Fanconi anemia, complementation group I; FANCI HLA + Fanconi anemia, complementation group J; FANCJ HLA + granulomatous disease, chronic, X-linked; CDGX HLA + HBB sickle cell anemia; beta-thalassemia 17

2 2 10 1 56 6 3 3 2 2 4 16 188

10

1 1 3 1 18 3 1 1 2 1 2 11 92

273,800 310,200 160,900 300291 131,750 227,650 227,645 227,646 603,467 614,082 609,053 609,054 306,400 603,903 613,985

ITGA2B, DMD DMPK IKBKG COL7A1 FANCA FANCC FANCD2 FANCF FANCG FANCI BRIP1 CYBB HBB

RPS19, RPS20, RPS24, RPL35A, RPS10

105,650 606,129 610,629 612,528 613,308

608,958 300,100 115,197 233,700

ADA ABCD1 MYBPC3 NCF1

# Cycle 119 1 7 1 3

Omim

Gene

# Patient 60 1 3 1 1

Table 5.1  Preimplantation HLA Testing (PGT-HLA) with and without PGT-M

2 4 12 103

2 1

2

6

29

1

1 8

2

14

# Transfers 73 1 2 1 2

3 3 15 159

3 2

3

9

42

1

2 10

4

20

# Embryo transferred 108 1 2 1 2

0 1 7 35

0 1

1

2

14

1

1 3

1

8

Pregnancy 25 1 1 1 1

0 0 1 3

0 0

0

0

1

0

0 1

1

0

SAB 3 0 0 0 0

0 1 6 32

0 1

1

2

13

1

1 2

0

8

Delivery 22 1 1 1 1

0 1 6 32

0 2

1

2

13

1

1 4

0

8

Birth 22 1 2 1 1

184 5  Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)

HLA + Immunodeficiency with hyper-IgM, type 1; HIGM1 HLA + Krabbe disease HLA + myelodysplastic syndrome; MDS HLA + neutropenia, severe congenital, 1, autosomal dominant; SCN1 HLA + Shwachman–Diamond syndrome; SDS HLA + thrombotic thrombocytopenic purpura, congenital; TTP HLA + thrombocythemia 1; THCYT1 HLA + Wiskott–Aldrich syndrome; WAS HLA + polycystic kidney disease 1; PKD1 HLA+ pyruvate kinase deficiency of red cells HLA + hyper-Ige recurrent infection syndrome, autosomal recessive Total

16 1 2 5 9 2 2 1 1 2 1 485

11 1 1 3 4 1 1 1 1 1 1 239

308,230 245,200 614,286 202,700 260,400 274,150 187,950 301,000 173,900 266,200 243,700

CD40LG GALC GATA2 ELANE SBDS ADAMTS13 SH2B3 WAS PKD1 PKLR DOCK8 291

2 0 1 1 0

3 2

10 1 1 4

424

2 0 2 1 0

3 4

15 2 1 4

125 43%

2 0 1 0 0

2 1

9 1 1 4

1 0 1 0 0 112 89.6%

13 10.4%

2 1

8 1 1 3

1 0 0 0 0

0 0

1 0 0 1

117

1 0 1 0 0

2 1

8 2 1 3

5.1 Thalassemia 185

186

5  Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)

from 6  months after birth. Bone marrow transplantation is the only option for radical treatment. We performed PGT for 54 of over 400 different mutations described in the beta-globin gene, located on chromosome 11 (11p15.5), causing congenital anemia of variable severity [3, 4]. Prenatal diagnosis has been applied widely for over four decades resulting in reduction of new cases of thalassemia up to 70% in many populations, including large countries in the Eastern Mediterranean region such as Turkey and Iran [5, 6]. Considerable progress has been achieved also in the treatment of the disease by bone marrow transplantation [7], the application of which is still limited to the availability of HLA-matched stem cells. PGT is clearly an attractive option for couples with thalassemia children, which has already been provided for over two decades [8]; PGT-HLA was belatedly but now concomitantly offered in the same framework. Thalassemia has been one of the major indications for PGT, from the very beginning of introduction of PGT-M.  It was first performed for couples who had previously undertaken prenatal diagnosis but had to terminate the pregnancy with an affected fetus on repeated attempts [8]. Then, it was offered as a primary option to the patients with infertility problems and to those who could not accept the risk for prenatal diagnosis and termination of pregnancy [8–10]. This was then followed by PGT-HLA for the couples with existing thalassemia children requiring HLA-compatible bone marrow transplantation [11–16]. Again, the objective of PGT in these cases was not only to have a thalassemia-free child but also to ensure that the resulting baby could serve as an HLA-compatible donor for bone marrow transplantation for the affected siblings. Reports of the outcomes of hundreds PGT-­ HLA cycles demonstrated the practical utility of PGT-M together with testing for the maternal and paternal chromosome 6 identity to the sibling chromosome 6 HLA patterns [11–17]. HLA genes were tested simultaneously, using the short tandem repeats in the HLA region. A multiplex heminested PCR system used closely linked polymorphic short tandem repeat (STR)

markers located throughout the HLA region: D6S426, D6S291, Ring 3 CA, TAP1, G51152, D6S2447, LH1,DN, D6S273, 9  N-2, TNF a,b,c,d; 62, MIC A, MIB, D6S276, D6S439, D6S1624, D6S265, D6S510; D6S248, RF, MOG a,b,c,d, D6S 258, D6S306, D6S464, D6S299, D6S461 (described in detail elsewhere [11, 17] and shown in Fig. 5.1). The choice of alleles and markers was based on information provided concerning the presence of maternal and paternal matching or non-matching chromosome 6 alleles. For each family, heterozygous alleles and markers (haplotypes) were selected that were not to be shared by parents. That is, haplotype analysis for the father, mother, and affected child was performed for each family prior to PGT-HLA. This allowed a strategy for detecting and avoiding misdiagnosis due to preferential amplification and allele dropout (ADO), potential recombination within the HLA region (see below), and a possible aneuploidy or uniparental disomy of chromosome 6. All of these pitfalls could adversely affect diagnostic accuracy of HLA typing of the embryo. In these cases, 30 PCR cycles were performed. A denaturation step began at 95 °C for 20 s, annealing at 62–50 °C for 1 min and elongation at 72 °C for 30 s [11, 17]. Twenty minutes of incubation at 96  °C was performed before starting cycling, and after cycling, 10 min of elongation at 72 °C was performed. Annealing temperature for the second round was programmed at 55 °C. Figure 5.2 presents PGT-HLA for carrier parents with different mutations, IVSI-5 and Cd8. Using PB1 and PB2 analysis, six mutation-free oocytes were identified prior to testing of embryo, used mainly for selection of paternal mutation-free embryos (embryos #1, #3, #5, and #9). HLA typing was also done in PB1 and PB2 to identify those oocytes with maternal HLA match. Embryos #2 and #6 appeared to be also HLAmatched to the affected sibling and were transferred, resulting in the birth of a healthy child who was confirmed to be thalassemia-free as well as HLA-matched to the affected sibling with thalassemia. The chance to identify unaf-

5.1 Thalassemia

187

Fig. 5.1 Polymorphic markers in HLA region applied for preimplantation HLA typing

Telomere D6S461 D6S276* D6S299 D6S464* D6S105

HLA-F

D6S306* D6S1624*

HLA-A

D6S1615 D6S258

HLA-C

Class I

D6S248*

HLA-E

MOG a,b,c,d

RF D6S265 D6S510

HLA-B

MIB MICA

Class III

TNF a,b,c,d

6p21.3

62 82-1 9 N2 D6S273* D6S1666 D6S1629 DN LH 1

HLA-DR

DQ-CAR II DQ-CAR

Class II

HLA-DQ

G51152

D6S2447 D6S2443 D6S2444 TAP 1 Ring 3CA D6S1568 D6S1560 D6S1618 D6S439 D6S291 D6S1583 D6S1610

Centromere

fected embryos fully matched to siblings with thalassemia is 18.75% (see below). The same applies to other autosomal recessive conditions. HLA-matched and thalassemia-free embryos were transferred back to the patient, based on the information about the mutation testing and polymorphic markers, whereas those predicted to be mutant or with insufficient marker information were subjected to confirmatory analysis. Non-­ matched unaffected embryos were frozen for future use by the couple. Of 54 different alpha- and beta-globin gene mutations tested (see the list of mutations in Chap. 4), 978 unaffected embryos (1.79 embryos per transfer on the average) were transferred in 546 (79.9) of 684 cycles performed, yielding 244

(44.6%) unaffected pregnancies and birth of 240 healthy children free of thalassemia (see Table 4.3 in Chap. 4). This is one of the three major indications in our overall PGT-M experience. A total of 188 of these cycles also involved PGT-HLA, allowing detecting and transferring unaffected HLA-matched embryos in 103 (54.8%) of them. A total of 159 (1.54 on the average) embryos predicted to be either unaffected carriers or normal and HLA-identical to the affected siblings, which is not significantly different from the expectation [15]. This resulted in 32 unaffected HLA-identical pregnancies and the birth of 32 healthy children. Umbilical cord blood was collected at birth from these children and transplanted or pend-

5  Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)

188

a

HLA Markers: D6S276 D6S258 D6S248 Mog A RF D6S1629 Ring 3CA D6S1560 D6S291

Family pedigree

144 135 283 188 270 190 155 148 116

117 133 250 182 258 188 159 166 114

HLA

b

144 135 283 188 270 190 155 148 116

PGT Embryo #

HBB

HLA

1

2

144 135 260 168 130 188 159 156 116

143 139 Cd8 N 114 120 6 7 120 140

1.1

HBB 144 135 260 168 130 188 159 156 116

3

CORD BLOOD TRANSPLANT

143 Cd8 114 6/7 120

139 IVSI-5 116 6 150

143 Cd8 114 6 120

143 N 114 6 120

139 143 N N 120 114 7 6 140 120

143 Cd8 114 6 120

144 135 260 168 130 188 159 156 116

144 135 283 188 270 190 155 148 116

144 135 260 168 130 188 159 156 116

117 133 250 182 258 188 159 166 114

144 137 260 168 280 186 161 163 124

144 135 283 188 270 190 155 148 116

NORMAL Non-match

CARRIER Full match

2.1

4

139 143 N N 120 114 7 6 140 120 117 133 250 182 258 188 159 166 114

NORMAL Non-match

1.2

2.2 PGT

5

139 IVSI-5 116 6 150 144 137 260 168 280 186 161 163 124

AFFECTED Non- match

ET

139 143 IVSI-5 N 116 114 6 6 150 120

HLA 144 135 283 188 270 190 155 148 116

Beta-Globin Markers: BSTR Mutation site BTG HPFH D11S1997

HBB

144 135 260 168 130 188 159 156 116

143 143 Cd8 N 114 114 6 6 120 120

6

139 139 N IVSI-5 120 116 6 7 140 150 144 135 283 188 270 190 155 148 116

144 137 260 168 280 186 161 163 124

144 137 260 168 280 186 161 163 124

CARRIER Non- match

8

9

143 Cd8 114 6 120

143 N 114 6 120

143 143 Cd8 N 114 114 6 6 120 120

143 Cd8 114 6 120

143 N 114 6 120

144 135 283 188 270 190 155 148 116

144 135 260 168 130 188 159 156 116

117 133 250 182 258 188 159 166 114

144 135 260 168 130 188 159 156 116

117 133 250 182 258 188 159 166 114

144 135 260 168 130 188 159 156 116

CARRIER Full match

CARRIER Non- match

CARRIER Non- match

ET

Fig. 5.2  Preimplantation HLA matching combined with PGT for thalassemia. (a) Family pedigree with HLA haplotype analysis based on parental (1.1 and 1.2) and affected child’s (2.1) genomic DNA testing. HLA marker order is presented on the upper left for the father and upper right for the mother. Color  bars represent the matching paternal and maternal HLA haplotypes, and the  black - bars non-matching haplotypes. (b) Maternal mutation and linked polymorphic markers were first assessed by sequential multiplex polar body (PB) analysis. Two oocytes (#4 and #5) had affected alleles, while the remaining six (#1, #2, #3, #6, #8, and #9) were normal (data not shown). Based on blastomere results, one embryo was affected (#4), two were homozygous normal

(#1 and #3), and five were carriers of paternal (#2, #6, #8, and #9) or maternal (#5) mutations. As seen from HLA typing below (see bottom panel), embryos #2 and #6 are also fully HLA-matched to the sick sibling (2.1). (Bottom panel) HLA typing by short tandem repeats (STRs) along with mutation analysis was performed on blastomeres from eight embryos, two of which (#2 and #6) were predicted to be HLA-matched to that of the affected sibling (2.1), although carrying the paternal mutation (also see above). Prenatal testing confirmed these results, and a healthy baby girl with HLA type matching that of the sick sibling was born. Cord blood stem cells were collected during the delivery and frozen for the stem cell transplantation

ing, resulting in a hematopoietic reconstitution. Figure 5.3 demonstrates a unique case of PGTHLA for a couple with two thalassemia children, resulting in preselection and transfer of unaffected embryos matched to each of the affected children. HLA typing showed that one of the embryos was a match for one of the affected siblings, another embryo a match for the other affected sibling (embryos #4 and #9),

and three were not matched (embryos #6, #7, and #8), including one with a single chromosome 6 (embryo #6). Aneuploid oocyte with trisomy 22 in the resulting embryo was detected by PB1 and PB2 analysis and excluded from further analysis with respect to the causative gene or HLA type, and other embryos showed monosomy 11 (embryo #2), or monosomy 6 (embryo #6). Two unaffected embryos, also

5.1 Thalassemia

189

aneuploidy-free and HLA-matched to each of the affected children (embryos #4 and #9), were transferred, resulting in the birth of a healthy HLA-matched baby. Similar experience was reported from other large series: 626 PGT-HLA cycles for 312 couples were performed (122 HLA only and 504 with PGT-M), resulting in 128 thalassemia-free children [18, 19]. Stem cells of 66 of these children were used for cord blood or bone marrow transplantation, which resulted in bone marrow reconstitution in all HLA Markers: D6S1583 RING D6S2447 LH1 TNF A MIB RF MOG D6S1624

a

156 161 151 169 100 121 301 159 178

HLA

154 155 160 167 94 127 252 167 180

139 180 7 IVS2-1 130 170 165

139 180 6 N 128 168 117

HBB

but two of them (transplantation treatment of the remaining 57 siblings pending). PGT may also have increasing impact on decisions made by well-treated B-thalassemia patients. The life expectancy of the patients with hemoglobin disorders has been dramatically improved with the increasing success rate of radical treatment by stem cell transplantation ­ [20]. However, the further impact of this treatment will depend on the availability of HLAidentical donors.

1.1

151 159 151 177 100 123 264 167 172

158 155 147 169 104 100 246 182 178

1.2

144 197 6 N 128 168 117

HLA

Family pedigree

139 180 6 IVS1-5 128 162 159

HBB Markers: BSTR HBG HPFH Mutation BRSA D11S1338 D11S1241

HBB

CORD BLOOD TRANSPLANT

PGT

b

2.1

PGT Embryo 1

HBB

2

4

2.2

6

2.3

7

8

9

0

HLA

Aneuploidy Testing By FISH or PCR

13,13; 16,16 18,18 21,21; 22,22,22

11, 0 13,13; 16,16; 18,18; 21,21; 22,22,

Trisomy 22 Monosomy 11

0 13,13; 16,16; 18,18; 21,21; 22,22,

NORMAL MATCH TO 2.2

ET

Fig. 5.3  PGT-HLA in a couple with two thalassemic children requiring HLA-matched bone marrow transplantation. (a) Family pedigree with HLA haplotype analysis based on parental (1.1 and 1.2) and affected children’s (2.1; 2.2) genomic DNA testing. HLA marker order is presented on the upper left for the father and right for the mother. Paternal and maternal matching HLA haplotypes to the affected children (2.1; 2.2) are shown in different colors. Maternal and paternal mutations and the linked markers are also presented accordingly. (b) HLA typing by short tan-

0; 6 13,13; 16,16; 18,18; 21,21; 22,22,

13,13; 16,16; 18,18; 21,21; 22,22,

AFFECTED Monosomy 6

NORMAL NON-MATCH

13,13; 16,16; 18,18; 21,21; 22,22,

NORMAL NON-MATCH

13,13; 16,16; 18,18; 21,21; 22,22,

CARRIER MATCH to 2.1

ET

dem repeats (STRs) along with mutation analysis was performed on blastomeres from seven embryos, one of which (#4) was predicted to be a carrier and an HLA match to the affected sibling 2.2, and another (#9) also a carrier and match to the affected sibling 2.1. Three others (embryos #6–#8) were non-matched, while embryo #2 was matched but had inconclusive results of mutation testing due to lack of maternal chromosome 11. Both carrier matched embryos were transferred, with the birth of a thalassemia-free child matched to one of the affected siblings

190

5.2

5  Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)

Immunodeficiency

prior to genetic testing, allowing simultaneous PGT-A (see Chap. 3). One or two Severe congenital immunodeficiency (SCID) is a immunodeficiency-­ free and aneuploidy-free large group of conditions, all of which require embryos were preselected for transfer; other PGT-HLA and stem cell transplantation. Without unaffected embryos were cryopreserved for poscompatible bone marrow transplantation, affected sible use by the patients. A total of 171 unaffected neonates cannot survive. Analogous to manage- and HLA-matched embryos were identified for ment of beta-thalassemia, HLA-matched stem transfer in 114 (85.4%) of 135 PGT cycles percell transplantation improves or completely formed for 74 couples carrying the abovemenreplenishes the immune system; thus, PGT-HLA tioned immunodeficiency mutations, resulting in is an obvious management option for inherited 54 unaffected (47.3% per embryo transfer) pregforms of SCID that will ensure the birth of unaf- nancies, which yielded the birth of 54 healthy fected HLA-matched children and provide poten- children free of immunodeficiency. The mean tial stem cell donor progeny for the affected number of embryos transferred was 1.5. siblings. A total of 135 PGT cycles were perExperience presented above demonstrates formed for 74 couples at risk for producing practical application of PGT-HLA for provision immunodeficiency progeny (Table  5.2). These of HLA-compatible stem cell transplantation include 26 for incontinentia pigmenti (IP); 24 for treatment for affected siblings with immunodefihyper-IgM type 1 immunodeficiency (HIGM1); ciency. It may be expected that with introduction 14 for X-linked chronic granulomatous disease of a widespread preconception carrier screening (CGD); 10 for hypohidrotic ectodermal dysplasia programs couples at risk for producing offspring with immunodeficiency (HED-ID); 9 for with congenital immunodeficiency could be Wiskott–Aldrich syndrome (WAS); 9 for ataxia– identified prospectively, so to offer PGT-M to telangiectasia (AT); 7 for type 1 X-linked agam- avoid the birth of affected children, while those maglobulinemia; 6 for Omenn syndrome (OMS); with affected immunodeficiency children will 5 for immunodysregulation, polyendocrinopathy, also benefit from PGT-HLA as a means for and enteropathy X-linked (IPEX); 5 for autoso- obtaining HLA-matched donor progeny to mal recessive severe combined immunodefi- achieve the effective stem cell transplantation ciency; 5 for X-linked severe combined treatment. The examples of application of PGT-­ immunodeficiency (SCIDX1); 5 for chronic HLA for immunodeficiency listed in Table  5.2 granulomatous disease; 5 for severe congenital are presented below. neutropenia 1 (SCN1); and single cycle for each Omen syndrome (OMS) is a rare autosomal of the other five types of immunodeficiency listed recessive disease with a prevalence of 1 in over in Table 5.2 [21]. 50,000. This severe primary immunodeficiency PGT cycles were performed using a standard disease is characterized by generalized erithroIVF protocol coupled with micromanipulation dermia, protracted diarrhea, repeated infections, procedures for the polar body, or embryo biopsy, hepatomegaly, and leukocytosis with eosinoas described elsewhere [17]. The majority of philia and elevated immunoglobulin E. samples were obtained by blastocyst biopsy, In OMS there is absence of B cells and excess using laser-assisted technology. To avoid misdi- production of highly restricted T lymphocytes, agnosis due to preferential amplification, multi- caused by mutation in recombinase-activating plex nested PCR was performed, involving genes RAG1 and RAG2 that are located on chrosimultaneous detection of the mutations together mosome 11p. RAG1 and RAG2 encode lymphoid-­ with up to three or more highly polymorphic specific proteins responsible for the process of markers closely linked to the genes tested. With variable, diversity, and joining (V (D) J) segment introduction of next generation sequencing recombination required for generation of the T(NGS) technology (Illumina, USA), whole-­ and B-cell repertoire (MIM 603554). No cure genome amplification (WGA) was performed exists other than stem cell transplantation.

Neutropenia, severe congenital, 1, autosomal dominant; SCN1 (de novo) Immunodeficiency 21; IMD21 Hyper-Ige recurrent infection syndrome, autosomal recessive Hemophagocytic lymphohistiocytosis, familial, 3; FHL3 Platelet disorder, familial, with associated myeloid malignancy; FPDMM Total

Disease Ataxia–telangiectasia; AT Agammaglobulinemia, X-linked, type I Immunodeficiency with hyper-IgM, type 1; HIGM1 Granulomatous disease, chronic, X-linked; CGD Immunodysregulation, polyendocrinopathy, and enteropathy, X-linked; IPEX Ectodermal dysplasia, hypohidrotic, with immunodeficiency Incontinentia pigmenti; IP Omenn syndrome (OMS) Severe combined immunodeficiency, autosomal recessive Wiskott–Aldrich syndrome; WAS Lymphoproliferative syndrome, X-linked, 1; XLP1 Severe combined immunodeficiency, X-linked; SCIDX1 Granulomatous disease, chronic, autosomal recessive

1 1 1 1 135

1 1 1 1 74

GATA2 DOCK8 UNC13D RUNX1

114 (84.4%)

0 0 1 1

4

5

614,172 611,432 608,898 601,399

8 21 5 4 9 2 7 4

# Transfers 5 7 18 13 5

10 26 6 5 9 1 5 5

IKBKG 3 IKBKG 14 RAG1 2 RAG2 2 WAS 5 SH2D1A 1 IL2RG 3 NCF1; 2 NCF2 ELANE 3

300,291 308,300 603,554 603,554 301,000 308,240 300,400 233,700 233,710 202,700

# Cycle 9 7 24 14 5

# Gene Patient ATM 5 BTK 4 CD40LG 12 CYBB 10 FOXP3 4

OMIM 607,585 300,755 308,230 306,400 304,790

Table 5.2  Preimplantation Genetic Testing (PGT) for Congenital Immunodeficiency

171 (1.5)

0 0 2 1

4

10 33 12 7 17 2 10 4

# Embryo transferred 6 13 26 19 5

54 (47.3%)

0 0 1 1

4

3 7 1 2 6 1 3 1

Pregnancy 4 3 8 6 3

5 (9.2%)

0 0 0 0

1

1 0 0 0 0 0 0 0

SAB 1 0 1 1 0

54

0 0 1 1

3

4 8 2 2 6 1 3 1

Birth 3 4 7 5 3

5.2 Immunodeficiency 191

192

5  Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)

The phenotypic variability among patients may be the result of different mutations in RAG1 and RAG2 ranging from missense to splice mutations to deletions. Despite specific therapy for dermatitis and lymphadenitis using immunosuppression or replacement therapy with intravenous immunoglobulins, persistent viral, bacterial infections and chronic diarrhea resulting in inanition may be responsible for death. As mentioned, the only cure is HLA-identical bone marrow transplantation. One OMS case is presented in Fig. 5.4. A couple at risk for producing progeny with OMS had two previous children, including a younger daughter who had severe OMS resulting in death. The child was double heterozygous. The inherited paternal R396C mutation represented a sequence change from arginine to cysteine at codon 396, caused by a single C to T transition (CGG to TGG). The maternal mutation (c.256_57 del AA) was in 86 codon, the deletion of lysine causing a frameshift that resulted in a premature termination signal 32 codons downstream. The older heterozygous unaffected daughter inherited only the paternal A396C mutation in the RAG1 gene. PGT-M was performed using PB1 and PB2; HLA typing was by embryo biopsy, resulting in transfer of two unaffected and aneuploidy-­ free embryos that yielded birth of healthy twins. In this case the affected sibling had died by the time PGT-HLA was completed; thus, there was no opportunity for stem cell transplantation treatment. However, the couples with previous OMS children will definitely be potential candidates for performing PGT-HLA to provide also an identical HLA donor progeny for stem cell transplantation. Ataxia–Telangiectasia (AT) is a progressive, neurodegenerative childhood disease that affects the brain and other body systems (MIM 208900). A weakened immune system makes patients susceptible to recurrent respiratory infections. The disease presents between 1 and 4 years of age as a delayed development of motor skills, poor balance, and slurred speech. Telangiectasia appears in the corners of the eyes or on the surface of the

ears and cheeks. Patients with AT may develop cancer, often acute lymphocytic leukemia or lymphoma. Other features include mild diabetes mellitus, premature graying of the hair, difficulty swallowing, and delayed physical and sexual development. Currently symptomatic and supportive treatment includes high-dose vitamin regimens, physical therapy, and gamma globulin injections to supplement a weakened immune system. Still, prognosis is very poor, and patients usually die in their teens. More than 500 unique mutations in the ataxia–telangiectasia mutated (ATM) gene are associated with AT. All result in absence of serine–protein kinase coded by the ATM gene located on chromosome 11q22.3 (MIM 607585). Sequence analysis of ATM detects as many as 90% mutations; PGT-M in the 10% can be tested by linkage analysis, based on intragenic markers connoting a haplotype with the mutation, albeit of unclear nature. The PGT cycle shown in Fig. 5.5 was performed for a couple with one affected child who had died in early infancy. The mother was a carrier with two ATM sequence changes in cis: involving exon 38 (5419A > G, K1807E) and exon 48 (6784G > C, A2262P). Because it was not known which of these two mutations was responsible for AT, both were tested in PB1 and PB2 and in embryo. Maternal mutation K1807E was identified by Bsm AI digestion, creating fragments of 96 and 92 bp in the PCR product of the mutant gene. In contrast, maternal mutation A2262P was not cut by HaeIII restriction digestion but created two fragments of 35 and 99 bp in the normal gene. Because the paternal mutation in the ATM gene was not identified, it was traced using four closely linked markers (Fig. 5.5a). Thus, sequential PB1 and PB2 testing was performed to identify the mutation-free oocytes and then tested the resulting embryos for paternal mutation by linkage analysis. Simultaneously testing was done for aneuploidy (Fig.  5.5c). The transfer of two unaffected carrier embryos resulted in a singleton pregnancy and the birth of a healthy baby boy, confirmed to be an unaffected carrier for the maternal mutation.

5.2 Immunodeficiency

193

103 N 159 139

a Family pedigree

103 R396C 163 139

1.1

Markers order: D11S4083 RAG1 (TG)n D11S4102

99 101 N AA del 161 161 139 137

1.2 34 yo

PGT

2.1

2.2

103 101 R396C AA del 163 161 139 137

103 R396C 163 139

2.3 103 N 159 139

99 N 161 139

2.4 103 R396C 163 139

99 N 161 139

99 N 161 139

b PGT

Predicted Genotype:

Predicted Karyotype:

1

2

103 101 R396C AA del 163 161 139 137

103 101 R396C AA del 163 161 139 137

Affected

Affected

13, 13; 16, 16; 18, 18; 21, 21; 22, 22; XY

13, 13; 16, 16; 18, 18; 21, 21; 22, 22; XY

3

103 R396C 163 139

4

99 N 161 139

Carrier

13, 13; 16, 16; 18, 18; 21, 21; 22, 22; XY

103 R396C 163 139

99 N 161 139

Carrier

5

103 N 159 139

99 N 161 139

Normal

13, 13; 16, 16; 18, 18; 21, 21; 22, 22; XX

13, 13; 16, 16; 18, 18; 21, 21; 22, 22; XX

ET

ET

Fig. 5.4  PGT for OMS with aneuploidy testing. (a) Family pedigree with the mutation and haplotype analysis of parents (1.1 and 1.2) and children (2.1 affected and 2.2 healthy heterozygous carrier of paternal mutation). (b) (Top) Embryo biopsy analysis involving mutation analysis of five embryos, including two affected (embryos #1 and #2), two carriers of paternal mutation (embryos #3 and

#4), and one free of both paternal and maternal mutations (embryo #5). (b) (Bottom) Aneuploidy testing, showing normal chromosomal sets for all five embryos, two of which were transferred (embryos #4 and #5) resulting in the birth of healthy twins (2.3 normal and 2.4 heterozygous carrier of the paternal mutation)

PGT for AT was first reported for a Saudi family having three affected children [22]. The disease was caused by a deletion of more than two-thirds of the AT gene, detected by amplification of one of the deleted exons (exon 19). Of three embryos available for biopsy and testing, one was deletion-free and transferred, resulting in an unaffected pregnancy. Fanconi anemia complementation group A (FANCA), similar to FANCC mentioned above [1, 2], is an autosomal recessive disorder causing

bone marrow failure with increased predisposition to leukemia. Bone marrow transplantation is the only treatment, restoring hematopoiesis in FANCA patients. However, because any modification of the conditioning is too toxic for these patients, leading to a high rate of transplant-­related mortality, the HLA-identical stem cell transplantation from a sibling is particularly valuable to avoid late complications due to severe GVH. Couples at risk for producing a progeny with FA (Table 5.1) included carriers of mutations in

194

a

5  Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA) Markers order: D11S1325 D11S1781 D11S1343 ATM D11S2179

Family pedigree

151 150 116 Mut

153 143 124 N N 157

151 150 116 Mut

170

1

151 150 110 K1807E A2262P 172

153 143 124 N N 157

2.1

Affected

PGT by Sequential Polar Body Analysis

1.2

40 yo

151 150 110 K1807E A2262P 172

170

b

1.1

151 150 110 K1807E A2262P 172

151 150 110 N N 163

2

2.2

4

2.3

6

2.4 PGT

Carrier

9

10

11

Oocyte 151 150 110 K1807E A2262P 172

Predicted Genotype:

c

Affected

PGT by Blstomeres Analysis

1

Predicted Karyotype:

151 150 110 N N 163

Affected

Normal

2

Embryo

Predicted Genotype:

151 150 110 K1807E A2262P 172

151 150 110 K1807E A2262P 172

Affected

4

Normal

6

151 150 110 K1807E A2262P 172

151 150 110 K1807E A2262P 172

151 150 110 N N 163

Affected

9

10

Affected

11

SNR

153 143 124 N N 157

151 150 110 K1807E A2262P 172

Carrier

13, 13; 16, 16; 18, 18; 21, 21; 22, 22; XY

151 150 116 Mut 170

151 150 110 N N 163

Carrier 13, 13; 16, 16; 18, 18; 21, 21; 22, 22; XY

ET

153 143 124 N N 157

151 150 110 K1807E A2262P 172

Carrier 13, 13; 16, 16; 18, 18; 21, 21; 22, 22; XY

153 143 124 N N 157

151 150 110 N N 163

Normal 13, 13; 16, 16; 18, 18; 21, 21; 22, 22; XX

153 143 124 N N 157

151 150 110 K1807E A2262P 172

Carrier 13, 13; 16, 16; 18, 0; 21, 21; 22, 0; XX

151 150 116 Mut 170

151 150 110 K1807E A2262P 172

Affected 13, 13; 16, 16; 18, 18; 21, 21; 22, 22; XY

ET

Fig. 5.5  PGT for AT with aneuploidy testing. (a) Family pedigree showing the results of mutation and haplotype analysis in the parents (1.1 and 1.2) and the affected child (2.1). (b) Results of PB analysis of seven oocytes, only two of which (oocytes #4 and #9) were free of mutation, based on mutation and marker analysis. The remaining five oocytes were affected, containing both maternal mutations tested. (c) (Upper panel) Results of mutation and linked marker analysis of six embryos originating from the above oocytes (no sample was available from the

embryo originating from oocyte #2). Five of these six embryos were either normal (embryo #9) or carriers (embryos #1, #4, #6, and #10), while the remaining one embryo (embryo #11) was affected, inheriting both maternal and paternal mutations. (c) (Bottom panel) Results of aneuploidy testing for chromosomes 13, 16, 18, 21, 22, X, and Y, showing one double monosomy 18 and 22 in heterozygous unaffected embryo #10. Carrier embryos #1 and #6 were transferred resulting in birth of anunaffected child

the FANCC, FANCD2, FANCF, FANCI, FAMCCJ, and FANCA including a case with different maternal and paternal mutations. The maternal allele involved an ATG to AAG substitution in exon 1, resulting in methionine to lysine amino acid substitution; the paternal allele involved a 14 bp deletion involved in exon 2, representing a frameshift mutation. The paternal mutation was detected on the basis of the size difference in capillary electrophoresis of the PCR product, whereas the maternal mutation was detected by

NlaIII restriction digestion. This digestion cuts the normal sequence but leaves the mutant sequence uncut. In the other couple, only the paternal mutation was known, representing a T1131A mutation, due to ACT to GCT substitution in exon 34,  creating a restriction site for Fsp4HI. In addition, another restriction enzyme, TspRI, was used to cut the normal sequence on the opposite end. When a mutation was not identified, unaffected embryos were chosen by linkage analysis, based on haplotype constituted by

5.2 Immunodeficiency

195

five closely linked polymorphic markers. Overall, 65 unaffected HLA-matched embryos were transferred in 46 of 76  cycles, resulting in 19 unaffected pregnancies and 18 FA-free and HLA-­ ­ matched neonates, representing potential donors for their older siblings (Table 5.1). Of special interest is a case of PGT-HLA involving a consanguineous couple carrying the identical FANCG deletion mutation that led to their affected child who required stem cell transplantation treatment (Fig. 5.6). Following embryo testing by mutation and linked STR analysis, two HLA-matched and disease-free (normal and carrier) embryos were transferred, resulting in a twin pregnancy. At 15 weeks gestation, cSMART analysis of the pregnancy plasma determined

fetal DNA fractions of 14.2% and 6.6% for twin 1 and twin 2, respectively. The maternal plasma FANCG mutation analysis was consistent with presence of a carrier and a normal fetus. Additional retrospective studies of the WGA products from the transferred embryos, using single molecule sequencing, confirmed the FANCG genotypes of transferred embryos and the HLA match to the older sick sibling. PGT-­ HLA results were confirmed after delivery of twins, whose bone marrow was transplanted for treatment of the affected child [23]. X-linked adrenoleukodystrophy (X-ALD) affects the nervous system and the adrenal cortex, with the three main phenotypes. One of them manifests between ages 4 and 8 as attention defi-

Family pedigree Markers order: HLA Markers D9S1788 order: D9S1845 D6S1583 D9S1878 RING D9S1805 LH FANCG D6S273 D9S163 TNF-A D9S1804 TNF-D D9S1791 D6S62 D9S50 MOG D6S306

Family pedigree 36 yo N / c.260 del G

N / c.260 del G

HLA

HLA

HLA MATCH AFFECTED c.260 del G

1

CARRIER

46,XY

3

2

NORMAL

AFFECTED

3

CARRIER

46,XY

46,XY

FET

FET

EMBRYO #3

46, XY

4

CARRIER

46,XX

EMBRYO #4

CARRIER

5

PGT

5

FET

Baby

2

2

3

HLA HLA NIPT confirmed PGT predictions for both twins

HLA

PGT

NORMAL

NORMAL or EUPOLID CARRIE

5

6

7

NORMAL AFFECTED

46,XY

46, XX

10

CARRIER CARRIER AFFECTED

RECOMBINANT

46,XY 46,XX

EMBRYO #11

Fig. 5.6  Family pedigree of consanguineous family, carrying the identical FANCG deletion mutation who have an affected child, requiring stem cell transplantation treatment. PGT-M combined with PGT-HLA, based on testing by mutation and linked STR analysis, identified two HLA-matched and disease-free (normal and carrier)

8

46, XY, DEL(8)

NA

11

12

CARRIER

CARRIER

13

14

CARRIER

46, XY,DEL(8) 46,XY, DEL1 47,XX, +16

EMBRYO #12

46, XY, DEL(1)

CARRIER

46,XY

EMBRYO #13

17

CARRIER

46,XX

47, XY, +16

embryos (embryos #11 and #13) which were transferred, resulting in a twin pregnancy. At 15 weeks gestation, the diagnosis was confirmed by NIPT using cSMART analysis (see the text). HLA results were confirmed after delivery of twins, whose bone marrow was transplanted for treatment of the affected child

196

5  Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)

cit disorder, followed by progressive impairment of cognition and behavior, vision, hearing, and motor function, leading to total disability within 2 years. The other type of adrenomyeloneuropathy manifests in the late 20s as progressive paraparesis, sphincter disturbances, and hearing loss. The third presents with primary adrenocortical insufficiency by 7–8 years of age. Regardless of other symptoms, 99% of patients have elevated very long chain fatty acids (VLCFA). There are more than 200 different ABCD1 mutations, which may be detected by PCR and direct sequencing. A few large deletions are best identified by Southern blot analysis. Carrier screening and prenatal diagnosis are available, amenable also for PGTM.  PGT-M was performed for two couples, involving the testing for G343D mutation, representing a sequence change from aspartic acid to glycine, caused by a single (G to A) sequence change in the nucleotide 1414 (G1414A) of the ABCD gene. PGT was based on Fok I restriction digestion, which creates two fragments in the PCR product of a normal gene, leaving the mutant one uncut. Five PGT cycles were performed, resulting in only one unaffected HLA-­matched embryo suitable for transfer. However, this failed to produce a clinical pregnancy (Table 5.1). Hyperimmunoglobulin M syndrome (HIGM) is a rare immunodeficiency characterized by normal or elevated serum IgM levels, as well as absence of IgG, IgA, and IgE, that result in an increased susceptibility to infections. This is manifested in the first few years of life and associated with a high frequency of autoimmune hematologic disorders; gingivitis, ulcerative stomatitis, fever, and weight loss are often present. HIGM is caused by mutation in the CD40 ligand gene (CD40LG), located on chromosome Xq26. This leads to a defective CD40 ligand expression resulting in failure of T cells to induce IgE synthesis in interleukin-4-treated B cells. Although regular administration of intravenous immunoglobulins is used for treatment, best results have been obtained by HLA-matched bone marrow transplantation. This makes PGT-HLA the method of choice for those who lack a suitable HLA match among their relatives. A total of 16

PGT-HLA cycles were performed for 11 couples with HIGM (Table 5.1), with transfer of 15 unaffected HLA-matched embryos in 10  cycles, yielding 9 clinical pregnancies and birth of 8 unaffected HLA-matched children. Among the mutations tested were C218X in exon 5 of CD40 ligand gene (CD40LG), C218X mutation in exon 4 c.437_38 ins A, and ins T in exon 4 c.397. Maternal mutations were analyzed by PB1 and PB2, followed by HLA and aneuploidy testing in embryo biopsy. A CYS218STOP mutation in exon 5 was detected by restriction digestion, given the restriction site for Cac 81 was eliminated. Thus, there were two fragments in PCR product from the normal gene, whereas the mutant gene product was uncut. For higher accuracy, another restriction enzyme (Mnl I) was applied, creating three fragments in the mutant PCR product compared to two fragments in the normal gene. Figure 5.7 presents the case of PGT-HLA for a couple at risk for producing offspring with HIGM. Of 15 oocytes tested by PB1 and PB2, 5 oocytes with conclusive results appeared to be free of maternal mutation, but only 1 was a maternal HLA match (embryo #2). In addition, three of five oocytes with the maternal mutation were HLA-matched (embryos #11, #13, and #15). However, embryos #13 and #15 were affected and a non-paternal match; only the maternal mutant chromosome was detected in embryo #11. Overall, only embryos #2 was predicted to be maternal mutation-free by PB analysis, a normal female, and also a paternal match. The transfer of this single embryo resulted in a singleton pregnancy, confirmed to be unaffected and HLA-­ matched by amniocentesis, yielding the birth of an unaffected HLA-matched baby girl. The first stem cell transplantation was performed using cord blood stem cells obtained from the child described. Unfortunately, no engraftment was achieved. A second transplantation was performed 1  year later, using bone marrow mixed with the remaining portion of the cord blood sample. This resulted in successful engraftment and reconstitution of the sibling’s bone marrow, thus a total cure.

