Morphological assessment of embryo viability.

114

Morphological Assessment of Embryo Viability Michael Abeyta, BS, TS (ABB)1

Barry Behr, PhD, HCLD2

1 Department of Obstetrics and Gynecology, Stanford Fertility and

Reproductive Medicine Center, Palo Alto, California 2 Department of Obstetrics and Gynecology, Stanford University Medical Center, Palo Alto, California

Address for correspondence Barry Behr, PhD, HCLD, Department of Obstetrics and Gynecology, Stanford University Medical Center, 900 Welch Road, Suite 14, Palo Alto, CA 94304 (e-mail: [email protected]).

Abstract

Keywords

► ► ► ►

morphology embryo implantation in vitro fertilization

Morphological assessment is discussed in the context of significant literature at all stages of in vitro development, beginning with the oocyte and culminating at the blastocyst stage. Current evidence is used to debate the inclusion of commonly observed morphological features in grading schemes. The biological rationale behind observed phenomena such as multinucleation and fragmentation are also explored. Current limitations as well as technological advancements that increase our ability to assess viability are highlighted. Particular attention is paid to the relationship between developmental timing and assessment schemes. Failure to standardize assessment timing and inclusion criteria is glaring weaknesses of the literature that currently make consensus unattainable. Mounting evidence suggests that the future of static assessment is very likely to be influenced by information gathered from preimplantation genetic screening and other invasive techniques as well as from continuous monitoring tools such as time lapse.

Choosing the optimal number of embryos to transfer back to the uterus is a highly variable and complex endeavor. Whatever the stage and timing of embryo transfer may be, the embryo selection process is at a crossroads of societal, economic, scientific, and moral debate. Although medical and socio-economic concerns pressure the drive toward single embryo transfer (SET), this is often juxtaposed with the logistical considerations of the infertile couple, who may lack the time or financial resources to attempt multiple cycles and thus desire an “instant family” (twins) or are willing to risk high-order multiple gestation to maximize their chance of achieving a pregnancy. Interwoven with these factors is the competitive pressure among clinics to increase success rates and overall efficiency, thereby fueling the desire to optimize the embryo selection process. Some countries impose limitations on the number of embryos that can be created, shifting the emphasis toward selection of the highest quality oocytes, while in countries without such limitations, supernumerary oocytes and embryos have allowed for selection at the other extreme, the blastocyst stage. No matter the developmental stage, by and large,

Issue Theme Selecting the Best Embryo; Guest Editor, Valerie L. Baker, MD

morphological assessment remains the primary method used to distinguish embryo quality and ultimately, viability. The main advantage of morphological assessment lies in the noninvasive nature. Embryos can be monitored with minimal impediment to the process of development, yet this type of assessment is far from ideal. Embryo grading is riddled with subjectivity. Even within the same clinic embryologists often struggle to establish consistency in grading.1–3 Inter- and intraobserver variability can vary enough to affect decision making, directly impacting IVF program success.2 Subjectivity aside, static morphologic assessment is merely a snapshot from the dynamic process of embryo development. While an interphase cell is morphologically static, and embryos do follow fairly predictable patterns of cleavage, time-lapse imaging has shown that during the cleavage stage, fragmentation percentage varies over time as some fragments can be reabsorbed into cells.4,5 The blastocyst stage is similarly dynamic as the embryo contracts and expands to facilitate the hatching process.4 From a single static observation, random chance alone can therefore cause a poor quality embryo to be mistaken for a good one, and vice versa.

Copyright © 2014 by Thieme Medical Publishers, Inc., 333 Seventh Avenue, New York, NY 10001, USA. Tel: +1(212) 584-4662.

DOI http://dx.doi.org/ 10.1055/s-0033-1363553. ISSN 1526-8004.

This document was downloaded for personal use only. Unauthorized distribution is strictly prohibited.

Semin Reprod Med 2014;32:114–126

The solution to these limitations may lay in a combination of approaches, ranging from dynamic assessment (timelapse) to cumulative scoring of embryos. This article will discuss the present and future tools at our disposal for the morphological assessment of embryos at the various stages of development, and analyze the relationship between these morphological assessment techniques and viability.

The Oocyte Without the stripping away of the cumulus little can be gathered about the appearance of the oocyte itself, as the zona pellucida, oolemma, and presence or absence of a polar body can be difficult to visualize. Yet, the vast majority of the cellular machinery driving early embryo development comes from the oocyte,6,7 which suggests that increasing our understanding of the oocyte and its follicular environment would enhance our ability to predict those oocytes that would result in top quality, viable embryos. The cumulus, coronal layer, and even follicular size and fluid content at the time of retrieval have been the target of research all aimed at understanding the retrieved oocyte. The very first opportunity for morphological assessment is not of the oocyte itself but rather the developing follicle on ultrasound. Studies of follicle size clearly show that larger follicles correlate with a higher likelihood of oocyte maturity.8–10 However, there is disparity in design and follicle size groupings among these studies, making it difficult to interpret other outcomes as clearly. One notable study by Bergh et al looked at the influence of large and small follicles on fertilization, cleavage, and pregnancy rate, separating intracytoplasmic sperm injection (ICSI) and in vitro fertilization (IVF) cycles.8 Interestingly, the pregnancy rates were lower when well developing embryos from small follicles were transferred in IVF cycles. This was not seen in ICSI cycles, where small and large follicles that produced top quality embryos created pregnancies at equal rates. The results of this study suggest that ICSI provides a selection advantage over IVF by eliminating immature oocytes from consideration for transfer because they are not routinely injected. In conventional insemination cycles, some of the immature follicles fertilize, and although they are capable of producing morphologically “normal” embryos, the study suggests that they have reduced viability. Rosen et al classified follicle size groupings and looked at fertilization and day 3 morphology as outcome measures.9 Although cell number on day 3 was similar among the groupings, they saw a progressive decline in fertilization and a corresponding increase in fragmentation as follicle size decreased in both ICSI and IVF cycles. Both studies demonstrate the influence of stimulation on oocyte and embryo quality. The cumulus oocyte complex (COC) retrieved at follicular aspiration is composed of the oocyte and its supporting cumulus/granulosa cell layers. The coronal layer is in direct contact with the oocyte via gap junctions11 and relays important signals to the oocyte from the hormone producing granulosa cells.12 Visually, the degree of expansion of this cell layer is a predictor of oocyte maturity, where few layers of

