Chapter 27 – Preimplantation Genetic Testing


In 2017, the World Health Organization published new nomenclature for preimplantation genetic diagnosis (PGD). They renamed the procedure preimplantation genetic testing (PGT), with PGT-M for monogenic diseases and PGT-SR for structural rearrangements. Preimplantation genetic screening (PGS) was renamed PGT-A (aneuploidy). In this chapter, PGT-M and PGT-SR will be referred to as PGT.

Chapter 27 Preimplantation Genetic Testing

Joyce C. Harper and Sioban B. SenGupta

1 Introduction

In 2017, the World Health Organization published new nomenclature for preimplantation genetic diagnosis (PGD). They renamed the procedure preimplantation genetic testing (PGT), with PGT-M for monogenic diseases and PGT-SR for structural rearrangements. Preimplantation genetic screening (PGS) was renamed PGT-A (aneuploidy). In this chapter, PGT-M and PGT-SR will be referred to as PGT.

PGT is performed for couples who are at risk of transmitting a specific inherited disorder. The reproductive options for these couples are to remain childless, have no genetic testing on any pregnancy (reproductive chance), undergo prenatal or preimplantation genetic diagnosis, have gamete donation or adopt. The couples who opt for PGT have already been diagnosed with their disorder. They may have had an affected child, have a known family history or been diagnosed as an adult. There are a growing number of people who have had direct to consumer genetic testing or preconception genetic risk screening and have identified that they are at risk of transmitting a genetic disease to their children [1]. Most couples going through PGT are fertile, and may have been through prenatal diagnosis and termination of an affected pregnancy. PGT is not an easy option as the couple has to go through in vitro fertilization (IVF); it is expensive and the success rates are only comparable to routine IVF patients. The added problem is that there are cases where all of the embryos are affected as can be seen from the ESHRE PGD Consortium data where for all indications there are a number of cycles that reached PGT but do not have a transfer procedure [2].

PGT was first performed in 1989 in a series of couples who were at risk of transmitting a sex-linked disease to their children [3]. The biopsy was performed by removing one cell from cleavage stage embryos. A polymerase chain reaction (PCR) method was used which was able to detect a segment of the Y chromosome. Soon after this, other centres around the world successfully applied PGT for a variety of single gene and chromosomal disorders [4].

Biopsy has been performed to remove polar bodies or embryonic cells from cleavage or blastocyst stage embryos. The diagnosis has been performed using fluorescent in situ hybridization (FISH), PCR and more recently by array comparative genomic hybridization (a-CGH), next generation sequencing (NGS) and single nucleotide polymorphism arrays.

The technology in PGT has been applied as an adjunct to embryo selection methods used in IVF to select the chromosomally ‘best’ embryo for transfer; PGT-A [5]. PGT-A has been applied to patients of advanced maternal age, repeated implantation failure, repeated miscarriages, severe male factor infertility and more recently, good prognosis (Table 27.1).

Table 27.1 Differences between PGT-M/PGT-SR and PGT-A

Aims Identify genetically normal embryos

Achieve a genetically normal pregnancy/birth
Increase in IVF delivery rate
Indication Monogenic disorder

X-linked disease

Known chromosome abnormality
Advanced maternal age

Repeated implantation failure

Repeated miscarriage

Severe male factor

Good prognosis
Fertility Often fertile Infertile or subfertile or recurrent pregnancy loss
Undiagnosed or inconclusive results Never transfer these embryos Can transfer these embryos
Prenatal Diagnosis Indicated Indicated for the same risk factors as natural conceptions

Adapted from [10]

2 Embryo Biopsy

For two decades, the prevalent method of embryo biopsy has been cleavage stage biopsy where usually one blastomere is biopsied from the cleavage stage embryo on day 3 of development [3]. The initial method used acid Tyrodes to drill a hole in the zona pellucida and aspiration to remove the blastomeres. To date, there is only one study examining the effects of biopsy but it is generally thought that the technique may have a negative effect on implantation and development.

For many years this method remained unchanged. It was only in the ESHRE PGD Consortium data collection for cycles performed in 2004 that the laser was used more often than acid Tyrodes. The other change was the introduction of Ca2+Mg2+ free biopsy media that reduced the junctions between blastomeres and made the biopsy easier [6].