5.2 Immunodeficiency

a

197

Family pedigree

126 186 175 148 111 274 167 116

130 159 181 159 103 270 165 114

HLA 126 186 175 148 111 274 167 116

144 167 171 163 101 270 175 120

HLA

b

1.1

182 N 118 234 170

144 167 171 163 101 270 175 120

1.2 186 N 120 232 176

174 M 124 228 172

XY

XX

130 188 175 148 111 276 165 114

HLA 174 M 124 228 172

182 N 118 234 2.2170

2.1

PGT

XY

186 N 120 232 176

?

?

?

?

1.3

1.4

1.5

1.6

10

11

126 186 175 148 111 274 167 116

XX

1.7

144 167 171 163 101 270 175 120

HLA

PGT 1

2

3

4

5

6

7

9

13

15

2.2 X-chromosome Markers : DXS1187 CD40 (CA)n DXS8094 DXS1062

HLA Markers: D6S276 Mog a D6s265 D6S510 TNF a D6S273 LH1 D6S291 Predicted Genotype

186 N 120 232 176

130 159 181 159 103 270 165 114

Y 130 188 175 148 111 276 165 114

Normal male Non-match

182 N 118 234 170

186 N 120 232 176

126 144 186 167 175 171 148 163 111 101 274 270 167 175 116 120 Normal female Match

ET

182 N 118 234 170

186 N 120 232 176

182 N 118 234 170

174 M 124 228 172

126 130 186 188 175 175 148 148 111 111 274 276 167 165 116 114

130 130 159 188 181 175 159 148 103 111 270 276 165 165 114 114

Normal female Non-match

Carrier Non-match

182 N 118 234 170

174 M 124 228 172

126 130 186 188 175 175 148 148 111 111 274 276 167 165 116 114 Carrier Non-match

186 N 120 232 176

186 N 120 232 176

182 N 118 234 170

186 N 120 232 176

Y

126 130 186 188 175 175 148 148 111 111 274 276 167 165 116 114

130 159 181 159 103 270 165 114

144 167 171 163 101 270 175 120

126 130 186 188 175 175 148 148 111 111 274 276 167 165 116 114

Normal female Non-match

Normal male Non-match

Y

Normal male Non-match

182 N 118 234 170

126 186 175 148 111 274 167 116

186 N 120 232 176

130 188 175 148 111 276 165 114

Normal female Non-match

174 M 124 228 172

126 186 175 148 111 274 167 116

174 M 124 228 172

144 167 171 163 101 270 175 120

X0 Match

130 159 181 159 103 270 165 114

Y 144 167 171 163 101 270 175 120

Affected Male Non-matched

174 M 124 228 172

130 159 181 159 103 270 165 114

Y

144 167 171 163 101 270 175 120

Affected Male Non-matched

Fig. 5.7  Preimplantation HLA typing combined with PGT for X-linked hyperimmunoglobulin M syndrome. (a) Family pedigree, with marker order  for parents and sffected sibling. CD 40 gene haplotype assignment is based on genomic DNA testing. Paternal and maternal matching HLA haplotypes are shown in different colors. (b) (Upper panel) PCR analysis of blastomeres removed from 12 embryos showed that all but three embryos (#11, #13, and #15) were predicted to be unaffected. (b) (Lower

panel) HLA typing was performed simultaneously with mutation analysis of all blastomeres. Embryo #2 was predicted to be a normal female and to have the same HLA profile as the affected sibling (2.1). The transfer of this embryo resulted in pregnancy and the birth of a healthy unaffected HLA-matched baby girl (2.2). Cord blood stem cells were collected at birth for stem cell transplantation resulting in a total cure

Wiskott–Aldrich syndrome (WAS) is a lethal X-linked immunodeficiency in which lymphocyte dysfunction and thrombocytopenia result in severe infections, bleeding, and increased risk of lymphoproliferative malignancies. Supportive therapy may increase survival rate, but the only way to avoid early mortality is bone marrow transplantation. WAS is caused by a mutation in the WAS gene mapped to the Xp11.22–11.23 region, resulting in actin polymerization, T lymphocytes exhibiting severe disturbance of the actin cytoskeleton. The gene has 12 exons that

encode a 502 amino acid cytosolic protein, expressed exclusively in hematopoietic cells. PGT was performed for five couples at risk for producing a progeny with WAS. One couple had two affected sons carrying the missense Leu39Pro mutation in exon 1 of WAS gene, which was due to a single nucleotide (CTT to CCT) substitution at position 150. This leads to substitution of leucine by proline at position 39. Mutation testing was done using Scr FI restriction digestion, cutting the mutant and leaving the normal gene product intact. A total of 9  cycles

5  Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)

198

were performed, resulting in transfer 17 unaffected embryos in 9  cycles, yielding 6 clinical pregnancies and 6 unaffected children born (Table 5.2). X-linked hypohidrotic ectodermal dysplasia with immunodeficiency (HED-ID) is a congenital disorder of the teeth, hair, and eccrine sweat glands, inherited as an X-linked recessive condition. HED-ID is caused by, approximately, two dozen different mutations in the IKK-gamma gene (IKBKG or NEMO) located in Xq28. The gene consists of ten exons and codes for a scaffold protein that binds IKK-alpha and IKK-beta, being essential for forming a functional IKK

a

Family pedigree

HLAMarker: MOG a RF D6S1568 TNF 9N2 D6S273 LH1 D6S291

168 243 131 102 134 272 149 117

186 258 137 111 130 276 164 124

139 137 134 156 N 165

HLA

complex. The disease is characterized by susceptibility to microbial and streptococcal infections, dysgammaglobulinemia, poor polysaccharide-­ specific antibody responses, and depressed antigen-­ specific lymphocyte proliferation. Intravenous immunoglobulins and prophylactic antibiotics may be useful in improving clinical status, but bone marrow transplantation is required to prevent early mortality. One of the ten PGT-HLA cycles performed for HED-ID is presented in Fig.  5.8. The mother was a carrier of a L153R mutation, resulting from T to G change (CTG-  >  CGG) in exon 4 of the NEMO (IKBKG) gene. Leucine

1.1

1.2

XX

2.1

HLA

168 252 133 102 132 270 164 115

168 252 133 102 132 270 164 115

170 270 135 109 132 272 151 117

128 117 140 156 M 167

XY

168 243 131 102 134 272 149 117

b

130 143 134 154 N 165

X-Markers order: DXS 9929 DXS8103 DXS1684 DXS8087 NEMO Gene DXYS154

HLA

2.2

PGT 168

128 117 140 156 M 167

243 131 102 134 272 149 117

X Y

168 252 133 102 132 270 164 115

HLA

139 137 134 156 N 165

128 117 140 156 M 167

XX

PGT

Embryo #

2

3

4

6

8

12

13

16

17

18

20

21

24

25

26

27

NEMO

HLA

Carrier Match

ET

Carrier Recombinant

XXY Non-match

Fig. 5.8  PGT for the mutation in NEMO gene with preimplantation HLA typing. (a) Family pedigree showing maternal and paternal matching HLA haplotypes in different colors. Marker order for testing NEMO gene is located next to the maternal haplotypes. Paternal (1.1), maternal (1.2), and affected sibling (2.1) NEMO gene haplotype assignment is based on genomic DNA testing. (b) (Upper panel) Results of blastomere DNA analysis from 16 embryos showing that 3 embryos were affected (embryos #17, #20, and #21), one with an extra X chromosome,

Affected Match

Normal Match

ET

suggesting the XXY genotype, with the remaining being either carriers or unaffected. (b) (Lower panel) HLA typing was performed simultaneously with mutation analysis of all blastomeres, showing that two of the unaffected embryos (#12 and #26) were also HLA-matched to the affected sibling. The transfer of these embryos resulted in pregnancy and the birth of a healthy unaffected HLA-­ matched baby girl (2.2). Cord blood stem cells were collected at birth for stem cell transplantation

5.3  Preimplantation HLA Matching Without PGT-M

was replaced with arginine at position 153. Because of the presence of a closely linked pseudogene having a normal sequence at the position of the mutation and thus co-amplified with the transcribed gene, a special design was developed to avoid misdiagnosis. A total of 16 embryos were analyzed, of which 6 were derived from oocytes free of maternal mutation based on PB1 and PB2 testing; however, none of these was a maternal HLA match. Figure 5.8 shows that of 16 resulting embryos for which embryo biopsy results were available both for mutation analysis and HLA typing, 3 were affected males (embryos #17, #20, and #21; only the latter being HLA-matched). There were four female carriers, two of which were non-matched (embryos #3 and #4); one was HLA-recombinant (embryo #13); one was HLA-matched (embryo #12). The remaining seven embryos were unaffected, including two male non-matched embryos (embryos #16 and #24), the former containing extra maternal X-chromosome; five normal female embryos were present, of which only one (embryo #26) was HLA-matched. While the normal embryos that were not HLA-matched to the affected sibling were frozen for future use by the couple, embryo #26 together with embryo #12, a normal female carrier, were transferred and resulted in a singleton pregnancy and the birth of an unaffected child confirmed to be HLA-­matched to the affected sibling. Cord blood from this child was collected and transplanted to the affected sibling, resulting in a complete cure. The presented data show the usefulness of PGT for SCID. PGT provides couples at risk with the option to avoid the affected pregnancy and have a progeny free of SCID. If there is already an affected child in the family, PGT-HLA makes it also possible to have access to the HLA-­identical stem cell transplantation through selection and transfer of those unaffected embryos which are also HLA-matched to the sibling. Because the finding of the HLA-identical stem cell donor is the key for achieving the success in stem cell transplantation [20], a complete cure was observed in the above cases of stem cell transplantation in siblings with immunodeficiency.

199

5.3

Preimplantation HLA Matching Without PGT-M

Preimplantation HLA matching without testing for a causative gene was first performed for leukemia and Diamond–Blackfan anemia (DBA). Both conditions require bone marrow or cord blood transplantation treatment [24]. The latter condition was sporadic and did not require mutation testing. The sole indication was for HLA typing. Mutation analysis may, however, be required for patients at risk of producing ­offspring with DBA, caused by mutations in the gene encoding ribosomal protein S19 on chromosome 19 (19q13.2); other genes are mapped to chromosome 8 (8p23.3-p22). Still, the majority of DBA is sporadic with no mutation detected, such as in two cases performed in our first reported experience [24]. There is no difference in performing preimplantation HLA testing without PGT-M, except for limiting the analysis of day 5 embryo biopsy material only to HLA matching to the sibling requiring stem cell transplantation, using a multiplex heminested PCR system. A haplotype analysis of the father, mother, and affected child was performed in each family prior to preimplantation HLA typing, using a set of polymorphic STR markers located throughout the HLA region, as shown above in Fig. 5.1. This allowed detecting and avoiding misdiagnosis due to preferential amplification and ADO, potential recombination within the HLA region, and a possible aneuploidy or uniparental disomy of chromosome 6, which may also affect the diagnostic accuracy of HLA typing of the embryo. Our experience includes a total of 119 clinical cycles for 60 couples, with preselection of 108 HLA-matched embryos for transfer (Table  5.1). The proportion of embryos predicted to be HLA-­ matched to the affected siblings was 21.5%, not significantly different from the expected 25% (Table  5.3). The transfer of 108 HLA-matched embryos transferred in 73 clinical cycles resulted in 25 singleton clinical pregnancies and 22 HLA-­ matched children born. These results suggest that testing of an available number of embryos per cycle allows preselecting a sufficient number of the HLA-matched embryos for transfer to achieve

200

5  Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)

Table 5.3  Chances for detection of disease-free and HLA-matched embryo in preimplantation HLA typing (PGT-HLA)

The relevance of aneuploidy testing for chromosome 6 for accuracy of diagnosis is seen also from the results of HLA typing in the other HLA match only – ¼ (25%) cycle, resulting in birth of a baby who was HLA-­ Autosomal recessive or X-linked free + HLA match – matched to the sibling with ALL (Fig. 5.10). Two ¾ × ¼ = 3/16 (18.75%) of ten embryos tested (of which only eight Autosomal dominant free + HLA match – ½ × ¼ = 1/8 (12.5%) embryos are shown in Fig. 5.10) appeared to have Autosomal recessive or X-linked free + HLA match + only maternally derived chromosomes 6, one aneuploidy-free – ¾ × ¼ × ½ = 3/32 (9.4%) with only one maternal chromosome (embryo Autosomal dominant free + HLA match + aneuploidy-­ #1) and the other with two maternal chromofree – ½ × ¼ × ½ = 1/16 (6.25%) somes, thus representing uniparental maternal disomy of chromosome 6 (embryo #2). In addia clinical pregnancy and birth of an HLA-­ tion, crossing over between D6S291 and class II matched progeny. HLA alleles was evident, making this embryo The usefulness of detecting recombination unacceptable for transfer. Of the remaining within the HLA region is demonstrated in embryos, only two were HLA-matched to the Table 5.4, describing the results of HLA typing affected sibling, which were transferred resulting of one of the cycles resulting in the birth of an in the birth of an HLA-matched baby. HLA-matched child to a sibling with acute lymPresented data show the utility and reliability phoid leukemia (ALL). Of 10 embryos tested of preimplantation HLA matching for families simultaneously for 11 alleles within the HLA having affected children with bone marrow disorregion in this family, crossing over between ders who may wish to have another child. An D6S2426 and Ring alleles was observed in HLA-matched donor of stem cells would provide embryos #4, #7, and #9. Of the remaining seven the option for  transplantation treatment of the embryos, three were fully matched (embryos #2, affected sibling. As seen from our data, HLA-­ #6, and #8), whereas the other four were HLA-­ matched embryos were preselected and incompatible to the affected sibling, as seen from ­ transferred in all cycles, resulting in clinical the haplotypes of the mother, father, and affected pregnancies and the birth of HLA-matched chilchild, presented in Table 5.4. Two recombination dren in almost every second transferred cycle. events were  detected in  preimplantation HLA Results also demonstrate the prospects of typing undertaken for DBA (Fig. 5.9). One  was applying this approach to a variety of conditions, a maternal recombination seen in the embryo #8 in which an HLA-compatible donor is required for results, whereas HLA testing of the other six stem cell transplantation. This provides a realistic embryos revealed three HLA matches to the option for couples who would like to have another affected sibling Another one was a double  recom- child, for they may now potentially provide an bination detected in embryo #16 with both mater- HLA-matched progeny for an affected sibling. In nal and paternal crossing over (both in Ring addition to leukemia and sporadic forms of DBA, allele). One more embryo was with  trisomy 6 the method may be applied for in other circum(embryo #5) with an additional maternal chromo- stances. Couples having affected children with difsome 6; thus, this and the other two embryos (#8; ferent cancers may be awaiting an HLA-compatible #16) were unacceptable for transfer. Two HLA donor without definitive prospects. However, with matched embryos  were transferred back to the progress in allogenic bone marrow transplantation patient and resulted in birth of an HLA-matched for treatment of leukemia, the number of preimbaby (Fig.  5.9). Cord blood collected from this plantation HLA typing cycles for this condition is baby was transplanted to the affected sibling, decreasing in contrast to overall increased numresulting in a complete cure. bers of PGT-HLA.

Embryo no. 1 2 130/118 144/139 3/2 1/32 16,167 148/163 AD 27 27/18 118/114 114/124 94/102 110/102 275/273 273/271 ADO/1 11/10 181/208 181/191 162/162 160/155 144/130 140/144 Non-­ Match match

3 130/118 3/2 163/167 ADO/27 118/114 94/102 275/273 ADO/1 181/208 162/162 144/130 Non-­ match

4 130/118 3/2 163/167 FA 118/114 94/102 275/273 ADO/1 181/208 162/162 140/130 Non-match recombinant

5 130/139 3/32 163/163 FA 118/124 94/102 275/271 ADO/10 181/191 162/155 144/144 Non-­ match

6 144/139 1/32 148/163 27/18 114/124 110/102 273/271 11/10 181/191 160/155 140/144 Match

ADO allele dropout, FA failed amplification. HLA matched alleles are shown in bold

HLA genes and STRs D6S276 HLA A D6S510 HLA B MIB TNF a D6S273 HLA-DRB1 G51152 Ring 3CA D6S426 Predicted genotype 7 130/118 FA 148/167 ADO/27 118/114 94/102 275/273 ADO/1 181/208 162/162 144/130 Non-match recombinant

8 144/139 1/32 148/163 27/18 114/124 110/102 273/271 11/10 181/191 160/155 140/144 Match

9 130/139 3/32 163/163 ADO/18 118/124 94/102 275/271 ADO/10 181/191 162/155 144/130 Non-match recombinant

10 130/139 3/32 163/163 ADO/18 118/124 94/102 275/271 FA 181/191 162/155 144/144 Non-­ match

Table 5.4  Preimplantation HLA typing resulting in the birth of an HLA-matched baby for the affected sibling with acute lymphoid leukemia Father 144/130 1/3 148/163 27/57 114/118 110/94 273/275 11/7 181/181 160/162 140/144 NA

Mother 118/139 2/32 167/163 27/18 114/124 102/102 273/271 1/10 208/191 162/155 130/144 NA

Affected baby 144/139 1/32 148/163 27/18 114/124 110/102 273/271 11/10 181/191 160/155 140/144 NA

5.3  Preimplantation HLA Matching Without PGT-M 201

5  Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)

202

a

Markers order: D6S426 147 RING 159 DRB1 1*11 TNF 110 HLA B B51 HLA C C4 D6S276 118 D6S265 179 HLA A 2 D6S258 147

Family pedigree

143 155 1* 01 94 B 51 C 4 144 181 A 2 133

1.1

139 157 1*01 100 B 35 C4 144 172 A3 135

1.2

129 161 1*07 94 B 57 C4 135 176 A 24 137

PGT 2.1

b

147 159 1*11 110 B51 C4 118 179 A2 147

PGT

Embryo 1

147 159 1*11 110 B51 C4 118 179 A2 147

139 157 1*01 100 B35 C4 144 172 A3 135

Match

5

147 159 1*11 110 B51 C4 118 179 A2 147

129 161 1*07 94 B 57 C4 135 176 A 24 137

8

139 157 1*01 100 B 35 C4 144 172 A3 135

Trisomy6

147 159 1*11 110 B51 C4 118 179 A2 147

147 159 1*11 110 B51 C4 118 179 A2 147

10

139 157 1*07 94 B 57 C4 135 176 A 24 137

Recombinant

ET

147 159 1*11 110 B51 C4 118 179 A2 147

139 157 1*01 100 B 35 C4 144 172 A3 135

11

139 157 1*01 100 B 35 C4 144 172 A3 135

Match

ET

Fig. 5.9 Preimplantation HLA typing for Diamond– Blackfan anemia, resulting in the birth of an HLA-­ matched child. (a) Family pedigree with marker order and haplotypes of the mother, father, and affected child. HLA matching haplotypes are shown in  different colors. (b) Results of HLA typing of biopsied blastomeres from eight embryos (other eight embryos which were also tested are not shown). Embryos #1, #10, #11, #12, and #18 are HLA-matched to the affected sibling (see a panel). Embryo #8 is a maternal non-match due to maternal

5.4

2. 2

139 157 1*01 100 B 35 C4 144 172 A3 135

Limitations and Future Prospects of PGT- HLA

Presented data demonstrate that PGT-HLA became a practical option, available for wider application in order to further improve the radical treatment for congenital and acquired bone marrow failures by stem cell transplantation. Despite the high rate of preferential amplification and ADO in PCR analysis of single or small number of biopsied embryonic cells, a potential recombination within the HLA region, and a high rate of mosaicism, the approaches described above

147 159 1*11 110 B51 C4 118 179 A2 147

139 157 1*01 100 B35 C4 144 172 A3 135

Match

12

147 159 1*11 110 B51 C4 118 179 A2 147

16

139 157 1*01 100 B35 C4 144 172 A3 135

Match

147 159 1* 01 94 B51 C4 144 181 A2 133

129 159 1*01 100 B35 C4 144 172 A3 135

Double recombinant

18

147 139 159 157 1*11 1*01 110 100 B51 B35 C4 C 4 118 144 179 172 A2 A 3 147 135

Match

recombination in the Ring allele, as well as embryo #16, which is both a paternal and maternal non-match, due to double recombination in the paternal and maternal Ring alleles. Embryo #5 is also a non-match due to an extra maternal chromosome, suggesting trisomy 6.  Two HLA matched embryos were transferred back to the patient and resulted in birth of an HLA-matched baby, whose cord blood was transplanted to the affected sibling, resulting in a total cure

appear to be highly accurate in preselecting HLA-matched embryos for transfer. As mentioned, one of the major limitations of PGT-HLA is a relatively high frequency of recombination in the HLA region, with a few possible hot spots. This may affect the accuracy of preimplantation HLA typing and the outcome of the whole procedure. In our experience involving the analysis of 1713 embryos tested for HLA, 1634 (95.5%) were nonrecombinant, 52 (3%) with maternal, and 27 (1.5%) with paternal recombination. In fact, the prevalence of recombination is even higher based on family evaluation, performed prior to PGT in 114 fami-

5.4  Limitations and Future Prospects of PGT- HLA

a

Markers order: D6S291 TAP 1 G51152 LH 1 D6S273 D6S62 TNF A D6S1624 D6S510 MOG D6S306

Family pedigree

124 207 181 171 272 197 99 178 163 167 106

114 207 181 158 270 200 103 176 148 174 104

203

1.1

1.2

116 218 187 149 268 195 97 184 167 178 108

122 207 183 151 279 189 93 178 148 186 106

PGT 2.1

b

114 207 181 158 270 200 103 176 148 174 104

PGT 1

122 207 183 151 279 189 93 178 148 186 106

Monosomy6

2

122 207 183 151 279 189 93 178 148 186 106

116 218 187 149 268 195 97 184 167 178 108

UNIPARENTAL DISOMY

4

114 207 181 158 270 200 103 176 148 174 104

116 218 187 149 268 195 97 184 167 178 108

Match

ET

2. 2 114 207 181 158 270 200 103 176 148 174 104

116 218 187 149 268 195 97 184 167 178 108

5

124 207 181 171 272 197 99 178 163 167 106

116 218 187 149 268 195 97 184 167 178 108

Maternal match

6

114 207 181 158 270 200 103 176 148 174 104

116 218 187 149 268 195 97 184 167 178 108

7

122 207 183 151 279 189 93 178 148 186 106

Paternal match

124 207 181 171 272 197 99 178 163 167 106

10

9

122 218 187 149 268 195 97 184 167 178 108

Maternal Recombination

114 207 181 158 270 200 103 176 148 174 104

116 218 187 149 268 195 97 184 167 178 108

Match

124 207 181 171 272 197 99 178 163 167 106

122 207 183 151 279 189 93 178 148 186 106

Non- match

ET

Fig. 5.10  Preimplantation HLA typing for acute lymphoid leukemia (ALL), resulting in the birth of an HLA-­ matched baby. (a) Family pedigree with marker order and haplotypes of the mother, father, and affected child. Matching maternal and paternal haplotypes are shown in different colors. (b) Results of HLA typing of embryo biopsy sample  from eight embryos. Embryos #4 and #9 are HLA-matched to the affected sibling (see a panel). Embryo #1 is a maternal non-match with no paternal chro-

mosome present (monosomy 6). Embryo #2 is also a non-­ match due to only maternal chromosomes present (uniparental disomy). Embryo #7 is both a paternal and maternal non-match, the latter being due to maternal recombination in the Ring allele. The other two embryos are a non-match, embryo #6 and embryo #10, while  other two embryos (embryos # 4 and # 9) were full match and transferred, resulting in birth of HLA match baby 

lies. The 6.1% recombination rate in siblings requiring HLA-compatible bone marrow transplantation suggests that preimplantation HLA typing may be unrealistic for identifying the HLA match for these siblings. Therefore, haplotype analysis prior to initiation of the actual cycle is required, so that the couples may be informed about their possible options. For example, in one of our cases performed for thalassemia, the fact that the child was recombinant became available only after PB1 analysis, without which maternal haplotypes could not be established. While paternal haplotypes may be identified through sperm typing, testing for maternal haplotypes requires PB analysis, or

maternal somatic cell haploidization, which may be performed by somatic cell nuclei transfer and fusion with matured oocytes [25]. As shown in Fig. 5.11, preparatory testing allowed identifying the siblings with maternal recombination; thus, if so it could have been unrealistic to identify the exact match. Therefore, couples should have been informed that only relatively close matches may be identified, warranting discussions with the pediatric hematologist on acceptable HLA profiles. Another important limitation is that most patients requesting preimplantation HLA typing are of relatively advanced reproductive age. Thus outcome of the procedure has not yet been suffi-

5  Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)

204

D6S306 MOG MIB 9N2 LH1 TAP RING D6S1618 D6S439 D6S1583

Telomere

100 167 123 133 151 205 155 128 128 149

104 182 119 135 167 207 161 143 123 142

HLA

HLA-F

Class I

104 182 119 135 167 207 161 143 123 142

HLA-E HLA-C HLA-B

Oocyte #

Class III

PGT IVSI-1 / IVSI-110

HLA-A

X

N / IVSI-110

IVSI-1 / N

1

114 172 106 131 173 216 159 128 126 149

HLA

104 182 123 135 173 216 159 128 126 149

114 172 106 131 164 207 161 145 123 153

HLA

N/N 114 172 106 131 173 216 159 128 126 149

2

3

IVSI-110

N

4

HLA HLA-DR

Class II

HLA-DQ

HBB

N

N

Embryo #

HLA HBB

N/N

ET

N/IVSI-110

IVSI-1 / N

N/N

ET

Fig. 5.11  Maternal recombination detected in thalassemia major sibling in preimplantation HLA typing combined with PGT. (Top panel) Family pedigree with HLA haplotype analysis based on parental and affected child’s genomic DNA testing. HLA marker order is presented on the upper left for the father, who was a heterozygous carrier of thalassemia gene IVS 1–1, and on the upper right for the mother, a heterozygous carrier of thalassemia mutation IVS1–110. Paternal and maternal HLA haplotypes are shown in different colors: paternal in blue/yellow and maternal in red/green. As seen from the HLA haplotypes of the affected child in need for transplantation, for whom HLA matching is performed, the maternal HLA contribution is recombinant (red and green instead of the expected red or green) between HLA-DR and HLA-B genes, shown schematically on chromosome 6 (on the far left). (Middle panel) Sequential multiplex polar body analysis for maternal mutation, linked polymorphic mark-

ers, and HLA haplotypes, showing that the oocyte #2 is affected (IVS1–110), while oocytes #1, #3, and #4 are normal, with no recombination in the HLA cluster. (Bottom panel) Embryo biopsy results revealed two heterozygous carrier embryos (embryos #2 carries maternal mutation IVS1–110 and embryo #3 carries paternal mutation IVS1–1) and two homozygous normal ones (#1 and #4). HLA typing (presented by respective colors) shows that neither of these embryos is fully HLA-matched to the sick sibling. Two embryos, #1 and #4, predicted to be homozygous normal, and partially HLA-matched, were transferred back to the mother, yielding a singleton pregnancy and the birth of a thalassemia-free baby, who may still be considered for possible bone marrow transplantation, as there is no probability of producing a completely HLA-matched offspring for the affected sibling with recombinant HLA haplotypes

ciently high, and many patients still undergo two or more attempts before they become pregnant and deliver an HLA-identical offspring. Concomitant PGT-A is thus useful for improving the reproduc-

tive outcome of PGT-HLA.  This also minimizes risk of delivering a child with chromosomal disorders, providing reassurance for patients concerned about their pregnancy outcomes [26].

5.4  Limitations and Future Prospects of PGT- HLA

PGT-A is currently offered as an integral component of preimplantation HLA typing to the patients of advanced reproductive age in our experience resulting in an increasing number of preimplantation HLA typing cycles combined with or without PGT-M. Although the chances of preselecting unaffected HLA-matched embryos that could be also euploid are very low (see Table 5.3), our preliminary results of the reproductive outcome comparison between the groups of PGT-HLA with and without PGT-A showed a significant difference in results [27] (Table 5.5). Despite transferring a lower number of embryos, the pregnancy rate was higher in PGT-A group, suggesting its utility in preimplantation HLA typing. Avoiding transfer of HLA-identical embryos that are chromosomally abnormal and destined to be lost anyway either before or after implantation is clearly desirable. Alternatively, incidental transfer of aneuploid embryos in the absence of chromosomal testing is likely to lead to implantation and pregnancy failures in PGT-­ HLA cycles or compromise pregnancy outcome through spontaneous abortions. Although more data are needed to further prove the impact of PGT-A on the outcomes of PGT-HLA, the presented data suggest that approximately half of the aneuploidy-free embryo transfers following PGT-HLA resulted in pregnancy and the birth of HLA-matched chilTable 5.5  Outcome of preimplantation HLA typing with and without aneuploidy testing

Patient/cycle Total embryos Matched embryos Non-matched embryos Transfers No. of embryos transferred Pregnancy Birth

HLA 11/25 224 48

HLA plus aneuploidy testing 14/27 204 21 (36)

Total 25/52 428 69 (84)

176

168

344

21 (84%) 33 (1.6)

13 (48%)

34

19 (1.4)

52

6 7 (53.8%) (28.5%) 6 6

13 (38%) 12

205

dren. This compares to 28.5% pregnancy rate following the transfer of HLA-matched embryos not tested for aneuploidy. As shown by a comparison of the number of cycles performed with or without aneuploidy testing in Table  5.5, despite unavailability of aneuploidy-free embryos for transfer in over half of the PGT-HLA cycles combined with PGT-A, comparable numbers of pregnancies and births of HLA-matched children were observed, indicating a clinical relevance of avoiding chromosomally abnormal embryos from transfer in PGT-HLA [27]. This may be due to the potential of PGT-A to identify at least 50% of chromosomally abnormal embryos in patients of advanced reproductive age, despite lowering the probability of detecting the embryos for transfer by half. In fact, the mean number of embryos for transfer was approximately 1.0 on the average, which also reflects the lower probability of identification of HLA-matched unaffected embryos free of aneuploidy, taking into consideration the average number of available embryos with results, which is usually much lower in women of advanced reproductive age (under ten embryos on the average in our experience). With one in two embryos expected to be aneuploid, one in four HLA-­ matched, and three in four unaffected in autosomal recessive conditions, the overall probability of finding a suitable embryo for transfer could not be expected to be higher than one in ten embryos (see also Table 5.3). Thus, with the availability of only less than ten embryos on the average having conclusive results in our material, only one HLA-matched unaffected euploid embryo may be expected to be available for transfer, assuming also that not all embryos develop to the status acceptable for transfer. However, with present tendency of limiting the transfer to only one blastocyst to avoid multiple pregnancies, availability of a single euploid embryo for transfer is usually sufficient to obtain a clinical pregnancy and birth of an HLA-­identical progeny for stem cell transplantation into the affected siblings. The usefulness of PGT-A is also obvious for the diagnostic accuracy improvement, as an error in the number of chromosomes may lead to mis-

206

5  Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)

diagnosis in testing for the causative gene and HLA. Thus, in addition to avoiding chromosomally abnormal embryos from transfer, testing for the copy number of chromosomes may become an important requirement for achieving the accuracy of PGT-HLA. In fact, the follow-up analysis of the mutant oocytes and embryos and the pregnancy outcomes in our experience did not find any misdiagnosis, suggesting an extremely high specificity and sensitivity using present PGT-­ HLA technology. Nonetheless, PGT-A will have less utility when only a few embryos are available for testing. To overcome this, two or more cycles are initiated to collect a sufficient number of embryos for analysis. But meaningful batching is not always possible because some older patients may not be able to produce additional oocytes. The only possibility in such cases is to offer the couple the option of HLA testing for the women’s younger sister, so that the sister’s HLA-matched donor oocytes could potentially be used for PGT-­ HLA cycle. Our experience showed the usefulness of this option as demonstrated by the PGT-HLA case below. The patient was a 43-year-old female partner who requested PGT-HLA because of her risk for producing affected child with 4  bp deletion c.1115_1118  in FANCA. She had two affected children with FANCA. Both affected babies had the same HLA haplotypes. In the initial cycles using the patient’s own oocytes, she had one successful clinical pregnancy with unaffected HLA-­ matched fetus; however, the pregnancy resulted in spontaneous abortion. As there was little hope that she could produce a sufficient number of the oocytes for the further PGT-HLA cycles, and taking into consideration that she had a younger cousin (see Fig. 5.12), the option for PGT-HLA cycle with her cousin’s oocytes was offered. HLA testing of her cousin showed an exact HLA match and noncarrier of FANCA; thus, the further PGT-­ HLA attempts were performed with donor eggs from patient’s cousin. A total of 16 oocytes were retrieved, of which eight resulted in blastocyst and tested for mutation, HLA, and aneuploidy. The patient became pregnant after the second frozen transfer, delivering two healthy HLA-­

matched baby girls at 35 weeks, who as predicted were potential stem cell donors for both affected siblings in the family (Fig.  5.12). This is the world’s first case of PGT-HLA using donor eggs from relatives, which resulted in obtaining unaffected HLA-matched progeny for HLA-matched stem cells transplantation, using the PGT-HLA cycle involving a sibling as an HLA-matched egg donor. Patients should be fully aware of the limits of the expected successful outcome of the above testing. Preselection and transfer of the HLA-­ matched unaffected embryos is appropriate in only 13.7% of the embryos tested, even lower than may have been predicted. Despite such a relatively moderate success rate, the number of PGT-HLA requests has been increasing overall.

5.5

Practical Implications of PGT-HLA

PGT-HLA provides an important prospect of PGT application to stem cell therapy. Because of limited availability of the HLA-matched donors even among family members, this approach appears to be attractive for couples with children requiring HLA-matched bone marrow transplantation. It is well-known that to achieve an acceptable engraftment and survival in stem cell therapy requires the finding of an HLA-identical stem cell transplant [20]. However, there remain a large number of patients for whom no HLA-­ matched family member exists; thus, the search is extended to haplotype-matched unrelated donors. This has allowed successful application of stem cell transplantation to some individuals without a matched related donor, despite resulting in severe complications in more than half of the patients [28]. For example, the experience of bone marrow transplantation for hemoglobinopathies comprises currently thousands of patients, showing 68% probability of cure in the world’s largest center [7, 20]. Success rate is reported as 87%, 85%, and 80% for Class 1 (patients with regular iron chelation therapy who have neither hepatomegaly nor liver fibrosis), Class 2 (with regular/

5.5  Practical Implications of PGT-HLA

129 155 135 252 187 289 144 278

131 164 112 246 190 267 137 249

HLA

1.1 106 136 162 159 N 129

113 136 160 159 N 134

2.3

FANCA

207

137 155 131 244 199 252 139 261

133 161 102 248 202 289 133 254

HLA

1.2 117 133 162 159 DEL 129

106 143 160 157 N 132

1.3 110 104 136 139 164 164 159 159 DEL N 129 132

2.2

2.1

FANCA

FANCA

PGT

3.1 Affected MARKERS ORDER: D16S3028 D16S413 D16S3023 D16S3026 FANCA D16S3407

: HLA Markers ORDER D6S1568 RING D6S2443 D6S273 D6S62 RF D6S258 D6S248

A MATCH

3.2 Affected

3.3 Normal 106 143 160 157 N 132

113 136 160 159 N 134

106 143 160 157 N 132

113 136 160 159 N 134

137 155 131 244 199 252 139 261

137 155 131 244 199 252 139 261

137 155 131 244 199 252 139 261

131 164 112 246 190 267 137 249

137 155 131 244 199 252 139 261

131 164 112 246 190 267 137 249

NORMAL or CARRIE

HLA

3.4 Normal

117 110 133 136 162 164 159 159 DEL DEL 129 129

131 164 112 246 190 267 137 249

129 159 108 253 212 255 135 279

PGT

117 110 133 136 162 164 159 159 DELDEL 129 129

131 164 112 246 190 267 137 249

131 164 112 246 190 267 137 249

EUPOLID

FET

Baby

Patient - 4 cycles (40-41 yo)

2(14)

9

10

1

0

Donor 4 cycles (30-31 yo)

7(20)

20

15

4

2

Fig. 5.12  PGT-HLA with donor egg from a sibling. Upper panel shows family pedigree with HLA haplotype analysis based on parental (2.1 and 2.2), egg donor (2.3), and affected siblings (3.1 and 3.2) genomic DNA testing. HLA and FANCA marker order is presented on the upper left for the father and egg donor(ED) and upper right for the mother. Affected haplotypes harboring deletion 1115_18 in FANCA gene are printed in red. Normal haplotypes are colored in black. HLA matching maternal and egg donor’s alleles are depicted in pink, and matching paternal alleles are colored in blue. Non-matched paternal and maternal alleles are black. Egg donor’s non-match

haplotype is printed in green. Donor is normal for FANCA. Affected siblings genotypes are presented in left lower panel. HLA-matched normal twins delivered after PGD-HLA are shown in right lower panel. Results of PGT-HLA cycles performed in this case are summarized in Table (at the bottom of lower panel), showing the failure of PGT-HLA with patient’s own oocytes, while PGT-­ HLA with the donor eggs from her younger sister resulted in a twin matched pregnancy and birth of two children as potential donors for stem cell transplantation of their affected children with Fanconi anemia

irregular chelation, borderline hepatomegaly, and fibrosis), and Class 3 (with irregular chelation, hepatomegaly, and fibrosis) patients under age 17, respectively. This may have wider implication for congenital bone marrow failures, depending primarily on the availability of HLA-matched donors.