Abeyta, Behr

tightly packed cells are usually a sign of immaturity and multiple expanded layers correlate with maturity.13 In addition to predicting maturity based on the morphology of granulosa cells, there is also data to suggest a correlation with oocyte competence.14 Research has sought to understand the signaling and metabolic activity of these supportive cells with the hope of determining the molecular fingerprint of a good quality oocyte.15–17 One recent study was able to correlate telomere length in cumulus cells with oocyte and embryo quality.18 Another study found that increased levels of oxidative stress in granulosa cells adversely affected IVF success.17 If molecular markers such as these could be related back to COC morphology, it would allow for the noninvasive selection of the best oocytes for insemination. The possibility of such a link has yet to be explored in a prospective study. Studies have assessed many of the physical characteristics of the stripped oocyte, including the overall size and shape, the zona pellucida shape and thickness, oolemma texture and color, and polar body size, shape, and degree of fragmentation.13,19–23 Giant oocytes (approximately 30% larger than normal oocytes) have been shown to be chromosomally abnormal and should therefore be excluded from injection and uterine transfer.24 Another study proved the source of the chromosome abnormalities from fertilized giant eggs to be digynic triploidy.25 In addition, a meta-analysis of 14 studies, including 3,688 ICSI cycles found a significant reduction in fertilization when oocytes displayed the presence of vacuoles, a large perivitelline space, refractile bodies, or large first polar body.23 It is estimated that more than 50% of retrieved oocytes contain one or more of these abnormalities, on average.26,27 The mechanisms leading to these morphological abnormalities are largely speculative, but the strong correlation with outcomes led the Alpha Scientists/European Society of Human Reproduction and Embryology (ESHRE) special interest group workshop to conclude that the ideal oocyte should have a uniform zona pellucida enclosing a spherical, uniform, and translucent cytoplasm which is free of inclusions and has a size-appropriate polar body.28 In addition to the morphology, the mere presence or absence of the polar body and the timing of extrusion has predictive value.29,30 Corroborating the theories from the Bergh study on follicle size, De Vos et al found that when in vitro matured metaphase I (MI) oocytes were injected, they fertilized at a lower rate, but produced similar quality embryos to metaphase II (MII) oocytes. However, when forced to transfer the embryos from in vitro matured MIs, only one pregnancy occurred from 15 transfers.29 Shu et al found that in patients who had no mature oocytes at the time of cumulus removal, the injection of MI oocytes that failed to progress to MII, along with those that converted to MIIs had severely reduced fertilization, pregnancy, and implantation rates.30 The two pregnancies (from 46 cycles) both resulted in spontaneous abortions. Twenty of the cycles resulted in complete failed fertilization. The inefficiency of pregnancies in these two studies demonstrates the severely compromised viability of oocytes with asynchronous nuclear and/or cytoplasmic maturity.31 They also suggest that the time between retrieval and cumulus stripping can play an important diagnostic role. Seminars in Reproductive Medicine

Vol. 32

No. 2/2014

115


Morphological Assessment of Embryo Viability


Abeyta, Behr

While waiting for several hours after retrieval before stripping may increase the percentage of mature oocytes, perhaps immediate stripping provides an additional selection tool, separating the most viable oocytes (MIIs) from the compromised oocytes (MIs that may later convert to MIIs). Further research is necessary to determine if additional time with cumulus cells intact would increase the viability of MI oocytes that convert to MII in comparison with in vitro matured oocytes without cumulus. One of the most intriguing advancements in IVF over the last decade was the adaptation of polarized light microscopy (PolScope) for the study of the MII oocyte.32 Through a combination of polarizing optical hardware and digital imaging software, birefringence of highly structured macromolecules can be observed noninvasively.33 Two structures of the oocyte that exhibit birefringence are the meiotic spindle and the zona pellucida. Historically, visualization of the meiotic spindle required destruction of the oocyte,34–36 but the Polscope allows for visualization of changes in real time and direct correlation to downstream outcomes.37 Clinically, the presence or absence of the meiotic spindle at the time of ICSI has helped to partially explain fertilization failure,38–40 and knowing the location of the spindle can aid in avoiding oocyte damage.41 The size, shape, and maximum birefringence of the spindle have all been suggested in the literature as important indicators of oocyte competence.38,42 However, viewing of the spindle image changes as the oocyte is rolled around. It can be difficult to find the optimal position for data collection of each egg. This can create variability in data collection and has made it difficult to establish clear thresholds or design multicenter studies. Assessment of the zona pellucida with polarized light microscopy has been limited up to this point due to the minimal importance of the zona in ICSI cycles. However, Held et al recently showed that zona pellucida birefringence parameters were inversely related to blastocyst development in bovine oocytes and could be correlated back to COC morphology scores.43 This work points toward a potential role for zona pellucida birefringence imaging in clinical IVF.