The main issue with cleavage stage biopsy is that high levels of chromosomal mosaicism are seen at this stage [7]. Chromosomal mosaicism within the preimplantation embryo makes PGT highly problematic as the blastomere removed may not be representative of the rest of the embryo. This biological phenomenon has to be taken into account, especially in diagnosis examining chromosomes as it could lead to false positive and negative results.

Polar body biopsy was first reported by Verlinsky and colleagues where they originally biopsied only the first polar body (pre-conception diagnosis) [8]. It was soon realized that both the first and second polar body were required for an accurate diagnosis. The main limitation of polar body biopsy is that it only allows identification of the maternal genes and chromosomes. Therefore it cannot be applied if the male is carrying a chromosome abnormality or dominant single gene disorder. Polar body biopsy has been used in countries whose laws forbid ‘embryo’ biopsy, such as Germany and more recently Italy. The polar bodies can be removed simultaneously or sequentially; both procedures having advantages and disadvantages. Simultaneous removal requires fewer manipulations, but it may be difficult to distinguish between the first and second polar body (which might be necessary for some diagnoses) and the first polar body may degenerate. Sequential biopsy requires two biopsy stages but has the advantage of knowing which polar body is removed.

Blastocyst biopsy is now the most common method of embryo biopsy [2]. Following pioneering work from Muggleton Harris and others, it was applied clinically ten years ago. Blastocyst biopsy can be performed in two ways. A hole can be drilled on day 3 and the embryos left in culture so that some of the trophectoderm cells herniate, which can be biopsied on day 5. The problem with this method is that inner cell mass may herniate instead of trophectoderm. In the second method, the hole is drilled on the morning of day 5, away from the inner cell mass to ensure that only trophectoderm cells herniate. The blastocyst can be returned to culture for a few hours to allow herniation and if required, trophectoderm can be gently aspirated. The cells are cut from the embryo using a laser. The blastocyst will collapse but usually rapidly reforms, sealing the hole in the trophectoderm where the cells have been removed.

One of the problems with blastocyst biopsy is that it gives a relatively short time for the diagnosis as transfer has to occur by day 6, giving just 24 hours for the diagnosis compared to 48–60 hours after cleavage biopsy. The introduction of vitrification as a method to cryopreserve blastocysts, and the reported high survival rates even after blastocyst biopsy has resulted in a shift to blastocyst biopsy and vitrification, allowing an unlimited amount of time for the diagnosis and batching of samples, consequently making the diagnosis cheaper.

Mosaicism is also seen at the blastocyst stage but at a much lower level [9]. Detection of mosaicism and its clinical significance is still being studied (see PGT-A section).

Blastocyst biopsy usually gives five cells that are analysed together which makes diagnosis easier and reduces the misdiagnosis rate. Since only approximately 50–60% of embryos reach the blastocyst stage, biopsy at this time results in fewer embryos to process which is more time and cost effective. This is especially relevant in comparison to polar body biopsy where at least 2–4 times the number of samples will have to be analysed.

Biopsy is an invasive procedure. In the case of blastocyst biopsy it removes some of the embryo or cells that will go onto make the placenta. Biopsy of the blastocoelic fluid and the use of spent culture media are being studied [10,11]. Whether there are two polar bodies, 1–2 blastomeres or 5 trophectoderm cells, the major advances in PGT over recent years have been the methods of diagnosis.

3 Diagnosis

3.1 Molecular Diagnosis

The polymerase chain reaction (PCR) is designed to enrich a DNA sample for one specific fragment, amplifying it to a level at which it can be visualized and subjected to further genetic analysis. It has become one of the most important methods in genetic testing and refinement of the PCR protocol for single cell analysis has proven highly successful. The sensitivity of PCR with fluorescently labelled primers is suitable for single cell analysis, allows multiple targets to be amplified simultaneously, and has reduced the problems of contamination and the time taken for the genetic analysis to be performed.

Allele dropout (ADO), the phenomenon where only one of the two alleles present in a cell is amplified to a detectable level, generally affects 5–20% of single cell amplifications (although in some instances the frequency is higher) and is a problem that is yet to be fully understood. This is an important consideration in the diagnosis of dominant disorders or recessive diseases where only one mutation can be detected. Multiple ADO events may appear to have occurred in cells that are monosomic for the chromosome being tested by PCR. Similarly contamination of DNA from cumulus cells, sperm or external sources will confound diagnosis.