In fact, due to a small number of children per family, less than one-third of patients are able to find an HLA-identical sibling. This may be improved by 3% using an extended family search for a matched related donor with one or two identical ancestral haplotypes [29]. In the remaining patients, the only resort is the identification of a

208

5  Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)

matched unrelated donor, which might be maximized by establishing national registries. These registries overcome to some extent the genetic heterogeneity in the populations, which may mitigate against the low frequency of unique and thus improves donor selection process. Stem cell transplantation obtained from umbilical cord blood may also provide the potential for further expanding the donor pool to patients without a suitable family match [30]. This has been performed for thousands of children and adults, allowing for a greater degree of HLA disparity in choosing donor cord blood units, with the potential of increasing the likelihood that a suitable unit can be identified for a particular patient. However, the advantages of this alternative source of stem cells can be more fully realized in the setting of the availability of matched related donor cord blood units, such as from siblings, providing advantages of earlier transplantation, lower risk of complications, and lower treatment-related mortality. For conditions such as FA, HLA-identical sibling donors of stem cells are the only solution, allowing successful treatment in up to 85% of cases. This is the reason why, for over two decades, these families were offered the option of prenatal diagnosis combined with HLA typing on cells derived from chorionic villus sampling or amniocentesis [31]. Of more than 80 pregnancies conceived during this time, only 1 resulted in 1 successful use of cord blood transplant in 1988 [32], opening the era of an alternative to bone marrow transplantation. However, because the probability of having an unaffected child who may also be an HLA match for an affected sibling is very low (see Table 5.3), these families often went through multiple cycles of pregnancy before conceiving an unaffected HLA match. Thus, PGT-HLA provides the only reasonable approach, because a sufficient number of embryos may be tested at a time, increasing the chances to identify an appropriate match. In addition, PGT-HLA allows identification of the match before pregnancy, obviating risk for termination of pregnancy on the basis of HLA type alone. Presented data show that couples undergoing PGT-HLA may be expected to require a repeated

cycle to be able to preselect and transfer HLA-­ matched embryos. Even with the probability of selecting only one HLA-matched embryo from five tested, an acceptable pregnancy rate was observed despite transferring only one embryo on the average. Data provide a realistic option for couples desiring to establish a pregnancy potentially providing an HLA match progeny for the treatment of an affected family member(s). However, PGT-HLA also raises important ethical, legal, and social issues, which are discussed in detail in the corresponding section. Despite some remaining ethical issues, there is an increase in the attractiveness of PGT-HLA for couples with affected children requiring HLAcompatible stem cell transplantation, providing a practical option especially for those couples who wish to have another child. This and other new indications cited above make PGT a realistic alternative to conventional prenatal diagnosis. Couples are provided the important prospect not only to avoid an inherited risk without facing termination of pregnancy but also to establish that pregnancy with genetic parameters, which may benefit the affected member of the family. The present experience of PGT-HLA has resulted in birth of over a hundred unaffected HLA-matched children, whose HLA-identical stem cells have already been used for transplantation therapy in affected siblings [11–19, 33, 34]. The world’s largest two experiences include preimplantation HLA typing in approximately 1000 cases, with the accuracy rate per transfer of 99.4% [11, 14–16, 18, 19]. The majority of cases were performed in combination with PGT for various genetic disorders, including thalassemia, sickle cell disease, FA, WAS, X-ALD, HIGM1, HED-ID, Krabbe disease, inherited form of DBA, and X-linked chronic granulomatous disease (CGD), involving the preselection of unaffected children who were also HLA-identical to the affected sibling. The introduction of PGT-A as well expands this practical application of PGT-­ HLA to patients of advanced reproductive age, improving their chances to become pregnant and deliver an HLA-matched progeny for stem cell transplantation in the affected siblings. This also makes possible applying this approach to HLA-­

5.5  Practical Implications of PGT-HLA HLA Markers order: D6S1629 D6S1560 TAP1 D6S2447 D6S273 TNF-A MIB RF MOG D6S306

185 159 217 151 270 102 119 295 160 108

172 153 207 168 272 109 121 276 172 116

HLA

209

139 180 Cod39 118 163 117

187 159 207 170 274 115 110 252 182 104

143 197 N 114 163 117

1.1 Cod 39/ N

HBB

185 159 217 151 270 102 119 295 160 108

189 149 205 162 270 100 112 228 174 110

HLA

139 143 180 199 Cod39 IVS1-110 118 118 163 168 117 165

1.2

N / IVS1-110

2.1

2.2

Cod 39/ IVS1-110

N /N

HBB

DOB 1/21/2006

HBB Markers order : BSTR HBG Cod39 IVS1-110 BTG D11S1338 D11S1241

143 143 197 199 N IVS1-110 116 118 172 168 113 165

HLA 185 159 217 151 270 102 119 295 160 108

PGT

189 149 205 162 270 100 112 228 174 110

HBB 189 149 205 162 270 100 112 228 174 110

143 197 N 114 163 117

HLA

143 197 N 116 172 113

HBB

Sequential Polar Bodies Analysis Oocyte # 1 Predicted Genotype:

4

143 199 IVS1-110 118 168 165

HBB

AFFECTED

5

143 197 N 116 172 113

143 197 N 116 172 113

NORMAL

NORMAL

6

7

143 199 IVS1-110 118 168 165

143 197 N 116 172 113

AFFECTED

NORMAL

8

9

143 197 N 116 172 113

143 197 N 116 172 113

NORMAL

NORMAL

11

12

143 199 IVS1-110 118 168 165

143 199 IVS1-110 118 168 165

AFFECTED

AFFECTED

Blstomeres Analysis Embryo # 1 Predicted Genotype:

HBB

HLA

4

139 143 180 199 Cod39 IVS1-110 118 118 163 168 117 165 172 153 207 168 272 109 121 276 172 116

0

AFFECTED; monosomy6

5

143 197 N 114 163 117

143 197 N 116 172 113

143 197 N 114 163 117

185 159 217 151 270 102 119 295 160 108

189 149 205 162 270 100 112 228 174 110

172 153 207 168 272 109 121 276 172 116

NORMAL; MATCH

0

0

MONOSOMY 6 & 11

6

7

139 143 180 199 Cod39 IVS1-110 118 118 163 168 117 165 172 153 207 168 272 109 121 276 172 116

189 149 205 162 270 100 112 228 174 110

AFFECTED; NON-MATCH

ET

143 197 N 114 163 117

143 197 N 116 172 113

185 159 217 151 270 102 119 295 160 108

187 159 207 170 274 115 110 252 182 104

NORMAL; NON-MATCH

8

143 197 N 114 163 117 185 159 217 151 270 102 119 295 160 108

9

11

12

143 197 N 116 172 113

139 180 Cod39 118 163 117

143 197 N 116 172 113

143 143 197 199 N IVS1-110 114 118 163 168 117 165

143 197 N 114 163 117

0

189 149 205 162 270 100 112 228 174 110

172 153 207 168 272 109 121 276 172 116

189 149 205 162 270 100 112 228 174 110

185 159 217 151 270 102 119 295 160 108

185 159 217 151 270 102 119 295 160 108

0

NORMAL; MATCH

ET

CARRIER; NON-MATCH

189 149 205 162 270 100 112 228 174 110

CARRIER ; MATCH

MONOSOMY 6 & 11

FROZEN

Fig. 5.13  PGT-HLA in couple with adult thalassemic son, requiring HLA-matched bone marrow transplantation. Upper panel: family pedigree with HLA haplotype analysis based on parental (1.1 and 1.2) and affected child (2.1) genomic DNA testing. HLA marker order is presented on the upper left for father and right for mother. Paternal and maternal matching HLA haplotypes to the affected child (2.1) are shown in different colors. Maternal (IVS1–110) and paternal (Cod39) mutations and the linked markers are also presented accordingly. Medium panel: sequential PB1 and PB2 analysis for maternal mutation IVS1–110, with 5 linked markers in 9 oocytes, predicting 4 mutant and 5 normal oocytes, 2 of them originated from heterozygous PB1 and mutant PB2, represent-

ing good candidates for transfer. Lower panel: HLA typing by short tandem repeats (STRs) along with mutation analysis was performed on embryo biopsy samples from all nine embryos, two of which (#4 and #8) was predicted to be mutation-free and also HLA match to the affected sibling 2.1. Another matched carrier of maternal mutation (embryo # 11) was frozen. Both normal matched embryos were transferred, resulting in a singleton pregnancy and birth of thalassemia-free child matched to the affected siblings. The bone marrow of this child was transplanted to the thalassemic sibling (2.1), with a successful hematopoietic reconstitution and total cure of the thalassemic sibling

compatible stem cell transplantation not only for children but also for older affected siblings that was not expected to be realistic. The first such case has been done in our experience, resulting in successful stem cell engraftment with neither acute nor chronic GVHD in the 14-year-old sibling with thalassemia [35] (Fig. 5.13).

In conclusion, results demonstrate increasing attractiveness of PGT-HLA for couples with affected children requiring HLA-compatible stem cell transplantation. That no embryo is discarded based on the results of PGT-HLA is important as all unaffected embryos are frozen for future use. Thus, couples at risk of having

210

5  Preimplantation Genetic Testing (PGT) for Human Leukocyte Antigens (HLA) (PGT-HLA)

children with congenital bone marrow disorders will benefit from information provided to them about presently available options not only of avoiding the birth of an affected child but also of selecting a suitable stem cell donor for their affected siblings.

References 1. Verlinsky Y, Rechitsky S, Schoolcraft W, Strom C, Kuliev A. Designer babies-are they reality yet? Case report: simultaneous preimplantation genetic diagnosis for Fanconi anemia and HLA typing for cord blood transplantation. Reprod Biomed Online. 2000;1:31. 2. Verlinsky Y, Rechitsky S, Schoolcraft W, Strom C, Kuliev A.  Preimplantation diagnosis for Fanconi anemia combined with HLA matching. JAMA. 2001;285:3130–3. 3. Online Mendelian Inheritance in Man (OMIM). John Hopkins University. 2001. http://www.ncbi.nlm.nih. gov/Omim. 4. Online Human Gene Mutation Database (HGMD). CELERA. 2004. http://archive.uwcm.ac.uk/uwcm/ mg/search/1119297.html. 5. Modell B, Kuliev A. History of community genetics: the contribution of hemoglobin disorders. Community Genet. 1998;1:3–11. 6. Canatan D.  Hemoglobinopathy prevention program in Turkey. Thalassemia Rep. 2011. https://doi.org/10.481/thal.2011.s2.e4. 7. Lucarelli G, Andreani M, Angelucci E.  The cure of thalassemia by bone marrow transplantation. Blood. 2002;16:81–5. 8. Kuliev A, Rechitsky S, Verlinsky O, et  al. Preimplantation diagnosis of thalassemia. J Assist Reprod Genet. 1998;15:219–25. 9. Kuliev A, Rechitsky S, Verlinsky O, et  al. Birth of healthy children after preimplantation diagnosis of thalassemias. J Assist Reprod Genet. 1999;16:207–11. 10. Kanavakis E, Vrettou C, Palmer G, et  al. Preimplantation genetic diagnosis in 10 couples at risk for transmitting beta-thalassemia major: clinical experience including initiation of six singleton pregnancies. Prenat Diagn. 1999;19:1217–22. 11. Rechitsky S, Kuliev A, Tur-Kaspa I, Morris R, Verlinsky Y.  Preimplantation genetic diagnosis with HLA matching. Reprod Biomed Online. 2004;9:210–21. 12. Van de Velde H, Georgiou I, De Rycke M, et al. Novel universal approach for preimplantation genetic diagnosis of β-thalassemia in combination with HLA matching of embryos. Hum Reprod. 2004;19:700–8. 13. Kahraman S, Karlilaya G, Sertyel S, Karadayi H, Findicli N, Oncu N. Clinical aspects of preimplantation genetic diagnosis of single gene disorders com-

bined with HLA typing. Reprod Biomed Online. 2004;9:529–32. 14. Kuliev A, Rechitsky S, Verlinsky O, Tur-Kaspa I, Kalakoutis G, Angastiniotis M, Verlinsky Y.  Preimplantation diagnosis and HLA typing for hemoglobin disorders. Reprod Biomed Online. 2005;11:362–70. 15. Kuliev A, Rechitsky S. Preimplantation HLA typing for stem cell transplantation treatment of genetic and acquired bone marrow failures. Hemat Med Oncol. 2016;1(2):46–9. 16. Kuliev A.  Practical preimplantation genetic diagnosis. 2nd ed. New York, London, Heidelberg: Springer; 2013. 17. Kuliev A, Rechitsky S, Verlinsky O. Atlas of preimplantation genetic diagnosis. 3rd ed. London: CRS Press, Taylor and Francis; 2014. 18. Kahraman S. PGD for HLA: clinical outcomes of HLA compatible transplantation following PGD.  Reprod Biomed Online. 2013;26(Suppl 1):S9–10. 19. Umay KB, Gavaz M, Kumtepe ÇY, Yelke H, Pirkevi Çetinkaya C, Çetinkaya M, Kahraman S. Successful hematopoietic stem cell transplantation in 62 children from healthy siblings conceived from preimplantation HLA matching: a clinical experience of 327 cycles. Reprod Biomed Online. 2019;39(Suppl. 1):e13–4. 20. Gaziev J, Lucarelli G.  Stem cell transplantation for thalassaemia. Reprod Biom Online. 2005;10:111–5. 21. Rechitsky S, Pakhalchuk T, Prokhorovich M, San Ramos G, Verlinsky O, Kuliev A.  Preimplantation genetic testing for inherited immunodeficiency. Hematol Transfus Int J. 2018;6:218–20. 22. Hellani A, Lauge A, Ozand P, Jaroudi K, Coskun S.  Pregnancy after preimplantation genetic diagnosis for Ataxia Telangiectasia. Mol Hum Reprod. 2002;8:785–8. 23. Rechitsky S, Kuliev A, Leigh D, et al. Single molecule sequencing: a new approach for preimplantation testing and noninvasive prenatal diagnosis confirmation of fetal genotype. Molecular Diagn 2020;22:220–7. 24. Verlinsky Y, Rechitsky S, Sharapova T, Morris R, Tharanissi M, Kuliev A. Preimplantation HLA typing. JAMA. 2004;291:2079–85. 25. Tesarik J, Mendoza C. Somatic cell haploidization: an update. Reprod Biomed Online. 2003;6:60–5. 26. Rechitsky S, Pakhalchuk T, Goodman A, San-Ramos J, Zlatopolsky Z, Kuliev A.  First systematic experience of combined PGD for single gene disorders and/ or Preimplantation HLA typing with 24-chromosome aneuploidy testing. Fertil Steril. 2015;103(2): 503–12. 27. Rechitsky S, Kuliev A, Sharapova T, et  al. Preimplantation HLA typing with aneuploidy testing. Reprod Biomed Online. 2006;12:81–92. 28. Baker S, Wagner J.  Advantages of umbilical cord blood transplantation. Abstracts of international seminal on preimplantation HLA typing & stem cell transplantation, Limassol, Cyprus, 2004. p.  7–8. www. pgdis.org.

References 29. Costeas PA.  Bone marrow donor registry and availability of HLA matched donors. Abstracts of international seminal on preimplantation HLA typing & stem cell transplantation, Limassol, Cyprus, 2004. p. 9. www.pgdis.org. 30. WHO News and activities. Cord blood banking. Bull World Health Organ. 1998;76:313–4. 31. Auerbach AD.  Preimplantation genetic diagnosis combined with HLA typing: the Fanconi anemia experience as a model. Abstracts of international seminal on preimplantation HLA typing & stem cell transplantation, Limassol, Cyprus, 2004. p. 13. www. pgdis.org. 32. Gluckman E, Devergie A, Schaison G, et  al. Bone marrow transplantation in Fanconi anemia. Br J Haematol. 1980;45:557–64. 33. Goussetis E, Kokkali G, Petrakou E, et al. Successful hematopoietic stem cell transplantation in 2 children

211 with X-linked chronic granulomatous disease from their unaffected HLA-identical siblings selected using preimplantation genetic diagnosis combined with HLA typing. Successful hematopoietic stem cell transplantation in 2 children with X-linked chronic granulomatous disease from their unaffected HLA-­ identical siblings selected using preimplantation genetic diagnosis combined with HLA typing. Biol Blood Marrow Transplant. 2010;16:344–9. 34. Kakourou G. PGD for HLA (ESHRE study). Reprod Biomed Online. 2018;36(Supplement 1):e4–5. 35. Kuliev A, Packalchuk T, Verlinsky O, Rechitsky S. Preimplantation diagnosis: efficient tool for human leukocyte antigen matched bone marrow transplantation for thalassemia. Thalassemia Rep. 2011;1:e1.. https://doi.org/10.4081/thal.2011.e1

6

Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic Testing for Chromosomal Disorders (PGT-A and PGT-SR)

It is well known that chromosomal abnormalities originate predominantly from female meiosis. As demonstrated by DNA polymorphism studies performed in families with aneuploid spontaneous abortions or liveborn babies with autosomal trisomy syndromes, these abnormalities derive mainly from meiosis I [1–3]. It is generally considered that the age-related increase of trisomies is probably determined by the age-related reduction of meiotic recombination, resulting in premature separation of bivalents and chromosomal nondisjunction; however, other mechanisms are also involved. Meiosis II errors were also postulated to derive from meiosis I, as a result of the increased meiotic recombination rate, which may lead to a separation failure of bivalents [4]. In this chapter we will focus on the analysis  of chromosome error dynamics from polar bodies to blastocyst, providing  the unique information on biological and clinical significance of the detected chromosomal instability, with special emphasis on mosaicism and sub-chromosomal variations. These contribute to strategies that must be developed for PGT-A  and PGT-SR, which is covered in clinical detail in Chap. 7.

6.1

Meiotic and Mitotic Abnormalities

One of the approaches to preimplantation genetic testing for aneuploidy (PGT-A) is direct testing of the outcome of the first and second meiotic © Springer Nature Switzerland AG 2020 A. Kuliev et al., Practical Preimplantation Genetic Testing, https://doi.org/10.1007/978-3-030-43157-0_6

divisions, using the first and second polar body (PB1; PB2): PB1 is extruded following maturation of oocytes and represents a by-product of meiosis I, while PB2 is a by-product of meiosis II and is extruded following the exposure of oocytes to sperm or ICSI.  The frequency and types of chromosomal errors detected by this approach are different from what was described in the traditional studies of meiotic chromosomes in metaphase II oocytes, according to which chromosomal anomalies in oocytes originate mainly from the errors of whole bivalents as a result of chromosomal nondisjunction [5]. In contrast, direct testing of meiotic outcome showed not only a higher prevalence of meiotic errors but also significant contribution of chromatid, rather than chromosomal, errors. The discrepancy may be due to the poor quality of meiotic chromosome preparations in earlier studies and also the lack of testing of the corresponding chromosome sets extruded in PB1, without which the resulting oocyte karyotype could not be reliably evaluated, particularly in the cases of missing chromosomes or chromatids. This was demonstrated in the study of simultaneous testing of metaphase II oocytes with their corresponding PB1, which showed that the normal chromosome pattern is represented by paired chromatids for each chromosome; an addition of one or both chromatids in either oocyte or PB1 reflects an exactly opposite pattern in the corresponding metaphase II oocytes or PB1, suggesting a high accuracy of the oocyte genotype 213

6  Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic…

prediction [6–10]. This is of practical relevance in preselection of aneuploidy-free oocytes in IVF patients of advanced reproductive age [11–17]. As polar bodies are extruded as a normal process, their removal is the least invasive approach for preselection of zygotes with the higher developmental potential in the efforts to improve the efficiency of assisted reproduction technology (ART), traditionally based on selection of embryos by morphological parameters which are of a limited value for this purpose [18–22]. Of limited value were also a pronuclear morphology scoring and microtubule and microfilament organization [23–25], as well as PB1 grading [26, 27]. The present standard methodology for PGT-A is based on next-generation sequencing (NGS) coupled with blastocyst biopsy procedures, described in Chaps. 2 and 3. Table 6.1 shows that, approximately, half of aneuploidies originate from the meiosis errors (9,812 of 20,986 oocytes tested), of which almost a third (2,921) are from both meiosis I and meiosis II, close to one-third (2,983) from only meiosis I and over another one-third (3,908) from only meiosis II (Table  6.2). The frequency of aneuploidies of meiosis origin increases with increasing maternal age, from up to 20% in patients of 35 years of age to over 40% in patients older than 40 (Fig. 6.1). This is in agreement with a study of 684 cycles from infertility patients that showed 55% aneuploidy rate deriving from meiosis errors [28, 29]. The meiosis I errors were 39% in the younger than 38-year-old patients and 58% in 44-year-old patients [30]. It is not clear to what extent the Table 6.1  Aneuploidies of meiotic origin Oocytes Couples Cycles with results Euploid 2,830 3,953 20,986 11,174 (53.2%)

Aneuploid 9,812 (46.8%)

Table 6.2  Types of meiotic aneuploidies Types MI + MII MI MII Total Aneuploid

Number 2,921 2,983 3,908 9,812

% 29.8 30.4 39.8 100

reported meiotic aneuploidies are related to IVF treatments involving aggressive hormonal stimulation (see Chap. 7); preliminary data on testing of donated oocytes from young fertile women suggest that the actual prevalence may be much lower, although more data are needed [31]. Table 6.2 shows comparable proportions of detectable aneuploidies originating from meiosis I (31%) and meiosis II (34%), in contrast to the well-established concept of female meiosis I origin of the majority of aneuploidies [2, 32]. As mentioned, one-third of the chromosomally abnormal oocytes originates from sequential meiosis I and meiosis II errors, suggesting that these meiosis II errors may be associated with the preceding meiosis I errors (Tables 6.2 and 6.3). This is in agreement with the concept of a possible relationship of meiosis II errors with the increased meiotic recombination rate [4]. 100

% abnormal oocytes

214

80

c

60

b 2,108

40

a 20

1,715 1,853

2,101

1,955

2,500

2,721 2,217

1,121

0 35

36

37

38

39 Age

40

41

42

43+

Fig. 6.1  Aneuploid oocytes in relation to maternal age. A number of oocytes tested for each age group are shown under the curve evidencing the increase of the overall frequency from 20% in the age group of 35 to close to 60% in the age group of 43 and over Table 6.3  Frequency and types of meiosis I and meiosis II errors Origin of Aneuploidies Euploid Aneuploid  Disomy  Nullisomy  Complex Total abnormal Total

Meiosis I No. 13,097

% 69.0

Meiosis II No. % 13,635 66.0

1,514 3,136 1,271 5,921 19,018

26.0 53.0 21.0 31.0 100

2,721 39.0 2,875 41.0 1,342 20.0 6,938 34.0 20,573 100

6.1  Meiotic and Mitotic Abnormalities

215 50 40 % abnormal oocytes

However, half of meiosis II errors are still observed independent from meiosis I, emphasizing clinical significance of testing the outcome of both meiotic divisions. This is also obvious from the age dependence of isolated errors of meiosis I and meiosis II, as well as sequential meiosis I and meiosis II errors (Fig. 6.2). This is even stronger than that for isolated meiosis I and meiosis II errors, more than doubled in 40-year-old patients compared to 35-years-old patients. Schematic representation of the types of aneuploidies originating in MI and MII are presented in Fig.  6.3, showing at least two times higher frequency of nullisomy compared to disomy ­following MI (approximately 2:1 ratio). This is in contrast to a comparable distribution of nullisomy/disomy following MII (Fig.  6.4). The other important phenomenon is that there is a significant predominance of chromatid errors over chromosomal ones (72.3% chromatid

30 20 10 0

406

35

379

433

36

517

37

MI

a

568

39 Age

353

40

MII

512

41

337 n

42

43+

MI & MII

Fig. 6.2 Relationship of different meiotic errors to maternal age, based on the analysis of 822 cycles. Upper curve includes oocytes with M1 errors irrespective of having or not having sequential abnormality in MII. Middle curve includes errors originating in MI with sequential meiosis II errors, which do not include isolated MI errors. Lower curve includes errors originating only in MII

Abnormal 31.1%

5.2%

382

38

Normal 68.9%

1.1% b

Fig. 6.3  Chromosome (chromatid) segregation errors in Meiosis I. Upper panel, center: Primary oocyte containing diploid set of chromosomes with the doubled amount of chromatin (4n) prior to maturation. Upper panel, right: Normal segregation of homologues in the first meiotic division, resulting in the extrusion of the first polar body (PB1) (smaller circle) containing one of the homologues. Accordingly, the resulting secondary (metaphase II) oocyte contains the remaining homologue with two chromatids. Upper panel, left: Meiotic errors leading to the extrusion of PB1 containing abnormal set of chromosomes. Lower panel, a: Chromosomal nondisjunction, leading to segregation of both homologues to MII oocyte, so that the extruded PB1 will not contain any material,

25.5%

46.7%

c

d

COMPLE 21.5%

e

resulting in a disomic oocyte. Lower panel, b: Chromosomal nondisjunction, leading to segregation of both homologues to PB1 (smaller circle), which will result in a nullisomic oocyte. Lower panel, c: Chromatid malsegregation, leading to an extra chromatid extrusion with PB1, which results in the lack of one chromatid in MII oocyte. Lower panel, d: Chromatid malsegregation, leading to a single chromatid extrusion with PB1, which results in the extra chromatid material in MII oocyte. Lower panel, e: Chromatid or chromosome malsegregation involving different chromosomes, resulting in complex errors, involving different types of errors of different chromatids or chromosomes in MII oocyte

6  Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic…

216

error rate, 46.7% missing and 25.5 extra chromatids, compared to 6.3% chromosome errors, 5.2% missing and 1.1% extra chromosomes) (Fig. 6.3). Therefore, similar to chromatid errors, missing chromosomes were more frequent than extra chromosomes, which is in agreement with a higher frequency of trisomies over monosomies in postimplantation embryos, detected in spontaneous abortions. As seen from Fig.  6.3, only 25.5% of meiosis I aneuploidies were of disomic type, compared to 46.7% of nullisomic type. Although the observed excess of nullisomies following MI may be attributable to technical errors, such as hybridization failure, it is also possible that a meiosis I mechanism exists that prevents extra chromosome material extrusion if

Abnormal 33.7%

66.3%

meiotic errors occur during oocyte maturation. In support of the biological nature of this phenomenon could be the age dependence of missing chromatids (Fig. 6.5) and missing chromosomes (Fig.  6.6), suggesting that the observed anomalies may be due to the overall disturbances of the meiosis process with advanced reproductive age. Although overestimate cannot be excluded, the true nature of the majority of MI and MII aneuploidies is confirmed by follow-up studies of the embryos resulting from these oocytes [12]. In contrast to the well-established concept of female meiosis I origin of chromosomal abnormalities, the above results show that the observed errors 80 75 70 65 60 55 50 45 40 35 30

b a n = 631

708

684

603

244

35–36

37–38

39–40

41–42

43+

% missing chromatids

a

b

c 39%

41%

20%

Fig. 6.4  Meiosis II errors, based on the PB2 analysis. Upper panel (center): Secondary (metaphase II) oocyte containing haploid set of chromosomes (2n) prior to fertilization. Upper panel (right): Normal segregation of chromatids in the second meiotic division (66.3%), resulting in the extrusion of the second polar body (PB2) (smaller circle) containing one of the chromatids. Accordingly, the resulting maternal contribution to zygote contains the remaining sister chromatid. Upper panel (left): Abnormal segregation of chromatids in the second meiotic division (33.7%), involving the abnormal segregation of chromatids, showed in the lower panel. Low panel (a): Chromatid nondisjunction leading to the extrusion of PB containing no chromatid material, so both chromatids will be left in oocyte, resulting in disomic oocyte (41%). Low panel (b) similarly, chromatid nondisjunction leading to the extrusion of PB containing both chromatids, which results in nullisomy of this chromosome in maternal pronucleus (39%). Low panel (c): Complex chromatid malsegregation, leading to different errors of different chromatids (20%)

Fig. 6.5  Prevalence of missing chromatids following MI in relation to maternal age. Numbers of oocytes tested for each age groups are shown under curve, evidencing the increase of the prevalence from 45% in the age group of 35–70% in the age group of 43 years and older 10 8

b

6 4

a

2 0

n = 631

708

684

603

244

35–36

37–38

39–40

41–42

43+

% missing chromosomes

Fig. 6.6  Prevalence of missing chromosomes following MI in relation to maternal age. Numbers of oocytes tested for each age groups are shown under curve, evidencing the increase of the prevalence from 4% in the age group of 35, to 8% in the age group of 43 years and older

6.1  Meiotic and Mitotic Abnormalities

originate from both meiosis I and II, as seen from the patterns of segregation illustrated in Figs. 6.3 and 6.4. The results are of clinical significance, suggesting that the genotype of the resulting zygote cannot be predicted without testing of the outcomes of both meiotic divisions. While testing of meiosis I errors alone could only reduce aneuploidy rates in the resulting embryos by two-­ thirds, over one-third of these oocytes will be still aneuploid following the second meiotic division. However, testing for only MI errors could still improve the implantation and pregnancy rates in poor-prognosis IVF or ICSI patients, by applying ICSI selectively to the oocytes with aneuploidy-­ free PB1. On the other hand, only close to half of the abnormalities deriving from the second meiotic division may be detected following MI. As seen from Fig. 6.3, the majority of aneuploidies originating from meiosis I are represented by chromatid errors, in contrast to the expected chromosomal nondisjunction, as suggested by previous traditional studies cited. However, chromosomal errors are still observed in 6.3% of oocytes, thus the abnormalities in MII oocytes are not solely of chromatid origin [34–40]. Although both chromatid and chromosomal errors are involved in producing metaphase II oocyte abnormalities, the chromatid/chromosome error ratio is as high as 10:1. There is little doubt that both of these meiosis I errors lead to aneuploidy in the resulting embryos, as demonstrated by the follow-up study of the embryos resulting from these oocytes [7, 11, 17]. However, differences in the effect of chromatid and chromosomal errors on the pre- and postimplantation development cannot be excluded. As mentioned, in contrast to meiosis I errors, there is no difference in the frequency of missing or extra chromatid errors following the second meiotic division (Fig.  6.4). Overall, 6,938 (33.7%) of 20,573 oocytes tested had meiosis II errors, of which 39% had an extra chromatid, 41% a missing chromatid, and 20% complex errors [17]. While at least 95% of aneuploidies originate from maternal meiosis, the remaining aneuploidies originate from paternal meiosis and mitotic errors. Standard PGT-A methodology is currently based on NGS following blastocyst biopsy. This is also

217

more practical than PB analysis for detection of all aneuploidies, because not all oocytes reach the embryo transfer stage. In addition, selection occurs against a significant proportion of aneuploidies, which may not survive to blastocyst [42]. On the other hand, the comparison of the spectrum of aneuploidies in oocytes and embryos shows some inconsistency between the expected and observed frequency of some types of aneuploidies in embryos as predicted by meiosis testing. For example, the predicted lower rate of monosomies is confirmed by lack of autosomal monosomies in spontaneous abortions, due to incompatibility with postimplantation development except for monosomy 21; thus, the predicted embryo trisomy predominance is not in agreement with data on the embryo chromosomal profile. In our current practice of PGT-A, based on the NGS analysis of 2922 blastocysts, 56.0% of embryos were aneuploid, comprised of 13.0% monosomy, 13.0% trisomy, 8.0% numerical mosaic, 14.0% segmental mosaic, and 8.0% complex errors (Fig. 6.7). It is of interest that no age dependence was revealed for monosomies in embryos (Fig. 6.8), suggesting that the rate of monosomies detected in embryo by PGT-A may be of artifactual nature. A possible explanation for this discordance is that the majority of monosomies detected in embryos are derived from mitotic errors, assuming technical causes are excluded. In fact, a significant proportion of the cleavage-stage monosomies appeared to be euploid after their reanalysis [43, 44]. Embryo monosomies that are not detected after implantation are either eliminated before implantation or have no biological significance, reflecting the poor viability of the monosomic embryos and their degenerative changes. However, the majority of pre-­zygotically derived monosomies, as well as some of post-­zygotic origin, may still survive until the b­ lastocyst stage, therefore leading to implantation failure or pregnancy loss. In one relevant study, the progression and survival of different types of chromosome abnormalities were followed up in 2,204 fertilized oocytes [45]. A variety of chromosome abnormalities was detected, including many types of errors not recorded later in development. However, these appeared to be tolerated until activation of the embryonic genome, after which there

6  Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic…

218

Fig. 6.7 Inconsistency of predicted and observed aneuploidies in oocytes and embryos. (Upper figure) Distribution of different types of aneuploidies predicted by oocyte testing suggested predominance of embryo trisomies (green sector in the circle). (Lower figure) Distribution of different types of aneuploidies by embryo testing, showing similar rates of monosomies (red sector in the circle) and trisomies (orange sector in the circle), opposite to prediction by oocyte testing

MI chromosomal abnormalities

Missing chromatid

21.8%

Extra chromatid Missing chromosome Extra chromosome

1.0%

Complex

4.5%

48.7%

25.0%

Reproductive Genetics Institute

Mosaic Complex Trisomy

Euploid

8%

Av. Age 36.7

%abnormal oocyte

443

15

a

b 482

352

410

c

368

10 5 0 35–36

37–38

39–40 Age

13%

* 2% Failed Amplification

30

20

44%

13% Monosomy

25

N = 14,922*

Segmental (full and mosaic) 14% 8%

41–42

43+

Fig. 6.8  Prevalence of monosomies at the cleavage stage in relation to maternal age. Numbers of embryos with monosomies for each age group are shown under curve, evidencing the lack of age dependence

were declines in frequency. However, many aneuploid embryos still successfully reach the blastocyst stage, even if some chromosome errors present during ­preimplantation development are not seen in later pregnancy. Of special interest is the group of complex aneuploidies, detected in one-fifth of abnormalities following meiosis I and II (21% and 20%, respectively), as well as in embryos (8%). The abnormalities are represented by different types of errors, involving more than one chromosome, or errors in both MI and MII of the same or different chromosomes that may result in balanced (normal) embryos. This is in agreement with other reported data [29, 42] and

6.1  Meiotic and Mitotic Abnormalities

may represent the phenomenon of aneuploidy rescue, similar to the well-known trisomy rescue mechanism. The resulting balanced chromosome set of the oocyte after complementary errors in meiosis I and meiosis II is shown in Fig. 6.9. The mechanism underlying formation of such balanced zygotes is not yet understood. The fate of the embryos resulting from such balanced oocytes is also not clear but may also result into abnormal (mosaic) status, uniparental disomy, and imprinting ­disorders. The observed aneuploidy rescue mecha-

219

nism in female meiosis cannot ensure the chromosomal normalcy of the resulting embryos and, hence, is not assurance of embryo transfer. The fact that the meiotic error of one chromosome may affect the segregation of other chromosomes, resulting in complex error, has long been described in 39,X female mice [41]. This was also observed in human embryo follow-up from meiosis I errors through meiosis II and cleavage (see below). Overall, the high prevalence of complex errors may indicate generalized disturbances in the meio-

a

b

c

Fig. 6.9  Array CGH analysis of PB1, PB2, and resulting oocyte showing abnormal chromosome sets in PB1 and PB2, resulting in a normal chromosome status in the resulting oocyte (performed in collaboration with Antony Gordon, Bluegnome, Cambridge, UK). (a) PB1 analysis shows extra chromatids 2, 7, 8, 14, 19, and 21 and missing

chromosomes 3, 6, and 17. (b) PB2 analysis shows extra chromatids, 3, 6, and 17 and missing chromatids 2, 7, 8, 14, 19, and 21. (c) Normal set of the resulting oocyte, which suggests the complete balancing of the karyotype in the resulting oocyte

220

6  Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic…

sis process, which may be due to the age-related effect on the recombination frequency, spindle formation errors that are also reported to increase with age, loss of chromosome cohesion, and mitochondrial and organelle dysfunction [4, 32, 33, 46–50]. However, the actual mechanism underlying correlation with maternal age is still not understood. The most recent data show a strong correlation between female aging and premature resolution of centromeric cohesion, which is characterized by depletion of the meiosis-specific alpha-kleisin subunit Rec8 from oocyte chromosomes [51]. Together with the involvement of other meiosisspecific proteins, this event may explain the prevalence of premature loss of centromeric cohesin in oocytes of older females. Analysis of the timing and mechanisms of cohesin depletion during female aging show that cohesin is gradually depleted from oocyte chromosomes during the prolonged arrest at prophase of meiosis I, long before oocytes are recruited for growth.

6.2

Chromosome-Specific Meiotic Error Origin and Its Impact on Embryo Viability

Analysis of chromosome-specific patterns has shown that chromosomes 15, 16, 21, and 22 are much more frequently involved in female meioTrisomy

sis errors than other 20 chromosomes, representing over one-third of all oocyte aneuploidies. This is followed by chromosomes 19 and 20, with errors of other 18 chromosomes being of lower frequency [52]. These data are in agreement with our data obtained in PGT-A at the blastocyst stage (Fig.  6.10) (examples of trisomies and monosomies detected by NGS are presented in Figs.  6.11 and 6.12). It was also previously demonstrated that despite differences in chromosome-­ specific aneuploidy rates, age dependence was observed for each of these chromosome errors, almost doubling between the age 35 and 43 years (Fig.  6.13), again suggesting overall disturbance of the meiosis process with advanced reproductive age [32]. Chromosome-specific origin of errors was also not similar: chromosomes 16 and 22 errors originated more frequently in meiosis II (44.4% and 41.5% meiosis II errors vs 32.0% and 34.3% meiosis I errors, respectively). Chromosome 13, 18, and 21 errors were more frequently from meiosis I (40.1%, 48.3%, and 41.4% in meiosis I vs 36.3%, 34.6%, and 36.7% in meiosis II, respectively). It is of note that the proportion of oocytes with errors of both meiosis I and meiosis II origin were not significantly different for errors of different chromosomes, except for chromosome 18 errors (Table 6.4). These data are opposite to the chromosome-­ specific meiosis origin observed in spontaneous

Mono

40

22

35

16

21

30 25 15

20

13

19

15 10 5

N = 14,922

0 1

3

5

7

9

11

13

15

17

19

21

X

Fig. 6.10  Distribution of trisomies and monosomies in our series of 14,922 blastocysts testing by NGS (see description in the text)

6.2  Chromosome-Specific Meiotic Error Origin and Its Impact on Embryo Viability

Fig. 6.11  Trisomies detected in our PGT-A practice by NGS.  Trisomic chromosomes are circled in red, with examples of rare trisomies (such as trisomies 1, 3, 6, and

221

7) on the left and most frequent ones, such as trisomies 15, 16, and 21 on the right (see also Fig. 6.10)

Fig. 6.12  Monosomies detected in our series of PGT-A by NGS Monosomies are circled in red

6  Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic…

222

abortions and liveborn children [32, 46] and may indicate poor viability of embryos resulting from the oocytes with the chromosome 16 and 22 errors of the second meiotic division, possibly incompatible with implantation and postimplantation development. At present, there is no explanation for the possible biological differences of aneuploidies depending on the meiotic origin, except for loss of heterozygosity or higher homozygosity of the embryos originating from meiosis II errors for the genes located on these chromosomes. This may lead to imprinting of paternal or maternal genes on chromosome 16 or 22. Although established imprinting genes are not known in these chromosomes, case reports exist

%abnormal oocyte

20

22 21

15

13 18 16

10

5 1,264

0 35–36

1,532

1,771

1,451

37–38

39–40

41–42

728

43+

Age

Fig. 6.13  Frequency of each chromosome-specific error in relation to maternal age. Numbers of oocytes tested for each chromosome are shown under the curves, all of which shows an increase with age, particularly high for chromosomes 21 and 22 aneuploidies

of possible imprinting on chromosome 16 affecting fetal development or associated with cancer [53–56]. The other discrepancy is related to the meiotic origin of chromosome 18 errors, which predominantly originate from meiosis I (Table  6.4), opposite to that in liveborn children [57]. Whatever may be the explanation for the above phenomena, these data provide evidence for viability differences depending upon not only the chromosome involved but also the meiotic origin of the error.

