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In the United States alone there are an estimated 6.7 million women of reproductive age with impaired fertility, which is defined as failure to conceive after a year or more of trying. And the problem is growing: the median age for first marriage in the U.S. has steadily increased, and women today often choose to delay childbirth until their late 30s or 40s in order to pursue and advance their careers. The downside for women deciding to postpone childbirth is an increased likelihood of encountering difficulty conceiving.
When other methods of assisted reproduction—such as hormone treatments to increase the number of eggs a woman releases with each cycle, or artificial insemination directly into the uterus—fail or are not applicable, in vitro fertilization (IVF) becomes one of the few options left for couples who want a genetically related child. But the process is far from easy. Long...
Doctors use several criteria to select embryos with the best chance of developing, but the success rate is still only about 30–40 percent for women under 35.
Once eggs are mature and ready to be ovulated, they are collected from the woman’s ovaries and incubated in vitro with sperm. Doctors use several criteria to select embryos with the best chance of developing, but the success rate is still only about 30–40 percent for women under 35 years of age, and much lower for women in their 40s. As a result, many couples often undergo several rounds of unsuccessful IVF treatment, which extracts huge emotional and financial costs and poses health risks for women due to repeated hormone treatment. Currently, the most commonly chosen option for increasing the likelihood of successful pregnancy from one cycle of in vitro treatment is to implant multiple embryos, which often results in multiple pregnancies, increasing the risk of serious health complications for the mother and the growing babies. Therefore, there has been a strong impetus to devise better methods for choosing the single best embryo the first time around. In the next few years, a number of promising new methods are likely to be tested in clinical trials.1
By sight and by genes
Since 1978, when the first IVF baby was born, doctors have had a limited set of tools to aid in choosing the best embryo. Visual assessment of embryo morphology is currently the most traditional and popular method. Using a simple light microscope, physicians assess several parameters at different developmental stages to gain valuable information about the quality of embryos.2 On the first day, fertilized eggs can be graded according to the morphology of their two pronuclei, which separately contain the mother’s and father’s contributions of genetic material. (See illustration above.) Embryologists examine the size of the pronuclei and the number and localization of nucleoli contained within each. In healthy embryos, pronuclei should be evenly sized and contain similar numbers of nucleoli grouped in one region. On the second and third days, once the pronuclei have fused and the fertilized eggs have begun cleavage divisions, physicians can assess the number and shape of the resulting blastomeres and the degree of fragmentation—the quantity of small cell fragments that can be created during divisions. Finally, on the fifth or sixth day, when the embryos have developed into blastocysts—hollow spheres with a mass of cells protruding inward at one end—physicians can evaluate the size and shape of the trophoblast, or outer sphere; the hollow cavity; and the protruding cell mass that will develop into the fetus, to help select the best embryo for transfer. Depending on the protocol employed by the fertility clinic, embryos may be evaluated at one or several of these developmental stages.
Even when expertise in visual embryo grading is available, some low-graded embryos have been experimentally shown to have high potential of developing to term.
Although morphological assessment is inexpensive and easy to implement in the clinic, it has drawbacks. The visual grading of embryos is subjective and requires considerable training and expertise. Moreover, even when such expertise is available, some embryos scored with a low grade have been experimentally shown to have a high potential of developing to term.
More recently, additional selection criteria based on genetic, proteomic, and metabolomic analyses of the embryo have been developed to help detect disorders that might either affect the viability of the embryo, or result in serious illness in the child if the pregnancy is successful. Preimplantation genetic diagnosis (PGD) is commonly used by parents who know they carry a genetic disorder that could be passed on to the child. The procedure begins with the harvest of one or two cells from the blastomere stage (day 2 or 3 after fertilization) or several cells from the trophoblast (day 5 or 6), either of which can be harvested, in most cases, without causing harm to the embryo. The harvested cells are prepared for analysis by polymerase chain reaction (PCR) or fluorescence in situ hybridization (FISH) to diagnose specific genetic mutations and chromosomal abnormalities, or to sex the embryos for patients carrying X chromosome–linked diseases. More recent forms of PGD include comparative genomic hybridization or single nucleotide polymorphism microchip arrays.3,4 Both of these approaches allow analysis of the entire embryonic genome and therefore provide a fuller characterization of the embryo.