Zygote Scoring The formation of pronuclei (PNs) is a regimented process that provides a predictable window for morphological viewing between 12 and 20 hours.44–46 It is critical that assessment takes place during this window as it is the only noninvasive measure we have of normal fertilization. Haploid and triploid embryos are known to cleave and sometimes progress as far as the blastocyst stage with excellent morphology. Therefore, in the absence of this assessment, abnormally fertilized oocytes could be selected for embryo transfer. Beyond assessing fertilization, many attempts have also been made to correlate PN morphology and timing with developmental potential. Scott and Smith originally introduced a PN stage scoring system which took into account the relative position and size of the PNs, the positioning of the nucleoli, and the appearance of the cytoplasm (presence or absence of a halo) at 16 to 18 Seminars in Reproductive Medicine

Vol. 32

No. 2/2014

hours postinsemination. A second observation at 24 to 26 hours postinsemination factored in developmental progression (nuclear membrane breakdown and cell division).47 The cumulative score from all categories ranged from 7 to 25. An average, or corrected score (CS) per patient was calculated by adding the scores of all embryos transferred and dividing by the number of embryos transferred. In this initial study, when the CS was 15, the pregnancy rate was 71%, compared with just 8% with CS 14. This system was subsequently revised twice into what is known today as a “z-score,” which only includes a single observation at fertilization assessment and categorizes zygotes into four groups based on developmental potential.47,48 Tesarik and Greco45 similarly proposed a scoring system for the detailed analysis of PN morphology. They argued that previous work by Tesarik and Kopecny49 had shown the biological relevance of nucleolar precursor body (NPB) dynamics and therefore zygote scoring should provide a reliable indication of downstream developmental competence. This scoring system differed from the work of Scott in that it was based solely on the synchronization of events between the two PNs rather than cumulative scores from multiple features. They recognized that NPB polarization is a progression and therefore it was the synchronized movement toward polarization that was important for a single static observation. On the basis of retrospective analyses of embryos with identifiable implantation outcomes from day 3 transfers, they characterized several distinct patterns of PN morphology as either normal or abnormal (described by Tesarik and Greco45). From pattern 0 zygotes, which displayed synchronous progression of NPBs, they found a significantly decreased likelihood of developmental arrest and a significant increase in both good morphology embryos and implantation rate. Attempts to validate PN grading systems based on the criteria above have had mixed results. Although several studies have demonstrated PN scoring to be predictive of blastocyst formation and quality,48,50,51 others have found either no difference versus day 2/day 3 embryo quality46,52–54 or even inferior results with PN scoring compared with scoring on other days.55 Another study found that subclassification of the type “0” pattern from the original scoring system of Tesarik and Greco further improves selection.56 In summary, the results from use of PN scoring systems are inconclusive. However, this is not unexpected considering the lack of consensus on several study criteria. First of all, many of the studies were retrospective analyses. Second, there are multiple grading systems being used, with varying degrees of overlap. Furthermore, there is a glaring lack of control for timing of observation. Although many studies noted above have shown polarization of nucleoli to be favorable and Nagy et al demonstrated that in ICSI cycles, the polarization status changes over time,46 currently no studies go as far as proving that NPB polarization status at different times postinsemination are better, worse, or equal. In addition, data showing a delay in PN formation with IVF versus ICSI57 warrant the creation of optimization curves for PN progression with ICSI versus conventional insemination. Another area of debate is the role of polarization in the human embryo relating to the


116

location of sperm entry, positioning of the PNs, NPBs, and polar bodies.7,44,58–61 Until these questions are definitively answered, there cannot be consensus on a PN scoring system.

Early Cleavage Timing of the first cell division was introduced as an independent marker of viability by Shoukir et al and Sakkas et al.62,63 In the original study, embryos were assessed at 25 hours postinsemination. Those embryos in which cleavage to the two-cell stage had occurred were designated as “early cleavage” (EC) embryos. This was based on earlier work by other groups reporting that the earliest that human embryos reach the two-cell stage is between 20 and 27 hours postinsemination.64–66 When comparing transfers where one or more embryos contained an EC embryo to those transfers without any EC embryos, they found the implantation rate to be threefold higher in the EC group and pregnancy rate doubled that of the non-EC group. On the opposite extreme, Van Blerkom demonstrated the relationship between delays in PN progression with delayed or failed progression to the two-cell stage.67 These studies beg the question of whether EC is actually optimal cleavage, and non-EC is actually delayed cleavage. The dilemma is determining the most appropriate time for observation and how best to capture these results. Using a time-lapse imaging system, Meseguer et al proposed an optimal first cleavage window of 25 to 28 hours and showed that embryos cleaving before this optimal window failed to implant.68 Montag et al then demonstrated how a single static observation within this time frame would not only select the 42.3% of optimally cleaving embryos (from their dataset) but also an additional 27.1% of embryos that cleaved earlier than 25 hours that would fail to implant according to Meseguer et al.69 As with PN scoring systems, differences in timing between ICSI and IVF embryos should be considered with EC.

Cleavage Stage Assessments When IVF was in its infancy, Edwards et al64,70 began observing and documenting the characteristic patterns of embryo development at the cleavage stage. Today, there are several morphological characteristics that are routinely documented at the cleavage stage. Among those most often identified as predictive are multinucleation, cell number, fragmentation, blastomere size, and symmetry. Although we have known for quite some time that these are key indicators, our knowledge of the factors influencing each continues to evolve.

Multinucleation Once the first mitotic division takes place, subsequent visualization of nuclei by routine microscopy is only possible at interphase. At this stage of the cell cycle, there should only be one nucleus visible per cell. A multinucleated blastomere (MNB) is one that contains multiple nuclei. Multinucleation is more frequently documented on day 2 than day 3, but it is unclear whether the actual frequency of multinucleation is