6.3

 itotic Errors in Embryos M in Relation to Meiosis Errors

As shown above, approximately half of meiosis II errors are observed in the oocytes with prior errors in meiosis I. As a result of such sequential errors, one-third of the resulting zygotes may have been considered normal (euplolid), provided that the preceding errors in meiosis I and meiosis II have no effect on the further preimplantation developments of the corresponding embryos. Based on this observation, it could have been postulated that a pooled testing of both polar bodies could be acceptable, as it will show the normal results in the cases of reciprocal errors in PB1 and PB2 or show an abnormality if any error occurs in the first or second meiotic divisions. However, follow-up testing of these embryos showed that only 18% embryos, deriving from the apparently balanced zygotes, were euploid for all the chromosomes analyzed; the

Table 6.4  Origin of chromosome 13, 16, 18, 21, and 22 aneuploidies (based on information from both meiosis I and meiosis II errors in testing 8,602 oocytes) Chromosome 13 16

Total abnormal (%) 1,086 (12.6) 1,531 (17.8)

18

1,098(12.8)

21

2,151(25.0)

22

2,736(31.8)

Meiosis I origin 436 (40.1)a 490 (32.0) P = 0.000 530 (48.3) P = 0.000 891 (41.4) NS 939 (34.3) P = 0.001

Meiosis II origin 394 (36.3)a 679 (44.4) P = 0.000 380 (34.6) NS 790 (36.7) NS 1,135(41.5) P = 0.003

Chi-square analysis, comparison to a statistically significant P value 1)

32.2% 27.1%

8% Duplication 30%

N = 14,922

blastocysts suggested a possible difference in specific chromosome involvement when comparing whole chromosome and segmental mosaicisms [74]. With the overall mosaicism rate 17.5% (a 2.46% mean rate of mosaicism per chromosome), trisomy was more frequently detected as whole chromosome mosaicism, whereas monosomy was more frequently seen in segmental mosaicism. Aneuploidy and mosaicism displayed different patterns of distribution in various chromosomes. In a recent PGDIS position statement, based on the analysis of the follow-up of, approximately, 500 transferred mosaic embryos, none appeared to lead to live births with detected neonatal abnormalities [75]. However, the implantation ability of a mosaic embryo depended not on the specific chromosome involved but the proportion of euploid cells present. If the vast majority of cells are chromosomally normal, these will dominate over aneuploid cells, thus facilitating

Deletion 62%

Mosaic 20- Mosaic 5050% 90%

Full

ability to implant and lead to a chromosomally normal live birth. The converse would apply if the vast majority of cells are aneuploid. The latter mosaic embryos either fail to implant or are ­destined to be lost early in pregnancy. Thus, low-­ level mosaic embryos have a significant potential to reach term and could be considered for transfer in the absence of euploid embryos (see Fig. 3.8 in Chap. 3). It may be suggested that the majority of low-level mosaics may have actually been a false positive, as has recently been demonstrated in the extended in  vitro culture of mosaic embryos to the day 12, 71% of which appeared to be normal both in trophectoderm and inner cell mass [76]. Figures 6.16 and 6.17 present the prevalence and distribution of sub-chromosomal variations in our experience of NGS-based PGT-A cases. The study involved NGS testing of 29,376 trophectoderm samples and included the analysis of deletions and duplications in all 24 chromosomes as

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6  Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic…

Fig. 6.19  Segmental abnormalities detected in our PGT-A practice by NGS (segmental anomalies are circles in red). The size of sub-chromosomal variations are shown in blue

small as 3 MB and as large as 139 MB (Figs. 6.18 and 6.19). Of 56.9% aneuploid embryos, 14.4% had deletions and duplications, 62.0% deletion, 30.0% duplication, and 8.0% multiple sub-chromosomal variants (examples of embryos with segmental abnormalities are demonstrated on Fig. 6.18). Mosaicism was detected in 59.3% of the variations (examples of embryos with mosaic segmental variations are shown in Fig. 6.19). As expected, larger chromosomes contained a higher proportion of deletions and duplications, with the exception of chromosome 6. The majority of duplications were observed in chromosomes 20 and Y; deletions were mainly in chromosomes 2, 12, 17, and 22. Segmental abnormalities represented de novo findings of mitotic origin, showing no maternal age dependence. Reproducibility in the follow-up testing in pregnancy losses will be useful for consideration on their biological and clinical significance. Much research will still be needed to understand whether mosaicism is constitutional or secondary to technical and biological artifacts. This was addressed in

a recent study of embryos donated for research that focused on retesting of embryos with segmental variations of different biopsy sites among 55 blastocysts or in inner cell mass (ICM) in additional 19 embryos with segmental aneuploidies [77]. The study showed that contrary to whole chromosome aneuploidies which were uniformly confirmed, segmental aneuploidies were not present in different blastocyst sections in a significant proportion of cases (46.03%), nor in the ICM (47.37%), suggesting their limited predictive value in PGT-A cycles. Thus, mosaicism restricted to segmental variations suggests potential viability of these embryos and may warrant transfer in the absence of euploid embryos. While the vast majority of chromosome abnormalities, if undetected, will cause implantation failure or miscarriage in the early first trimester, only a very small proportion will persist during pregnancy and as clinical chromosomal syndromes in the newborn. Comparing frequency of chromosome abnormalities in blastocysts and miscarriage samples based on high-resolution copy number varia-

6.7  Possible Additional Tests to Preselect Euploid Embryos with Higher Potential for Pregnancy

tion sequencing (CNV-Seq) is useful in understanding the clinical significance of mosaicism and segmentals in preimplantation development [78]. Retrospective analysis of clinical samples comprising 1454 blastocysts from PGT-A cycles and 1810 miscarriages detected 711 euploid (49%) and 743 aneuploid (51%) embryos. This included trisomies/monosomies (26%), segmental imbalances (44%), mosaics (9%), and complex aneuploidies (21%) comprising trisomies/monosomies with additional segmentals or mosaics. By comparison, CNV-Seq analysis of 1810 miscarriage samples identified 881 (48.7%) euploid and 929 (51.3%) aneuploid fetuses. Fetal aneuploidies detected included trisomies (47.5%), 45, X (9.9%), segmental imbalances (8.5%), mosaicism (13%), polyploidy (9%), and complex aneuploidies (12%). Comparing these two data sets showed that the vast majority of embryo segmental imbalances were nonpathogenic, whereas the vast majority of segmental imbalances in fetal POCs were associated with genes important for normal fetal development. By extrapolating from the type and frequency of the different chromosomal aneuploidies detected in embryos and in miscarriage POCs, findings suggested that a significant proportion of embryos with segmental aneuploidies or high-level mosaicism are lost in the early first trimester of fetal development. This suggests that PGT-A using a higher-resolution methodology may provide an improvement in reproductive outcomes by reducing the incidence of unexplained miscarriages following the transfer of embryos that were identified as “euploid” by earlier lower-resolution methods. In the other study, chromosomal copy number data was obtained from oocytes and embryos of 635 IVF patients from PGT-A program, with a total of 3541 samples, including 452 oocytes, 1762 cleavage-stage embryos, and 1327 blastocyst stage embryos, all tested by aCGH [79]. Segmental abnormalities, involving loss or gain of chromosomal fragments, were found in 10.4% in oocytes, 24.3% at the cleavage stage, and 15.6% in blastocyst. While some segmental errors were clearly of meiotic origin, most appeared to arise during the first few mitoses following fertilization, with decrease of their prevalence at the blastocyst stage. This suggested elimination of some of them before the blastocyst stage. It was also shown that varia-

229

tions were not entirely random but tended to occur within distinct chromosomal regions, identified as hot spots corresponding to known fragile sites. Although the origin of sub-chromosomal duplications and deletions remain unclear, it is possible that they originate in the first mitotic divisions following fertilization due to relaxed cell cycle control. This permitted DNA double-strand breaks to persist through cell division. This is also in agreement with finding of the paternal origin in more than half of segmental variants [73], with paternal chromosomes known to be characterized by delayed DNA replication.

6.7

Possible Additional Tests to Preselect Euploid Embryos with Higher Potential for Pregnancy

6.7.1 Mitochondrial DNA It has been proposed that evaluating the mtDNA copy number has a predictive value for success of euploid embryo transfer: the lower the mtDNA copy number, the better the outcome [80]. This mitoscore was examined in 205 blastomeres obtained from day 3 euploid embryos and in 65 blastocyst biopsies in patients undergoing single euploid embryo transfer, with known implantation and pregnancy outcomes. An increased amount of mtDNA in euploid embryos was believed related to poor implantation potential. Speculation exists that an embryo with a high mitoscore was responding to “crisis,” perhaps undergoing reparative damage. This was consistent with results in another report, which was based on 340 blastocysts and 39 cleavage-stage embryos [81]. A significant increase in embryo mtDNA quantity was also observed with advancing female age and in aneuploid embryos, whereas euploid embryos capable of ­implantation tended to contain lower mtDNA quantities than those failing to implant. The mtDNA content threshold established during these studies could have represented a novel biomarker, independent of embryo morphology and aneuploidy and capable of identifying euploid blastocysts that are not able to lead to clinical pregnancies after transfer. However, there are alternative

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6  Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic…

explanations. In a preimplantation embryo, its entire quantity of mtDNA is gained during oocyte maturation, after which the level of mtDNA is reduced with each successive cell division. According to the frequently evoked “bottleneck theory,” not all cells will have the same amount of DNA after a given division. Indeed, the authors themselves observed a wide range of mtDNA content in the biopsied materials, confirming unequal distribution of mtDNA during preimplantation development. Thus, the mtDNA copy number in a biopsied cell is not necessarily representative of the copy number of all cells in its corresponding embryo. Actually, criticism could be raised concerning the methodology used in the above studies. Introduction of a correction factor showed no correlation between DNA content and implantation [82]. To investigate the factors underlying such differences in the relationship of high mtDNA levels to poor implantation potential, we performed a blinded retrospective analysis of the reproductive outcomes of euploid embryo transfers, depending

on mtDNA levels, performed by identical methodology in two different groups of patients in two different clinics. Two independent PGT clinics performed this retrospective reanalysis of mtDNA levels following single blastocyst transfers (group A, n = 49; group B, n = 70) or double blastocyst transfers (group A, n = 36; group B n = 29). Levels of mtDNA were correlated with implantation, miscarriage, and live birth. Aneuploidy testing was performed by NGS and the data used to calculate the mtDNA/gDNA ratio (mtDNA ratio), expressed as the number of mapped mtDNA sequencing reads x 100%/number of mapped autosomal sequencing reads. Only embryos identified as euploid and suitable for transfer were analyzed. Implantation rates for single embryo transfers were 43% (group A) and 55% (group B) and for double embryo transfers 54% (group A) and 62% (group B). Overall, the average mtDNA ratio was 0.018% (0.002–0.14%). For the single embryo transfer groups, there was no significant difference with respect for the average mtDNA ratio and for

0.10 P = 0.827

mtDNA ratio (%)

0.08

0.06

0.04

0.02

0.00 Implanted (N = 50)

not implanted (N = 62)

Upper panel

Fig. 6.20  mtDNA copy number does not predict implantation (upper), miscarriage (middle), or embryo quality (bottom)

6.7  Possible Additional Tests to Preselect Euploid Embryos with Higher Potential for Pregnancy

231

RGI-SET & age 0.10 P = 0.662

mtDNA ratio (%)

0.08

0.06

P = 0.095

0.04

0.02

Miscarriage 0.00 Middle panel

Ectopic 23–37

Implanted (N = 33)

23–37

Not implanted (N= 44)

38–44

Implanted (N =17)

38–44

Not implanted (N =18)

RGI-SET & embryo quality 0.10 P = 0.407

0.08

mtDNA ratio (%)

P = 0.309 0.06

0.04

0.02

Miscarriage Ectopic

0.00

Quality 4,5,6 is high, 3 is low Bottom panel

Fig. 6.20 (continued)

High quality Implanted (N =10)

High quality Not implanted (N =13)

Low quality Implanted (N = 40)

Low quality Not implanted (N = 49)

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6  Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic…

age to two-cell embryo (t2) and subsequent divisions t3, t4, t5, t6, t7, t8, and t9+ were recorded in the EmbryoViewer® workstation. Time of all mitotic events was expressed as hours post-­ ICSI. The kinetics of euploid embryos was evaluated according to their pregnancy outcome. Results showed that implanted euploid embryos progress more rapidly for each embryonic event or cleavage time. When median values were compared, a difference of 0.3–3.28h was found between implanted and non-implanted euploid embryos, statistically significant differences also being found for pronuclei appearance (tPNa) (p = 0.0065) and the time to achieve 9 cells (t9) (p = 0.0132). At each embryonic event, including cleavage time, implanted euploid embryos were 6.7.2 Morphokinetic Assessment faster. In addition, pronuclei appearance and last cleavage (t9) were significantly different between Other approaches to maximize the likelihood of a implanted and non-implanted euploid embryos, euploid embryo succeeding have included time-­ suggesting that morphokinetic criteria may be lapse imaging to track the dynamics of embryo useful in prioritization of euploid embryos for development. This approach was investigated for transfer.  The usefullness of time of morulation possible use in distinguishing the implantation and trophectoderm quality as  predictors of a potential among euploid embryos that otherwise euploid blastocyst’s reproductive competencen show similar morphology. Only a few studies com- was recently demonstrated in a large  collaborabining aneuploidy diagnosis and time-lapse imaging tive study, involving different centers under speare available. Available findings suggest that once an cific culture conditions [84]. embryo has been identified as being euploid, morIn addition to mitoscore and morphokinetic phokinetic markers may be additionally be used to parameters, other predictors of euploid embryos predict implantation potential of embryos. continue to be explored. As mentioned in Chap. The large study in which both data on aneu- 2, the potential value of cell-free DNA (cfDNA) ploidy and morphokinetic parameters were avail- profiles in spent media or blastocoele fluid is able was based on 143 patients (mean age: 35.07) being tested. These could provide independent applying for PGT-A due to advanced maternal data relevant to preselection of euploid embryos. age (>37) or a history of abnormal fetal karyo- It may be expected that a comprehensive use of type [83]. Cases included in the analysis had at all potential factors for preselection of embryos least two euploid embryos, all cultured in a time-­ could have high potential to result in pregnancy. lapse incubator (EmbryoScope™). Good or top-­ This could require artificial intelligence (AI) to quality blastocysts were biopsied and analyzed avoid a subjective interpretation of the results either by NGS (PGM platform, ThermoFisher) or [85]. Application of AI is currently being tested by array CGH (Illumina). Embryo morphokinet- for its value in NGS-based PGT-A in attempt of ics of 57 patients whose embryos had a negative avoiding operator subjectivity (www.coopergeoutcome (no pregnancy) were compared to these nomics.com). parameters of 86 patients with an ongoing pregnancy. Morphokinetic variables for all cleavage Conclusions: Tests to Preselect Euploid events up to the expanded blastocyst stage were Embryos annotated. All relevant events (fertilization, The introduction of next-generation technolocleavages, morula, and blastocyst formation) gies – SNP array, array CGH, qPCR, and NGS in were checked on a daily basis, and time of cleav- PGT-A – is of practical relevance in removing the the mtDNA ratio profile of the pregnant and nonpregnant women (Fig.  6.20). Among both pregnant groups, there was no obvious difference in mtDNA between the miscarriage subgroup and the group with a continuing, successful pregnancy. Further, there was no obvious trend between the mtDNA ratio and twin delivery, singleton delivery, miscarriage, or failure to implant. Thus, this study could not support the findings suggesting that high mtDNA ratios are an indicator of implantation failure. Results show that mtDNA ratio of trophectoderm biopsy samples cannot be used as an additional parameter for preselection of euploid embryos for transfer.

6.8  Characterizing Chromosomal Rearrangements

more than half of the tested oocytes or embryos that are aneuploid and lack the developmental competence and embryo potential to implant. However, the use of these sophisticated methods requires the critical step of DNA amplification from limited amounts of starting material obtained from an embryo biopsy. Amplification may also introduce artifacts that can be misinterpreted as mosaicism and segmental aneuploidy. Comparison of the types of chromosomal aneuploidies and the prevalence of each chromosome-­ specific error in oocytes and embryos further suggest that most chromosomal aneuploidies in embryos originate from female meiosis, predisposing to further sequential post-­zygotic errors, which may explain the high rate of chromosomal instability in preimplantation embryos. This may also be revealed to requirements for detection of the origin of aneuploidies, having important value in preselection of the embryo for single embryo transfer. Additional data on the outcome of mosaic embryo transfer and the follow-up of their clinical outcome will help determine recommendations for clinical practice. Preselection and prioritization of euploid embryos for transfer could apply these parameters to help identify those single euploid embryos with the highest potential to result in pregnancy and birth of a healthy offspring. Further evidence of significant impact of preselection of euploid embryos on reproductive outcome has been demonstrated by the results of the presently available randomized controlled trials, to be described in Chap. 7.

6.8

Characterizing Chromosomal Rearrangements

PGT for structural rearrangements (PGT-SR) was introduced only in 1996 but soon became one of the most practical applications of PGT.  Major impact has occurred in the clinical management of balanced translocation carriers. Initially, PGT-SR was done for maternally derived translocations and involved testing PB1 [86], specifically PBIbased metaphase chromosomes. However, PB1 alone does not always provide complete informa-

233

tion. Thus, PB1 is combined with PB2 analysis or performed using a single blastome and trophectoderm cells. FISH was then applied, but at present next-generation technologies (array CGH: NGS) are preferred. Although FISH is no longer used routinely, it is still useful when neither array CGH nor NGS is applicable for identification of fragments smaller than 5 Mb. FISH is also useful in distinguishing normal from balanced rearrangement, following methods for visualization of chromosomes in PBs and individual embryonic cells (see below).

6.8.1 Polar Body Approach This approach was introduced to perform PGT-SR based on PB1 never forming an interphase nucleus and consisting only of metaphase chromosomes. PB1 chromosomes are recognizable when isolated 2–3 h after in  vitro culture, with degeneration beginning 6–7 h after extrusion [87]. Therefore, centromeric and whole chromosome painting to determine the number of chromatids or the chromosome segment-specific probes was applied for testing of maternally derived chromosomal translocations in PB1 [87, 88]. The method resulted in a significant reduction in the number of spontaneous abortions in patients carrying translocations, yielding ­unaffected pregnancies and births of healthy children. The method appeared to be sensitive to malsegregation and/or recombination between chromatids, thus requiring follow-up analysis of PB2 in order to predict accurately the meiotic outcome following the second meiotic division to identify a possible chromatid exchange, as presented in Chap. 3. To follow up accurately the outcome of chromatid exchange, PB2 oocytes were transformed into metaphase chromosomes via electrofusion with foreign one-cell human embryo; however, the proportion of metaphases did not exceed 64% even after enucleation of the recipient one-cell stage mouse embryo, as described in Chap. 3 and reported elsewhere [89–91]. Confirmatory testing was possible in two of three spontaneously aborted embryos, showing the presence of de novo translocations different from the expected meiotic outcomes. Thus, application of this

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6  Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic…

approach was not practical, being replaced by nuclear conversion of the biopsied embryonic cells [92, 93].

6.8.2 K  aryotyping of Embryos via Nuclear Conversion To generate karyotypes, single embryonic cells were initially fused with enucleated or intact mouse zygotes at a known pronuclear stage in the S-phase of the cell cycle. Resulting heterokaryons were fixed at the metaphase of the first cleavage division, as described in Chap. 3 and reported elsewhere [88, 92]. This methodology was applied in 133 PGT-SR cycles, resulting in metaphase or S-period premature chromosome condensation (S-PCC) in 63.0% and 12.7%, respectively, from 1,451 cells in which an attempt was made to visualize chromosomes. Thus, chromosomal analysis was performed in 75.7% of cases, involving both normal and balanced embryos for transfer in 103 (77.4%) cycles. However, to avoid human embryonic cells fusing with mouse oocytes and to simplify the process of conversion from interphase nuclei to metaphase, a chemical conversion method was introduced that appeared to be more robust and highly reproducible, as described in detail in Chap. 3 and reported elsewhere [94]. This method was applied in 94 PGT-SR conversion cycles, involving chromosome analysis of 877 embryos. Metaphases were obtained from 672 embryos  – a conversion rate of 71%. In agreement with previous reports, detection rates of embryos suitable for transfer depended on the type of the translocation present, with a higher rate of unbalanced embryos found in reciprocal than in Robertsonian translocations. The proportion of embryos suitable for transfer did not significantly differ depending on the parental origin in reciprocal translocations, but the rate of unbalanced embryos was significantly lower in Robertsonian translocations of paternal origin. This is in agreement with the results of PGT-SR performed by the use of the next-generation technologies (see below), although neither of these differences affected the pregnancy rates, which

were comparable in reciprocal and Robertsonian translocations irrespective of parental origin of either type of rearrangements. This may be due, in part, to the current improvements in embryo culture systems with the shift toward single embryo transfer. In our cases, an average of 1.2 embryos per transfer was sufficient for yielding robust clinical pregnancies rates. In addition to the providing possibility of testing for small inversions and insertions, the conversion technique improved the accuracy of PGT-SR by distinguishing balanced from normal embryos, performed now also by other approaches, described in Sect. 6.8.4. Overall, the conversion methods allowed distinguishing 126 balanced from 112 normal ones in reciprocal translocations of maternal origin and 102 balanced from 105 normal embryos of paternal origin. In Robertsonian translocations, 33 balanced were distinguished from 27 normal embryos of maternal origin and 40 balanced from 27 normal embryos of paternal origin.

6.8.3 P  GT-SR Using Next-­ Generation Technologies As mentioned in Chap. 3, despite accuracy and exceptional utility of PGT-SR by specific FISH probes or karyotyping, their application is presently limited to specific cases and circumstances. The current standard technologies for PGT-SR involve array CGH and NGS, which further improve accuracy of testing and also allow a combined PGT-A. Examples of PGT-SR by NGS are presented in Figs. 6.21 and 6.22. Our present PGT-SR experience using aCGH and NGS is listed in Table  6.8. Overall, 331 PGT-SR cycles were performed, including 149 by aCGH and 182 by NGS, for 242 balanced translocation carriers (118 by aCGH and 124 by NGS) (Table  6.9). A total of 1,857 blastocysts were tested, 1,809 (97.41%) of which were with the results of PGT-SR and PGT-A. As seen from Table 6.9, a total of 568 (31.8%) blastocysts were identified as suitable for transfer, which were balanced/normal and also euploid: 313 (37.8% suit-

6.8  Characterizing Chromosomal Rearrangements

235

Fig. 6.21  NGS-based PGT-SR for maternal translocation t(11:20)(q23; q11.2). Embryo 1 has derivative chromosome 11 (pointed by red arrows); embryo #2 has a derivative chromosome 11 (pointed by red arrows), together

with monosomy 16 (circled in red). Embryo #3 has a derivative chromosome 20 (pointed by red arrows) and also trisomy 16 (mosaicism) (circled in blue)

able embryos for transfer) of these embryos were analyzed using aCGH and 255 (25.9%) using NGS. As seen from the details of the remaining abnormal embryos in Table 6.9, 362 (29.1%) of the embryos were balance/normal and could have been selected for transfer if not tested for aneuploidy and, thus, destined to be lost during implantation and postimplantation development, including 138 (27.0%) embryos tested by aCGH and 223 (30.6%) tested by NGS. The largest group, 179 patients, were carriers of reciprocal translocation, 74 of maternal and 85 of paternal origin (Tables 6.10). Of 53 Robertsonian translocation carriers, 27 were

with translocations of maternal and 26 of paternal origin. Only ten patients carry inversions, three maternal and seven paternal in origin. The lowest rate of suitable embryos for transfer was detected in PGT-SR performed for carriers of reciprocal translocations, 29.1%, irrespective of parental origin, compared to 39.0% performed for carriers of Robertsonian translocations, which was higher in PGT-SR for Robertsonian translocations of paternal origin (44.8%) compared to 33.8% PGT-SR for Robertsonian translocations of maternal origin. Reproductive outcome of the above PGT-SR will be presented below.

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6  Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic…

Fig. 6.22  NGS-based PGT-SR for maternal translocation t(11:22)(q23.1;q11.2). Of the three tested embryos by NGS shown, embryo #4 has a derivative chromosome 11 (pointed by red arrows) and also segmental mosaicism for monosomies 1(q32.3 q44) (circled in red); embryo #7 has

an extra derivative chromosome  11 (pointed by red arrows); and embryo #8 also with a derivative chromosome 11 (pointed by red arrows), plus monosomy  15 (circled in red)

6.8.4 PGT-SR Developments to Distinguishing Noncarrier Balanced Embryos from Normal

serve as a reference for distinguishing balanced translocation from normal blastocysts. Thus, utility of the method is limited because the requisite reference may not be available. The more universal approach is a specially designed NGS technology termed mate-pair sequencing (MPS). This involves a high depth mate-pair sequencing to identify breakpoint regions and Sanger sequencing to define the exact breakpoint needed for designing specific primers required for identifying normal and carrier embryos. Application of this technique has resulted in birth of a child free of a balanced translocation, avoiding to transmission of the rearrangement to the next generation (Fig. 6.23) [97].

In addition to the conversion methods described above by karyotyping, a few sophisticated approaches based on next-generation technologies have been developed for distinguishing balanced embryos from the normal ones. One of such technologies involves the use of single nucleotide polymorphism microarray, which was validated in 148 embryos and showed concordance with the karyotype [95, 96]. However, this method requires availability of both the unbalanced embryo, and  parental DNA necessary to

6.8  Characterizing Chromosomal Rearrangements

237

Table 6.8  List of chromosomal rearrangements for which PGT was performed by the next-generation technologies (NGS and aCGH) # CYCLES 1 1 1 1 1 1 1 1 1 1 2 3 1 1 3 1 1 2 1 1 2 1 1 1 1 2 1 1 3 1 2 1 2 2 1 1 3 1 1 1 1 2 1 1 3 1 1 2

Karyotype 46,XY,inv(19)(p13.3q13.1) 46,XX,inv(18)(p11.32q12.2) 46,XY,inv(14)(q24.1;q32.1) 46,XY,inv(11)(p14q24.2) 46,XX,inv(8)(p23.q21.2) 46,XX,inv(6)(p12.2p25) 46,XX,inv(6)(p21.3q15) 46,XY,inv (5) (q31.1q33.1) 46,XY,inv(5)(p15.3;q13.3) 46,XX,inv(4)(p15.3q24) 46,XY,inv(1)(p22.3q42.1) 46,XX,t(4;6)(p14;q16.2) 46,XX,t(11;15)(q23;q15) 46,XX,t(3;21)(q12;q11.2) 46,XX,t(1;3)(p36.3;p25) 46,XX,t(7;19)(q22;q13.4) 46,XY,t(6;7)(q27;q22) 46,XX,t(18;22)(p11.2;q13.1) 46,XY,t(1;17)(q21;p13.1) 46,XX,t(8;18)(p21.1:p11.2) 46,XX,t(5;13)(q22;q32) 46,XX,t(17;18)(p13;q23) 46,XY,t (5;21) (p13.1;p11.1) 46,XX,t(3;17)(q21;p11.2) 46,XX,t(4;7)(q27;q31.2 46,XX,t(2;18)(q32.2;q21.1) 46,XX,t(12;16)(q22;p13.1) 46,XX,t(7;13)(p11.2;q21.2 46,XX,t(11;22) (q23;q11.2) 46,XY,t(2;7)(p25;p21) 46,XX,t(9;16)(q34.3;p13.12) 46,XX,t(2;16)(q21.1;p12.2) 46,XX,t(3;5)(q12;p13.2) 46,XY,t(3;9)(q26.2;q13) 46,XX,t(4;13)(p16;q12) 46,XX,t(16;17)(p11.2;p13) 46,XY,t(18;22)(p11.3;q13) 46,XX,t(11;13)(q22;q31) 46,XX,t(6;10)(p25.3;p14) 46,XX,t(7;20)(q31;q13.3) 46,XX,t(11;12)(q24.2;q24.1) 46,XY,t(10;22)(q25.2;q11.2 46,XX,t(5;6)(q33.1q13) 46,XY,t(1;3)(p36.3;p25) 46,XY,t(4;11)(q33;q24.2) 46,XX,t(1;14)(q42.13;q32.33) 46,XX,t(5;16)(q12;p13.1) 46,XX,t(2;20)(q33;p12)

Transl type inv inv inv inv inv inv inv inv inv inv inv Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip

# Balanced euploid 0 1 1 2 1 8 2 5 4 1 1 6 5 1 2 1 1 3 1 0 1 4 3 1 1 0 0 3 3 1 3 2 1 4 0 1 2 0 0 1 3 3 0 1 1 1 2 3

FET 0 1 1 1 1 1 2 3 1 1 1 3 1 1 2 1 1 2 1 0 1 2 1 1 1 0 0 1 2 1 2 1 1 3 0 1 1 0 0 1 1 2 0 1 1 1 1 3

# Embryos transferred 0 1 1 2 1 1 2 3 1 1 1 4 1 1 2 1 1 2 1 0 1 2 2 1 1 0 0 1 3 1 2 2 1 4 1 2 0 0 1 2 2 0 1 1 1 2 3

Pregnancy 0 0 1 0 0 1 1 2 1 1 1 1 1 1 1 1 0 1 0 0 1 1 1 0 1 0 0 1 1 1 1 1 0 1 0 0 1 0 0 1 1 0 0 1 0 1 1 1

Birth 0 0 1 0 0 1 1 2 1 1 1 1 1 1 1 1 0 1 0 0 1 1 2 0 1 0 0 1 1 1 1 1 0 1 0 1 1 0 0 1 1 0 0 1 0 1 1 1

Age 42 32 42 43 34 36 35 35 34 36 36 33 33 32 26 36 30 31 35 41 32 30 32 30 30 38 28 33 33 33 31 38 41 38 32 30 29 30 29 31 31 27 25 31 36 39 31 31

(continued)

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Table 6.8 (continued) # CYCLES 1 1 1 1 1 1 1 1 1 1 2 1 1 1 1 3 3 1 1 1 1 2 1 1 1 1 1 1 1 1 1 1 1 2 2 1 1 1 1 1 1 1 1 1 2 1 1 1

Karyotype 46,XY,t(12;14) (p11.2;q32.5) 46,XX,t(16;17)(p11.2;p13) 46,XX,t(2;16)(p11.2;p12) 46,XY,t(11;19)[p15.1;q13.3) 46,XY,t(12;14) (p11.2;q32.5) 46,XY,t(11;22)(q23:q11) 46,XY,t(1;6)(q25;q15) 46,XY,t(1;2)(q44;q32.2) 46,XY,t(11;13)(q21;q14.1) 46,XX,t(4;8)(p16.1;p23.1) 46,XX,t(6;14)(q25.3;q32.3) 46,XY,t(3;21)(p24.3;p11.2) 46,XY,t(10;17)(q23.3;q12) 46,XY,t(11;22)(q23;q11.2) 46,XX,t(8;11)(q24.1;q23.3) 46,XY,t(2;9)(p25.1;p22) 46,XX,t(13;14)(q12;q24.3) 46,XX,t(8;11)(p12;p14) 46,XX,t(4;9)(q21;q34) 46,XX,t(11;14)(q22;q23) 46,XY,t(9;16)(q32;q23) 46,XX,t(1;7)(p36.1;q32) 46,XY,t(3;4)(q25.1;p12) 46,XY,t (11;22)(q23;q11.2) 45,X/46,X,r(X)(p22.1q26) 46,XX,t(11;22)(q23.3;q11.2) 46,XY,t (15;21) (q15;q22.3) 46,XY,t(11; 18) (q14; q23) 46,XY,t(6;12)(q16.2;q22) 46,XY,t(2;14)(p15;p12) 46,XY,t(11;22)(q23;q11.2) 46,XX,t(7;18)(q11.1;q11.1) 46,XY,t(2;3)(q23;q23) 46,XY,t(7;20)(p21.3;p13) 46,XX,t(16;22)(p13.3;q11.2) 46,XX,t(16;22)(p13.3;q11.2) 46,XX,t(4;14)(q34;q24.2) 46,XY,t(11;22)(q23;q11.2) 45,X,dic(Y;15)(q11.2;p11.2) pat,t(2;17)(p25.2;p13.1)mat 46,XY,t(3;7)(p25;p14.2) 46,XX,t(4;9)(p15.31;p23) 46,XY,t(18; 20)(q21.2; q13.1) 46,XY,t(17;22)(p13;q11.2) 46,XX,t(11;20)(q23; q11.2) 46,XX,t(5;9)(q33.1;q32) 46,XX,t(7;17)(q11.23;q25.1) 46,XY,t(12;16)(q24.31;p11.2) 46,XX,t(6;18)(p21.3;p11.2)

Transl type Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip

# Balanced euploid 1 3 1 1 2 3 2 1 1 1 3 1 2 4 2 2 2 1 3 0 3 2 1 1 0 2 4 2 4 3 1 1 0 0 0 1 0 6 2

# Embryos FET transferred

Pregnancy Birth

2 1 0 1 1 1 1 1 1 2 1 1 1 1 2 2 1 1 0 1 2 1 1 0 1 2 1 2 2 1 1 0 0 0 1 0 2 1

2 1 0 2 1 1 1 1 1 2 1 1 2 1 2 2 1 1 0 1 2 1 1 0 1 2 1 2 2 1 1 0 0 0 1 0 2 1

1 0 1 1 1 1 0 0 0 1 1 1 1 1 1 1 1 1 0 1 1 1 0 0 1 1 1 1 2 0 1 0 0 0 1 0 1 1

1 0 1 1 1 1 0 0 0 1 1 1 1 1 1 0 0 1 0 1 1 0 0 0 1 1 1 1 1 0 1 0 0 0 0 0 1 1

Age 51 30 34 37 24 36 31 35 37 32 32 36 37 27 34 32 40 34 25 29 28 29 34 34 34 28 27 32 30 33 37 34 38 33 38 37 38 33 28

Recip Recip Recip Recip Recip Recip Recip Recip Recip

0 4 3 0 2 0 3 3 1

0 1 1 0 1 0 1 1 1

0 1 2 0 1 0 1 1 1

0 1 1 0 1 0 1 1 0

0 1 2 0 1 0 1 1 0

35 34 34 34 24 37 33 34 34

6.8  Characterizing Chromosomal Rearrangements

239

Table 6.8 (continued) # CYCLES 1 2 1 2 1 1 1 1 1 2 2 2 1 1 3 1 1 1 1 1 1 1 1 3 1 2 1 2 1 1 2 1 2 1 2 1 1 1 2 2 1 1 1 1 1 2 1 1 1

Karyotype 46,XY,t(11;22)(q23.3;q11.21) 46,XY,t(8;14)(q11.2;q24.3) 46,XX,t(10;12)(q24;p13) 46,XX,t(10;12)(q24;p13) 46,XY,t(1;9)(q21;q13) 46,XY,t(2;7)(q37.3;q32) 46,XX,t(8; 11) (q21.2; p13) 46,XX,t(3;11)(p23;p15) 46,XX,t(4;11)(q27;q23.1) 46,XX,t(5,13)(p15.1;q22) 46,XX,t(10;12)(q22.1;q13.3) 46,XX,t(11; 22)(q23.3; q11.2) 46,XY,t(1;8)(q23.1,p21.3) 46,XX,t(11;22)(q23;p11) 46,XY,t(3;10)(p21.1;p15) 46,XY,t(16;18)(12.1;p11.3) 46,XX,t(10;21)(p13;q11.2) 46,XY,t(4;12)(q31.1;q24.1) 46,XX,t(3;16)(p14.2;p13.1) 46,XY,t(2;11)(p13;q13.1) 46,XX,t(7;8)(p21;q12) 46,XX,t(8;15)(q24.1;q22.1) 46,XY,t(3;16)(p21.3; p13.3) 46, XX,t(2;5)(q37;q31) 46,XY,t(3;15)(p23;q22.3) 46,XX,t(12;22)(q23.3;q11.2) 46,XY,t(1;5)(p13;q33) 46,XX,t(12;22)(q24.31;q11.2) 46,XY,t(3;13)(q23;q12.3) 46,XX,t(3;19)(q21;p13.3) 46,XX,t(2;6)(p22;q22.2) 46,XY,t(9;15)(q13;q11) 46,XX,t(15;18)(q11.2;p11.2) 46,XX,t(11;22)(q23.1;q11.22) 46,XX,t(11;22)(q23.1;q11.22) 46,XY,t(7;10)(q11.21;q21.2) 46,XY,t(1;10)(p13;q11.1) 46,XX,t(11;12)(p11.2;q15) 46,XY,t(7;13)(p11.2;q22) 46,XX,t(7;8)(p13;q23.1) 46,XX,t(7;20)(p15;q13.1) 46,XY,t (11; 21) (p14; q22.3) 46,XY,t(1;3)(q25.3;q28) 46,XX,t(4;17) (q31.1;q23.1) 46,XY,t(9;12)(q22.1;q24.1) 46,XX,t(3;15)(q26.2;q23) 46,XY,t (3;4) (q25.3;q31.3) 46,XX,t(2;13)(p21;q22) 46,XX,t(5;18)(p13;p11.2)

Transl type Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip

# Balanced euploid 3 2 2 5 2 2 2 2 0 2 1 4 0 2 3 1 4 2 2 4 2 0 0 7 5 3 1 2 2 0 4 3 1 3 4 5 3 2 4 2 0 2 9 3 1 4 2 2 0

FET 2 1 1 2 2 1 1 1 0 1 1 1 0 2 2 1 1 1 1 1 1 0 0 2 2 1 1 2 1 0 2 1 1 1 2 2 1 2 1 1 0 1 1 2 1 3 1 1 0

# Embryos transferred 2 1 2 3 2 1 1 2 0 2 1 1 2 3 1 2 1 2 2 2 0 0 3 2 1 1 2 1 0 3 2 1 2 3 2 2 2 1 1 0 1 1 2 1 3 1 1 0

Pregnancy 1 1 1 2 1 1 1 0 0 1 1 1 0 1 2 1 1 0 0 1 1 0 0 1 1 1 0 1 1 0 1 1 1 1 2 1 1 1 1 1 0 1 1 1 1 2 1 1 0

Birth 1 1 1 2 1 1 1 0 0 1 0 1 0 1 2 1 1 0 0 1 1 0 0 1 2 1 0 1 1 0 1 1 1 1 1 2 2 1 1 1 0 1 1 1 1 2 1 1 0

Age 25 30 35 36 34 33 34 36 33 37 28 32 37 34 34 33 36 35 41 30 35 30 36 27 32 31 30 35 36 44 39 31 39 33 35 31 33 32 34 37 40 33 26 36 34 36 29 32 36

(continued)

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Table 6.8 (continued) # CYCLES 1 2 1 1 2 1 4 1 1 1 1 1 1 1 1 1 1 2 4 2 1 2 2 3 1 3 1 1 1 1 1 2 1 2 1 1 2 1 1 1 1

1 1 4 2

Karyotype 46,XY,t(4;9)(q33;p24.3) 46,XY,t(16;18)(q22;p11.1) 46,XX,t(8;9)(p21;q13) 46,XY,t(6;18)(q13;q23) 46,XY,t(7;11)(q35;p15.4) 46,XY,t(9;20)(q34.3;q11.2) 46,XY,t(1;9)(q41;p21) 46,XX,t(11;20)(q13.1; q13.1) 46,XX,t(10;13)(p13;q31) 46,XX,t(7;14)(q22;q13) 46,XY,t(2;11) (p24;q13.3) 46,XY,t(6,12)(q15;p11.2) 46,XX,t(12;17)(p12.1,q25.3) 46,XY,t(1;6)(p13.3;q14) 46,XX,t(3;18)(q13.3;q21.3) 46,XY,t(10;14)(q11.2;q13) 46,XX,t(11;22)(q23.3;q11.21) 46,XY,t(2;7)(q11.2; p13) 46,XY,t(4;15) (q25; q26.1) 46,XX,t(1; 17) (q42.1; q23) 46,XY,t(11;22)(q23;q11.2) 46,XY,t(11;22)(q23;q11.2) 46,XY,t(5;21)(p10;p10) 46,XX,t(19;20)(q13.1;p13) 46,XX,t(1;8)(q23.1;q22.1) 46,XY,t(18;21)(q12.2;q22.1) 46,XY,t(5;14)(p14;q31.1) 46,XY,t(11;18)(q14;q23) 46,XX,t(5;13)(p12;q32) 46,XY,t(2;17)(q13;q25) 46,XX,t(8;13)(p21.1;q22) 46,XX,t(8;13)(p21.1;q22) 46,XX,t(8;17)(p10;p10) x2/46,XX(48) 46,XX,t(1;6)(p36.3;p21.1) 46,XX,t(7;15)(q11.2;q26.1) 46,XY,t(10;15)(q21.2;p13) 46,XX,t(11;22)(q23;q11.2) 46,XX,t(3;14)(p21.3;q32.3) 46,XY,t(1;19) (p36.11;p13.3) 46,XY,t(18;21)(q12.1;q21.2) 46,XX,der(5) t(5;20) (q35.1;p11.2), del (10) (p13p15), der (20) t(5;20) (q35.1;p11.2) in (20;10) (p11.2;p13p15) 46,XY,t(1;9)(q42.13;q32 46,XX,t(7;9)(p21.2;p23) 46,XX,t(7;9)(p21.2;p23) 46,XY,t(1;4)(q24.3;p15.2)

Transl type Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip Recip

# Balanced euploid 1 7 2 3 4 9 3 4 4 1 4 1 7 1 0 3 3 2 9 0 0 2 2 1 3 4 2 1 0 1 3 4 6

FET 1 3 1 2 2 1 3 1 1 1 1 1 2 1 0 1 1 1 3 0 0 1 1 1 1 2 1 1 0 1 1 1 3

# Embryos transferred 1 6 1 2 2 2 4 1 1 1 1 1 2 1 0 1 1 1 3 0 0 1 2 1 2 4 1 1 0 1 2 1 4

Pregnancy 0 3 1 1 2 1 3 1 1 1 1 0 2 0 0 1 1 1 2 0 0 1 1 0 1 1 1 1 0 1 0 1 1

Birth 0 2 1 1 2 1 2 1 3 1 1 0 2 0 0 1 1 1 2 0 0 1 1 0 2 2 0 1 0 1 0 1 1

Age 29 37 36 31 32 31 30 35 36 28 30 40 29 40 33 34 24 29 37 39 33 33 31 37 29 31 30 38 34 30 28 29 41

Recip Recip Recip Recip Recip Recip Recip Recip

1 0 1 3 2 2 4 2

1 0 1 2 1 2 1 1

1 0 1 3 1 2 1 2

0 0 1 1 1 1 1 1

0 0 1 1 1 1 1 1

38 27 39 34 28 30 32 30

Recip Recip Recip Recip

2 1 8 3

1 1 4 3

1 1 5 3

1 1 2 2

1 1 2 1

33 31 35 27

6.8  Characterizing Chromosomal Rearrangements

241

Table 6.8 (continued) # CYCLES 3 1 1 1 1 2 1 1 1 1 2 1 1 1 1 1 4 1 1 1 2 1 1 3 3 1 1 1 1 2 2 1 4 1 1 2 1 1 1 2 1 1 1 1 1 1

Transl Karyotype type 46,XY,t(3;15)(p14;q25) Recip 46,X,t(X,4)(p11.4;q31.3) Recip 46,X,der(Y) t(Y;13)(q11.23;q21.3 Recip 46,XY,t(Y; 15) (q11.22; q11.2) Recip 46,Y,t(X;19)(p10;p10) Recip 46,XX,t(4;5;11)(p15.2;q15;p13) Recip (3 chr) 46,XY,t(1;2;2;3) Recip (p13;p13:q24.3:p13) (3 chr) 46,XY,t(3;14)(p23;q13),t(8;13) Recip (q22.3;q14.3) (3 chr) 45,XY,der(14;15)(q10;q10) Rob 45,XY,der(13;14)(q10;q10) Rob 45,XX,der (13;14)(q10;q10) Rob 45,XX,der(13;14)(q10;q10)) Rob 45,XX,der(13;14)(q10;q10) Rob 45,XX,der(13;14)(q10;q10) Rob 45,XY,der(13;21)(q10; q10) Rob 45,XY,der(13;14)(q10;q10) Rob 45,XX,der(6;14)(p10;p10) Rob 45,XX,der(15;21)(q10;q10) Rob 45,XX,der(13;14)(q10;q10) Rob 45,XY,t(1;10)(q21;q26.3) Rob 45,XX,der(13;22) (q10; q10) Rob 45,XX,der(13;14)(q10;q10) Rob 45,XY,der(13;21) (q10; q10) Rob 45,XY,der(13;14)(q10;q10) Rob 45,XY,der(14;21)(q10;q10) Rob 45,XY,der(13;14)(q10;q10) Rob 45,XY,der(14;21)(q10;q10) Rob 45,XY,der(13;14)(q10;q10) Rob 45,XX,der(13;14)(q10;q10) Rob 45,XY,der(13;14)(q10;q10) Rob 45,XX,der(13;14)(q10:q10) Rob 45,XX,der(14;21)(q10;q10) Rob 45,XX,der(14;21)(q10;q10) Rob 45,XY,der(13;15)(q10;q10) Rob 45,XY,der(13;14)(q10;q10) Rob 45,XY,der(13;14)(q10;q10) Rob 45,XY,der (13;14)(q10;q10) Rob 45,XY,der(13;14)(q10;q10) Rob 45,XY,der(13;14)(q10;q10) Rob 45,XY,der(14;21)(q10;q10) Rob 45,XY,der(13; 14) (q10;q10) Rob 45,XX,der(14;22)(q10;q10) Rob 45,XY,der,(13; 14) (q10;q10) Rob 45,XX,der(13;14)(q10;q10) Rob 45,XX,der(13;14)(q10;q10) Rob 45,XY,der(14;15)(q10;q10) Rob

# Balanced euploid 6 3 1 1 2 2

FET 3 1 1 1 1 1

# Embryos transferred 3 1 1 1 1 1

Pregnancy 1 1 0 0 1 0

Birth 1 1 0 0 1 0

Age 38 37 35 35 27 34

1

1

1

0

0

35

0

0

0

0

0

33

1 4 2 1 2 0 2 3 4 3 2 4 7 4 2 9 5 2 0 3 0 1 7 0 2 4 1 8 0 2 0 4 0 0 1 0 1 4

1 2 1 1 2 0 1 2 2 2 1 2 3 1 1 4 2 1

1 2 1 1 2 0 2 2 2 2 2 3 5 2 1 4 2 1

1 1 1 1 1 0 1 2 2 1 0 1 1 1 1 1 1 1

1 1 1 1 1 0 2 2 2 1 0 1 1 2 1 2 1 1

1 0 1 3 0 1 2 1 2 0 1 0 2 0 0 1 0 1 1

1 0 1 3 0 1 2 1 2 0 1 0 2 0 0 1 0 1 1

1 0 1 2 0 1 1 0 1 0 1 0 1 0 0 0 0 0 1

1 0 0 2 0 1 1 0 0 0 1 0 1 0 0 0 0 0 1

21 33 31 34 35 41 38 31 43 30 32 35 32 31 32 29 33 32 41 34 34 34 32 40 37 30 37 34 40 40 32 33 42 34 32 37 34 29 (continued)

242

6  Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic…

Table 6.8 (continued) # CYCLES 1 1 1 1 1 1 1 1 1 1 1 1 1 1 2

Karyotype 45,XX,der(13;14)(q10;q10) 45,XX,der(13;14)(q10;q10) 45,XX,der(13;15)(q10;q10) 45,XY,der(22;13)(p12;p12) 45,XY,der(13;14)(q10;q10) 45,XX,der(13;14)(q10;q10) 45,XX,der(13;14)(q10;q10) 45,XY,der(13; 14) (q10; q10) 45,XY,der(13;14)(q10;q10) 45,XY,der (13; 14) (q10;q10) 45,XX,der(13;14)(q10;q10) 45,XX,der(13;14)(q10;q10) 45,XX,der(13;14)(q10;q10) 45,XX,der(13; 14)(q10;q10) 45,XX,der(14;21)(q10;q10) TOTAL

Transl type Rob Rob Rob Rob Rob Rob Rob Rob Rob Rob Rob Rob Rob Rob Rob

# Balanced euploid 1 3 0 0 2 5 2 0 2 6 4 4 1 1 2 567

A similar approach, termed nanopore long-­ read sequencing, also discriminates carrier from noncarrier embryos through a high-resolution breakpoint mapping followed by breakpoint PCR [98]. This method was applied to a couple in which the female partner had a reciprocal translocation (46,XX,t(7:13)(p13;q12.3), for whom a sequencing library was prepared from genomic DNA.  After sequence alignment, breakpoints were accurately predicted on chromosomes 7 and 13; PCR primers were designed, which were applied for PGT-SR of nine biopsied blastocysts. Using PGT-SR, two euploid, two mosaic, and four unbalanced carrier embryos were detected. Thus, following application of breakpoint PCR, carrier embryos could be discriminated from noncarrier embryos. Preselection and transfer of one euploid noncarrier blastocyst resulted in a clinical pregnancy. Both these approaches enable accurate high-resolution breakpoint mapping directly on balanced reciprocal translocation carriers, providing the option of transferring euploid noncarrier embryos.