The same methods of genetic analysis can be used to screen preimplantation embryos of older women and those who have suffered repeated miscarriages and implantation failures, or of couples that experience severe male-factor infertility with no diagnosed genetic cause. Such preimplantation genetic screening can also be done at different stages in early embryonic development.
Each of these methods provides information regarding the embryo’s risk for inherited diseases or de novo chromosomal abnormalities, but they require removing a sample from the developing embryonic mass. Newer methods of assessing an embryo’s viability, which have recently seen some popularity in clinics, are metabolomic and proteomic analyses. These methods rely solely on sampling the culture medium in which the embryo is growing and do not disturb the cell mass.5 By collecting and analyzing the culture medium surrounding each embryo, researchers have been able to observe changes in pyruvate or glucose concentration, which can be used as proxy measurements for metabolism. The usefulness of these data in predicting the embryo’s quality, however, is still a matter of debate. Other variables, such as oxygen consumption and amino-acid turnover in the surrounding medium, have proved to be more reliable indicators of viability. Recently, the analysis of single metabolites has been replaced by approaches that examine a profile of metabolites and proteins secreted by the embryo. This allows for a broader examination of an embryo’s metabolism and gene-expression patterns. Although many reports show a relationship between metabolic status of the embryo and its viability, suggesting the potential value of such analyses for use in IVF clinics, the technique may be difficult to implement in a clinical environment because of the cost of equipment and the expertise required to perform the analyses.
© TAMI TOLPAEven with these procedures in place, success rates have not markedly improved, and researchers in the field have continued to investigate noninvasive methods of scoring an embryo in the first few days after fertilization. The window for collecting information is short. After day 5, the embryo begins to prepare for attachment to the uterine wall and must be implanted or frozen. Most existing methods can accurately assess the embryo by day 3, though some take until day 5 or 6. It is important to minimize the time outside the body of the mother, as some researchers report concerns that keeping the embryo in culture might increase the chances that its genome will be affected epigenetically by the culture environment. Therefore, it would be desirable to have an alternative method of embryo evaluation that would provide a higher success rate, more detailed information about the developmental status and potential of the embryo at earlier stages, and most importantly, would be quantitative and objective.
The next generation of selection
My lab (Zernicka-Goetz), started working on new methods for embryo selection as an offshoot of our studies on early mouse development. In 2005 it became apparent to me that the best way to better understand what happens in the first days of embryo development would be to film the event in living embryos without disturbing them in any way. I passed this idea on to a new PhD student of mine, Emlyn Parfitt, and to Marcus Bischoff, a postdoc who had previously used imaging to analyze the development of worm embryos and who had just joined the lab of my friend Peter Lawrence at the University of Cambridge. Emlyn and Marcus began filming the development of the mouse embryo and, with the help of sophisticated software, tracked the fate of each cell from its “birth,” through cell division, until its fate was established by the blastocyst stage. Their 4-D analysis of these movies was absolutely captivating as they revealed how the embryo’s first cell divisions and movements relate to the establishment of the complex blastocyst structure.6 It was a very important step forward because it had not previously been possible to quantitatively track the development and fate of each cell in living mammalian embryos, and the information they gleaned paved the way for many future experiments.
© PROFESSOR PIETRO M. MOTTA ET AL./SCIENCE SOURCEOther labs were also developing time-lapse culture methods, and in 2010 Renee Reijo Pera's lab at Stanford University published a study tracking human embryogenesis using time-lapse image analysis.7 Their group recorded the growth of embryos up to the blastocyst stage, tracking those embryos that developed normally as well as those whose growth appeared to stall. Poor-quality embryos could be determined visually as those in which the final step of cell division—cytokinesis, or the cinching of the membrane that separates the two daughter cells—took longer than usual. Moreover, low-quality embryos underwent the second cleavage division either significantly earlier or significantly later than good-quality embryos. In both high- and low-quality embryos, this second division results in three cells because only one of the two cells from the first cleavage divides. (See illustration above.) Finally, Reijo Pera’s group found that embryos that developed successfully, at least to the blastocyst stage, had their second and third divisions (which result in three and then four cells) in rapid succession, or almost synchronously. The study not only confirmed but also quantified earlier observations, which had suggested that the timing of the first divisions were correlated with higher-quality human embryos.2
It was around 2007 when we first realized that imaging development immediately after fertilization might help us establish a method to determine embryo quality. This work stemmed from our earlier studies, which inspired our curiosity because they suggested a correlation between the site of the sperm entry into the egg and subsequent developmental processes.8 A collaboration with Chris Graham, my former mentor at the University of Oxford, and Adrian Thomas’s team, also at Oxford, led us to discover that fertilization initiates a series of rapid cytoplasmic movements that proved key in predicting subsequent developmental events. To detect these movements, we filmed embryos with rapid time-lapse imaging (1 frame every 10–20 seconds) and analyzed the images using a computational method based on particle image velocimetry (PIV), which was developed to measure the dynamics of movements such as airflow over the wings of a plane or blood flow in an artificial heart. PIV follows patterns of contrast (white and black pixels) between sequential pairs of images and calculates displacement vectors for them. These vectors are used to create a map that covers the entire zygote and quantifies how the cytoplasm moves over time. (See illustration above, right.)