Abeyta, Behr

greater on day 2 than day 3. It has been suggested that observation of MNBs on day 2 may be easier due to the larger size of blastomeres and the increased visibility when fewer cells and fragments are present, as is often the case on day 2 relative to day 3.71 Munné and Cohen performed fluorescence in situ hybridization analysis on blastomeres from discarded/arrested embryos and found several patterns of multinucleation.72 The patterns illustrated three possible mechanisms for the origin of multinucleation73–75: Mitotic replication without cytokinesis would explain instances where each MNB nucleus has the same genetic constitution as their mononucleated sibling blastomeres. Fragmentation of nuclei without previous deoxyribonucleic acid (DNA) replication would explain how a MNB would have the sum total of chromosomes in the multiple nuclei as found in mononucleated sibling blastomeres. Finally, a defective mitotic spindle, potentially associated with multiple rounds of DNA replication was theorized to explain instances where the number of a particular chromosome was double, triple, or quadruple the copy number of mononucleated cells, and the replicates of this same chromosome were randomly distributed among the MNB nuclei. Van Royen et al linked MNB occurrence with stimulation issues that affect the oocyte such as decreased cycle length and increased follicle stimulating hormone total dose.71 It has also been suggested that subpar laboratory conditions can cause multinucleation.76 However, multinucleation in human embryos appears to be a common phenomenon both in vivo and in vitro. Hertig et al observed the presence of MNBs in four out of six early embryos recovered from the female genital tract,77 and several researchers have documented MNB in at least one embryo from roughly two-thirds of all cycles assessed in vitro.73,78 Tesarík et al used thymidine labeling techniques in combination with electron microscopy to analyze the ultrastructure of MNBs.74 On the basis of the observation of chromatin aggregates outside of the nuclei, they postulated that embryos use lysosomes to actively expel extrachromosomes as a repair mechanism. Low levels of ribonucleic acid synthesis from MNBs at the eight-cell stage suggest irreparable developmental arrest of these blastomeres but may allow the sibling mononucleate blastomeres to continue on. Whether or not repair mechanisms exist, it is clear that when present, multinucleation should always be noted. Among the observed negative consequences of multinucleation are increased rate of aneuploidy,79–82 impaired cleavage,71 increased fragmentation,71 and decreased implantation rates.71,83,84 The abundance of literature suggests that embryos with MNBs should be excluded from consideration for transfer when other mononucleated embryos are available in sufficient numbers.82

Cell Number (Cleavage Rate) It is universally accepted that on day 2 after insemination that an embryo should be at the two- to four-cell stage and that by day 3, an embryo should reach the six- to eight-cell stage. However, the amount of time individual embryos take to reach these milestones varies, and as such, many researchers Seminars in Reproductive Medicine

Vol. 32

No. 2/2014

117




Abeyta, Behr

have sought to understand the importance of this variability as it relates to the implantation potential of the embryo. Guerif et al found that although there was a significant correlation between day 2 cell number and blastocyst formation, no relationship existed between day 2 morphological score and implantation among blastocyst SETs after controlling for blastocyst morphology.85 They offered as an explanation a possible growth discordance between early stages compared with after the activation of the embryonic genome, with the latter being more reflective of embryo viability.86 Several studies have shown blastomere number on day 2 to be an independent predictor of implantation potential, with four cells being the optimal cell number.87–89 Holte et al found cleavage rate to be the most powerful independent morphological predictor of implantation by multiple regression analysis.88 In agreement with many earlier studies,87,90–94 a superior viability of four cells on day 2 was observed, while also demonstrating a more pronounced decline in implantation rate for cell numbers below four than above it. Related to the issue of cleavage synchrony, Scott et al retrospectively found that all viable pregnancies in a controlled dataset of day 2 transfers came from two and four cell embryos.89 Pregnancies did occur, albeit at a significantly reduced rate in embryos that were at 3 and > 4 cells 42 hours postinsemination. However, zero made it to delivery. In this study, they found day 2 parameters to be more predictive than morphological scoring on any other day in regard to take home baby rate. The decreased viability of three cell embryos on day 2 may partially be described by abnormal cell division, as witnessed through the use of time-lapse monitoring. Wong et al recently described a prediction model for development to the blastocyst stage based on early cell cycle parameters.95 Among the important factors was length of time at the three-cell stage. It is known that normal cleavage from two to four cells goes through a brief three cells intermediate before cleavage to four cells.87 A prolonged period at the three-cell stage may indicate reduced blastocyst formation95,96 and implantation potential.68,97 It has also been postulated that blastomeres are most fragile immediately before cytokinesis.87,98 Embryos with odd cell numbers are presumably in that fragile state. An increased likelihood of damage during transfer may explain the reportedly reduced implantation rates found when transferring odd numbers compared with even. In addition, some triploid embryos have been shown to abnormally cleave directly from one to three blastomeres, followed by a long delay (8.7–12.7 hours) before cleaving again to four cells.95 This demonstrates a potential link between abnormal cytokinesis patterns and aneuploidy. Quite possibly, several three cell embryos transferred in the aforementioned studies fall into this category. The importance of cell number extends to day 3 as well. However, there does appear to be a slightly more liberal range of optimal cell number on day 3 as opposed to day 2. Alikani et al found that the optimal range for blastocyst development was between seven and nine cells.99 Racowsky et al found seven to eight cells correlated with highest blastocyst viability Seminars in Reproductive Medicine

Vol. 32

No. 2/2014

(rather than formation), with < 7 and > 8 cells showing significantly lower viability.100 These and other studies demonstrate that both highly advanced and delayed cleavage rates may be suboptimal on day 3.90

Cell Symmetry Uniformity of blastomere size and the spatial relationship between blastomeres make up the symmetry component of many cleavage stage grading systems.101,102 A normal mitotic cleavage pattern is expected to ultimately result in an even number of blastomeres, with an equal distribution of cytoplasm (and therefore evenly sized blastomeres). At the cleavage stage, each successive round of cell division leads to smaller cells because there is no net gain in material.103 It is important to realize that slight asynchrony in the timing of divisions, which is normal,95,104 can be randomly captured by a static observation. Therefore, an important distinction must be made between even cell numbers of unequal size and odd cell numbers with unequal size. The latter is a typical observation before cytokinesis, where one larger cell will soon divide, creating cells of equal size to the others that have already undergone that same round of cell division.105 However, when there are larger cells in an embryo with an even number of blastomeres, it is likely due to either an unequal distribution of cytoplasm among daughter cells, individual blastomere dysfunction/arrest, or fragmentation.106 This would explain the findings from several researchers of reduced developmental and implantation potential of asymmetric embryos,87–89,93,103,106–108 as well as an increased likelihood of aneuploidy.109–112