FET 1 1 0 0 2 1 1 0 1 1 2 1 1 1 2 287

# Embryos transferred 1 2 0 0 2 1 2 0 1 1 2 2 1 1 2 337

Pregnancy 1 1 0 0 1 1 1 0 1 1 2 1 0 0 2 191

Birth 1 1 0 0 1 1 1 0 1 1 2 2 0 0 1 174 delivery 191 babies

Age 26 31 37 30 31 34 31 34 21 35 32 36 37 39 34

6.8.5 Reproductive Outcome of PGT-SR PGT-SR has become a preferred option for carriers of balanced rearrangements to have an unaffected child of their own without fear of repeated spontaneous abortions or affected children. Thus, an increasing number of PGT-SR cycles are being performed, with the current experience now including many thousands of clinical cycles worldwide, resulting in hundreds of clinical pregnancies and births of unaffected children. This experience provides the proof that PGT-SR improves pregnancy rates and reduces risk of spontaneous abortions. Our data comparing pregnancy outcomes before and after PGT-SR demonstrates a considerable reduction of spontaneous abortions from 87.8% to 17.8% and increased take-home baby rates from 11.5% to 81.4% after PGT-SR. Prior to PGT-SR, the same patients had a highly unfavorable reproductive history of 654 pregnancies: 496 (75.8%) spontaneous abortions, 49 therapeutic abortions, 9 stillbirths, and 25 offspring with unbalanced translocations [90, 91].

Maternal, paternal Maternal, paternal

Origin Maternal, paternal Maternal, paternal

TOTAL

NGS

Maternal, paternal Maternal, paternal Maternal, paternal

Subtotal aCGH Maternal, paternal NGS Maternal, paternal Subtotal

NGS

aCGH

Subtotal

NGS

Test type aCGH

All types tested aCGH

Inversion

Robertsonian

Translocation type Reciprocal

11 149

10

118

242

331

182

8

7

124

71 3

30

41

249

144

# Cycles 105

53 3

19

34

179

98

# Patient 81

1857

1005

852

44

29

355 15

123

232

1458

853

Total studied 605

# With results 589 (97.3%) 836 (98%) 1425 (97.7%) 224 (96.5%) 117 (95.1%) 341 14 (93.3%) 29 (100%) 43 (97.7%) 827 (97%) 982 (97.7%) 1809 (97.4%)

Table 6.9  Results of PGT-SR for all types of translocations by aCGH and NGS

255 (25.9%) 567 (31.3%)

313 (37.8%)

11 (37.9%) 18 (41.8%)

37 (31.6%) 133 7 (50%)

96 (42.8%)

416 (29.1%)

206 (24.6%)

Balanced euploid (suitable for fet) 210 (35.6%)

514 (62.1%) 728 (74.1%) 1242 (68.6%)

18 (58.6%) 25 (55.8%)

Abnormal Total abnormal 379 (64.3%) 630 (75.5%) 1009 (70.8%) 128 (57.1%) 80 (68.3%) 208 7 (50%)

223 (30.6%) 362 (29.1%)

11 (64.7%) 16 (64%) 138 (27%)

39 (48.7%) 97 5 (71.4%)

58 (46%)

174 (27.6%) 249 (24.6%)

Balanced aneuploid 75 (19.7%)

262 (35.9%) 468 (37.6%)

6 (35.9%) 8 (32%) 206 (48.2%)

12 (15%) 32 2 (28.5%)

20 (15.6%)

237 (37.6%) 421

Unbalanced euploid 184 (48.5%)

250 (34.3%) 410 (33%)

160 (31.1%)

1 (5.8%) 1 (4%)

29 (36.2%) 69 0

40 (31.2%)

219 (34.7) 339

Unbalanced aneuploid 120 (31.6%)

6.8  Characterizing Chromosomal Rearrangements 243

53 3 7

10 118 125

243

Subtotal aCGH NGS

Subtotal aCGH NGS

Total

All types

Robertsonian

Inversion

# Patient 81

99 180 34 19

Test type aCGH

NGS Subtotal aCGH NGS

Translocation type Reciprocal

331

11 149 182

71 3 8

144 249 41 30

# Cycles 105

Balanced euploid 210 (35.6%) 206 (24.6%) 416(29.1%) 96 (42.8%) 37 (31.6%) 133 7 (50%) 11 (37.9%) 18 (41.8%) 314 (37.8%) 254 (25.9%) 567 (31.3%)

Table 6.10  PGT-SR Outcome for all types of translocations by aCGH and NGS

287∗ Numerous fet from the same cycle

12 134 153

62 5 7

125 213 41 21

# Transferred 88

337 (1.17)

13 170 167

74 5 8

136 250 51 23

# Embryos 114

191 (66.5%)

7 101 90

39 3 4

73 145 26 13

Pregnancy 72

174 (91%)/191

7/7 92/106 82/85

36/41 3/3 4/4

66/69 131/143 24/29 12/12

Delivery/#children born 65/74

244 6  Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic…

6.8  Characterizing Chromosomal Rearrangements B. Clinical PGT*

A. Breakpoint (BP) identification*

BP1

245

IVF

Balanced translocation

Blastocysts BP2

̘

BP1

Biopsy

Mate pair sequencing TE cells

a g c t t g t a g

̘ g g c c a a t a c

̘ BP primer design

WGA/CNV-Seq BP2

PCR of WGA product with BP primers

̘

Diagnostic primers

Unbalanced

Balanced

Normal Transfer

Carrier

*Prepared by David Cram

Fig. 6.23  Mate-pair sequencing for distinguishing normal from balance rearrangements (Prepared by David Cram)

The reproductive outcome is extremely poor for carriers of structural rearrangements, with more than three quarters of their pregnancies resulting in spontaneous abortions. This is explained by unfavorable meiotic outcomes. Outcomes vary depending on the type of translocations and their origin. To investigate the meiotic outcome of translocations in relation to the type and origin, segregation patterns from 130 patients carrying balanced translocations were analyzed (Figs. 6.24 and 6.25). Meiotic outcomes were inferred either from PB1 and PB2 (Fig. 6.24) or from embryo analysis (Fig. 6.25). Meiotic outcome detection rates by each of these methods are comparable, except for chromatid exchanges detected only by sequential PB1 and PB2 analysis (16.4%). Complex errors were more frequent in PB analysis (17.3% vs 4.2%); 3:1 segregation was found more frequently in embryo analysis (7.7% vs 23%).

Segregation patterns for paternally and maternally derived translocations showed similar tendencies, predominantly represented by alternate segregation (35 and 34%, respectively) and adjacent I segregation (28 and 34%, respectively), with less frequent adjacent II segregation (9.1 and 11.4%, respectively). These meiotic outcomes explain the high proportion of unbalanced embryos predicted in 77.6% embryos obtained from maternally derived reciprocal translocations; only 22.4% were suitable for transfer: 7.8% balanced, 6.9% normal, and 7.7% balanced/ normal. On the other hand, unbalanced embryos were predicted in 75% embryos obtained from paternally derived reciprocal translocations, leaving 25% embryos suitable for transfer, including 7.1% balanced, 7.3% normal, and 10.6% balanced/normal. Testing embryos for maternally derived Robertsonian translocations resulted in 64.4% unbalanced embryos. The remaining

6  Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic…

246 35 30 35 25

34.1

34.1 28

20

18 23.1

15 10

9.1 11.4

5 0.7

0.9

4.2

1.6

0 Alternate

Adjacent 2

Adjacent 1

4:0 seg

3:1 seg

Complex

% from 143 Blastomeres (female carriers) % from 317 Blastomeres (male carriers)

Fig. 6.25  Distribution of segregation patterns. Bar graph demonstrating the distribution of inferred segregation modes for female reciprocal translocations compared to male reciprocal translocations

35

28 23.1

18.2

17.8 14.4

9.1

8.2

7.7

0 Alternate

Adjacent 1

17.3

16.4

Adjacent 2

3:1 seg

0.7

4:0 seg

0 CE (1)

4.2 0 CE (both)

Complex

% from 439 1st PBs % from 143 Blastomeres

Fig. 6.24  Distribution of segregation patterns. Bar graph demonstrating the distribution of observed segregation modes for female reciprocal translocations by PB1 analysis versus inferred segregation patterns by blastomere analysis

35.6% embryos were suitable for transfer, including 13.2% balanced, 10.8% normal, and 11.6% balanced/normal. Similarly, testing of embryos for paternally derived Robertsonian translocations resulted in identification of 53.4% unbal-

anced embryos, the remaining 46.6% suitable for transfer including 10.4% balanced, 7% normal, and 29.2% balanced/normal. Overall clinical pregnancies were obtained in 37% of transfer cycles, with 26% overall delivering healthy chil-

6.8  Characterizing Chromosomal Rearrangements

dren. The data on the meiotic outcome may explain the observed 85% spontaneous abortion rate in patients prior to undertaking PGT-SR procedure, now reduced to 17% after PGT-SR. This demonstrates the tremendous positive impact of PGT-SR on the clinical outcome of pregnancies in couples carrying translocations. In some rearrangements, prediction of the unbalanced embryos presents a challenge. An example occurs in couples in which there is mosaicism for a translocation or in which both parents have a highly complex translocation. For example, we performed PGT-SR a couple in which the male partner had mosaicism for a balanced reciprocal translocation 46,XY,t(10;11) (q23;q23/46,XY). Of six embryos tested, three were unbalanced; three were normal/balanced, two of which were transferred resulting in an unaffected pregnancy. In an unusual case, PGT-SR was performed for a consanguineous couple, in which both the female patient and her husband carried balanced Robertsonian translocation (der(13;14)(q10;q10)) in homozygous form (44,XY,der(13;14;q10;q10) x 2). Two PGT cycles were performed, with unaffected embryos identified for transfer in each cycle. It is understood that no normal embryos can be formed in these cycles; thus, only balanced embryos could have been selected for transfer. Two such embryos were available for transfer in the first cycle and two in the second cycle; however, neither resulted in clinical pregnancy, presenting difficulty in interpreting possible effect of this complex rearrangement. Although application of the conversion technique to visualize chromosomes in single blastomeres improves the accuracy of the diagnosis by analysis of metaphase chromosomes, a high frequency of mosaicism in the cleavage-stage embryos still presents problems for diagnosis. Follow-up analysis of unbalanced embryos included those in which chromatid malsegregation or recombination had been identified by analysis of PB1 and subsequently tested at PB2 to infer a balanced or normal embryo. Mosaicism rate was 41%, and different cell lines were present, including normal or balanced. If investigated only by embryo biopsy, misdiagnosis could have

247

occurred. Therefore, PGT-SR strategy for maternally derived translocations may benefit from PB1 and PB2 testing, deferring nuclear conversion in embryonic cells until necessity. We have observed that unbalanced chromosome complements do not adversely affect the potential of an embryo to reach the blastocyst stage in extended culture. Of 250 unbalanced embryos cultured for 1 week, 78 (31%) reached blastocyst. Thus, a chromosomal rearrangement may not be lethal in preimplantation development but is eliminated either during implantation or postimplantation development [99]. This explains the extremely high spontaneous abortion rate in couples carrying such translocations. Our data are in agreement with previous reports suggesting as much as sixfold reduction of spontaneous abortions in PGT-SR cycles for translocations [100, 101]. The introduction of the current standard technologies, using next-generation technologies, is highly accurate, as shown by follow-up testing of embryos predicted to be abnormal and confirmatory prenatal diagnosis of the ongoing pregnancies following PGT-SR.  These also allow combination with PGT-A, further improving reproductive outcome of PGT-SR, as shown by increased implantation and pregnancy rates and reductions in spontaneous abortions. Of 893 PGT-SR cycles for translocations analyzed in our series, 331 were performed with the use of next-­ generation technologies that employed 24-­chromosome aneuploidy testing (Table 6.10). This resulted in 287 embryo transfers (86.7%), with the transfer of 337 balanced/normal/euploid embryos (1.1 average embryos per transfer), 191 (66.5%) pregnancies, and birth of 19 unaffected babies in 174 deliveries. This demonstrates the reduction of spontaneous abortion rates to as low as 8.9%, compared to 87.8% without PGT-SR, and increase of pregnancy rate to 66.5, compared to 11.5% without PGT-SR, following the transfer of practically a single embryos on the average. Thus, introduction of the next-generation technologies concomitant with aneuploidy testing leads to an almost twofold increase of pregnancy rate and fourfold reduction of spontaneous abortion rate.

248

6  Origin of Aneuploidy and Strategies Underlying Clinical Application of Preimplantation Genetic…

In conclusion, carriers of structural rearrangement carry extremely high risk for reproductive failure. These patients benefit from PGT-SR, and if of advanced maternal age almost obligatorily, only this approach is practiced. Otherwise, achieving pregnancy by natural conception may require more years than a patient has available for reproduction. PGT-SR permits such couples to establish unaffected pregnancies within a year, in contrast to an average of 5 years by natural cycle. Multiple unsuccessful attempts for a liveborn and prenatal genetic diagnosis and potential pregnancy termination in multiple ongoing pregnancies can be eschewed to pregnancy termination. The current technologies allow not only to insure the acceptable pregnancy outcome for carriers of structural rearrangement but also allow avoiding the balanced offspring, not to postpone the problem to the next generation.

References 1. Sherman SL, Peterson MB, Freeman SB, et  al. Nondisjunction of chromosome 21  in maternal meiosis I: evidence for a maternal age-dependent mechanism involving reduced recombination. Hum Mol Genet. 1994;3:1529–35. 2. Hassold T, Merril M, Adkins K, Freemen S, Sherman S. Recombination and maternal age-dependent nondisjunction: molecular studies of trisomy 16. Am J Hum Genet. 1995;57:867–74. 3. Peterson MB, Mikkelsen M.  Nondisjunction in trisomy 21: origin and mechanisms. Cytogenet Cell Genet. 2000;91:199–203. 4. Lamb NE, Freeman S, Savage-Austin A, et  al. Susceptible chiasmate configurations of chromosome 21 predispose to nondisjunction in both maternal meiosis I, and meiosis II.  Nat Genet. 1996;14:400–5. 5. Pellestor F, Andreo B, Armal F, Humeau C, Demaille J.  Mechanisms of non-disjunction in human female meiosis: the co-existence of two modes of malsegregation evidenced by the karyotyping of 1397 in-vitro unfertilized oocytes. Hum Reprod. 2002;17:2134–45. 6. Dyban A, Fredine M, Severova E, Cieslac J, Wolf G, Kuliev A, Verlinsky Y.  Detection of aneuploidy in human oocytes and corresponding first polar bodies using FISH. Seventh international conference on early prenatal diagnosis. Jerusalem; 1994 (Abstract #97). 7. Verlinsky Y, Cieslak J, Freidin M, et al. Pregnancies following pre-conception diagnosis of common

aneuploidies by fluorescent in-situ hybridization. Hum Reprod. 1995;10:1923–7. 8. Munné S, Daily T, Sultan KM, Grifo J, Cohen J. The use of first polar bodies for preimplantation diagnosis of aneuploidy. Hum Reprod. 1995;10:1014–120. 9. Dyban A, Fredine M, Severova E, et  al. Detection of aneuploidy in human oocytes and corresponding first polar bodies by FISH.  J Assist Reprod Genet. 1996;13:72–7. 10. Pujol A, Boiso I, Benet J, et  al. Analysis of nine chromosome probes in first polar bodies and metaphase II oocytes for the detection of aneuploidies. Eur J Hum Genet. 2003;11:325–36. 11. Verlinsky Y, Cieslak J, Ivakhnenko V, et al. Birth of healthy children after preimplantation diagnosis of common aneuploidies by polar body FISH analysis. Fertil Steril. 1996;66:126–9. 12. Verlinsky Y, Cieslak J, Ivakhnenko V, et  al. Preimplantation diagnosis of common aneuploidies by the first and second polar body FISH analysis. J Assist Reprod Genet. 1998;15:285–9. 13. Verlinsky Y, Cieslak J, Ivakhnenko V, et  al. Prepregnancy genetic testing for common age-­ related aneuploidies by polar body analysis. Genet Test. 1998;1:231–5. 14. Verlinsky Y, Cieslak J, Ivakhnenko V, et al. Prevention of age-related aneuploidies by polar body testing of oocytes. J Assist Reprod Genet. 1999;16:165–9. 15. Verlinsky Y, Cieslak J, Ivakhnenko V, et  al. Chromosomal abnormalities in the first and second polar body. Mol Cell Endocrinol. 2001;183:S47–9. 16. Verlinsky Y, Cieslak J, Kuliev A. High frequency of meiosis II aneuploidies in IVF patients of advanced maternal age. Reprod Technol. 2001;10:11–4. 17. Kuliev A, Zlatopolsky Z, Kirillova I, Spivakova J, Cieslak-Janzen G.  Meiosis errors in over 20,000 oocytes studied in the practice of preimplantation aneuploidy testing. Reprod Biomed Online. 2011;22:2–8. 18. Gianaroli L, Magli MC, Ferraretti AP.  The in  vivo and in vitro efficiency and efficacy of PGD for aneuploidy. Mol Cell Endocrinol. 2001;183:S13–8. 19. Munne S.  Preimplantation genetic diagnosis of numerical and structural chromosome abnormalities. Reprod Biomed Online. 2002;4:183–96. 20. Preimplantation Genetic Diagnosis International Society (PGDIS). Guidelines for good practice in PGD: program requirements and laboratory quality assurance. Reprod Biomed Online. 2008;16:134–47. 21. ESHRE Preimplantation Genetic Diagnosis (PGD) Consortium. Best practice guidelines for preimplantation genetic diagnosis/screening (PGD/PGS). Hum Reprod. 2011;26:14–46. 22. Preimplantation Genetic Diagnosis International Society (PGDIS). 10th international congress on preimplantation genetic diagnosis. Reprod Biomed Online. 2010;20:S1–42. 23. Van Blercom J, Davis P, Alexander S.  Differential mitochondrial distribution in human pronuclear embryos leads to disproportionate inheritance

References between blastomeres: relationship to microtubular organization, ATP content and competence. Hum Reprod. 2000;15:2621–33. 24. Magli C, Capoti A, Resta S, et al. Prolonged absence of meiotic spindles by birefringence imaging negatively affects normal fertilization and embryo development. Reprod Biomed Online. 2011;23:747–54. 25. Ebner T, Yaman C, Mose M, Sommergruber M, Feichtinger O, Tews G.  Prognostic value of first polar body morphology on fertilization rate and embryo quality in itracytoplasmic sperm injection. Hum Reprod. 2000;15:427–30. 26. Balaban B, Urman B, Isiklar A, Alatas C, Aksoy S, Mercan R. The effect of polar body morphology on embryo quality, implantation and pregnancy rates. Fertil Steril. 2001;76(Suppl1):S8. 27. Miller KF, Sinoway CE, Fly KL, Falcone T.  Fragmentation pf the polar body at the time of ICSI does not predict fertilization or early embryo development but may be associated with improved pregnancy and implantation. Fertil Steril. 2001;76(Suppl1):S201. 28. Verlinsky Y, Munne S, Cohen J, et al. Over a decade of preimplantation genetic diagnosis experience – a multi-center report. Fertil Steril. 2004;82:292–4. 29. Magli MC, Gianaroli L, Crippa A, Grugnetti C, Ruberti A, Ferraretti AP.  Causes of aneuploidy  – polar body based PGD.  Reprod Biomed Online. 2009;18(Suppl 3):S3. 30. Gianaroli L, Magli MC, Lappi M, Capoti A, Robles F, Ferraretti AP.  Preconception diagnosis. Reprod Biomed Online. 2009;18(Suppl 3):S5. 31. Fragouli E, Escalona E, Guttieres Mateo C, et  al. Comparative genomic hybridization of oocytes and first polar bodies from young donors. Reprod Biomed Online. 2009;19:228–37. 32. Hassold T, Hall H, Hunt P.  The origin of human aneuploidy: where we have been, where we are going. Hum Mol Genet. 2007;16:R203–8. 33. Lamb NE, Feingold E, Savage-Austin A, et  al. Characterization of susceptible chiasmate configurations that increase the risk for maternal nondisjunction of chromosome 21. Hum Mol Genet. 1997;6:1391–401. 34. Kuliev A, Cieslak J, Verlinsky Y. Frequency and distribution of chromosomal abnormalities in human oocytes. Cytogenet Genome Res. 2005;111:193–8. 35. Angel R.  First meiotic division nondisjunction in human oocytes. Am J Hum Genet. 1997;65:23–32. 36. Gutierrez-Mateo C, Benet J, Colls P, et  al. Aneuploidy study of human oocytes first polar body comparative genomic hybridization anf metaphase II fluorescence in situ hybridization analysis. Hum Reprod. 2004;19:2859–68. 37. Fragouli E, Alfarawati S, Katz-Jaffe M, et  al. Comprehensive chromosome screening of polar bodies and blastocysts from couples experiencing repeated implantation failure. Fertil Steril. 2009; https://doi.org/10.1016/j.fertnstert.

249 38. Geraedts J, Montag M, Magli C, et  al. Polar body array CGH for prediction of the status of the corresponding oocyte. Part I: clinical results. Hum Reprod. 2011;26:3172–80. 39. Magli C, Montag M, Koster M, et  al. Polar body array CGH for prediction of the status of the corresponding oocyte. Part II: technical aspects. Hum Reprod. 2011; https://doi.org/10.1093/humrep/ der295. 40. Gabriel AS, Thornhill AR, Ottolini CS, et al. Array comparative genomic hybridization on first polar bodies suggests that non-disjunction is not the predominant mechanism leading to aneuploidy in humans. J Med Genet. 2011;48:433–7. 41. Hunt P, LeMaraire R, Embury P, Sheean L, Mroz K.  Analysis of chromosome behaviour in intact mammalian oocytes: monitoring the segregation of a univalent chromosome during female meiosis. Hum Mol Genet. 1995;4:2007–12. 42. Capalbo A, Bono S, Spizzichino L, et al. Sequential comprehensive chromosome analysis on polar bodies, blastomeres and trophoblast: insights into female meiotic errors and chromosomal segregation in the preimplantation window of embryo development. Hum Reprod. 2013;28:509–18. 43. Colls P, Escudero T, Cekleniak N, Sadowy S, Cohen J, Munne S. Increased efficiency of preimplantation genetic diagnosis for aneuploidy by testing 12 chromosomes. Reprod Biomed Online. 2009;19:532–8. 44. Uher P, Baborova P, Kralickova M, Zech MH, Verlinsky Y, Zech N.  Non-informative results and monosomies in PGD: the importance of a third round of re-hybridization. Reprod Biomed Online. 2009;18:530–46. 45. Fragouli E, Alfarawati S, Spath K.  The origin and impact of embryonic aneuploidy. Hum Genet. 2013;132:1001–13. 46. Sherman SH, Freeman SB, Allen EG, Lamb NE.  Risk factors for nondisjunction of trisomy 21. Cytogenet Genome Res. 2005;11:273–80. 47. Battaglia DE, Goodwin P, Klein NA, Soules MR.  Influence of maternal age on meiotic spindle assembly in oocytes from naturally cycling women. Hum Reprod. 1996;11:2217–22. 48. Eichenlaub-Ritter U, Vogt E, Yiu H, Gosden R.  Spindles, mitochondria and redox potential in ageing oocytes. Reprod Biomed Online. 2002;5:117–24. 49. Chatzimeletiou K, Morrison EE, Prapas N, Prapas Y, Handyside AH. Spindle abnormalities in normally developing and arrested human preimplantation embryos in  vitro identified by confocal laser scanning microscopy. Hum Reprod. 2005;20:672–82. 50. Angel E, Antonarakis SE. Genomic imprinting and uniparental disomy in medicine: clinical and molecular aspects. New York: Willey Liss; 2002. 51. Herbert M. How and when do oocyte chromosomes fall apart during female ageing?. Reprod Biomed Online. 2019; 39(Suppl. 1):e8–e9.

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52. Handyside A, Montag M, Magli C, et  al. Multiple meiotic errors caused by predivision of chromatids in women of advanced maternal age undergoing in  vitro fertilization. Eur J Hum Genet. 2012;20:742–7. 53. Abu-Amero S, Monk D, Apostolidou S, Stanier P, Moore G.  Imprinted genes and their role in human fetal growth. Cytogenet Genome Res. 2006;113:262–70. 54. Nashmyth K, Peters JM, Uhlman F.  Splitting the chromosome: cutting the ties that bind sister chromatids. Science. 2000;288:1379–84. 55. Yuan L, Liu J, Hoja M, Wilbertz J, Nordqvist K, Hoog C. Female germ cell aneuploidy and embryo death in mice lacking the meiosis-specific protein SCP3. Science. 2002;296:1115–8. 56. Mummert S, Lobanenkov V, Feinberg AP. Association of chromosome arm 16q loss with loss of imprinting of insulin-like growth factorII in wilms tumor. Genes Chromosomes Cancer. ­ 2005;43:155–61. 57. Fisher JM, Harvey JF, Morton NE, Jacobs PA. Trisomy 18: studies of the parent and cell division of origin and effect of aberrant recombination on nondisjunction. Am J Hum Genet. 1996;56:669–75. 58. McCoy RC, Newnham LJ, Ottolini CS, et  al. Tripolar chromosome segregation drives the association between maternal genotype at variants spanning PLK4 and aneuploidy in human preimplantation embryos. Hum Mol Genet. 2018;27:2573–85. 59. Kim NH, Chung HM, Cha KY, Chung KS. Microtubule and microfilament organization in maturing human oocytes. Hum Reprod. 1998;13:2217–22. 60. Barrit J, Brenner C, Cohen J, Matt D. Mitochondrial DNA rearrangement in human oocytes and embryos. Mol Hum Reprod. 1999;5:927–33. 61. Perez G, Flaherty S, Barry M, Matthews C. Preliminary observations of polar body extrusion and pronuclear formation in human oocytes using timelapse video cinematography. Hum Reprod. 1997;12:532–41. 62. Kahraman S, Kumpete Y, Sertyel S, et al. Pronuclear scoring and chromosomal status of embryos in severe male infertility. Hum Reprod. 2002;17:3193–200. 63. Gianaroli L, Magli MC, Ferraretti AP, et  al. Pronuclear morphology and chromosomal abnormalities as scoring criteria for embryo selection. Fertil Steril. 2003;80:837–44. 64. DeBaun MR, Niemitz EL, Feinberg AP. Association of in  vitro fertilization with Beckwith-Wiedemann syndrome and epigenetic alterations of LIT1 and H19. Am J Hum Genet. 2003;72:156–60. 65. Gicquel C, Gaston V, Maldenbaum J, et al. In vitro fertilization may increase the risk of Beckwith-­ Wiedemann syndrome related to the abnormal imprinting of the KCNQ1OT gene. Am J Hum Genet. 2003;72:1338–41. 66. Maher ER, Brueton LA, Bowdin SC, et al. Beckwith-­ Wiedemann syndrome and assisted reproduction technology (ART). J Med Genet. 2003;40:62–4.

67. Niemitz EL, Feinberg AP. Epigenetics and assisted reproductive technology: a call for investigation. Am J Hum Genet. 2004;74:599–609. 68. Halliday J, Oke K, Breheny S, Algar E, Amor JA. Beckwith-Wiedemann syndrome and IVF: a case– control study. Am J Hum Genet. 2004;75:526–8. 69. Lucifero D, Chaillet JR, Trasler M.  Potential significance of genomic imprinting defects for reproduction and assisted reproductive technology. Hum Reprod Update. 2004;10:3–18. 70. PGDIS position statement on chromosome mosaicism and preimplantation aneuploidy testing at the blastocyst stage, 2016 PGDIS Newsletter, July 19, 2016 (www.pgdis.org). 71. Munne S, Sandalinas M, Escudero T, et  al. Some mosaic types increase with maternal age. Reprod Biomed Online. 2002;4:223–32. 72. Silber S, Sadowy S, Lehahan K, Kilani Z, Gianaroli L, Munne S.  High rate of chromosome mosaicism but not aneuploidy in embryos from karyotypically normal men requiring TESE. Reprod Biomed Online. 2002;4(Suppl 2):20. 73. Hornak M, Horak J, Kubichek D, et  al. The incidence and origin of chromosome aneuploidies in high quality karyomapping SNP profiles. Reprod Biomed Online. 2019;39:e24. 74. Nahuda G, Chen J, Butler R, et  al. Frequencies of chromosome specific mosaicism in trophectoderm biopsies detected by next generation sequencing. Fertil Steril. 2018;199:857–65. 75. PGDIS.  Position statement on transfer of mosaic embryos in preimplantation genetic testing for aneuploidy. Reprod Biomed Online. 2019;39:e1–4. 76. Popovich M, Dhaemens L, Thelman J, et  al. Expanding in vitro culture of human embryos demonstrated the complex nature of diagnosis chromosomal mosaicism from single trophectoderm biopsy. Hum Reprod. 2019:1–12. https://doi.org/10.1093/ humrep/dez012. 77. Girardi L, Romanelli V, Fabiani M, et al. Segmental aneuploidies show mosaic pattern predicting value compared to high whole chromosome aneuploidy representativeness. Reprod Biomed Online. 2019;39:e18–9. 78. Cram D.  Mosaicism and segmentals on POCs and prenatal diagnosis. Reprod Biomed Online. 2018;36(Suppl 1):e1–e42. 79. Babariya D, Fragouli E, Alfarawati S, Spath K. The incidence and origin of segmental aneuploidy in human oocytes and preimplantation embryos. Hum Reprod. 2017;32:2549–60. 80. Diez-Juan A, Rubio C, Rodrigo L, et al. Mitochondrial DNA content as a viability score in human euploid embryos: less is better. Fertil Steril. 2015;104:534–41. 81. Fragouli E, Spath K, Alfarawati SK, et  al. Altered levels of mitochondrial DNA are associated with female age, aneuploidy, and provide an independent measure of embryonic implantation potential. PLoS Genet. 2015;11:e1005241. https://doi.org/10.1371/ journal.pgen.1005241.

References 82. Victor AR, Brake AJ, Tyndall J, et al. Accurate quantitation of mitochondrial DNA reveals uniform levels in human blastocysts irrespective of ploidy, age, or implantation potential. Fertil Steril. 2017;107:34. e3–42.e3. 83. Çolakoğlu YK, Çetinkaya CP, Ünsal E, Çetinkaya M, Kahraman S.  Impact of a morphokinetic selection on the outcome of euploid embryo. Reprod Biomed Online. 2019;39:e22. 84. Laura Rienzi, Danilo Cimadomo, Arantxa Delgado, Maria Giulia Minasi, Gemma Fabozzi, Raquel del Gallego, Marta Stoppa, Jose Bellver, Adriano Giancani, Marga Esbert, Antonio Capalbo, Jose Remohì, Ermanno Greco, Filippo Maria Ubaldi, Marcos Meseguer, (2019) Time of morulation and trophectoderm quality are predictors of a live birth after euploid blastocyst transfer: a multicenter study. Fertility and Sterility 112 (6):1080–93.e1 85. Zaninovich N, Elemento O, Rozenwaks Z. Artificial intelligence: its applications in reproductive medicine and the assisted reproductive technologies. Fertil Steril. 2019;12:28–9. 86. Munne S, Morrison L, Fung J, et  al. Spontaneous abortions are reduced after preconception diagnosis of translocations. J Assit Reprod Genet. 1998;15:290–6. 87. Verlinsky Y, Kuliev A, editors. Preimplantation diagnosis of genetic disorders: a new technique for assisted reproduction. New York: Wiley Liss; 1993. 88. Verlinsky Y, Kuliev A.  Atlas of preimplantation genetic diagnosis. New  York/London: Parthenon; 2000. 89. Verlinsky Y, Evsikov S.  Karyotyping of human oocytes by chromosomal analysis of the second polar body. Mol Hum Reprod. 1999;5:89–95. 90. Verlinsky Y, Cieslak J, Evsikov S, Galat V, Kuliev A.  Nuclear transfer for full karyotyping and preimplantation diagnosis of translocations. Reprod Biomed Online. 2002;5:302–7. 91. Kuliev A, Cieslak-Jansen J, Zlatoposlsky Z, Kirilllova I, Illlevitch Y, Verlinsky Y. Conversion and non-conversion approach to preimplantation diagnosis for chromosomal rearrangements in 475 cycles. Reprod Biomed Online. 2010;21:93–9.

251 92. Verlinsky Y, Evsikov S.  A simplified and efficient method for obtaining metaphase chromosomes from individual human blastomeres. Fertil Steril. 1999;72:1–6. 93. Willadsen S, Levron J, Munne S, et al. Rapid visualization of metaphase chromosomes in single human blastomeres after fusion with in-vitro matured bovine eggs. Hum Reprod. 1999;14:470–4. 94. Shkumatov A, Kuznyetsov V, Cieslak J, et  al. Obtaining metaphase spreads from single blastomeres for PGD of chromosomal rearrangements. Reprod Biomed Online. 2007;14:498–503. 95. Treff NR, Tao X, Schileings W, Bergh PA, Scott RT, Levy B.  Use of single nucleotide polymorphism microarrays to distinguish between balanced and normal chromosomes in embryos from a translocation carrier. Fertil Steril. 2011;96:e58–65. 96. Treff NR, Thompson K, Rafizadeh M, et  al. SNP array-based analysis of unbalanced embryos as a reference to distinguish between balanced translocation carriers and normal blastocysts. J Assist Reprod Genet. 2016;38:1115–9. 97. Kuliev A, Zlatopolsky Z, Wang L, Yao Y, Cram D, Rechitsky S.  Evolution of PGD for translocations. Abstracts of 15th international conference on preimplantation genetics, Bologna, Italy, 2016. 98. Chow JFC, Cheng HH, Lau EYL, Yeung WSB, Ng EHY.  Selective transfer of euploid non-carrier embryos with the use of long-read sequencing in preimplantation genetic testing for reciprocal translocation. Reprod Biomed Online. 2019 (in press). 99. Evsikov S, Cieslak J, Verlinsky Y. Survival of unbalanced translocations to blastocyst stage. Fertil Steril. 2000;74:672–6. 100. Munné S, Sandalinas M, Escudero T.  Outcome of preimplantation genetic diagnosis of translocations. Fertil Steril. 2000;73:1209–18. 101. Fisher J, Escudero T, Chen S, et  al. Obstetric outcome of 100 cycles of PGD of translocations and other structural abnormalities. Reprod Biomed Online. 2002;4(Supplement 2):26.