My lab observed mouse embryos just after the sperm had entered the egg, leaving on its surface a protruding cap called the fertilization cone at the site of sperm entry. In the minutes and hours that follow, the cytoplasm of the zygote began to swirl at high speed and then slow down, oscillating within the cell in a pattern caused by contractions of the actomyosin network crisscrossing the cell. We also observed the protruding fertilization cone flatten just before a new cytoplasmic flow began, then slowly begin to protrude again prior to the next flow. The actomyosin contractions, and thus the cytoplasmic movements, are caused by oscillations in intracellular calcium levels released from the endoplasmic reticulum, which are first triggered by the fertilizing sperm. Whenever the level of calcium increases, the actomyosin network contracts, and cytoplasmic movement speeds up. Indeed, when we experimentally block the bursts of calcium, fast cytoplasmic movements do not occur.
We observed that newly fertilized mouse embryos displaying very frequent increases in cytoplasmic movements (i.e., very frequent calcium bursts) and low cytoplasmic speeds in the intervening periods (i.e., poor-quality actin cytoskeleton) developed to birth almost three times less frequently than embryos displaying more average values of these parameters. This allowed us to develop the first, and to our knowledge the only, noninvasive and quantitative way of assessing the quality of the actomyosin network and calcium oscillations in the cell, both of which are essential for proper embryonic development. Importantly, cytoplasmic movements very similar in character have also been discovered in activated human eggs by our Cardiff University collaborator, Karl Swann.10 We have filed a patent to help commercialize this technology and are currently testing the predictive potential of cytoplasmic movements in human embryos. In addition, the method can be applied with minimal technical training and, at least in mice, provided results as early as several hours after fertilization.
Looking to the future
Although tracking the speed of cell divisions using the Reijo Pera method or monitoring cytoplasmic flows remains to be proven predictive for assessing the ability of preimplantation human embryos to develop to birth, they offer great hope, and clinical trials of both methods are currently under way.
Research carried out over the past three decades has provided a broad repertoire of embryo selection methods. Some of them, such as visual evaluation of an embryo’s nuclei and genetic testing, focus on the nuclear component of the embryo, looking for potential chromosomal abnormalities. Others, such as the analysis of metabolites and secreted proteins or of cytoplasmic flows, concentrate on the cytoplasmic component, by investigating the quality of an embryo’s metabolism, its calcium homeostasis, and its cytoskeleton. We believe that the ultimate selection method should combine these approaches. This can be achieved, for instance, by following the timing of embryonic cell divisions, as their duration and synchrony can be affected by improper chromosome segregation, cytoskeletal properties, or energy levels. An alternative approach could combine preimplantation genetic screening with the analysis of an embryo’s metabolism or its cytoplasmic movements. Combining genetic testing with examination of cytoplasmic flows is especially promising. This combination would provide information about the embryo’s quality within several hours after fertilization, and would therefore significantly quicken the selection process, thus potentially shortening the time IVF embryos spend in culture before being transferred to mothers-to-be.
Anna Ajduk is an assistant professor at the University of Warsaw, Poland, and Magdalena Zernicka-Goetz is a professor of Mammalian Development and Stem Cell Biology and a Group Head at the University of Cambridge, U.K.
This article is adapted from a review in F1000 Biology Reports, DOI:10.3410/M3-15 (open access).
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