Fragmentation To summarize multiple early definitions, fragmentation has been described as the expulsion of anucleate, membranebound cytoplasm during EC events.90,113,114 Fragmentation is primarily recorded as a cumulative percentage, although work by Alikani et al suggests that the spatial arrangement of the fragments is a predictor of implantation potential and as such devised a unique scoring system factoring in both the size and relative distribution of the fragments.114 Fragmentation has been linked to increased risk of chromosomal abnormalities,112,115–118 which explains why countless studies have associated increasing degree of fragmentation with negative IVF outcomes such as decreased blastocyst formation and implantation rates.21,87,93,100,114 Despite such a strong inverse relationship, there remains little consensus on what threshold of fragmentation is the best predictor. Some studies have shown that < 10% fragmentation is an important cutoff,119–121 while others report < 15%114 or < 20%.92,94,122 The subjectivity of assessing fragmentation undoubtedly plays a role in the lack of consensus, but the biological mechanisms of fragmentation may also provide a clearer explanation and possibly expose the need for a revised definition of fragmentation. Several studies related to the biological basis of fragmentation have been presented in the literature, encompassing divergent mechanisms that suggest multiple origins of fragmentation. This would explain the seemingly inconsistent


118

fates of individual embryos with different patterns and timing of fragmentation.60 Fragmentation has been associated with depletion of essential organelles, such as mitochondria123,124 or proteins located in polarized domains.60 Similarly, membrane components such as high-density lipoprotein125–127 and E-cadherin128 have been shown to be related to fragmentation. In addition, telomere shortening primarily due to high levels of oxidative stress in dysfunctional mitochondria has been suggested as a cause of fragmentation.129 In mice, which have extremely long telomeres relative to humans and typically do not exhibit fragmentation, artificial shortening of telomeres induced signs of reproductive aging seen in humans, including an increased incidence of fragmentation.130,131 It is unclear whether apoptosis leads to fragmentation132,133 or is perhaps a downstream consequence of fragmentation,60,128 but the involvement of aberrant cell division in fragmentation has become increasingly clear,134 as multiple researchers have shown that fragmentation is directly tied to the process of cell division.135,136 Chavez et al determined that fragmentation patterns differ between meiotic errors and mitotic errors, with triploid embryos and those with meiotic errors typically exhibiting fragmentation just before first cytokinesis, while embryos with mitotic errors often fragment later.5 The phenomenon of fragment reabsorption into blastomeres has been demonstrated repeatedly,4,5,137 but contrary to the long-standing definition of fragments, Chavez et al have recently shown the localization of chromosomes within fragments, both unencapsulated and also encapsulated in what they have termed micronuclei.5 They postulate that fragment reabsorption by the same cell has the potential to restore euploidy, while fusion to another cell would result in complex abnormalities. This proposed mechanism would suggest that mechanical removal of fragments would result in inconsistent outcomes, depending on the composition of the fragments.60,114,138,139 An automated tracking algorithm has been created to provide objective analysis of fragmentation.5 This may enhance our ability to perform statistical analyses of fragmentation patterns quantitatively to refine and standardize grading criteria in the future.

Abeyta, Behr

A limited number of morula grading systems have been proposed and/or used for embryo selection.142–145 When comparing transfer of only top quality day 3 embryos versus top quality day 4 embryos, Tao et al demonstrated a significantly increased implantation rate on day 4 (46.4 vs. 21.4%) while simultaneously reducing the number transferred.142 Heavier emphasis was placed on the day 2/day 3 score and day 4 shape and fragmentation than the degree of compaction. Ebner et al modified Tao’s system, eliminating cleavage stage scores and further classifying the degree of compaction. They demonstrated that the loss of cell volume in morulae due to either fragmentation or exclusion of entire blastomeres significantly decreased pregnancy rates on day 5. In addition, day 4 embryos showing full compaction without fragmentation formed significantly more top-quality blastocysts than fully compact morulae with fragmentation.145 In another study, researchers retrospectively compared SETs on day 4 to day 5 SET resulting in similar ongoing pregnancy rates of 38.7 and 32.1%, respectively.143 The day 5 grading system was as described by Gardner and Schoolcraft,146 whereas the day 4 system was novel, distinguished by degree of compaction and presence or absence of anomalies (vacuolization, fragmentation, excluded cells). In a recent study of practical significance, Ivec et al sought to understand the viability of developmentally delayed morulae.147 Morula grading I to III was based on the degree of fragmentation, categorized < 5, 5 to 25, and > 25%, respectively. Compared with day 4 morulae, the day 5 morula group formed significantly less clinically usable blastocysts (35.2 vs. 74.8%; p < 0.01). There was a significant negative association between fragmentation category and usable blast formation on both days. Fragmentation was negatively correlated with inner cell mass (ICM) size and number of trophectoderm cells. Delayed morula formation was associated with reduced ICM volume. More data are needed before strong conclusions can be drawn regarding the value of morula transfer, but it does appear that degree of compaction and fragmentation are key components to any morphological grading system for day 4.

Day 5 and Day 6 Assessment Day 4 Grading Although not commonly assessed, normally developing embryos are expected to reach the morula stage on day 4 postinsemination. The morula is characteristically known for the loss of discernible individual cell borders. The visual loss of cell borders is part of the compaction process and is due to the formation of tight junctions between adjacent cells as part of the process of forming a transport epithelium.140 Quite routinely, what is referred to as early compaction is noted in day 3 embryos. This often occurs between the 8- and 16cell stage and has been proposed to be a positive predictor of implantation when combined with < 10% fragmentation,141 although Desai et al did not find a significant correlation.102 To make a clear distinction from the day 3 compacting embryo, the morula exhibits the presence of compaction in addition to having undergone a fourth round of cell division (16-cell stage).