7

Clinical Outcome of Preimplantation Genetic Testing

After three decades of application to clinical practice, PGT is no longer a research tool but an established procedure considered by patients as a realistic option for responsible reproduction without risk of having offspring with a genetic disorder. At-risk couples can achieve the desired family size with not much more difficulty than couples without known inherited risk. Available data of hundreds of thousands of PGT cycles indicate that the procedure is safe, accurate, and expanding, in particular for many monogenic disorders (PGT-M) never previously performed for traditional prenatal diagnosis. These include preimplantation gender determination for social reasons, late onset diseases with genetic predisposition, and preimplantation HLA typing. PGT is of special utility for at-risk couples who may otherwise not reproduce because of fear of an affected pregnancy leading to termination of a clinical pregnancy. Although the majority of PGT cycles are performed in the United States and Western Europe, increasing numbers are performed in Eastern Mediterranean and Asian countries. The majority of PGT cycles are performed for aneuploidy (PGT-A) and chromosomal rearrangements (PGT-SR), the ratio of PGT-A to PGT-M being, approximately 3:1.

© Springer Nature Switzerland AG 2020 A. Kuliev et al., Practical Preimplantation Genetic Testing, https://doi.org/10.1007/978-3-030-43157-0_7

7.1

Ovulation Stimulation for a PGT Cycle

This complex topic is covered elsewhere and is far beyond the scope of this volume. Still, comments on two areas are highly relevant. First, are there minimal criteria for ovulation stimulation for any ART cycle and for PGT in particular? Second, are there disorders for which ovulation stimulation protocols should differ compared to the general population of women undergoing ovulation stimulation?

7.1.1 Ovulation Stimulation and Aneuploidy Controlled ovulation ovarian  stimulation (COS) is essential for ART and PGT. Could this actually be deleterious with respect to oocyte development? Our experience shows that no justification exists to cancel a stimulation cycle solely because a minimal number of oocytes are not met. Also, age-adjusted aneuploidy rates are not altered by type of ovarian stimulation or number of oocytes recovered. Although numerous studies have been published, no further consensus was achieved. ESHRE and ASRM provide guidelines on requi-

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site requirements for ART or PGT centers but do not codify surgical instruments or are able to take into account proprietary details concerning culture media. Various ART registries exist, but none are available for PGT alone nor for PGT including neonatal outcome or complication of a stimulation cycle. Without taking into account these confounding variables, it is not possible to generalize conclusions. More recent reports have still not brought clarity. In a  study of 828 of  1122 IVF cycles no association was found between cumulative gonadotropin administration and aneuploidy [1]. Yet, in cycles stratified by stimulation not required past day 12, there was no association, whereas if stimulation was required past 12 days, there was an association. Some reports have found reduced aneuploidy with low stimulation, whereas others found higher aneuploidy in unstimulated cycles. Still, a reasonable ­ assumption is that ovulation stimulation is a safe and efficacious method to generate oocytes in women desiring ART and PGT [2].

7.1.2 Specific Disorder Characterized by Poor Ovulation Response A second area of uncertainty with respect to ovulation stimulation lies in women having a specific disorder characterized by poor ovulation response.

7  Clinical Outcome of Preimplantation Genetic Testing

ferent. The above is consistent with the 10–15% rate of premature ovarian insufficiency in carriers. Still, ovarian response reserve is usually sufficient.

7.1.2.2 Myotonic Dystrophy In this autosomal dominant disorder, females have been reported to have reduced ovarian reserve, reduced ovarian response, and lower-­ quality embryos [6, 7]. However, this was not confirmed [8]. It remains unclear whether women with myotonic dystrophy require a distinct approach during an ART stimulation cycle. The magnitude of gene expression most likely explains variability. 7.1.2.3 BRCA 1/2 As discussed in Chapters 1 and 4, PGT for adult-­ onset heritable cancers is increasing in utilization. Candidates may be known to have the mutation or may be offspring of an affected parent. In either case the goal is to select embryos to transfer. It was shown that the average number of oocytes per cycle is 6.5 oocytes in BRCA 1 carriers, 7.5 in BRCA 2 carriers, and 8.0 in controls [9], but no differences were also reported [10]. Although increased estrogens generated during stimulation potentially could extend a deleterious oncogenic effect, risks of cancer provocation seem low [11].

7.1.2.4 Carriers of Balanced Translocations 7.1.2.1 Fragile X Syndrome In Chap. 6 the magnitude of unbalanced embryos Females who are carriers of fragile X syndrome derived from female or male carriers was dis(FMR1) and have the full mutation (>200 CGG cussed at length. Overall, 75% of their embryos reports) usually do not show ovarian insuffi- are unbalanced. Poorer response in female carriciency. Thus, they respond to COS in the expected ers than in unaffected wives of male carriers was fashion. Paradoxically, premutation carriers (55– also reported [12]. In our experience female car199 CGG repeats) show ovarian insufficiency. riers had fewer oocytes (13.5 ± 3.5) than wives of Higher levels of FSH have thus been recom- male carriers (18.7  ±  6.6); fewer embryo were mended, especially in women having 80–99 also transferred (1.0 ± 0 vs 1.9 ± 0.5). However, repeats [3, 4]. A linear relationship has predict- this was not confirmed by others [13], while more ably been found for ovarian response. Fewer aggressive stimulation has been recommended to oocytes were retrieved in premutation (200 CGG) (14.1  ±  1.7) [5]. Although female carriers with a balanced Fertility and biopsy rates were, however, not dif- translocation showed disadvantageous outcomes,

7.2  Anomalies in ART/ PGT Pregnancies and in ART Pregnancies Without PGT

this is not necessarily because of ovulation stimulation. Other factors exist, in particular, the specific rearrangement. Irrespective, more oocytes/ embryos are needed than usual. Irrespective, multiple cycles may be required to batch embryos to reach a sufficient number of euploid embryos.

7.2

 nomalies in ART/ PGT A Pregnancies and in ART Pregnancies Without PGT

7.2.1 B  irth Defects in ART/PGT Pregnancies Risks are associated with any procedure and must be determined and balanced against benefit. In PGT risks potentially encompass two methodologies. First, do risks exist with ovulation ­stimulation and embryo culture, i.e., traditional IVF? Second, do risks exist due to embryo biopsy, which is unique to PGT but not occurring in ART without PGT? Clinical outcomes for PGT were evaluated in our center for 7126 PGT cycles and in 39 other centers (ESHRE PGD Consortium) for 15,885 PGT cycles [14]. These data were based on 4227 healthy children (1504 and 2723, respectively). Multiple gestations occurred in over one-third of cases. The congenital malformation rate was under 5%; half were major anomalies and the others minor. Rates were not different from general population prevalence. Similarly, no increase in anomalies was found in a systematic follow-up conducted at the world’s second largest PGT center [15]. Physical findings were recorded at birth and up to 2  months of age for 995 children born after PGT.  Comparison was made to 1507 children born after ICSI/ ART.  No differences were observed for preterm birth, mean birth weight, very low birth weight (12 months

7.3  Diagnostic Accuracy of PGT-M

of unprotected intercourse without conception) who achieve pregnancy without ART; the rate of birth defects is almost the same as that in couples requiring ART.  OR of 1.17 (95% CI 1.0–1.36) was reported for infertile couples who conceived naturally while awaiting ART [30]. OR of 1.29 was in offspring of subfertile women who conceived without ART (95% CI: 0.99–1.68). Women who previously had an ART but who conceived spontaneously in a later cycle also had offspring with increased birth defects (OR: 1.25; 95% CI: 1.01–1.56). At present, subfertility is generally accepted as the major factor in increased ART-related birth defects [31]. That is, the increased risk of birth defects in ART offspring reflects selection bias of subfertile couples, who without medical assistance would never achieve pregnancy.

7.2.5 G  enetic Counseling for Birth Defects in ART and PGT Data indicate ART pregnancies without PGT have a 30% increase in birth defects. That the reason is unclear is highly relevant to PGT.  The PGT-M group undergoing PGT is fertile and should not be at increased risk if subfertility is the underlying basis. If the increase is related to ovulation stimulation or embryo culture, an increased anomaly rate would be expected. The PGT-A group is most often not fertile; many in fact undergo PGT-A to achieve pregnancy. They could thus be at increased risk in either circumstance. Overall, one should be prepared to communicate pitfalls in data used to derive these opinions. Yet perfection cannot be achieved in experimental design because there is no ideal control group that cannot be constructed. It is useful to remind couples contemplating ART or PGT that the baseline anomaly rate in the general population is 2–3%. A 30% increase would constitute 3–4%. Both traditional IVF and ICSI/IVF show the same increased risk. No particular organ system seems disproportionately affected, except for genital anomalies with ICSI.

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7.3

Diagnostic Accuracy of PGT-M

In Chap. 4 the strategy and accuracy of PGT were covered and a representative spectrum of the 581 different disorders illustrated. Often the strategies necessary had to be devised expressly. De novo cases require single-sperm analysis or  sequential polar bodies (PB1 and PB2). Determination of haplotypes (mutant allele chromosome versus normal allele chromosome) is necessary in de novo cases. We have used these strategies in 6204 PGT-M cycles (Table 4.2). Both polar body (PB)-based approaches and embryo-based (blastocyst) approaches have been shown to be safe and accurate. We studied PGT accuracy in 9036 oocytes [32] tested by sequential first (PB1) and second (PB2) polar body removal; amplification efficiency was 97% with embryo transfer in 84.2% of cycles. Only two misdiagnoses occurred. One involved fragile X syndrome and one myotonic dystrophy. Both misdiagnoses was due to allele dropout (ADO), a known pitfall (Chap. 4). However, such an error is no longer expected to occur at the present time. In those two cases, insufficient number of linked markers were used to confirm haplotype and guard against ADO. Additional markers are now considered necessary for exclusion of ADO (Chaps. 3 and 4). Still, assuming that the same percent misdiagnoses occur in all 790 PB-based PGT transfer cycles, accuracy using this approach would be as high as 99.7% per transfer. The majority of our PGT cycles are no longer performed by polar body or cleavage-stage embryo biopsy. Still our fist evaluation of the outcome  of 2158 PGT cycless performed for 239 genetic conditions, yielding 735 (41.3%)  unaffected pregnancies and 688 healthy children [32], showed an extremely high accuracy rate of 99.4% per transfer. In addition to the above two misdiagnoses,  there were two more in blastomere based testing. Both misdiagnosis were due to undetected ADO, one in a cystic fibrosis (CF) and one in a beta-thalassemia case involving heterozygous embryos, erroneously diagnosed as unaffected carriers.

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A high accuracy rate was also reported in the world’s second large PGT-M experience [15]. The misdiagnosis rate in 1443 PGT cycles was 0.6%, one in myotonic dystrophy and three in Charcot–Marie–Tooth disease (CMT1A). Analogous to our  experience, these errors occurred while developing the linkage analysis strategy prior to actual DNA analysis of the embryo.

7.4

Diagnostic Accuracy and Predictive Value of PGT-A

More than half of preimplantation embryos are chromosomally abnormal from the onset. The meiotic and mitotic origin of chromosomal abnormalities was discussed in Sect. 6.1. Aneuploidy prevalence in oocytes and embryos is 25–90% depending on maternal age. Because euploid and aneuploid embryos are morphologically indistinguishable, the rationale underlying PGT-A has long been obvious. If implanted, aneuploid embryos are often lost during implantation (failed cycle) or lost post-implantation (miscarriage). There should thus be benefit in avoiding transfer of aneuploid embryos, improving pregnancy outcome in poor prognosis IVF patients and increasing the overall standard of medical practice. Given this, how accurate is a laboratory test result stating chromosomal complement of an embryo undergoing PGT-A? What is its predictive value? Prior to 2010 and development of 24-chromosome testing in embryos, predictive value of PGT for chromosome analysis could not be assessed because fewer than half of all chromosomes could be tested routinely. Transfer of an embryo aneuploid for other than chromosomes 13, 18, and 21 would not be known. This pitfall precluded determining positive or negative predictive value. Once array CGH and NGS were developed (2008–2010), all chromosomes could be evaluated. Further, culture to the blastocyst stage allowed 5–10 cells, facilitating PGT-A or PGT-M of high diagnostic accuracy [33].

7.4.1 Predictive Value of PGT-A Positive and negative prediction for PGT-A was reported in 2012 in 146 couples (mean maternal age 34.0  ±  4.4  years) [34]. A total of 255 embryos were biopsied and tested before transfer: 133 cleavage stage and 142 blastocyst stage. Aneuploidy testing was performed but results were not known at the time of transfer. Cleavage-­ stage embryos were scheduled for transfer on day 3, immediately after biopsy; embryos (blastocysts) scheduled for transfer on day 5 underwent first a laser-induced opening on day 3 and then trophectoderm biopsy on day 5. The embryo was transferred thereafter. The livebirth rate following transfer of 133 euploid embryos was 41% (positive predictive value of aneuploid embryo transfer). Of 99 embryos predicted to be aneuploid, 4 resulted in healthy euploid offspring. Although the 4% pregnancy rate (positive predictive value) was significantly lower than outcome of all embryos transferred (28%), the negative predictive value of 96% was less than the predicted 100%. Still, no errors occurred and accuracy was confirmed to be very high.

7.4.2 A  neuploidy and Error Origin of Chromosomal Abnormalities Neither origin of aneuploidy, chromosome involved, nor parental origin seems to alter predictive values. As discussed in Chap. 6, at least 95% of aneuploidies originate from errors in maternal meiosis; 5% originate from paternal meiosis or mitotic errors. Given aneuploidy predominantly of maternal origin, polar body analysis was applied initially to detect aneuploidies in embryos [32]. It was quickly realized that not all oocytes reach the embryo transfer stage; thus, selection occurs against a significant proportion of aneuploidies, some of which do not survive to become blastocysts. At present, PGT-A is based on NGS analysis of blastocysts. In 2922 embryos, 56.0% were aneuploid, com-

7.5  Mosaicism in PGT-A

prised of 13.0% monosomy, 13.0% trisomy, 8.0% numerical mosaicism, 14.0% segmental mosaicism, and 8.0% complex errors (Fig. 6.7). Comparing the spectrum of aneuploidies in oocytes and in embryos shows inconsistency between expected and observed frequencies of specific aneuploidies. Chromosomes 15, 16, 21, and 22 are relatively more frequently involved in female meiosis errors, resulting in trisomy more often than other autosomes. Relative frequency of these trisomies is followed by chromosomes 19 and 20; errors of the other chromosomes are less common [35]. Despite relative differences in chromosome-­specific aneuploidy rates, maternal age dependence is observed for every chromosome. Usually the rate nearly doubles between ages 35 and 43 years (Fig. 6.13), suggesting generalized disturbance of meiosis with advanced reproductive age [36]. The causes and spectrum of complex abnormalities are discussed in Chap. 6. Chromosome-specific errors of meiotic origin do not seem to alter predictive value of accuracy. Errors in chromosome 16 and 22 originate more frequently in meiosis II (44.4% and 41.5%, respectively) vs meiosis I (32.0% and 34.3%, respectively). Errors in chromosomes 13, 18, and 21 conversely occur more frequently in meiosis I (40.1%, 48.3%, and 41.4%, respectively) than meiosis II (36.3%, 34.6%, and 36.7%, respectively). The proportion of oocytes with errors of both meiosis I and meiosis II does not significantly differ among chromosomes, possibly except for chromosome 18 (Table 6.4).

7.5

Mosaicism in PGT-A

The greatest clinical confusion for PGT-A at present is related to mosaicism. This is of relatively recent realization, paradoxically the result of enhanced sensitivity of next-generation sequencing (NGS). Indeed, 24-chromosome array CGH a decade ago had been a great advance, enabling to show that predictive value for a euploid transfer was 41% [34]. Unrealized at the time, a salutary

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benefit of array CGH was that laboratory results were fundamentally dichotomized into whether a chromosome was euploid or aneuploid. With NGS, however, more than one cell line could be identified. The reason was that more or less than exactly 100% of expected quantity of DNA for a single chromosome would be evident, with NGS sensitivity much greater than array CGH. The rate of numerical mosaicism in our experience was 8.0%, and the rate of segmental mosaicism 14.0% (Fig. 6.7).

7.5.1 Numerical Mosaicism With NGS, results are based not on the number of individual cells, but on quantity of DNA. A blastocyst biopsy usually contains 5–10 cells. However, with both array CGH and NGS, the exact number of cells in a given biopsy is not known. Instead, laboratory results must be based on the quantity of normal (euploid) versus abnormal (aneuploid) DNA. For clinical purposes, the report would ideally be stratified into full euploid vs full aneuploid. However, quantity of DNA may not be exactly 0% aneuploid or 100% aneuploid. A range is thus given based on quantity of DNA serving as a surrogate for number of cells. An euploid result is considered 0–20% aneuploid DNA; an aneuploid result is 80–100%. Results that lie between full aneuploid and full euploid are defined as mosaicism. This could involve whole chromosomes (numerical) or partial deletions or duplications. It was shown that mosaic embryo transfer may result in successful pregnancies of mosaic embryos [37]. Trophectoderm biopsies showing whole-chromosome mosaicism – mosaic monosomic or mosaic trisomic – could result in liveborns. Presumably the (non-biopsied) inner cell mass from which embryonic organ differentiation arises was fully euploid. Thus, biopsied trophectoderm contained aneuploid cells, but only euploid cells were present in the inner cell mass. The Preimplantation Genetic Diagnosis International Society (PGDIS) recommends

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that mosaicism 20% or less aneuploid DNA be defined as euploid whereas if greater than 80% or more defined as aneuploid. Some centers may use different levels, e.g., ≤30 or ≥ 70%. There is no evidence for different chromosomes contributing disproportionately to the mosaicism observed and, hence, these do not differentially affect diagnostic accuracy of PGT-A. An NGS-­ based analysis from 1547 blastocysts [38] found 2.5% mosaicism per chromosome, with an overall mosaicism rate of 17.5%. When rates of chromosomes involved in whole-chromosome mosaicism and in segmental mosaicism were compared, disparate frequencies were found in only a few chromosomes. Overall, trisomy was more frequently detected as whole-chromosome mosaicism, whereas partial monosomy was more frequently detected as segmental mosaicism. Outcome following transfer of NGS-detected mosaic embryos [38–43] has been assessed clinically in 500 or more mosaic embryos. No live births reported have stated presence of anomalies, but anecdotal concerns exist. Ability of a mosaic embryo to result in a successful pregnancy depends on the proportion of euploid DNA.  A consistency of diagnosis comparing TE biopsy to ICM was demonstrated in 93 whole-chromosome trophectoderm-based aneuploid embryos donated for research [39]: in 93% of embryos, TE biopsy and the inner cell mass correlated. This also held with a second TE biopsy (98%). If most DNA at TE biopsy is chromosomally normal (e.g., 60–80% euploid), such an embryo is more likely to implant and result in a chromosomally normal live birth. That is, embryos having only 20–40% aneuploid DNA have a significant potential to reach term. By contrast, if aneuploid DNA constitutes 40–80% of trophectoderm DNA, the embryos more often fail to implant or are lost early in pregnancy. In such embryos ICM is more likely to show aneuploidy, whereas this is less likely for an embryo with 20–40% aneuploid DNA.

7  Clinical Outcome of Preimplantation Genetic Testing

7.5.2 Segmental Mosaicism Accuracy of PGT-A must take into account not only numerical chromosomal mosaicism but sub-­ chromosomal (segmental) mosaicism. In our experience totaling 5869 NGS-based PGT-A embryos, trophectoderm samples tested with NGS reveal deletions and duplications in all 24 chromosomes, some as small as 3 MB and others as large as 139 MB (Fig. 6.17). A total of 57% embryos were aneuploid; 14% of these had deletion/duplications, 62.0% only a deletion, and 30.0% only a duplication. A total of 8.0% had multiple sub-chromosomal variants, with mosaicism detected in 59% (Fig. 6.18). Larger chromosomes predictably had a higher proportion of deletions and duplications, excepting chromosome 6. Validation of segmental abnormalities was determined in the study of embryos donated for research that were retested [39]: segmental aneuploid cases were correlated in a second trophectoderm biopsy with the inner cell mass (ICM). Contrary to the over 90% consistency previously found in whole-chromosome aneuploidy, consistencies between ICM and trophectoderm segmentals occurred in only 47%. This suggests limited predictive value for segmental mosaicism. Nonetheless, mosaicism restricted to segmental variations may be warranted in the absence of euploid embryos. Comparing frequency of chromosome abnormalities between blastocysts and miscarriage samples can also be expected to help understand the clinical significance of mosaicism in preimplantation development  (see Chap. 6). Samples comprising 1454 blastocysts from PGT-A cycles revealed 711 euploid (49%) and 743 aneuploid (51%) embryos (see details in Chap. 6). The latter consisted of trisomies/monosomies (26%), segmental imbalances (44%), mosaicism (9%), and complex aneuploidies (21%) in addition to segmentals or mosaics. Copy number variation (CNV)-Seq analysis of 1810 miscarriage samples identifies 881 euploid (48.7%) and 929 aneuploid

7.5  Mosaicism in PGT-A

(51.3%) fetuses. Among fetal aneuploidies were trisomies (47.5%), monosomy X (9.9%), segmental imbalances (8.5%), mosaics (13%), polyploids (9%), and complex aneuploidies (12%). Comparing embryo and miscarriage data sets reveals that most segmental imbalances in embryos are not pathogenic. Those segmental imbalances associated with miscarriage presumably involve genes important for fetal development. By extrapolating from the type and frequency of aneuploidies differentially detected in embryos versus miscarriages, a significant number of embryos with segmental aneuploidies can be deduced to be lost in the early first trimester. Others that are not pathogenic may simply reflect DNA replication not yet completed in a given cell cycle. This would explain why larger chromosomes (e.g., 1, 2, 3) are more likely to show segmental mosaicism.

7.5.3 P  riority of Mosaic Embryos for Transfer Communicating to patients that one embryo is mosaic based on a percentage of DNA (and not actual number of cells) is initially disquieting, but cutoff values are now well established and codified. As stated, non-mosaic euploidy is diagnosed when the normal (euploid) DNA content is 80% or more (i.e., no more than 20% aneuploid DNA); non-mosaic aneuploidy is diagnosed when aneuploid DNA content is 80% or more (i.e., no more than 20% euploid DNA). Mosaicism is said to exist if a quantity of aneuploid or euploid DNA is between 21% and 79%. Observational follow-up of mosaic embryos has made clear that a high proportion of mosaic embryos have as noted developmental competence [41–43]. Although transfer of mosaic or mosaic segmental embryos can result in healthy pregnancies, reduced implantation and increased miscarriage rates do occur. Success is relatively good with 20–40% aneuploid DNA. However, transfer of mosaic embryos containing 40–80%

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aneuploid DNA is less likely to result in a liveborn. Even worse outcomes are achieved with the transfer of complex mosaics, namely, those in which more than one chromosome is involved. Finally, outcomes appear independent of the specific chromosomes involved. Several technical reasons could explain why a truly euploid embryo will not show 100% euploid DNA: 1. Method: Poor biopsy technique may lead to cell damage, partial destruction, and loss of cellular DNA. Physicians and clinics should appreciate that suboptimal biopsy technique can adversely affect subsequent analyses. If a consistently high level of mosaicism is identified in an embryo cohort stimulation cycle, consideration should be given to investigating the embryo biopsy method. If the biopsy is facilitated using a laser, contact points should be minimized and preferably made at cell junctions. 2. Analysis: Algorithms used for normalizing the chromosome mapping bins can potentially alter profiles, especially if bin counts used to normalize the profiles are variable or low. 3. Biases. This may occur in the library due to construction and starting DNA compromised in whole-genome amplifications. This could lead to under- or overrepresentation of whole chromosomes (numerical) or sub-­ chromosomal regions (segmental). Criteria for prioritization of embryo selection in the context of multiple mosaic embryos has been published by PGDIS [44]: 1. An euploid embryo should always be transferred rather than a mosaic embryo. 2. A second ovulation stimulation cycle is recommended. 3. If another cycle is not possible, a mosaic embryo can be transferred with appropriate counseling. This dilemma is not common, in approximately 1% of cycles a mosaic embryo being the only non-aneuploid option.

7  Clinical Outcome of Preimplantation Genetic Testing

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4. Transfer of a mosaic embryo should preferentially choose that with the lowest level of aneuploid DNA is preferred, namely, 20–40%. 5. Embryo mosaic for chromosomes associated with uniparental disomy should be avoided. Most commonly cited are chromosomes 6, 7, 11, 15, 19, and 20. However regions of imprinting exist in all chromosomes. 6. Prenatal diagnosis by amniocentesis or CVS is recommended. NIPT for 5–7 chromosomes or ultrasound does not suffice. Patients can be counseled that transfer of mosaic embryos appears to be a relatively safe option for couples, with low or minimal risk of adverse outcomes for the pregnancy. Irrespective, it is prudent not to state that no risk exists. Anecdotal reports can be expected to state otherwise.

7.6

Diagnostic Accuracy of PGT-SR

Reproductive outcome is very poor among carriers of structural rearrangements, as discussed in Sect. 6.8. This is the result of unfavorable meiotic segregation. Three-quarters of embryos are unbalanced. PGT-SR to exclude embryos with  unbalanced translocations  from transfer, is thus therefore, efficacious, in fact one of the first practiced applications of PGT.  From the onset in 1996, diagnostic accuracy has been high. Clinical and genetic outcome in PGT-SR varies depending on the type of translocations and parental origin. In our experience, reciprocal translocation segregation patterns for paternally and maternally derived reciprocal translocations show similar tendencies. Alternate segregation (35% and 34%, respectively) is more common than adjacent I segregation (28 and 34%, respectively). Least frequent is adjacent II segregation (9.1 and 11.4%). These meiotic outcomes explain the high proportion of unbalanced embryos derived from segregation in maternally derived reciprocal translocations  – 78%; only 22.4% of PGT-SR embryos are normal: 7.8% balanced, 6.9% normal, and 7.7% balanced/normal. Unbalanced embryos were also observed in 75%

of paternally derived reciprocal translocations, again leaving 25% embryos suitable for transfer: 7.1% balanced, 7.3% normal, and 10.6% balanced/normal. In Robertsonian translocations, embryos derived from maternally derived translocations show 64.4% unbalanced embryos, similar in percentage to reciprocal translocations. Of the 35.6% of embryos suitable for transfer, 13.2% were balanced, 10.8% normal cytogenetically, and 11.6% not tested to distinguish between balanced or normal cytogenetically. In embryos from paternally derived Robertsonian translocations, 53.4% were unbalanced; 46.6% were suitable for transfer (10.4% balanced, 7% normal, and 29.2% balanced/normal). Clinical pregnancies were produced in 37% of transfer cycles, 26% overall delivering healthy children. Initially breakpoint-specific FISH probes were used to distinguish unbalanced embryos from balanced embryos, including  those clinically normal and lacking a rearrangement (46,XX or 46,XY) and those clinically normal but having a balanced translocation. PGT-SR methods currently using array CGH or NGS or low-pass genome sequencing do not routinely distinguish between the notmal and balanced embryos,  because interpretation is based solely on quantity of DNA. Section 6.8 in Chap. 6 discusses our experience in using nuclear or chemical conversion technologies once utilized to distinguish cytogenetically normal from cytogenetically balanced embryos. Conversion technologies were effective but are rarely performed at present. Contemporary methods to distinguish between balanced cytogenetically normal are under investigation, namely, nanopore long-­read sequencing and mate-pair sequencing.

7.7

 linical Utility of PGT-A: C Randomized Clinical Trials Involving 24-Chromosome Aneuploid Testing

Clinical utility for the various types of preimplantation genetic testing has greatly advanced in the past decade. PGT-M and PGT-SR are now rarely questioned. PGT-A also has unequivocal

7.7  Clinical Utility of PGT-A: Randomized Clinical Trials Involving 24-Chromosome Aneuploid Testing

benefit, showing favorable benefit in selected indications. One reason for acceptance is that trophectoderm biopsy is technically easier and safer than removal of a single blastomere from an 8-cell cleavage-stage embryo or removal of polar bodies. The second reason is diagnostic accuracy proven with array CGH or NGS.  Ability to test all chromosomes is the diagnostic “game-­ changer” for PGT-A.  Prior application and guidelines governing questioning PGT-A to improve pregnancy rates are now obsolete because only 5–7 chromosome could be tested, whereas now the status of every chromosome is known. Two other advances concomitantly have greatly facilitated clinical utility: (1) Improved culture media enabled day 3 (cleavage-stage) embryos to remain in  vitro to day 5–6 (blastocyst stage). Blastocysts with over 100 cells were available, safely providing more cells at biopsy than the single blastomere from the 8-cell day 3 cleavage-­stage embryo can. (2) Cryopreservation methods allow biopsied embryos to be frozen subject to diagnosis results, to be thawed later without ostensible damage. Genetically normal embryos could then be transferred. Sequential manipulations were possible – polar body or cleavage-­stage biopsy followed by blastocyst biopsies, cryopreservation, thawing, and transfer. Embryo loss and damage appear to be surprisingly low in experienced hands, even if re-biopsy is necessary. Application of these technological advances alone contributed to PGT-A outcome, obvious already from observational study [45]. Outcome of 100 frozen embryo transfers tested by array CGH was reported. Trophectoderm biopsy was performed on day 5 or 6; inclusion criteria consisted of women ≥38 years, women having had recurrent miscarriages (≥2), or women having recurrent implantation failure (≥2 cycles). Most (86%) had normal ovarian reserve; 90% of male partners lacked evidence of infertility. Of 751 blastocysts, 47% were euploid. Of the 100 euploid blastocysts transferred, 97 survived for transfer. The biochemical pregnancy rate was 87%; clinical pregnancy rate was 73%, and livebirth rate was 71%. This study validated that 24-chromosome analysis was superior to cleavage-­stage biopsy using 5–7 chromosomes

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by FISH. This validation of analytical validity for PGT-A was followed by studies determining predictive value of transferring aneuploid or euploid embryos [34]. To reiterate, transferring a euploid transfer results in a liveborn in 41% of transfers; an aneuploid transfer results in 4% liveborns.

7.7.1 R  CTs Involving Polar Body Biopsy The first multicenter study using 24-­chromosome testing in PB1 and PB2 [46, 47] was conducted using array-based CGH analysis. Analysis was completed within 12 h on day 2 of preimplantation development, followed by fresh transfer. Accurate identification of the maternal contribution to chromosomal status of the zygote was achieved in more than 90% of cases; remaining cases showed amplification failure or a high signal-to-noise ratio. Follow-up testing of abnormal zygotes revealed 94% (130 of 138) concordance for the aneuploidies detected in PB1 and PB2. In one (1) of the eight aneuploid results, aneuploidy differed from prediction, whereas the remaining seven showed not aneuploid but euploid DNA.  Presumably euploid DNA was derived from PB2 or sperm. Although at least one zygote was predicted to be affected in 41 of 42 cycles, euploid zygotes were nonetheless available for transfer in 23  cycles. A total of 39 euploid embryos were transferred (1.6 per cycle on average), resulting in eight clinical pregnancies (33% per transfer) and seven unaffected children; implantation rate was 26% per embryo transferred. In conclusion, array CGH analysis of PB1 and PB2 for 24-chromosome testing was accurate, reliable, and feasible in sufficient time for fresh transfer.

7.7.2 RCTs Involving 24-Chromosome Testing of Blastocysts Since 2012 multiple RCTs have been conducted in blastocysts interrogated with 24-chromosome array or NGS, showing benefit, approximating a

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15–20% increase in pregnancy rates compared to embryos transferred solely based on morphological criteria. A significant benefit was observed in ages 35–39, but not universal in all age groups. The above observational and the predictive value studies set the stage for “gold standard” RCTs. The first RCT using 24-chromosome analysis was performed in a series of 112 women randomized into two groups [48]: transfer of a PGT-A embryo versus transfer of a morphologically normal embryo not biopsied or tested. A total of 425 blastocysts were biopsied and subjected to array CGH; 45% (191/425) were aneuploid. The control group not undergoing biopsy consisted of 389 blastocysts of normal morphology. Transfer of euploid embryos resulted in a 71% pregnancy rate versus 46% in non-biopsied controls (p > 0.017). In the other RCT, 72 cases of euploid embryo blastocysts were transferred, compared to 83 controls not subjected to PGT-A [49]. The sustained implantation rate in the former was 66%, compared to 48% in control cases that were morphologically normal but not tested for aneuploidy. Delivery rates were 85% versus 68%, respectively. Another RCT involved comparing two morphologically normal but untested embryos versus one embryo subjected to PGT-A [50]. Pregnancy rates were 61% for single embryo PGT-A and 65% for non-tested double embryo transfer. Twin rates were 0% versus 48%. Another RCTs involved cleavage-stage embryo biopsy [51]. In a meta-analysis of RCTs repeated at the time, significant benefit was shown in all [52]. Forrest plots were universally in the direction of benefit as well as in the eight reported observational studies [50, 53–59]. RCTs involving blastocysts and 24-­chromosome platforms thus clearly show outcomes preferable to cleavage biopsy and 5–7-chromosome FISH. Preimplantation genetic testing (PGT-A) using blastocysts and 24 chromosomes should now be considered standard practice for patients of advanced maternal age desiring PGT-A.  All patients undergoing ART should thus be informed of the option of PGT-­ A.  If PGT-M is undertaken, PGT-A should be considered nearly obligatory because there is no additional risk to the embryo. A single biopsy

7  Clinical Outcome of Preimplantation Genetic Testing

provides sufficient DNA for whole-genome amplification allowing both PGT-A and PGT-M. Additional studies will be needed to determine extent of clinical utility with respect to maternal age. What is the precise age range at which women would benefit? Extant data indicate optimal results lie in the 35–39 age quintile. For this quintile, 2014 SART cycles revealed a 52% pregnancy rate with PGT-A compared to 39% without PGT-A [60]. A comparable data set exists in the STAR trial [61], which was undertaken to determine whether universal (all ages) PGT-A was warranted to improve pregnancy rates. In the STAR trial, PGT-A was also significantly efficacious in the 35–39 quintile: 51% pregnancies with PGT-A compared to 37% without undergoing PGT-A. The rationale for universal PGT-A in other age groups is more arguable because the frequency of aneuploid embryos is lower below maternal age 35. The frequency in a 25-year-old woman is 1/476, compared to 1 in 204 for a 35-year-old and 1 in 65 for a 40-year-old. Universal PGT-A would indeed reduce frequency of liveborn trisomy and miscarriage to a lesser rate than in women a decade older. It is thus not surprising that in the STAR trial, with its limited sample size, universal benefit was not shown. A much larger sample would be needed to mitigate against less favorable power calculations given lower aneuploidy prevalence in younger women. Benefit of PGT-A to women over age 40 years in achieving pregnancy is also unclear. Of course, excluding chromosomal abnormalities, reducing miscarriage, and allowing single embryo transfer to avoid multiple gestation are major benefits and alone justify PGT-A.  However, damage from biopsy in a cohort of only 1 or 2 embryos in a woman aged 40–42 may outweigh benefit from transferring a known euploid embryo. Transferring the only two morphologically normal embryos without testing is not unreasonable, however, given there is low likelihood of multiple gestations because 75% or more embryos are expected to be aneuploid and unable to result in a liveborn trisomy. Thus rationale is defensible as well if a prior cycle similarly had few embryos and resulted in the pregnancy.

7.8  Ancillary Tests to Enhance PGT-A Pregnancy Rates

7.7.3 C  onfirming Clinical Utility of a PGT-A Verifying the chromosomal status of a miscarriage would be almost essential for quality control if readily achievable. If miscarriage occurred following a PGT-A transfer of a euploid embryo, this would be unexpected, albeit not rare. Approximately 5% of pregnancies undergoing PGT-A experience a miscarriage despite an ostensible euploid transfer. In such circumstances, the assumption has long been that the miscarriage was indeed euploid but due to nonchromosomal etiology (e.g., autoimmune disease or infection). Thus, no laboratory diagnostic error is assumed. This may or may not be correct but has been difficult to determine. Knowing the chromosomal status of such a miscarriage could, however, verify that the PGT-A result was indeed correct; miscarriage occurred for a nonchromosomal explanation. The converse would require attention to analytical validity. Guidelines by major organizations call for a karyotype or array CGH (chromosomal microarray) to determine cytogenetic status of miscarriages. The rationale is that future management requires stratification aneuploid recurrent miscarriages versus euploid recurrent miscarriages. Knowing this, however, is difficult because traditional analysis of products of conception required successful cell culture. However,  cultured cells are not required for chromosomal microarray (CMA), for which reason array CGH is now the preferred option. However, either cell culture or CMA requires recovery of products of conception, an unpleasant and not always successful task, because inadvertent inclusion of maternal tissue is a problem. Also,  patient compliance is low, and an informative result is obtained in only perhaps 50%. This impediment in obtaining informative results is expected soon to be mitigated through testing cell-free DNA in maternal plasma. Through venipuncture, studies of chromosomal status of any pregnancy ending in miscarriage can be determined cytogenetically, naturally including those following PGT-A. Cell-free DNA analysis in ongoing pregnancies is currently lim-

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ited to assessing liveborn trisomies (13, 18, 21), with the accuracy is high for 18 and 21 (98% and 99.9%, respectively) [62, 63]. The analytical approach is based on counting the total number of transcripts of a specific chromosome number in maternal plasma. One counts the total number of transcripts for all chromosomes  – maternal or fetal in origin. This is followed by comparing quantity of each individual chromosome. Total quantity of DNA is compared to that of a normal DNA reference standard. No attempt is made to separate maternal from fetal DNA transcripts. A small but quantifiable excess will be present if the embryo is trisomic. At present all chromosomes can actually be tested, but rare (non-liveborn) aneuploid trisomies are rarely sought and require different platforms. However, it is expected that routine cell-free DNA tests for all chromosome will soon be widespread. Cell-free DNA analysis for all pregnancies ending in miscarriage should be in fact even easier than doing so in ongoing pregnancies. The 4% fetal fraction required for cell-free DNA analysis of maternal blood in ongoing pregnancies [62, 63] will almost certainly be achieved in miscarriages because embryonic demise occurs 2–3 weeks before clinical manifestations of pregnancy loss. During this interval placental tissue is undergoing autolysis. Fetal DNA fragments released into the maternal circulation as a result of degradation result in a higher quantity of fetal DNA than in viable ongoing pregnancies.

7.8

Ancillary Tests to Enhance PGT-A Pregnancy Rates

PGT-A is contributing to ART pregnancy rates of 50–60%. Yet, increasing pregnancy rates further is still desirable. Various ancillary approaches provide additional information that could identify the optimal euploid embryo to transfer among an indistinguishable cohort of euploid embryos. Mitochondrial DNA  Mitochondrial DNA can be assessed from embryo biopsy. It is hypothesized that the lower the mtDNA copy number in a euploid embryo, the better the outcome. A

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“MitoScore” based on mtDNA copy number has thus been proposed. In one study quantity of mtDNA was determined in 205 blastomeres obtained from day 3 euploid cleavage-stage embryos and from 65 blastocysts [64]. Increased mtDNA in euploid embryos correlated inversely with implantation potential. A high mitoscore was, paradoxically, thought to reflect embryonic “crisis,” perhaps manifesting by reparative damage necessitated by need to cellular repair. Similar findings were made in another report based on 340 blastocysts and 39 cleavage-stage embryos [65]. However other studies have found no correlation of mitochondrial DNA content and ART success. If a mtDNA score was to be useful as a biomarker to distinguish among euploid blastocysts ostensibly suitable for transfer, pitfalls must be taken into account. In any preimplantation embryo, the total quantity of mtDNA is accumulated during oocyte maturation. Thereafter, the amount of mtDNA is unavoidably reduced with each successive cell division. Yet not all cells receive same amounts of mitochondrial DNA after a given cell division. Thus, a range of mtDNA content is observed in individual cells and, hence, trophectoderm biopsies. That is, stochastically unequal distribution of mtDNA occurs during preimplantation development. There is no guarantee of a single sample trophectoderm being representative of the copy number of the entire embryo, given that the “bottleneck” phenomena is without biological explanation. Section 6.7 provides additional discussion on experimental pitfalls that could impede mitochondrial DNA being a reliable biomarker, including our own data on the topic. Time-Lapse Cell Cycle Imaging  Likelihood of success for a euploid embryo could be judged prior to transfer based on the dynamics of embryo development [66]. This approach investigates the relative implantation potential among euploid embryos that are morphologically similar based on morphokinetic markers. Already classification exists (Gardner) for embryos of differing desirability. Cell cycle dynamics could potentially

7  Clinical Outcome of Preimplantation Genetic Testing

enhance transfer, based simply on cell cycle, which seems unlikely to be as accurate in excluding aneuploidy but morphokinetics in sync with PGT-A is of potential value in selecting among euploid embryos, as demonstrated in a recent collaborative study [67]. Further considerations are provided in Chap. 6.