A growing number of genes have been analysed by PCR–PGT and a variety of mutation detection strategies employed. Minisequencing [12] and real-time PCR assay with fluorescence resonance energy transfer (FRET) hybridization probes followed by melting curve analysis [13] are the most common methods used. The best protocols take into consideration the principal problems; amplification efficiency, contamination and allele dropout. It is now understood that an absence of amplification (PCR failure) should never be taken as an indication that an embryo is free of a mutation; amplification is unsuccessful in approximately 10% of isolated blastomeres regardless of their genotype.

In a multiplex PCR reaction, several loci can be investigated at once if the primers are labelled with different fluorescent tags and by designing primers to give PCR products of different size ranges. In all cases of molecular diagnosis, multiplex PCR of the mutation locus together with flanking linked polymorphic markers provides an additional means of determining the genotype of the embryo hence reducing the risk of misdiagnosis due to ADO.

Protocols based solely on the analysis of multiple microsatellite markers linked to and flanking the mutation site provide an indirect method for diagnosis by the identification of alleles in phase with the mutation. These protocols require the availability of DNA from at least two individuals in the family with the mutation so that the phase alleles can be identified by linkage analysis. Linkage based strategies without mutation detection are the most commonly used approach when reference samples for phase determination are available. In the case of de novo mutations, phase alleles can be identified using a combination of mutation detection and haplotyping of linked STR (short tandem repeat) markers by single sperm or polar body multiplex PCR [14].

There are a variety of methods aimed at non-specific amplification of the entire genome (whole genome amplification – WGA) [15]. Using these techniques a single genome can be amplified numerous times, thus providing sufficient DNA templates for many independent PCR amplifications including mutation detection and polymorphic markers that could test for ADO and contamination. WGA is required for a-CGH, SNP arrays and NGS. Methods include multiple displacement amplification (MDA) which is a non-PCR isothermal, strand-displacing amplification method. Other strategies involve fragmentation of DNA and ligation to linkers followed by PCR (PicoPLEX) or limited MDA followed by PCR for multiple annealing and looping-based amplification cycles (MALBAC).

PGT for mutations in the mitochondrial genome is complex due to random genetic drift during oogenesis resulting in heteroplasmy, where primary oocytes have a mixture of mitochondria with and without the inherited mutation. A bottleneck that operates at oogenesis determines the mutational load in primary oocytes. The size of the bottleneck appears to vary for different mitochondrial mutations and also between individuals. Determining a suitable threshold value for mutational load in embryos is difficult as the mutational load is also subject to random genetic drift in somatic tissue during development which will affect embryo survival and overall phenotype. PGT for mitochondrial mutations can be considered to be risk reducing rather than complete removal of the mutation, and couples need to be carefully counselled about the limitations of PGT for these disorders. PCR with restriction enzyme analysis has been used for the detection of specific mitochondrial mutations showing skewed meiotic segregation and the selection of embryos with a low mutational load. PGT for mutations in the mitochondrial genome requires extensive workup to allow semi-quantitative assessment of mutational load in single cells. Transplantation of nuclear DNA into enucleated donor oocytes or fertilized eggs may be clinically available in the future as a reproductive treatment option for females with mitochondrial mutations. The assessment of the mutational load by PGT is likely to still be needed to determine the extent of carry over of mitochondria during nuclear DNA transfer.

3.2 Examining Chromosomes

Fluorescent in situ hybridization (FISH) uses fluorescently tagged DNA probes that bind to their complementary sequence and can be visualized under a fluorescent microscope. The first chromosome analysis for PGT used FISH to identify the X and Y chromosomes. This method was rapidly applied to detect inherited chromosome abnormalities, such as translocations and non-inherited chromosome abnormalities for PGT-A.

The biopsied cells are spread, usually using HCl/Tween or methanol:acetic acid and FISH performed using appropriate probes. Early studies showed that the more probes used, the less efficient the procedure becomes. Today it is not advised to use FISH for PGT/PGT-A as more efficient techniques are available, such as a-CGH and NGS.

Only gold members can continue reading. Log In or Register to continue

Oct 26, 2020 | Posted by in GYNECOLOGY | Comments Off on Chapter 27 – Preimplantation Genetic Testing
Premium Wordpress Themes by UFO Themes