The blastocyst stage is initiated with the onset of cavitation. Formation of a blastocoel, or fluid filled cavity, coincides with cell differentiation into two distinct types, the ICM, which will form the fetus, and the trophectoderm, which will later form the amnion. As the blastocoel increases in size internal pressure builds, forcing the zona pellucida to stretch until it breaks. The expansion process culminates in complete escape (hatching) from the zona pellucida. An early attempt to evaluate blastocysts by Dokras et al placed emphasis on typical versus atypical developmental sequence.148 There were only three categories: BG1—normal developmental sequence (as described above) and distinct differentiation. BG2—atypical cavitation pattern but resulting in similar morphology to BG1. BG3—atypical/incomplete cavitation and incomplete cell differentiation. This system was not initially used for selection, but rather as a descriptive analysis. Gardner and Schoolcraft later expanded upon this concept by Seminars in Reproductive Medicine

Vol. 32

No. 2/2014

119




Abeyta, Behr

creating a grading system that described in detail both the degree of expansion and the quality and organization of the two differentiated cell types.146 The value of this grading system lies in the fact that the three different measurements can be quantified at any given time point and analyzed for individual importance in selection. Many researchers have subsequently validated the Gardner’s system, showing improved implantation rates when highly scored blastocysts are available for transfer.149,150 Balaban et al compared the systems of Dokras and Gardner and found that Gardner’s system led to significantly higher implantation rates and multiple pregnancies.149 Comparisons of PN and cleavage stage selection versus blastocyst selection have had mixed results, especially when comparing cumulative pregnancy rates,89,122,151 but as blastocyst culture has improved it has become clear that morphological selection at the blastocyst stage provides the highest implantation rates for fresh transfers.149,151,152 Several researchers have further scrutinized assessment of the ICM and trophectoderm in attempts to find the key characteristics of each. Richter et al found that both ICM size (larger) and shape (oval) were highly predictive of implantation,153 while two recent studies representing more than 1,800 single-blastocyst transfers found that the trophectoderm is significantly predictive of implantation and live birth in multivariate analyses.150,154 Further work is necessary to determine the relative weight of each characteristic and implications for miscarriage; particularly when top scoring blastocysts are not available. Although the majority of blastocyst transfers occur on day 5, it is currently common practice to extend culture until day 6 while waiting for preimplantation genetic screening (PGS) results or due to developmental delay of cavitation. The causes and consequences of delayed development are often considered, but difficult to interpret due to confounding variables such as culture conditions and issues with endometrial synchrony. Hill et al154 found that single morula transfers when no blastocysts were available on day 5 resulted in a 67% miscarriage rate and only 13% take home baby rate. Some have suggested that developmental delays lead to distorted sex ratios,155,156 higher rates of aneuploidy,156,157 and there are conflicting reports of postthaw viability.158–160 Nonetheless, viable pregnancies have occurred from day 6 transfers (with PGS) and frozen embryo transfers of day 6 frozen blastocysts with adequate efficiency to suggest this practice will continue.

Sequential Grading Systems A greater understanding of the morphological features that coincide with in vitro development of the human embryo has led to many refinements in embryo scoring and selection. However, there is a lack of consensus on the optimal days and features, which is why many researchers have sought to increase predictive power by creating sequential grading systems that combine the scoring of morphological features from multiple days. The graduated embryo score (GES) proposed by Fisch et al is an example of a cumulative, weighted scoring system. Their Seminars in Reproductive Medicine

Vol. 32

No. 2/2014

algorithm, a 100-point system that included zygote scoring and EC on day 1, and blastomere number, symmetry, and fragmentation on day 3, was found to correlate with higher blast formation rates.161 Applying this algorithm prospectively for embryo selection of day 3 and day 5 transfers, they found that GES performs better than a single day 3 observation.122 Neuber et al similarly found that a sequential assessment scoring system increased the prediction of good blastocyst formation, and later demonstrated improved selection over a static observation on day 3.162,163 Their system placed equal weight on PN alignment, EC, D2 4 cells, and D3 7 cells with low fragmentation scores. Prediction power increased with each milestone met, with embryos meeting all four criteria, the most likely to form good blastocysts (54.2% likelihood) and embryos meeting none of the criteria being significantly less likely (5.6%). In a prospective study, Rienzi et al randomly assigned patients to either day 3 or day 5 transfer.164 The day 3 transfer group had embryos chosen based on a sequential score of PN grading (described by Tesarik and Greco), day 2/day 3 cell number, blastomere size, fragmentation, and MNB status. The best features were given the lowest scores, and the embryos with the lowest cumulative scores were chosen for transfer in this group. For day 5 transfers, the embryos were chosen solely based on blastocyst morphology. With two embryos transferred in each study group, they found that the day 3 sequential scoring system performed as well as the blastocyst group, and had a significantly improved outcome compared with blastocyst transfer when including the cumulative fresh and frozen transfer data (per oocyte retrieval) due in part to the increased number of frozen embryos on day 3. Sela et al uniquely looked at all possible cleavage patterns for each cell number on days 2 and 3 and incorporated them into a sequential scoring system that also included zygote scoring, EC, day 2/day 3 cell number, and fragmentation.105 The purpose of assessing cleavage pattern was to determine the influence of nonsynchronous cleavage (three, five–seven cells) with varying degrees of cell size asymmetry. The scoring system was used to retrospectively compare embryos from 100% implantation transfers versus 0% implantation transfers. They found that certain combinations of cell size asymmetry that were associated with nonsynchronous cleavage reflected the normal developmental course and were therefore considered as part of a good cleavage pattern (►Fig. 1). Univariate and multivariate analyses both demonstrated the added benefit of including cleavage pattern in the scoring system. The area under the curve (AUC ¼ 0.707) and correct classification were highest when cleavage pattern was assessed on both day 2 and day 3. Although many researchers were able to demonstrate significantly improved selection with sequential scoring, others have questioned the usefulness of such systems. Racowsky et al tested seven candidate models for their ability to accurately select embryos for day 3 transfer.119 These models were composed of days 1–3 observations either alone or in combination. Discrimination using receiver operating characteristics curve showed that the AUC was lowest on


120


Abeyta, Behr

121

day 1 alone (AUC ¼ 0.683) and highest with combination of all three observations (median AUC ¼ 0.737), although this was only marginally better than day 2 (AUC ¼ 0.733) or day 3 (median AUC ¼ 0.732) alone. The final, best-fit model for day 3 observation included variables for age, donor egg status, cell number, fragmentation, and symmetry. It was suggested that day 3 observation alone was sufficient since this was the day of transfer and marginal increases from other days is offset by the additional time and potential detriment to embryos by increasing the number of observations. Finally, the argument was made that AUC 0.7 should be set as a benchmark for minimally acceptable discrimination. Failure to meet this benchmark led Guerif et al to conclude that the combined day 1 and day 2 evaluations of PN morphology, EC, cell number, and fragmentation were insufficient predictors of embryo viability.85 In a large prospective study based on 4,042 embryos, despite independent associations of each factor to blastocyst development, the combined AUC of 0.688 was weak, suggesting that the early parameters provided no additional benefit toward the selection of viable blastocysts over blastocyst morphology alone.