Cell-Free DNA in Culture Media and Blastocoel Fluid  Consisting of DNA fragments, usually 150  bp and higher, cell-free DNA is as stated ubiquitous in blood (plasma). These fragments originate from breakdown of nuclear DNA derived from all organs. Discussed already is that fetal trisomy can be detected on the basis of an increase in quantity of all DNA fragments. Cell-free DNA technology may similarly be applicable for preselection of the optimal euploid embryo on the basis of quantifying amount or quality of embryonic DNA in blastocoel fluid or spent culture media. Cell-free DNA in blastocoel fluid and in culture media probably originates from cells damaged at biopsy or undergoing apoptosis during cell division. Assuming that cells/DNA recoverable do not preferentially reflect cells undergoing de-selection, as the result, should reflect inner cell mass in blastocoel biopsy, and, hence, developing embryo.  Spent culture media could likewise correlate with embryonic status. Several groups [68–76] have shown the feasibility of assessing blastocoel fluid and culture media. Current results are promising, but cell-free DNA is associated with several pitfalls. First, a requisite amount of embryonic DNA is required. The fraction of embryonic DNA in blastocoel fluid or spent culture media necessary for accurate diagnosis is not known but presumably should be comparable to 4% required in maternal plasma. Second, DNA in media or blastocoel fluid must correlate with that of the embryo itself (inner cell mass). The ICM cannot be biopsied and the ongoing pregnancy maintained; however, correlated 90% concordance of trophectoderm with spent culture media (19 of 21) was reported [77]. Concordance of ICM with trophectoderm

7.10  Why Did Cleavage-Stage RCTs Before 2010 Fail to Show Benefit of PGT-A?

267

(TE) was similarly 86% (18 of 21) in aneuploid embryos not transferred. Correlation of blastocyst “karyotype” was 77% (16/21) with culture media and also 77% (16/21) with trophectoderm. These comparisons were based on 42 vitrified blastocysts derived from 22 couples donating an aneuploid embryo and from 8 couples donating a euploid embryo. On the other hand, it was also reported that only 35% of blastocoel samples could be amplified for PGT-A and only 27% for PGT-M [72]. As for the use of cell-free DNA as an ancillary screen to help select the optimal euploid embryo, it  seems promising. Its use in lieu of embryo biopsy cannot be recommended at present because sensitivity is much less than the near 100% with PGT-A following embryo biopsy. In circumstances in which technical skills are lacking for embryo biopsy, 80% concordance could be preferable to the only 50% provided on the basis of a morphologically normal embryos not tested with PGT-A.

without knowledge of chromosomal status of prior miscarriages. Euploid loss accounts for half of half of miscarriages. The other half are aneuploid. Once 24-chromosome noninvasive testing (NIPT) based on cell-free DNA in maternal plasma is available, determining status of a miscarriage will doubtless be routine. This will determine whether recurrent losses would benefit from PGT-A.  This is feasible because recurrent miscarriages can be stratified into consecutive aneuploid embryos or consecutive euploid embryos. PGT-A is already offered for couples having experienced recurrent miscarriage. Much more targeted would be restricting PGT-A to those in which a prior miscarriage had been known to be aneuploid. If so, PGT-A would constitute unequivocal treatment. If prior losses were euploid, PGT-A would not be helpful. Widespread commercial availability of 24-chromosome NIPT can be expected in the near future [79, 80].

Artificial Intelligence  Other proposals for maximizing pregnancy rates of euploid embryos also exist. One might cumulatively construct a weighted series of odds ratios relating to factors of presumptive contributory value. In addition to “machine learning,” “artificial intelligence” has attractions because correlative factors can be hypothesis-free. AI could gather unsuspected correlations that constitute non-hypothesis-based software. That is, certain relationships might not be suspected by designers of software (i.e., humans). Using agnostic AI correlations, subjective interpretation can be avoided. At present, no AI approaches are routine, but intrigue exists [78].

7.10 W  hy Did Cleavage-Stage RCTs Before 2010 Fail to Show Benefit of PGT-A?

7.9

PGT-A for Recurrent Miscarriages

Miscarriages are significantly decreased if euploid rather than aneuploid embryos are transferred. This is seen clinically in outcome of PGT-A cycles or PGT-M cycles accompanied by PGT-A. Logically, value for PGT-A could exist in couples experiencing recurrent miscarriages but

Optimism currently expressed with respect to accuracy and clinical utility of PGT-A has not always existed. During 2007–2010, PGT-A was in some circles not considered beneficial. There were two understandable explanations. (1) Optimal embryo biopsy techniques (trophectoderm) were not utilized at that time, instead restricted to cleavage stage biopsy. (2) Diagnostic technology was limited, not interrogating all 24 chromosomes but only 5–7. To appreciate disparate opinions that arose and may still persist, let us consider the status of PGT-A before the current era, i.e., prior to 2010.

7.10.1 Biopsy Damage in Cleavage-­ Stage Embryos If not performed well, any operative procedure can have a detrimental effect. As discussed in Chap. 2, cleavage-stage biopsy especially

268

requires surgical prowess in order to obtain a single blastomere from a 6–8 cell embryo. Loss of two cells seems particularly deleterious [81]. Failure to observe a positive effect on reproductive outcome after PGT may thus merely reflect embryo damage, exacerbated by operator inexperience especially with cleavage-stage embryos [82–84]. Blastocyst biopsy now applied is technically easier and results in more (5–10) cells.

7.10.2 Observational Studies and RCTs Involving ­Cleavage-­ Stage Embryos That Showed Benefit PGT-A was introduced in 1994, and extensive observational data were reported over the next decade. In the early 2000s, these was increasing experience and reports of thousands of patients satisfied with PGT-A for achieving an ART pregnancy. In 2005 two large observational series [85, 86], each comprising over 500 couples compared outcomes before and after PGT-A in individual patients. These studies in aggregate showed an almost fivefold increase in implantation rate, threefold decrease in spontaneous abortion rate, and more than a twofold increase in “take-home” baby rates. Clinical benefit of PGT-A also seemed clear for patients experiencing prior poor reproductive performance. A 2008 report continued to show salutary results [87]. Investigators reporting these studies had sought to perform RCTs, but their patients were reluctant to be randomized. Governmental research funding for RCTs in the United States and elsewhere may not be possible if funds are proscribed for “embryo research.” This impediment exists in the United States due to National Institutes of Health regulations. Still, investigations in the private sector centers were able to perform smaller studies. In the United States, such an example study included 57 patients undergoing PGT-A for nine chromosomes (13, 15, 16, 17, 18, 21, 22, X, Y) by FISH, whereas in a comparison group, PGT-A was not done [88]. Pregnancy rates were increased in PGT-A compared to controls in

7  Clinical Outcome of Preimplantation Genetic Testing

women with recurrent pregnancy loss (7/11; 63.6% versus 3/8; 37.5%), advanced maternal age (3/7; 43% versus 3/12; 25%), or prior failed cycles (2/10; 2% versus 0/9; 0%). Sample sizes were obviously small but reasons for benefit existed.

7.10.3 RCTs in Cleavage-Stage Embryos Not Showing Benefit In 2007 a randomized study was reported from a center that had apparently only recently introduced PGT-A [89]. Their conclusions were that the RCT not only failed to demonstrate improved ART pregnancy rates but in fact was damaging. Given controversy and ensuing controversy generated from this report [57–59], discussion seems necessary even while contemporary technology has rendered the protocol no longer applicable. The other involved a single blastomere biopsyand FISH for 5-chromosome specific probes. The implantation rate in euploid PGT-A transfers was 16.8%, whereas in the control (non-biopsied) group, the rate was 14.7%. Thus, benefit would seem to exist (2.1% absolute increase). However, in a third group, biopsy was performed but there was no result. At that time an experienced laboratory would expect to have a “no results” in almost 5% of cases; however, in the RCT under discussion, the rate was as high as 20%. “No result” embryos were, however, still transferred; their implantation rate was only 6%, lower than both the 14.7% in the non-biopsied controls and 16.8% in biopsied and known euploid embryos. This difference (14.7% v. 6%) presumably reflected embryo damage. The “no result” transfer was considered obligatory to adhere to the “intention to treat” experimental design. Once assigned to a given ovum (PGT or control), intention to treat dictates that transfer is required even if a prior step (e.g., knowledge of chromosomal status) had not been accomplished. The no-result (6%) embryos were to be lumped with the 16.8% PGT-A euploid group. The aggregate success rate of 11.7% was then compared to the 14.7% non-­ biopsied controls. Only this comparison showed

References

269

lack of benefit. The authors concluded that PGT-A did not achieve benefit [90]. Despite methodological shortcomings cited above and despite criticism [82–84], the ASRM Practice Committee later in 2007 recommended transferring embryos without aneuploidy testing [91]. Lack of efficacy due to a limited number of testable chromosomes was surely one factor; damage of cleavage stage biopsy was likely another, accentuated by smaller ART units doubtless being less likely for an operator to gain experience in embryo biopsy. At present new programs contemplating initiation of PGT-A, but lacking extant operator biopsy experience, should take advantage of training options offered by PGDIS and other organizations. While developing requisite skills, centers should eschew participation in RCTs until requisite skills are gained. By 2010, these concerns had largely dissipated. Aneuploidy testing for all 24 chromosomes had become routine, trophectoderm biopsy had replaced cleavage-stage blastomere biopsy, and vitrification had allowed safe cryopreservation that could be followed by thawing.

66% liveborn rate with only 1% twins. Since 2014, the twin rate in the United States has decreased for the first time in four decades. Between 2014 and 2018, the multiple gestation rate decreased 10% for women aged 30–34  years, 23% for women aged 35–39, and 23% for women aged over 39 years [93]. Although not stratified by presence or absence of PGT-A in database presented, it is likely that this reduction occurred in those women whose age involved women most frequently undergoing PGT-A to achieve pregnancy. PGT-A with SET was surely a contributing factor. Irrespective of advocacy of PGT that results in (50–60%) pregnancy, the prevalent practice in many venues remains embryo selection based solely on morphologic criteria. This is not likely to persist. SET will probably become obligatory. PGT-A with transfer of a single euploid embryo to maximize pregnancy and minimize multiple gestation is favored by insurance carriers and public funders, who increasingly strive to restrict the number of transferred embryos.

7.11 U  niversal Single Embryo Transfer to Avoid Multiple Gestations

1. Sekhon J. The cumulative dose of gonadotropins used for controlled ovarian stimulation does not influence the odds of embryonic aneuploidy in patients with normal ovarian response. J Assist Reprod Genet. 2017;34:749–58. 2. Hong H, Franasiak JM, Werner MM, et al. Embryonic aneuploidy rates are equivalent in natural cycles and gonadotropin-stimulated cycles. Fertil Steril. 2019;112:670–876. 3. Sullivan AK, Marcus M, Epstein MP, Allen EG, Anido AE, Paquin JJ, Yadav-Shah M, Sherman SL.  Association of FMR1 repeat size with ovarian dysfunction. Hum Reprod. 2005;20:402–12. 4. Ennis S, Ward D, Murray A.  Nonlinear association between CGG repeat number and age of menopause in FMR1 premutation carriers. Eur Hum Genet. 2006;14:253–5. 5. Avraham S, Almog B, Reches A, Zakar L, Malcov M, Sokolov A, Alpern S, Azem F.  The ovarian response in fragile X patients and premutation carriers undergoing IVF-PGD: reappraisal. Hum Reprod. 2017;32:1508–11. 6. Feyereisen E, Steffann J, Romana S, Lelorc’h M, Ray P, Kerbrat V, Tachdjian G, Frydman R, Frydman N. Five years’ experience of preimplantation genetic diagnosis in the Parisian Center: outcome of the first 441 started cycles. Fertil Steril. 2007;87:60–73.

Transfer of a single euploid embryo is sufficiently efficacious than multiple or single embryo transfer (SET) withouit PGT-A. A randomized trial showed that SET with a single vitrified euploid embryo was as efficacious as transfer of two non-­tested embryos (i.e., two morphologically normal embryos not subjected to PGT-A) [50]. Pregnancy rates (>24 weeks) were 61% vs. 65% (RR 0.9), respectively. More importantly, twin rates were 0% versus 48%, the latter clearly unacceptably high. A non-PGT-A alternative to avoiding multiple gestations is transfer in successive cycles of a single embryo without PGT-A. The CDC states in a brochure that transfer of a single blastocyst results in a 51% “take-home baby” rate with 1% twins; double embryo transfer results in a 60% livebirth rate with 27% twins [92]. One (1) fresh SET followed in another cycle by one frozen SET results in

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271 single-nucleotide polymorphism microarray– based comprehensive chromosome screening in infertile patients. Fertil Steril. 2011;96:638–40. 46. Geraedts J, Montag M, Magli C, et  al. Polar body array CGH for prediction of the status of the corresponding oocyte. Part I: clinical results. Hum Reprod. 2011;26:3172–80. 47. Magli C, Montag M, Koster M, et al. Polar body array CGH for prediction of the status of the corresponding oocyte. Part II: technical aspects. Hum Reprod. 2011;26:3181–5. 48. Yang Z, et al. Selection of single blastocysts for fresh transfer via standard morphology assessment alone and with array CGH for good prognosis IVF patients: results from a randomized pilot study. Mol Cytogenet. 2012;5:24. https://doi.org/10.1186/1755-8166-5-24. 49. Scott RT, Tao X, Ferry KM, Treff NR. A prospective randomized controlled trial demonstrating significantly increased clinical pregnancy rates following 24 chromosome aneuploidy screening: biopsy on day 5 with fresh transfer. Fertil Steril. 2010;94(Suppl):S2.. -0–05 50. Forman EJ, Tao X, Ferry KM, Taylor D, Treff NR, Scott RT Jr. Single embryo transfer with comprehensive chromosome screening results in improved ongoing pregnancy rates and decreased miscarriage rates. Hum Reprod. 2012;27:1217–22. 51. Rubio C, Bellver J, Rodrigo L, Castillón G, Guillén A, Vidal C, Giles J, Ferrando M, Cabanillas S, Remohí J, Pellicer A, Simón C.  In vitro fertilization with preimplantation genetic diagnosis for aneuploidies in advanced maternal age: a randomized, controlled study. Fertil Steril. 2017;107:1122–9. 52. Dahdouh EM, Balayla J, García-Velasco JA. Comprehensive chromosome screening improves embryo selection: a meta-analysis. Fertil Steril. 2015;104:1503–12. 53. Greco E, Bono S, Ruberti A, Lobascio AM, Greco P, Biricik A, et  al. Comparative genomic hybridization selection of blastocysts for repeated implantation failure treatment: a pilot study. Biomed Res Int. 2014;2014:457913. 54. Schoolcraft WB, Fragouli E, Stevens J, Munne S, Katz-Jaffe MG, Wells D. Clinical application of comprehensive chromosomal screening at the blastocyst stage. Fertil Steril. 2010;94:1700–6. 55. Sher G, Keskintepe L, Keskintepe M, Maassarani G, Tortoriello D, Brody S.  Genetic analysis of human embryos by metaphase comparative genomic hybridization (mCGH) improves efficiency of IVF by increasing embryo implantation rate and reducing multiple pregnancies and spontaneous miscarriages. Fertil Steril. 2009;92:1886–94. 56. Keltz MD, Vega M, Sirota I, Lederman M, Moshier EL, Gonzales E, et al. Preimplantation genetic screening (PGS) with comparative genomic hybridization (CGH) following day 3 single cell blastomere biopsy markedly improves IVF outcomes while lowering multiple pregnancies and miscarriages. J Assist Reprod Genet. 2013;30:1333–9.

272 57. Lee HL, McCulloh DH, Hodes-Wertz B, Adler A, McCaffrey C, Grifo JA. In vitro fertilization with preimplantation genetic screening improves implantation and live birth in women age 40 through 43. J Assist Reprod Genet. 2015;32:435–44. 58. Feichtinger M, Stopp T, Gobl C, Feichtinger E, Vaccari E, Madel U, et  al. Increasing live birth rate by preimplantation genetic screening of pooled polar bodies using array comparative genomic hybridization. PLoS One. 2015;10:e0128317. 59. Fishel S, Craig A, Lynch C, Dowell K, Ndukwe G, Jenner L, et  al. Assessment of 19,803 paired chromosomes and clinical outcome from first 150 cycles using array CGH of the first polar body for embryo selection and transfer. J In Vitro Fert Embryo Transf. 2011;1:1–8. 60. Society for Assisted Reproductive Technology. SART national summary report: Final CSR for 2016. https:// www.sartcorsonline.com/rptCSR_PublicMultYear. aspx?reportingYear¼2016. Accessed May 6, 2019. 61. Munné S, Kaplan B, Frattarelli JL, Child T, Nakhuda G, Shamma FN, Silverberg K, Kalista T, Handyside AH, Katz-Jaffe M, Wells D, Gordon T, Stock-Myer S, Willman S, STAR Study Group. Preimplantation genetic testing for aneuploidy versus morphology as selection criteria for single frozen-thawed embryo transfer in good-prognosis patients: a multicenter randomized clinical trial. Fertil Steril. 2019;112:1071. https://doi.org/10.1016/j.fertnstert.2019.07.1346. [Epub ahead of print]. 62. Bianchi DW, Parker RL, Wentworth J, Madankumar R, Saffer C, Das AF, Craig JA, Chudova DI, Devers PL, Jones KW, Oliver K, Rava RP, Sehnert AJ, CARE Study Group. DNA sequencing versus standard prenatal aneuploidy screening. N Engl J Med. 2014;370:799–808. 63. Norton ME, Jacobsson B, Swamy GK, Laurent LC, Ranzini AC, Brar H, Tomlinson MW, Pereira L, Spitz JL, Hollemon D, Cuckle H, Musci TJ, Wapner RJ. Cell-free DNA analysis for noninvasive examination of trisomy. N Engl J Med. 2015;372:1589–97. 64. Diez-Juan A, Rubio C, Marin C, Martinez S, Al-Asmar N, Riboldi M, Díaz-Gimeno P, Valbuena D, Simón C.  Mitochondrial DNA content as a viability score in human euploid embryos: less is better. Fertil Steril. 2015;104:534–41. 65. Fragouli E, Spath K, Alfarawati S, Kaper F, Craig A, Michel CE, Kokocinski F, Cohen J, Munne S, Wells D.  Altered levels of mitochondrial DNA are associated with female age, aneuploidy, and provide an independent measure of embryonic implantation potential. PLoS Genet. 2015;11:e1005241. https:// doi.org/10.1371/journal.pgen.1005241. 66. Çolakoğlu YK, Çetinkaya CP, Ünsal E, Çetinkaya M, Kahraman S. Impact of a morphokinetic selection on the outcome of euploid embryo. Reprod Biomed Online. 2019;38(suppl 1):e28–9. 67. Lee CI, Chen CH, Huang CC, et al. Embryo morphokinetics is potentially associated with clinical outcomes of single-embryo transfers in preimplantation

7  Clinical Outcome of Preimplantation Genetic Testing genetic testing for aneuploidy cycles. Fertil Steril. 2019;39:569–79. 68. Shamonki MI, Jin H, Haimowitz Z, Liu L.  Proof of concept: preimplantation genetic screening without embryo biopsy through analysis of cell-free DNA in spent embryo culture media. Fertil Steril. 2016;106:1312–8. 69. Xu J, Fang R, Chen L, Chen D, Xiao JP, Yang W, Wang H, Song X, Ma T, Bo S, et  al. Noninvasive chromosome screening of human embryos by genome sequencing of embryo culture medium for in  vitro fertilization. Proc Natl Acad Sci U S A. 2016;113:11907–12. 70. Sanchez T, Seidler EA, Gardner DK, Needleman D, Sakkas D.  Will noninvasive methods surpass invasive for assessing gametes and embryos? Fertil Steril. 2017;108:730–7. 71. Vera-Rodriguez M, Diez-Juan A, Jimenez-Almazan J, Martinez S, Navarro R, Peinado V, Mercader A, Meseguer M, Blesa D, Moreno I, et  al. Origin and composition of cell-free DNA in spent medium from human embryo culture during preimplantation development. Hum Reprod. 2018;33:745–56. 72. Capalbo A, Romanelli V, Patassini C, Poli M, Girandi L, Giancani A, Stoppa M, Cimadomo D, Ubaldi FM, Rienzi L. Diagnostic efficacy of blastocoel fluid and spent media as sources of DNA for preimplantation genetic testing in standard clinical conditions. Fertil Steril. 2018;110:870–9. 73. Gianaroli L, Magli MC, Pomante A, Crivello AM, Cafueri G, Valerio M, Ferraretti AP.  Blastocentesis: a source of DNA for preimplantation genetic testing. Results from a pilot study. Fertil Steril. 2014;102:1692–9. 74. Magli MC, Pomante A, Cafueri G, Valerio M, Crippa A, Ferraretti AP, Gianaroli L.  Preimplantation genetic testing: polar bodies, blastomeres, trophectoderm cells, or blastocoelic fluid? Fertil Steril. 2016;105:676–83. 75. Palini S, Galluzzi L, De Stefani S, Bianchi M, Wells D, Magnani M, Bulletti C.  Genomic DNA in human blastocoele fluid. Reprod Biomed Online. 2013;26:603–10. 76. Li P, Song Z, Yao Y, Huang T, Mao R, Huang J, Ma Y, Dong X, Huang W, Huang J, et al. Preimplantation genetic screening with spent culture medium/ blastocoel fluid for in  vitro fertilization. Sci Rep. 2018;8:9275. 77. Tobler KJ, Zhao Y, Ross R, Benner AT, Xu X, Du L, Broman K, Thrift K, Brezina PR, Kearns WG.  Blastocoel fluid from differentiated blastocysts harbors embryonic genomic material capable of a whole-genome deoxyribonucleic acid amplification and comprehensive chromosome microarray analysis. Fertil Steril. 2015;104:418–25. 78. Jiao J, Shi B, Sagnelli M, Yang D, Yao Y, Li W, Shao L, Lu S, Li D, Wang X.  Minimally invasive preimplantation genetic testing using blastocyst culture medium. Hum Reprod. 2019;34:1369–79.

References 79. Miyagi Y, Habara T, Hirata R, Hayashi N. Feasibility of artificial intelligence for predicting live birth without aneuploidy from a blastocyst image. Reprod Med Biol. 2019;18:204–11. 80. Pertile MD, Halks-Miller M, Flowers N, Barbacioru C, Kinnings SL, Vavrek D, Seltzer WK, Bianchi DW.  Rare autosomal trisomies, revealed by maternal plasma DNA sequencing, suggest increased risk of feto-placental disease. Sci Transl Med. 2017;9:pii: eaan1240. https://doi.org/10.1126/scitranslmed. aan1240. 81. Fiorentino F, Bono S, Pizzuti F, Duca S, Polverari A, Faieta M, Baldi M, Diano L, Spinella F. The clinical utility of genome-wide non-invasive prenatal screening. Prenat Diagn. 2017;37:593–601. 82. Cohen J, Wells D, Munné S. Removal of 2 cells from cleavage stage embryos is likely to reduce the efficacy of chromosomal tests that are used to enhance implantation rates. Fertil Steril. 2007;87:496–503. 83. Munné S, Gianroli L, Tur-Kaspa I, Magli C, Sandalinas M, Grifo J, Cram D, Kahraman S, Verlinsky Y, Simpson J.  Substandard application of pre implantation genetic screening may interfere with its clinical success. Fertil Steril. 2007;88:781–4. 84. Cohen J, Grifo JA.  Multicentre trial of preimplantation genetic screening report in the New England Journal of Medicine: an in-depth look at the findings. Reprod Biomed Online. 2007;15:365–6. 85. Munné S, Cohen J, Simpson JL.  In vitro fertilization with preimplantation genetic screening. N Engl J Med. 2007;357:1769–70. 86. Gianaroli L, Magli MC, Ferraretti AP, Tabanelli C, Trengia V, Farfalli V, Cavallini G.  The beneficial

273 effects of preimplantation genetic diagnosis for aneuploidy support extensive clinical application. Reprod Biomed Online. 2005;10:633–40. 87. Kuliev A, Verlinsky Y.  Impact of preimplanta tion genetic diagnosis for chromosomal disorders on reproductive outcome. Reprod Biomed Online. 2008;16:9–10. 88. Kuliev A. Clinical and technical aspects of preimplantation genetic diagnosis. Expert Rev Obstet Gynecol. 2008;3:591–3. 89. Werlin L, Rodi I, DeCherney A, Marello E, Hill D, Munné S.  Preimplantation genetic diagnosis as both a therapeutic and diagnostic tool in assisted reproductive technology. Fertil Steril. 2003;80:467–8. 90. Mastenbroek S, Twisk M, van Echten-Arends J, Sikkema-Raddatz BS, Korevaar JC, Verhoeve HR, Vogel N, Arts E, de Vries J, Bossuyt PM, et al. In vitro fertilization with preimplantation genetic screening. N Engl J Med. 2007;357:9–17. 91. The Society for Assisted Reproduction and Practice Committee of the American Society for Reproductive Medicine. Preimplantation genetic testing: a practice committee opinion. Fertil Steril. 2007;88:1497–504. 92. Centers for Disease Control and Prevention. Having healthy babies one at a time: How many embryos should I transfer to have one baby? https://www. cdc.gov/art/pdf/patient-resources/Having-HealthyBabies-handout-1_508tagged.pdf. Accessed November 27, 2019. 93. Sunderam S, Kissin DM, Zhang Y, Folger SG, Boulet SL, Warner L, Callaghan WM, Barfield WD. Assisted reproductive technology surveillance  — United States, 2016. Surveill Summ. 2019;68(4):1–23.

8

Social, Ethical, and Legal Aspects

Preventive approaches to congenital disorders always raise ethical problems, because the natural emphasis should be on treatment rather than on the avoidance of birth of children with congenital disorders. Unfortunately, the former option is at present usually unavailable for most genetic disorders. The most ethically acceptable preventive approaches are those that involve primary preventive measures, which are better accepted by society than secondary preventive measures involving pregnancy termination. As described in Chap. 1, the paradigmatic primary preventive measure is population-based fortification of the major foodstuffs by folic acid containing multivitamins, which has significantly reduced neural tube defects and congenital malformations overall. These primary preventive measures are ethically acceptable in any population because they provide the actual gain in infants free of congenital malformations, rather than avoiding birth of affected children. Still such programs have been introduced only in a few populations. Lack of similar preventive measures in most communities may be the reason that thousands of children with congenital disorders continue to be born, when otherwise they might have been born healthy.

© Springer Nature Switzerland AG 2020 A. Kuliev et al., Practical Preimplantation Genetic Testing, https://doi.org/10.1007/978-3-030-43157-0_8

8.1

Legality and Regulation of PGT

Preimplantation genetic testing (PGT) is also a primary preventive measure albeit applied on a family level. PGT allows genetically disadvantaged couples to produce unaffected children of their own. Otherwise these children might not be born at all because of fear of these couples to reproduce, undergo prenatal diagnosis, and terminate pregnancy [1, 2]. Any legal restrictions of these patients’ choices may only force them to achieve their goal by traveling to other countries where the regulations regarding PGT are more liberal. The available reviews on the status of PGT in different countries [3–8] show that the international legal practices range from explicit legalization (Netherlands, United Kingdom, France, Spain) and “lawless control” (Belgium, United States) to legal prohibition through restrictive laws (Italy, Germany, Austria, Switzerland). However, even in these countries, there is a tendency to ease legal restrictions. For example, there is little interdiction of PGT in Austria, neither through the Law on Reproductive Medicine nor through the Law on Genetic Engineering (an exception might arise if the polar body or blastomere biopsy were misinterpreted as “interference in the germ cell lineage”); the latter is explicitly prohibited [7]. In France, PGT is under the control of CNMBRDP (Commission nationale de médecine 275

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et de biologie de la reproduction et du diagnostic prenatal), a governmental commission that also regulates IVF [4, 5]. Agreement is required to perform embryo biopsy and genetic testing, evolution of which and indications for which are subject to follow-up by representatives on a regular basis. Only a few centers in France are allowed to perform PGT during the current initial 5 years. These centers are subject to renewal afterward. Regulations also fall under the “ethical” law, requiring forthcoming reexamination. In the United Kingdom, PGT is regulated by HFEA, also a governmental organization. HFEA provides licenses for performing PGT and approves any new indication for PGT.  PGT-A had actually been practiced for more nearly 20  years in the United Kingdom and in many other countries but was officially allowed by HFEA only much later. HFEA initially refused to allow preimplantation HLA typing without PGT-M and then changed its position. In Italy, PGT and many aspects of IVF were forbidden by the Act of Parliament almost for 7  years. Only three oocytes were allowed to be aspirated for fertilization in vitro, clearly following reflecting the opinions of the hierarchy of the Roman Catholic Church [9]. Prior to the law, Italy had been among the most active ones involved in the development and application of PGT for genetic and chromosomal disorders [2]. After the laws, it was difficult to perform PGT.

8.2

 GT in Venues Having P Restrictions

PGT has sometimes become ethically acceptable and practiced in countries with restrictive laws. An example is Germany, where no manipulations are allowed after conception. Methods involving only preconception diagnosis were practiced, i.e., using polar body (PB) diagnosis. The German Embryo Protection Act states that a fertilized, viable ovum is already an embryo in legal sense [10]; thus, no manipulation is allowed that could potentially damage the embryo, despite that approximately 120,000 abortions are performed annually in Germany. Removing and examining

a blastomere during PGT would destroy the cell and thus could be punishable with a prison sentence for up to 3 years or a fine. That is, PGT is an undisputable violation of the Act, reasoning that the life of a human being (i.e., a biopsied cell) is destroyed. In these circumstances, however, PB testing avoids violation of this law because no embryo is formed prior to fusion of male and female pronuclei (see Chap. 2). This approach may also resolve the ethical issues impeding PGT in Austria, Switzerland (despite the recent changes of the law for embryo biopsy restrictions), Malta, and other predominantly Catholic countries [7]. The same holds for other countries where no preventative measures have been allowed on religious grounds. For example, the law has evolved recently in Switzerland, citizens voting to introduce PGT into clinical practice. There is room for utilization of preconception diagnosis in Muslim communities, where PGT at preconception stage may be more attractive than embryo biopsy and is currently acceptable as a more preferred option over traditional prenatal genetic diagnosis. Preconception testing is, of course, restricted to maternally derived genetic abnormalities, as there has not been progress in sperm duplication technology that could allow sperm testing for paternally derived abnormalities prior to fertilization (see Chap. 2). On the other hand, legal restrictions on PGT have stimulated progress in the development of PGT techniques. This has especially been observed in Italy. After introductions of restriction on IVF and PGT stating no more than three oocytes were allowed and extant PGT prohibited, PB analysis was introduced to test the mature ovum prior to fertilization. Avoiding use of oocytes after fertilization of the oocytes (PB II) was still precluded. PGT has become routine in the increasing number of countries (United States, Belgium) in which no strict governmental regulations for PGT exist. It is, therefore, not surprising that the largest experiences in PGT-M and PGT-A have been accumulated in these countries. Guidelines and standards for PGT practice have been developed by Preimplantation Genetic Diagnosis International Society (PGDIS) and ESHRE and

8.4  PGT for Disorders with Genetic Predisposition

may be followed to achieve the required standards of PGT [11, 12]. Regulations have also been developed by national scientific societies, for example, Japan where no active PGT program is currently available. Regulations for PGT were developed by the National Society of Obstetrics and Gynecology and by the Japan Society of Human Genetics [8].

8.3

Ethical Issues Unique to PGT

Ethical issues underlying PGT have been evolving together with the development of the PGT technology and expansion of PGT indications. PGT was initially applied only to preexisting conditions, the only goal being avoiding birth of children with genetic disorders. However, PGT bypassed traditional prenatal genetic diagnosis of ongoing pregnancies and termination of pregnancy by making prevention of genetic disorders more ethically acceptable to many. Some couples had experienced the repeated pregnancy terminations before having a normal child, but others could not accept prenatal diagnosis and termination of pregnancy at all. Thus, PGT became an important alternative: (1) providing at-risk couples the opportunity of either going through prenatal diagnosis and termination of pregnancy, or (2) controlling their pregnancy outcome by testing the oocytes or embryos before implantation to verify that any pregnancy is unaffected from the onset. Accordingly, informing or not informing genetically disadvantaged couples about PGT availability affects their possible choices, an important ethical and legal issue. This was especially important for conditions in which there was low likelihood or a normal embryo/pregnancy. As mentioned in Chap. 6, analysis of meiotic outcome in carriers of a balanced translocation often leaves little statistical chance of a normal outcome. Prenatal genetic diagnosis to identify a balanced or normal fetus results in carriers of translocations having more than 80% prospect of losing their pregnancy by spontaneous abortions. PGT for such couples is clearly the most realistic option of having unaffected children of their own.

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In Chap. 4 PGT was shown to provide practical implications for progress in quality of life, life expectancy, and reproduction by patients having genetic disorders. For example, the life expectancy in some cystic fibrosis (CF) patients differs little compared to genetically normal individuals; procreation is possible with children being normal. Similarly, with success in stem cell transplantation, children with thalassemia may literally be cured; so-called ex-thalassemic patients having undergone PGT avoid the 50% risk of producing their own thalassemic children. Also in Chap. 4, PGT was shown applicable for homozygous or double heterozygous affected individuals with CF, thalassemia, and phenylketonuria (PKU); all should have their own unaffected children using PGT-M. A concern is that their success by PGT could generate the feeling that extreme variations of the genotype are rejected by society; thus, couples could face a complex decision of transferring back the embryos with affected genotypes. Some couples may elect to transfer the embryos carrying affected genes, such as for deafness or achondroplasia. These parents might themselves be affected and for reasons of verisimilitude choose to conceive only a disabled child. PGT introduces an additional option for the patients with a genetic disorder who are facing such decisions [13].

8.4

 GT for Disorders with P Genetic Predisposition

An important breakthrough from the ethical and social point of view was the PGT introduction for the diseases with genetic predispositions, as opposed to certainty of manifestation. PGT makes it possible to avoid the transfer of the embryos carrying genes predisposing to certain common, often adult-onset, disorders. PGT had long been applied to chromosomal disorders, autosomal recessive metabolic disorders, and other single gene disorders whose age of onset was birth or early childhood. So PGT for adultonset disorders with genetic predisposition was controversial or even believed unacceptable in the context of prenatal diagnosis. Of course,

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diagnosis had been possible by chorionic villus sampling (CVS) or amniocentesis for genes predisposing to an early-­onset autosomal dominant (e.g., Alzheimer disease (AD) as described in Chap. 4). But such couples would have to make a concerning decision of ongoing pregnancy termination. Prenatal genetic diagnosis was generally justified not on the basis of genetic predisposition alone but to the dire clinical manifestation of the disease. PGT technology then allowed genetic testing of human eggs and embryos before pregnancy, therefore making it realistic to establish only potentially normal pregnancies without the disease with early- or late-onset disorder with genetic predisposition. No mutant genes would be transmitted. This strategy was applied for Huntington disease (HD), a late-onset disease, causing death within 15  years after manifestation. A “nondisclosure PGT” (Chap. 4) was often the best option for these couples. Asymptomatic individuals at risk of carrying HD could be offered PGT to test embryos without this individual ever being informed about the specific test results, i.e., whether they themselves were affected. For example, a grandparent of the embryo could be known to be affected but not necessarily their son or daughter who may or may not have the mutation. Prenatal genetic diagnosis for late-onset common disorders is also considered controversial because individuals with a given mutation may never manifest the phenotype during their whole life span. This holds whether there is or is not current prospect for treatment. However, prevalence for dominant late-onset diseases with genetic predisposition of interest (e.g., HD or AD) is, thus, usually very high. Prevention by PGT is the only practical management because there is 50% lifetime risk of passing this genetic predisposition to each offspring. So desire not to avoid this dilemma in future generations is high [14, 15]. In addition, the affected partner may not survive to see this child grow up [16]. The surviving parent would then be the sole support for a child growing up and taking responsibility for his or her own future. On the other hand, it can be

8  Social, Ethical, and Legal Aspects

argued that this is not much different from circumstances in which one parent develops cancer or is killed in an automobile accident. Still PGT is preferable to children without testing because each child has 50% chance of inheriting autosomal dominant late-onset disorders with genetic predisposition. It is true that novel approaches mitigating or treating the clinical manifestation of these disorders in those with a mutant gene may be discovered in the forthcoming decades and should not be excluded, but neither can this be assumed. Given the above, PGT for common late-onset genetic disorders provides a novel nontraditional option for patients who may wish to avoid the transmission of the mutant gene predisposing their potential children to a genetic disorder. For some patients this would be the only reason for undertaking ART, namely, establishing from the onset a pregnancy free from risk. Individuals of reproductive age should be informed about this indication for this emerging new technology of PGT-M.  Their choice is between directing their own reproductive outcome or ongoing pregnancy late with potential pregnancy termination. To many this will seem ethically more acceptable rather than not. Results of PGT for late-onset disorder are discussed in Chap. 4. In our experience PGT for heritable breast cancer is accounting for the majority of cycles of heritable cancers (see Chap. 4). Breast cancer is not infrequently caused by BRCA1 and BRCA2 genes. Individuals with BRCA1 mutation have a 70% likelihood of breast cancer by age 70 years so PGT for breast and ovarian cancer has been also allowed by HFEA. Like for other conditions with genetic predisposition, the use of, PGT for cancers depended on the specific gene or allele, clinical features, and e­ fficacy of treatment, age of onset, penetrance, and individual family history PGT for inherited cardiac diseases is also increasing in number. Usually there is no family history that presages prenatal diagnosis by CVS or amniocentesis. Preventive management that could not affect premature or sudden death is rarely offered. The cumulative experience of PGT for inherited cardiac diseases (Chap. 4)

8.5  PGT for HLA Prediction and Stem Cell Transplantation

showed the results of PGT for familial hypertrophic and dilated cardiomyopathy. With PGT, couples with cardiac disease predisposing genes can reproduce without fear of having offspring with these at-risk genes. Again, ethical concerns in relation to PGT for these common disorders are still evolving.

8.5

 GT for HLA Prediction P and Stem Cell Transplantation

An important ethical discussions concerning use of PGT relate to preimplantation HLA typing. PGT for this indication is usually done for the benefit of a potential recipient that is not the embryo itself [13, 17, 18]. This has evoked moral outrage to some, whereas others justify the action as saving a moribund child’s life from an inexorably severe disease or death. Most Americans are supportive of using PGT to ensure that a neonate having been selected by PGT provides an exact HLA match with stem cells or tissue to an older sibling [19]. Attitudes may be different depending on whether the genetic testing in the embryo is done or not. Preimplantation HLA typing is usually performed in combination with PGT to exclude a causative gene such as Fanconi anemia (FA) [20]. Selecting both an unaffected embryo and HLA match (3  in 16) appears more acceptable than preimplantation HLA typing as a sole purpose. An example of the latter is PGT to provide umbilical cord stem cells to treat another sibling having leukemia (for which risk is low) or sporadic Diamond–Blackfan anemia (DBA) [21]. Such parents were initially denied permission for preimplantation HLA typing in the United Kingdom – a factor in the moral discussion being the desire of parents to have another child. At present, preimplantation HLA typing as the sole indiscretion is now allowed in the United Kingdom. Some dilemmas associated with preimplantation HLA typing are related to the actual indications for preimplantation HLA testing and are similar to those related to stem cell transplanta-

279

tion. Indeed, preimplantation HLA typing has the objective of identifying an HLA-identical stem cell transplant, which is the key to achieving acceptable engraftment and survival in stem cell therapy. No doubt indications will be modified with progress in treatment of bone marrow disorders. With current success in the cure rate of these disorders, acute lymphoid leukemia (ALL) and acute myeloid leukemia (AML), by chemotherapy, it could be argued that PGT may no longer be justified as an indication in a sizeable proportion of patients, that may be curative by chemotherapy [22]. At present, however, all conditions for which bone marrow or cord blood stem cell transplantation is required remain justifiable for preimplantation HLA typing. Another important issue is the applicability of preimplantation HLA typing for sporadic conditions, such as DBA, for which there are also inherited forms. Excluding inherited forms would require PGT for the mutations involved, excluding transplantation of compatible stem cells that might contain the same mutation as had by the affected sibling [23]. For example, mutations causing DBA include one in the gene encoding ribosomal protein S19 on chromosome 9 and another in the gene mapped to chromosome 8. Additional causative (and transmittable genes) mutations may exist in other sporadic forms, thus requiring PGT [24]. Without information from PGT, couples would have to make a decision about the need for undertaking transplantation without taking into account risk of serious GVH incurred by if stem cell transplantation were incompatible. Therefore, all known mutations causing the disease should be excluded by detailed mutation testing in parents and affected children. This should also be confirmed by the ongoing follow-up studies of the HLA-matched children born after preimplantation HLA typing. Controversy exists concerning the written consent form, to be signed by parents for the embryo. A similar situation arises when parents sign a consent form for umbilical cord blood stem cell collection and storage. It could be argued that parents do not actually need a baby and have it so as a means to save its older sibling; thus, the baby would then become a commodity

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280

in some peoples’ eyes. However, parents usually claim plausibly that another child is desired anyway. Regardless, the decision belongs to the parents. When only umbilical cord blood stem cells are being collected for the “designer babies” at birth, no harm exists for the donor baby. However, the same strategy could be used for organ donation and risk would arise. With the progress in differentiation of cord blood stem cells into the other types of cells [25], such possibilities cannot be entirely excluded. Preimplantation HLA typing also allows avoiding many ethical issues of reproductive and therapeutic cloning, providing in the views of most a more ethically acceptable option of selecting an HLA-matched progeny, than obtaining custom-made embryonic stem cells following somatic nuclear transfer and cloning.

8.6

PGT for Sex Selection

Prenatal genetic diagnosis or PGT for sex selection has long been controversial. Yet sex selection is de facto practiced in much of the world, and sex selection is considered acceptable when applied for balancing in a second or subsequent child [26, 27]. In some countries (e.g., India, Jordan), PGT is legally used for sex balancing, and it seems to be also well justified [28, 29] as part of reproductive autonomy, privacy in reproductive decisionmaking, and the moral superiority of preimplantation selection over adoration for sex selection of clinical pregnancy [30, 31]. It can also be argued that PGT for gender determination reinforces existing sexism and expectation of conformity to stereotypical gender norms, thus being inconsistent with the ideal of parents having unconditional love for each child [32, 33]. Despite such opposition and the opinion of the American Society for Reproductive Medicine [26], American College of Obstetricians and Gynecologists [34], and HFEA [35] that the creation of embryos to select sex or enhance gender variety in the family is an inappropriate way to allocate medical resources, the reality is that PGT for this purpose is steadily increasing. Thousands of PGT cycles are conducted annually in the

United States alone [36]. Although the majority of cases were performed for medical reasons or together with PGT for genetic conditions (X-linked recessive disorders) or aneuploidies [37–39], increasing numbers are performed for nonmedical reasons [40–42]. For example, one study performed to investigate moral attitudes and beliefs of the couples pursuing PGT solely for the purpose related to sex selection showed that the motivations for requesting gender determination include a rationale desire to limit family size, yet have a balance in sex of offspring, and concerns about parental age and financial concerns [43]. Nonetheless, one of the main desires was stated – to achieve a gender-balanced family; it was also shown that most couples (78%) were seeking sex selection in order to have a boy [41, 42].

8.7

Conclusions

PGT raises many ethical issues that are not unique to prenatal genetic diagnosis or assisted reproduction [13]. A persistent criticism concerns the selection of the embryos to fulfill certain genetic criteria complemented by destruction for other reasons. Selection of a few embryos for transfer from approximately a dozen available after hyperstimulation is a routine practice of IVF, the remaining embryos being either frozen or discarded. Such embryo selection is usually done routinely and based on morphological criteria. Routinely one identifies embryos with the highest developmental potential. This is practically enhancement albeit based on non-specific criteria. On the other hand, PGT improves upon embryo selection, by applying genetic tests; morphologically normal embryos may be chromosomally abnormal and destined to be selected against during preimplantation and post-­ implantation development. As described in Chap. 6, approximately half of oocytes and embryos from women of advanced reproductive age are chromosomally abnormal, indicating that it might not be acceptable practice simply to select embryos solely on morphological grounds. In other words, the burgeoning advent of PGT is a natural evolution of assisted reproduction, allow-

References

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Index

A ABCD gene, 196 Abnormalities chromosomal, 5, 40, 167, 213, 216, 223 aneuploidy and error origin of, 258–259 genital, 256 meiotic and mitotic, 213–220 segmental, 227, 260 Abortions, 1, 13, 81, 216, 233, 242, 246, 247 autosomal monosomies in, 217 spontaneous, 13, 45, 81, 99, 100, 107, 108, 161, 172, 175, 206, 213, 216, 217, 233, 242, 246, 247, 268, 277 aCGH, 147, 229, 234, 235, 237–243 PGT-SR, 243, 244 Acrylamide gel electrophoresis, 101, 157, 162 Acute lymphoid leukemia (ALL), 192, 200, 203, 279 preimplantation human leukocyte antigens typing for, 200, 201, 203 Acute myeloid leukemia (AML), 279 ADO, see Allele dropout (ADO) Adrenoleukodystrophy, X-linked, 195 Adrenomyeloneuropathy, 196 Aicardi–Goutieres syndrome, 110 ALL, see Acute lymphoid leukemia (ALL) Allele dropout (ADO), 16, 96, 102, 257 in blastomeres, 31 of dominant mutant allele, 34 in mutation analysis, 35 in PCR analysis, 202 in PGT for monogenic disorders (PGT-M), 33 rates, 32 Allele-specific amplification failure, 31 α-thalassemia mutations, 81 PGT for monogenic disorders (PGT-M), 103 preimplantation genetic testing (PGT), 81, 82 Alzheimer disease, 143–146 AML, see Acute myeloid leukemia (AML) Amniocentesis, 2, 9, 26, 166, 170, 196, 208, 278 Amplification, DNA, 157, 162, 233 of blastomere, 120 of polar bodies, 108 multiplex, 169 © Springer Nature Switzerland AG 2020 A. Kuliev et al., Practical Preimplantation Genetic Testing, https://doi.org/10.1007/978-3-030-43157-0

Amyloid precursor protein (APP) gene, 143, 144, 146 Anemia congenital, 186 Diamond–Blackfan anemia (DBA), 36, 199, 202, 279 preimplantation human leukocyte antigens typing for, 200, 202 Fanconi anemia (FA), 183, 207, 279 sickle cell, 7, 16 Aneuploid embryos, 42, 45, 107, 167, 205, 218, 229, 258, 260, 264, 267 Aneuploid oocytes, 188, 214, 218, 220, 225, 259 Aneuploidy, 216, 280 chromosomal, 21, 39, 133, 229, 233, 239 error origin of chromosomal abnormalities, 258–259 identification of origin of, 224 meiotic, 214, 224 mitotic, 224 originating from meiosis I, 217 ovulation stimulation and, 253–254 testing, 100 Aneuploidy rescue mechanism, 219 Aneuploidy testing, 100, 200 ataxia–telangiectasia (AT), 192, 194 Omen syndrome (OMS), 192, 193 outcome of preimplantation human leukocyte antigens typing, 205 Angelman syndrome, 224 ART, see Assisted reproduction technology (ART) Artificial intelligence (AI), 232, 267 Assisted reproduction technology (ART), 214 birth defects in genetic counseling for, 257 pregnancies, 255–257 without preimplantation genetic testing (PGT), 255–256 offspring, birth defects, 256 single embryo transfer (SET) policy in, 183 stimulation cycle, 254 Ataxia–Telangiectasia (AT), 192, 193 with aneuploidy testing, 192, 194 Ataxia–telangiectasia (ATM) gene, 192 Atrophy, spinal muscular, 7, 82, 88, 89, 110, 149 Autonomic neuropathy, hereditary sensory, 82 283

284 Autosomal dominant conditions, 4, 88, 126, 147, 254 Charcot–Marie–Tooth (CMT) disease, 92–95 primary torsion dystonia (PTD), 88–92 Autosomal monosomy detection, 223–224 in spontaneous abortions, 217 Autosomal-recessive de novo mutations (DNM), 122 Autosomal recessive disorders, 5, 6, 75, 113, 146, 149, 183, 190 cystic fibrosis (CFTR), 81–82 familial dysautonomia (FD), 82–88 spinal muscular atrophy (SMA), 81–82 Autosomal trisomy syndromes, 213 Axenfeld–Rieger syndrome, 172 B Bacterial Artificial Chromosomes (BAC), 42 Beckwith–Wiedemann syndrome, 224 Benign mutant haplotype, 118, 123 Beta-globin gene, 17, 186, 187 mutations in, 32, 80, 81, 136 β-thalassemias, 7, 36, 183, 189, 190 PGT for monogenic disorders (PGT-M) for, 103 B glucocerebrosidase (GBD) genes, 110 Biopsied single cells, 31 Biopsy blastocyst, 14, 15, 21, 23–24, 26, 36, 42, 75, 120, 172, 176, 190, 217, 228, 229, 259, 263, 268 blastomere, 21, 31, 36, 38, 40, 75, 110, 138, 268, 269, 275 conventional trophectoderm, 23 damage in cleavage-stage embryos, 267–268 embryo, 21–23 trophectoderm (see Blastocyst biopsy) Birth defects in assisted reproduction technology (ART) offspring, 256 pregnancies, 255 without preimplantation genetic testing (PGT), 255–256 in offspring of subfertile couples, 256–257 Blastocoel fluid, 27 aspiration of, 26 cell-free DNA in culture medium and, 266–267 Blastocyst 24-chromosome testing of, 263–264 formation, 16, 21, 22 Blastocyst biopsy, 14, 15, 21, 36, 42, 75, 120, 172, 176, 190, 214, 217, 228, 229, 259, 268 for genetic and chromosomal disorders, 21 for preimplantation genetic testing for aneuploidy (PGT-A), 26–27 procedure, developments, 23–25 Blastomere allele dropout (ADO) in, 31 analysis, 18, 19, 91, 100, 110, 111, 138, 148, 149, 178, 245 DNA amplification of, 120 nuclear conversion to metaphase, 41

Index Blastomere biopsy, 21, 31, 36, 38, 40, 75, 110, 138, 268, 269, 275 Blood group incompatibility, 155–158 BlueGnome, 40, 42 Bone marrow transplantation, 186, 189, 203, 206, 209 for hemoglobinopathies, 206 BRCA 1/2, 254 BRCA1 gene, 34 BRCA1 mutation, 34, 131, 132, 133, 138, 139, 278 BRCA2 mutation, 131, 132, 133, 138, 139 Breast cancer, 34, 39, 126, 131–133, 138, 278 C Cancer, 126–131, 133–139, 192 breast, 34, 39, 126, 131–133, 138, 278 Capillary electrophoresis, 89, 158, 164, 194 Cardiac disease, 147–154 with genetic predisposition, 150 inherited, 147, 148, 155, 278 Cardioencephalomyopathy, 153, 154 Cardiomyopathy, 151 dilated, 149, 151, 279 hypertrophic, 148, 149, 152, 153, 279 Carrier screening, 5, 6–10 on PGT for monogenic disorders (PGT-M) uptake, 8 Causative homeobox gene, 163–164 CD40 ligand gene (CD40LG), 196 Cell-free DNA (cfDNA), 232 analysis in pregnancy, 265 in culture medium and blastocoel fluid, 266–267 Cell haploidization, somatic, 20–21, 203 Centromeric cohesin, 220 Centrosomal regulator PLK4, 223 cfDNA, see Cell-free DNA (cfDNA) CFTR gene, 9, 32, 81, 109 CFTR mutations, 84, 103, 106, 107, 108, 111 preimplantation genetic testing (PGT) for, 81 prevalence of, 81 Charcot–Marie–Tooth (CMT) disease, 92–95, 258 Chorionic villus sampling (CVS), 2, 9, 99, 100, 103, 278 Chromatid, 213, 215 abnormal segregation of, 216 normal segregation of, 216 Chromatid errors, 215, 216, 217 Chromatid malsegregation, 39, 215, 216, 247 Chromatid nondisjunction, 216 Chromosomal abnormalities, 5, 40, 167, 213, 216, 217, 218, 223 aneuploidy and error origin of, 258–259 Chromosomal aneuploidies, 21, 39, 133, 229, 233, 239 Chromosomal anomalies, in oocytes, 213 Chromosomal errors, 213, 217 Chromosomal nondisjunction, 213, 215, 217 Chromosomal rearrangements, 233–248 karyotyping of embryos via nuclear conversion, 234 mate-pair sequencing, 245 next-generation technologies, 237–242 PGT for structural rearrangements (PGT-SR)

Index developments to distinguishing noncarrier balanced embryos, 236, 242 reproductive outcome, 242, 244, 246–248 using next-generation technologies, 234–236 polar body approach, 233–234 segregation patterns, 246 24-Chromosomal aneuploidy testing, randomized controlled trials (RCTs) involving, 262–265 24-Chromosome noninvasive testing, 267 Chromosome (chromatid) segregation errors, in meiosis I, 215 Chromosome-specific errors, 222 of meiotic origin, 259 in oocytes and embryos, 233 Chromosome-specific meiotic error, 220–222 24-Chromosome testing of blastocysts, randomized controlled trials (RCTs), 263–264 Chronic granulomatous disease, 36, 123, 190, 208 Cleavage-stage embryos, 258 biopsy damage in, 267–268 Cleavage-stage randomized controlled trials, 267–269 CMT1A, see Charcot–Marie–Tooth disease (CMT1A) CMT disease, see Charcot–Marie–Tooth (CMT) Disease CNV, see Copy number variants (CNV); Copy number variation (CNV) CNV-Seq, see Copy number variation sequencing (CNV-Seq) Coagulopathy, 76–77 Col4A5 gene, X-linked de novo mutation in, 177 Compound heterozygous conditions, 101–107 Congenital anemia, 186 Congenital anomalies in liveborns, 167 preconception prevention of, 2–4 Congenital disorders, 275 genetic history and avoidance of, 4–5 prevention of, 1 Congenital heart disease (CHD), 3 Congenital immunodeficiency, preimplantation genetic testing (PGT) for, 190, 191 Congenital malformations, 1, 2, 14, 21, 158–167 infants free of, 275 multiple, 158 rate, 255 risk for offspring with, 166 Controlled ovarian stimulation (COS), 253, 254 Conventional trophectoderm biopsy, 23 Copy number variants (CNV), 7–8 Copy number variation (CNV), 260–261 Copy number variation sequencing (CNV-Seq), 228–229 COS, see Controlled ovarian stimulation (COS) Craniofacial disorders, PGT for monogenic disorders (PGT-M) for, 158, 160 Creutzfeldt–Jakob Disease, 145, 146 Crouzon syndrome, 166, 167 Cryopreservation, 263 of PGT for monogenic disorders (PGT-M) embryos, 280–281 Currarino syndrome (CS), 163–164, 165 Currarino triad, see Currarino syndrome (CS)

285 CVS, see Chorionic villus sampling (CVS) C3422Y mutation, 166 Cystic fibrosis (CF), 7, 81–82, 103–107, 277 copy number variants (CNV) in, 8 mutations, 104–106 D DBA, see Diamond–Blackfan anemia (DBA) Deafness, PGT for monogenic disorders (PGT-M) for, 170, 173 Delta F508 mutation, 33, 106, 107, 108, 110 De novo mutations (DNM), 37, 113–126 autosomal-recessive, 122 Diamond–Blackfan anemia (DBA), 36, 199, 202, 279 preimplantation human leukocyte antigens typing for, 200, 202 Dilated cardiomyopathy, 149, 151, 279 Disomy, uniparental, 42, 99, 100, 101, 166, 186, 199, 224–225 DNA cell-free DNA (see Cell-free DNA (cfDNA)) mitochondrial, 26, 229–232, 265–266 polymorphism, 213 replication, 261 delayed, 229 DNA amplification, 157, 162, 233 of blastomere, 120 multiplex, 169 of polar bodies, 108 DNM, see De novo mutations (DNM) Donor egg from sibling, PGT-HLA, 206, 207 Down syndrome, 1–2 Dynamic mutations, 96, 140, 167–172 Dystrophy myotonic, 35, 49, 168, 169, 254 X-linked adrenoleukodystrophy (X-ALD), 195 DYT1 gene, 88, 90, 92 DYT1 mutation, 89, 90, 92 E Early-onset neurodegenerative disorders, 146 Electrophoresis acrylamide gel, 101, 157, 162 capillary, 89, 158, 164, 194 Embryonic lineage and differentiation, 167 Embryos aneuploid, 42, 45, 107, 167, 205, 218, 229, 258, 260, 264, 267 biopsy, 21–23 blastocyst formation of, 16 blastomere analysis of, 18 chromosome-specific errors in, 233 chromosome-specific meiotic error origin and impact on, 220–222 cleavage-stage, 258 euploid, 14, 25, 47, 107, 113, 133, 176, 177, 205, 218, 223, 227 with higher potential for pregnancy, 229–233

286 Embryos (cont.) karyotyping of, 234 monosomies, 217 mosaic, 226, 227, 233 priority of, 261–262 noncarrier balanced, PGT-SR developments to distinguishing, 236, 242 preimplantation, 15, 20, 226, 233, 258 triploid, 42 vitrified, 23 EmbryoScope™, 232 Endocrine disorders, 169 PGT for monogenic disorders (PGT-M) for, 171 Euploid embryos, 14, 25, 47, 107, 113, 133, 176, 177, 205, 218, 223, 227 with higher potential for pregnancy, 229–233 Expand long template (ELT) kit, 35 F FA, see Fanconi anemia (FA) Fabry disease, 111 Familial dysautonomia (FD), 6, 82, 85 Familial hyperinsulinism, 172 FANCA, see Fanconi anemia complementation group A (FANCA) FANCG mutation, 195 Fanconi anemia (FA), 183, 207, 279 Fanconi anemia complementation group A (FANCA), 193, 194, 206 FD, see Familial dysautonomia (FD) Fertilization, 20, 22, 39, 87, 99, 216, 223, 229 rate, 16 in vitro fertilization (IVF), 2, 14, 27, 49, 130, 155, 178, 190, 214, 224, 256, 276, 280 FFI, see Flour fortification initiative (FFI) Fibroblast growth factor receptor 2 (FGFR2) gene, 166, 167 mutation in, 168 Fibrosis, cystic, 7, 8, 81–82, 103–107, 277 FISH, see Fluorescent in situ hybridization (FISH) Flour fortification initiative (FFI), 2 Fluorescent in situ hybridization (FISH), 13, 233, 234 analysis in preimplantation genetic testing for aneuploidy (PGT-A), 38–40 Fluorescent PCR (F-PCR), 32, 33, 35 Folate deficiency, 2–3 Folic acid (FA) food fortification, 3, 4 supplementation, 2–4 Food fortification folic acid, 3, 4 multivitamin, 2–4 Fortification, food, 2–4 F-PCR, see Fluorescent PCR (F-PCR) Fragile X syndrome, 7, 96, 254 FRM1, preimplantation genetic testing (PGT) for, 98 G Gaucher disease (GD), 6, 110 GD, see Gaucher disease (GD) G343D mutation, 196

Index Gel electrophoresis, acrylamide, 101, 157, 162 Genetic conditions, for preimplantation genetic testing (PGT), 50–74 Genetic counseling, 136, 176, 177 for birth defects in ART and PGT, 257 Genetic disorders, 1, 4, 7, 13, 36, 107, 109, 110, 113, 253, 275 late-onset, 278 prevention of, 178, 277 X-linked, 101 Genital abnormalities, 256 Genotyping human leukocyte antigens (HLA), 122, 123 KEL, 156 oocytes, 102 single-sperm, 161 German Embryo Protection Act, 276 Germline mutations, 131, 138, 139, 140 Gertsmann-Straussler-Scheinker syndrome (GSS) (prion disease), 148 Gestations, 195 multiple, 255, 264, 269 single embryo transfer to avoid multiple, 269 GJB2 gene, 170, 174 Gonadal mosaicism carriers, 125 Granulomatous disease, chronic, 36, 123, 190, 208 GSS, see Gerstmann–Straussler–Sheinker syndrome (GSS) GVH, 193 H Haploidization, somatic cell, 20–21, 203 Haplotypes benign mutant, 118, 123 maternal, 87, 90, 99, 102, 107, 113, 169, 172, 176, 198, 203 paternal, 36, 38, 82, 87, 90, 99, 102, 108, 118, 119, 122, 125, 156, 169, 203 HCM, see Hypertrophic cardiomyopathy (HCM) HD, see Huntington disease (HD) HDN, see Hemolytic disease of the newborn (HDN) Health-related implications genetic conditions with, 155 blood group incompatibility, 155–158 congenital malformations, 158–167 HED-ID, see Hypohidrotic ectodermal dysplasia with immunodeficiency (HED-ID) Hemangioblastomas, 133 Hematologic disorders, 76–77, 196 Hemoglobin disorder, 75 Hemoglobin electrophoresis, 7 Hemoglobin H disease, 81 Hemoglobinopathies, 6, 75, 183 bone marrow transplantation for, 206 clinical outcome preimplantation genetic testing (PGT) for, 79–80 Hemolytic disease of the newborn (HDN), 155 Hereditary sensory autonomic neuropathy, 82 Heterokaryons, 40, 234 Heterozygotes, 7, 8, 81 compound, 81, 101, 107, 110 detection rate for, 6

Index Heterozygous carriers, 81, 87, 103, 125, 193, 204 Heterozygous conditions, compound, 101–107 Hexosaminidase A (HEX A) gene, 17, 110 Hexosaminidase B (HEX B) gene, 17 HFEA, 276, 278, 280 HIGM, see Hyperimmunoglobulin M syndrome (HIGM) HLXB9 mutation, 163, 164, 165 Holoprosencephaly (HPE), 158, 161 Holt–Oram syndrome (HOS), 147 Homocysteine, 3 HOS, see Holt–Oram syndrome (HOS) HPE, see Holoprosencephaly (HPE) HTT gene, 144 Human Genome Project, 158 Human leukocyte antigens (HLA) genes, 186 haplotype analysis, 186, 187, 188, 189, 193, 194, 199, 203, 204, 207, 209 identical embryos, 205 polymorphic markers in, 187 prediction, 279–280 Huntington disease (HD), 133, 140, 143, 144, 145, 278 Hydrops fetalis, 81 Hyperhomocysteinemia, 2 Hyperimmunoglobulin M syndrome (HIGM), 196, 197 X-linked, 197 Hyperinsulinemia, 6, 170 Hyperinsulinism, familial, 172 Hypertrophic cardiomyopathy (HCM), 148, 149, 152, 153, 279 Hypohidrotic ectodermal dysplasia with immunodeficiency (HED-ID), 198 Hypospadias, 256 I ICM, see Inner cell mass (ICM) ICSI, see Intracytoplasmic sperm injection (ICSI) IKBKAP gene, 82, 85, 86 mutation in, 86 Immunodeficiency, 190–199 lethal X-linked, 197 Inherited cardiac disease, 147, 148, 155, 278 Inherited paternal R396C mutation, 192 Inner cell mass (ICM), 22, 23, 227, 228, 259, 260, 266 Intracytoplasmic sperm injection (ICSI), 15, 16, 17, 22 In vitro fertilization (IVF), 2, 14, 49, 130, 155, 178, 190, 214, 224, 256, 276, 280 preimplantation genetic testing (PGT) without, 27 K Karyotyping of embryos, 234 KEL gene, 157 Kell genotype, 156, 157 L Laser-assisted hatching, 23, 24 Laser-assisted trophectoderm biopsy, 24

287 conventional, 25 non-laser-assisted, 25 Late-onset common disorders with genetic predisposition, 126 breast cancer, 131–133 cancer, 126–131, 133–139 prenatal genetic diagnosis for, 278 Lethal X-linked immunodeficiency, 197 Leukemia, 193, 199, 279 acute lymphoid leukemia (ALL), 192, 200, 203, 279 preimplantation HLA typing for, 200, 201, 203 acute myeloid leukemia (AML), 279 childhood, 183 Limb reduction defects (LRD), global estimate of reduction of, 3 LMNA gene, 149, 151 Lymphocytes, 41, 119, 121, 161, 197, 198 T lymphocytes, 190, 197 Lymphoma, 192 M Machado–Joseph disease, 169, 170 Mate-pair sequencing (MPS), 236, 245 Maternal G542X mutation, 107 Maternal haplotypes, 87, 90, 99, 102, 107, 113, 169, 172, 176, 198, 203 Maternal mutation, 81, 86, 106, 153, 188, 192, 194, 196, 204, 209 Maternal plasma FANCG mutation analysis, 195 MED-ID, 198 Meiosis, 213 meiosis I aneuploidies originating from, 217 chromosome (chromatid) segregation errors in, 215 Meiosis errors meiosis I errors, 214, 215, 217 meiosis II errors, 214, 215, 216 mitotic errors in embryos in relation to, 222–223 Meiotic abnormalities, 213–220 Meiotic aneuploidies, 214, 224 Meiotic errors, 213 chromosome-specific, 220–222 during oocyte maturation, 216 Meiotic origin, chromosome-specific errors of, 259 Mendelian disorders, PGT for monogenic disorders (PGT-M) cycles for, 75 Methyl tetrahydrofolate reductase (MTHFR) mutation, 3 Microarray analysis, 40–42 Miscarriages, preimplantation genetic testing for aneuploidy (PGT-A) for recurrent, 267 Missense Leu39Pro mutation, 197 Missing chromatids, 216, 218, 219 Missing chromosomes, 213, 216, 218, 219 Mitochondrial DNA (mtDNA), 229–232, 265–266 Mitotic abnormalities, 213–220 Mitotic aneuploidies, 224 Mitotic errors, in embryos in relation to meiosis errors, 222–223

288 Monosomies autosomal detection, 223–224 in spontaneous abortions, 217 cleavage-stage, 217 in postimplantation embryos, 216 prevalence of, 217, 218 ratio, discordance, 223 Morphokinetic assessment, 232–233 Mosaic embryos, 226, 227, 233 for transfer, priority, 261–262 Mosaicism, 35, 247 and sub-chromosomal variations, 226–229 numerical, 259–260 in preimplantation genetic testing for aneuploidy (PGT-A), 259–262 segmental, 260–261 Mosaic sub-chromosomal variation detection, 228 Mother-Fetus Kell incompatibility, 155 MPS, see Mate-pair sequencing (MPS) mtDNA, see Mitochondrial DNA (mtDNA) Multiple gestations, 255, 264, 269 single embryo transfer to avoid, 269 Multiple malformation, PGT for monogenic disorders (PGT-M) for, 159 Multivitamin fortification programs, 2–4 Muscular atrophy, spinal, 7, 82, 88, 89, 110, 149 Mutant allele, 18, 164 allele dropout (ADO) of dominant, 34 Mutant haplotype, benign, 118, 123 Mutation(s) α-thalassemia, 81 in beta-globin gene, 32, 80, 81, 136 BRCA1 mutation, 34, 131, 132, 133, 138, 139, 278 BRCA2 mutation, 139 in CFTR gene, 84 C3422Y mutation, 166 CYS218STOP, 196 Delta F508 mutation, 33, 106, 107, 108, 110 de novo mutations (DNM), 37, 113–126 autosomal-recessive, 122 dynamic, 167–172 FANCG deletion, 195 in FGFR2 gene, 168 G343D, 196 germline, 131, 138, 139, 140 in IKBKAP gene, 86 inherited paternal R396C, 192 missense Leu39Pro, 197 MTHFR, 3 sonic hedgehog (SHH), 158 T1131A, 194 thalassemia, 78 V717L mutation, 143, 145, 146 See also specific types of mutation MYBPC3 gene, 149, 151 Myotonic dystrophy, 35, 49, 168, 169, 254 N Nanopore long-read sequencing, 242 NEMO gene, 198

Index Neural tube defects (NTD) global estimate of reduction of, 3, 4 prevention of, 2, 4 Neurodegenerative disease, 140 Neurodegenerative disorder, 169 early-onset, 146 Neurofibromatosis (NF), 130 NF type I (NF1), 130 NF type II (NF2), 131 Neurologic disease, 140 Neurologic disorders, with genetic predisposition, 141–142 Neuromuscular disorders, preimplantation genetic testing (PGT) for, 93–94 Neuropathy, hereditary sensory autonomic, 82 Next-generation sequencing (NGS), 42–45 monosomies, 221 PGT-SR, 243, 244 segmental abnormalities, 227 NF, see Neurofibromatosis (NF) NIPGT, see Noninvasive PGT (NIPGT) Noncarrier vs. balanced embryos, PGT-SR developments to distinguishing, 236, 242 Noninvasive PGT (NIPGT), 26–27 Non-laser-assisted blastocyst biopsy procedure, 24, 25 NTD, see Neural tube defects (NTD) Nuclear conversion, karyotyping of embryos via, 234 Nullisomies, 215, 216 Numerical mosaicism, 259–260 O Omen syndrome (OMS), 190, 192 with aneuploidy testing, 192, 193 Oocytes, 15, 186, 194, 196, 199, 204, 207, 209 aneuploid, 188, 214, 218, 220, 225, 259 chromosomal anomalies in, 213 chromosome-specific errors in, 233 genotyping, 102 maturation, 216, 229–230 PB2, 233 pronuclear-stage, 17 Ophtalmologic disorders, PGT for monogenic disorders (PGT-M) for, 175 Ornithine transcarbamylase (OTC) deficiency, 98 Ovulation response, 254 BRCA 1/2, 254 Ovarian stimulation and aneuploidy, 253–254 for preimplantation genetic testing (PGT) cycle, 253–255 P PAH gene, 101, 102 PAH mutation, 112 Panethnic screening, 7 Paternal haplotypes, 36, 38, 82, 87, 90, 99, 102, 108, 118, 119, 122, 125, 156, 169, 203 PB, see Polar body (PB) PCC, see Premature chromosome condensation (PCC)

Index PCR, DNA analysis to avoid misdiagnosis in single-cell, 31–35 PEGT, see Pre-embryonic genetic testing (PEGT) Pelizaeus–Merzbacher Disease (PMLD), 98–99 Personal Genome Machine (PGM), 45 PGH, see Preimplantation genetic haplotyping (PGH) PGM, see Personal Genome Machine (PGM) PGT-A, see Preimplantation genetic testing for aneuploidy (PGT-A) PGT for monogenic disorders (PGT-M), 5, 7, 8, 9, 31, 38, 196 adult-onset, 9 allele dropout (ADO) in, 33 for beta and alpha-thalassemia, 103 for craniofacial disorders, 158, 160 for deafness, 170, 173 diagnostic accuracy of, 257 embryos, cryopreservation, 280–281 for endocrine disorders, 171 for multiple malformation, 159 nondisclosure, 140, 143 for ophtalmologic disorders, 175 preimplantation HLA matching, 199–202 preimplantation HLA testing with and without, 184–185 for pulmonary disorders, 83 for renal disorders, 97 requests for cystic fibrosis (CFTR gene), 9 traditional indications and strategies for, 75–82 uptake, carrier screening, 8 PGT for structural rearrangements (PGT-SR), 233 developments to distinguishing noncarrier from balanced embryos, 236, 242 reproductive outcome of, 242, 246–248 translocations by aCGH and NGS, 243 using next-generation technologies, 234–236 PGT-HLA, see Preimplantation genetic testing for human leukocyte antigens (PGT-HLA) PGT of structural rearrangements (PGT-SR), 13, 39 clinical and genetic outcome in, 262 conversion of interphase nuclei to metaphase for, 41 diagnostic accuracy of, 262 PGT-SR, see PGT for structural rearrangements (PGT-SR) Phenylalanine hydroxylase (PAH) gene, 101 R408W mutation in, 102 Phenylketonuria (PKU), 101–103 PITX2 gene, 172, 176 PKU, see Phenylketonuria (PKU) Plasma homocysteine, 2 Pleuropulmonary blastoma, 126, 130 PLP1 gene, 99 PMLD, see Pelizaeus–Merzbacher Disease (PMLD) Polar body (PB), 213, 214, 215 array CGH analysis of, 219 biopsy, 15 randomized controlled trials (RCTs) involving, 263 sampling, 15–16 sperm analysis, 20–21 testing as preconception testing strategy, 16–20

289 Polymerization, actin, 197 Polymorphic markers, 80, 188, 190, 195, 204 in human leukocyte antigens (HLA) region, 187 Polymorphic short tandem repeat (STR) markers, 186 Pre-embryonic genetic testing (PEGT), 16, 18, 20 Pregnancy, 183, 186, 206, 208 cell-free DNA analysis in 265 euploid embryos with higher potential for, 229–233 plasma, cSMART analysis, 195 singleton, 196, 199, 204, 209 twin, 195, 207 Pregnancy loss, 26, 217, 228, 265 Pregnancy rates, preimplantation genetic testing for aneuploidy (PGT-A), 265–267 Preimplantation embryos, 15, 20, 226, 233, 258 Preimplantation Genetic Diagnosis International Society (PGDIS), 259–260, 276 Preimplantation genetic haplotyping (PGH), 36 Preimplantation genetic testing (PGT) application of, 13, 14 cycle, ovarian stimulation, 253–255 for disorders with genetic predisposition, 277–279 ethical issues unique to, 277 for FRM1, 98 genetic conditions for, 50–74 for human leukocyte antigens (HLA) prediction, 279–280 for inherited cardiac diseases, 278 legality and regulation of, 275–276 legal restrictions on, 276 for sex selection, 280 for stem cell transplantation, 279–280 without in vitro fertilization (IVF), 27 Preimplantation genetic testing for aneuploidy (PGT-A), 205–206, 213, 276 breakthroughs in, 40 clinical benefit of, 268 clinical utility of, 262–265 fluorescent in situ hybridization (FISH) analysis in, 38–40 mosaicism in, 259–262 mosaic sub-chromosomal variation detection, 228 predictive value of, 258 pregnancy rates, 265–267 for recurrent miscarriages, 267 segmental abnormalities in, 227 Preimplantation genetic testing for human leukocyte antigens (PGT-HLA) for acute lymphoid leukemia (ALL), 200, 203 combined with PGT for thalassemia, 188 for Diamond–Blackfan anemia (DBA), 200, 202 with donor egg from sibling, 206, 207 limitations and future prospects of, 202–206 matching, 183 without PGT for monogenic disorders (PGT-M), 199–202 practical implications of, 206–210 testing with and without PGT-M, 184–185 in thalassemic children, 188, 189 with and without aneuploidy testing, 205

290 Premature chromosome condensation (PCC), 39 Prenatal genetic diagnosis, for late-onset common disorders, 278 Primary torsion dystonia (PTD), 88–92 Prion diseases, 145, 148 PRNP gene, 146, 148 Pronuclear-stage oocytes, 17 PSEN1 gene, 147 PTD, see Primary torsion dystonia (PTD) Pulmonary disorders, PGT for monogenic disorders (PGT-M) for, 83 Pyloric stenosis, 2 Q Quantitative PCR (qPCR), 42 R Randomized controlled trials (RCTs) cleavage-stage, 267–269 involving 24-chromosome testing of blastocysts, 263–264 involving polar body biopsy, 263 Recessive disorders, X-linked, 280 Renal disorders inherited forms of, 172 PGT for monogenic disorders (PGT-M) for, 97 Restriction fragment length polymorphism (RFLP), 101 RFLP, see Restriction fragment length polymorphism (RFLP) Rhesus disease (RhD), 155, 157, 158 Robertsonian translocations, 234, 246, 247, 262 R408W mutation, in phenylalanine hydroxylase gene, 102 S Sandhoff disease (SHD), 16, 17, 19 Sanger sequencing, 236 SCID, see Severe congenital immunodeficiency (SCID) Segmental abnormalities, 227, 260 Segmental imbalances, 261 Segmental mosaicism, 260–261 SET, see Single embryo transfer (SET) Severe congenital immunodeficiency (SCID), 190, 199 Sex selection, preimplantation genetic testing (PGT) for, 280 SHD, see Sandhoff disease (SHD) SHH mutation, see Sonic hedgehog (SHH) mutation Short tandem repeats (STRs), 32, 108, 156, 188, 189, 209 markers, 16–17 polymorphic, 186 Sibling human leukocyte antigens-identical, 207, 208 PGT-HLA with donor egg from, 206, 207 Sickle cell anemia, 6, 7, 16 Single-cell allele dropout (ADO) rates in, 31 genetic analysis, 31

Index Single-cell PCR, DNA analysis to avoid misdiagnosis in, 31–35 Single embryo transfer (SET) to avoid multiple gestations, 269 policy in assisted reproduction technology (ART), 183 Single-gene disorder, 5 concomitant PGT for, 107–113 Single nucleotide polymorphisms (SNPs), 32 Single-sperm genotyping, 161 Single-sperm testing, 87 Single-sperm typing, 118 SMA, see Spinal muscular atrophy (SMA) SMN1 gene, 88, 111 SNPs, see Single nucleotide polymorphisms (SNPs) Somatic cell haploidization, 20–21, 203 Sonic hedgehog (SHH) mutation, 158–163 Southern blot analysis, 196 S-period premature chromosome condensation (S-PCC), 234 Sperm analysis, 20–21 genotyping, 20 Spinal muscular atrophy (SMA), 7, 82, 88, 89, 110, 149 Stem cells, 186, 189 human leukocyte antigens-identical sibling donors of, 208 Stem cells transplantation, 103, 183, 190, 195, 208 preimplantation genetic testing (PGT) for, 279–280 Stenosis, pyloric, 2 STRs, see Short tandem repeats (STRs) Super Plex Single Cell Whole Genome Amplification Kit, 42 Survival motor neuron 1 (SMN1), 7 T T1131A mutation, 194 Tay-Sachs disease, 6 Testicular sperm extraction (TESE), 226 Thalassemia, 103, 183, 186–189, 209 α-thalassemia mutations, 81 PGT for monogenic disorders (PGT-M) for, 103 preimplantation genetic testing (PGT) for, 81, 82 β-thalassemias, 7, 183, 189, 190 PGT for monogenic disorders (PGT-M) for, 103 maternal recombination detected in, 203, 204 preimplantation HLA matching combined with PGT for, 188 3D-PZD, 21 Time-lapse cell cycle imaging, 266 T lymphocytes, 190, 197 Transplantation bone marrow, 186, 189, 203, 206, 209 stem cells, 103, 183, 190, 195, 208 Triploid embryos, 42 Trisomic chromosomes, 221 Trisomies age-related increase of, 213 autosomal trisomy syndromes, 213 ratio, discordance, 223

Index Trisomy rescue mechanism, 219, 225, 226 Trophectoderm biopsy, 259, 263 conventional, 23 procedure, 24–25 Trophectoderm cells, 23 U Uniparental disomy (UPD), 42, 99, 100, 101, 166, 186, 199, 224–225 V Variants of unknown significance (VUS), 174–178 VeriSeq™ PGT Kit, 45 Very long chain fatty acids (VLCFA), 196 Vitrified embryos, 23 V717L mutation, 143, 145, 146 Von Hippel–Lindau (VHL) syndrome, 133 Von Recklinghausen disease, 131 VUS, see Variants of unknown significance (VUS)

291 W WAS gene, 197 Whole-genome amplification (WGA), 26, 36, 42, 89, 190 Wiskott–Aldrich syndrome (WAS), 197 X X-linked adrenoleukodystrophy (X-ALD), 195 X-linked Alport disease, 172 X-linked de novo mutation, in Col4A5 gene, 177 X-linked disorder, 7, 95–98, 101 Fragile X Syndrome (FMR1), 96–98 ornithine transcarbamylase (OTC) deficiency, 98 Pelizaeus–Merzbacher disease (PMLD), 98–101 X-linked genetic disorders, 101 X-linked hyperimmunoglobulin M syndrome, 197 X-linked hypohidrotic ectodermal dysplasia with immunodeficiency (HED-ID), 198 Z Zona pellucida (ZP), 15, 21, 23