Discussion Although much of the focus has been on the results of the studies, the methodologies employed are glaringly inconsistent in three critical areas: grading criteria, design/statistical analysis, and timing of observation. Undoubtedly, these inconsistencies have contributed to the lack of consensus regarding the utility of individual and sequential scoring systems. Influential organizations such as the Society for Assisted Reproductive Technologies (SART) committee and the Alpha scientists have both attempted to increase uniformity in embryo grading. The SART committee has published a grading scheme used for reporting of day 3 transferred embryos to SART that will not only allow for direct comparison of clinics but also the creation of a vast dataset that would otherwise be impossible from custom tailored grading schemes.165,166 However, further refined, tightly controlled studies employing appropriate statistical analyses are necessary before we can definitively say what criteria should be included in any morphological assessment. When considering universal adoption of a grading system, we must be cognizant of interand intraobserver variability with each component of the system and the ultimate embryo fate decision. Research has shown that fragmentation percentage,3 localization of frag-

mentation,1 and cell symmetry3 are all prone to poor agreement among observers. Simpler systems with easily described criteria are ideally suited for widespread use. Yet to maximize the predictive value of a scoring system, we must find a means of accurately incorporating all independent measures that increase success rates. Noninvasive computer-assisted cell counting167 and fragmentation scoring programs5 may contribute to the standardization process. The gold standard for assessing the predictive power of any test criteria is a prospective randomized control trial. Unfortunately, very few of such studies have been performed in the arena of morphology grading criteria. The vast majority of cumulative scoring systems mentioned here were tested retrospectively, and compounding the issue, they were created based on retrospective analyses of individual grading criteria. In addition, the bias of what is perceived to be the ideal embryo often prevents those embryos perceived as lower quality from being transferred prospectively. To reach true significance, a study should be appropriately powered and include a multivariate analysis (particularly in multifactor grading systems) to account for the interaction among variables. A properly designed morphology study should also seek to create clear traceability of results back to individual features of the embryo. This can only be achieved if the embryos are cultured individually from start to finish. An elective SET provides clearest outcomes, whereas an all implantation versus no implantation study design introduces greater statistical vulnerability to random chance variables such as transfer technique or endometrial receptivity and to possible interactions between the transferred embryos. There is also a loss of large amounts of potentially useful data from the partial implantation groups. The evidence presented for the value of morphological assessment at the different stages clearly illustrates the importance of timing. Embryo development is a highly orchestrated sequence of events that begins in the oocyte and resumes at fertilization. Therefore, insemination time should set the clock for observation timing. However, a significant distinction must be made between fertilization with conventional insemination and ICSI. The direct injection of spermatozoa into the oocyte with ICSI allows for a more precise account of fertilization timing. Research by Nagy et al has shown that there are delays of PN formation and first cleavage with conventional insemination compared with ICSI embryos.168 This delay is attributed to the additional time required by the spermatozoa to reach and penetrate the oocyte.108 Therefore, assessments of PN formation should be delayed by Seminars in Reproductive Medicine

Vol. 32

No. 2/2014


Figure 1 Definition of good embryo cleavage patterns. Dark gray indicates a blastomere after the first even cleavage; white indicates a blastomere after the second even cleavage; and light gray indicates a blastomere after the third even cleavage. (Reprinted with permission from Sela et al. 105).


Abeyta, Behr

2 hours and EC by approximately 4 hours for embryos fertilized by conventional insemination.108 This distinction is overlooked in most if not all grading systems currently published. Accelerated and delayed embryo development on days 2 and 3 have both been shown to negatively impact implantation, further reinforcing the importance of appropriately incorporating timing into any grading system. The literature sorely lacks prospective studies focused on the optimization of the timing window for static morphological assessment as it relates to selection of viable embryos. Time lapse should help to create biologically relevant embryo assessment time frames. The technology possesses two key requirements to ideally perform the necessary studies: (1) the embryos are cultured individually from fertilization to selection for transfer and (2) the assessments are in essence continuous, eliminating both laboratory workflow issues and the bias of perceived best observation time. Yet the knowledge we gain directly from technologies such as time lapse, PGS, proteomics, and metabolomics do not operate in confinement. Overlap of these technologies in both cross-platform validation studies and in a combined prospective manner is where the greatest leaps in progress will occur in morphological assessment. Precise and accurate grading criteria are necessary attributes in the drive toward universal adoption of SET. Perhaps we have all the tools to make this a reality, but have yet to harness their full potential.

11 Downs SM. A gap-junction-mediated signal, rather than an

References

21

12

13

14

15

16

17

18

19

20

1 Arce JC, Ziebe S, Lundin K, Janssens R, Helmgaard L, Sørensen P.

2

3

4

5

6

7 8

9

10

Interobserver agreement and intraobserver reproducibility of embryo quality assessments. Hum Reprod 2006;21(8): 2141–2148 Baxter Bendus AE, Mayer JF, Shipley SK, Catherino WH. Interobserver and intraobserver variation in day 3 embryo grading. Fertil Steril 2006;86(6):1608–1615 Paternot G, Wetzels AM, Thonon F, et al. Intra- and interobserver analysis in the morphological assessment of early stage embryos during an IVF procedure: a multicentre study. Reprod Biol Endocrinol 2011;9:127 Mio Y, Maeda K. Time-lapse cinematography of dynamic changes occurring during in vitro development of human embryos. Am J Obstet Gynecol 2008;199(6):e1–e5 Chavez SL, Loewke KE, Han J, et al. Dynamic blastomere behaviour reflects human embryo ploidy by the four-cell stage. Nat Commun 2012;3:1251 Schatten G. The centrosome and its mode of inheritance: the reduction of the centrosome during gametogenesis and its restoration during fertilization. Dev Biol 1994;165(2):299–335 Mtango NR, Potireddy S, Latham KE. Oocyte quality and maternal control of development. Int Rev Cell Mol Biol 2008;268:223–290 Bergh C, Broden H, Lundin K, Hamberger L. Comparison of fertilization, cleavage and pregnancy rates of oocytes from large and small follicles. Hum Reprod 1998;13(7):1912–1915 Rosen MP, Shen S, Dobson AT, Rinaudo PF, McCulloch CE, Cedars MI. A quantitative assessment of follicle size on oocyte developmental competence. Fertil Steril 2008;90(3):684–690 Rodríguez-Fuentes A, Hernández J, García-Guzman R, Chinea E, Iaconianni L, Palumbo A. Prospective evaluation of automated follicle monitoring in 58 in vitro fertilization cycles: follicular volume as a new indicator of oocyte maturity. Fertil Steril 2010; 93(2):616–620

Seminars in Reproductive Medicine

Vol. 32

No. 2/2014

22

23

24

25

26

27

28

29

external paracrine factor, predominates during meiotic induction in isolated mouse oocytes. Zygote 2001;9(1):71–82 Albertini DF, Combelles CM, Benecchi E, Carabatsos MJ. Cellular basis for paracrine regulation of ovarian follicle development. Reproduction 2001;121(5):647–653 Coticchio G, Sereni E, Serrao L, Mazzone S, Iadarola I, Borini A. What criteria for the definition of oocyte quality? Ann N Y Acad Sci 2004;1034:132–144 Warriach HM, Chohan KR. Thickness of cumulus cell layer is a significant factor in meiotic competence of buffalo oocytes. J Vet Sci 2004;5(3):247–251 Feuerstein P, Cadoret V, Dalbies-Tran R, Guerif F, Bidault R, Royere D. Gene expression in human cumulus cells: one approach to oocyte competence. Hum Reprod 2007;22(12):3069–3077 Jungheim ES, Macones GA, Odem RR, et al. Associations between free fatty acids, cumulus oocyte complex morphology and ovarian function during in vitro fertilization. Fertil Steril 2011;95(6): 1970–1974 Karuputhula NB, Chattopadhyay R, Chakravarty B, Chaudhury K. Oxidative status in granulosa cells of infertile women undergoing IVF. Syst Biol Reprod Med 2013;59(2):91–98 Cheng EH, Chen SU, Lee TH, et al. Evaluation of telomere length in cumulus cells as a potential biomarker of oocyte and embryo quality. Hum Reprod 2013;28(4):929–936 Xia P. Intracytoplasmic sperm injection: correlation of oocyte grade based on polar body, perivitelline space and cytoplasmic inclusions with fertilization rate and embryo quality. Hum Reprod 1997;12(8):1750–1755 Kahraman S, Yakin K, Dönmez E, et al. Relationship between granular cytoplasm of oocytes and pregnancy outcome following intracytoplasmic sperm injection. Hum Reprod 2000;15(11): 2390–2393 Shen Y, Stalf T, Mehnert C, Eichenlaub-Ritter U, Tinneberg HR. High magnitude of light retardation by the zona pellucida is associated with conception cycles. Hum Reprod 2005;20(6): 1596–1606 Esfandiari N, Burjaq H, Gotlieb L, Casper RF. Brown oocytes: implications for assisted reproductive technology. Fertil Steril 2006;86(5):1522–1525 Setti AS, Figueira RC, Braga DP, Colturato SS, Iaconelli A Jr, Borges E Jr. Relationship between oocyte abnormal morphology and intracytoplasmic sperm injection outcomes: a meta-analysis. Eur J Obstet Gynecol Reprod Biol 2011;159(2):364–370 Balakier H, Bouman D, Sojecki A, Librach C, Squire JA. Morphological and cytogenetic analysis of human giant oocytes and giant embryos. Hum Reprod 2002;17(9):2394–2401 Rosenbusch B, Schneider M, Gläser B, Brucker C. Cytogenetic analysis of giant oocytes and zygotes to assess their relevance for the development of digynic triploidy. Hum Reprod 2002;17(9): 2388–2393 De Sutter P, Dozortsev D, Qian C, Dhont M. Oocyte morphology does not correlate with fertilization rate and embryo quality after intracytoplasmic sperm injection. Hum Reprod 1996;11(3): 595–597 Ebner T, Moser M, Tews G. Is oocyte morphology prognostic of embryo developmental potential after ICSI? Reprod Biomed Online 2006;12(4):507–512 ALPHA Scientists In Reproductive Medicine; ESHRE Special Interest Group Embryology. Istanbul consensus workshop on embryo assessment: proceedings of an expert meeting. Reprod Biomed Online 2011;22(6):632–646 De Vos A, Van de Velde H, Joris H, Van Steirteghem A. In-vitro matured metaphase-I oocytes have a lower fertilization rate but similar embryo quality as mature metaphase-II oocytes after intracytoplasmic sperm injection. Hum Reprod 1999;14(7): 1859–1863


122

Abeyta, Behr

30 Shu Y, Gebhardt J, Watt J, Lyon J, Dasig D, Behr B. Fertilization,

48 Scott L, Alvero R, Leondires M, Miller B. The morphology of

embryo development, and clinical outcome of immature oocytes from stimulated intracytoplasmic sperm injection cycles. Fertil Steril 2007;87(5):1022–1027 Eichenlaub-Ritter U, Schmiady H, Kentenich H, Soewarto D. Recurrent failure in polar body formation and premature chromosome condensation in oocytes from a human patient: indicators of asynchrony in nuclear and cytoplasmic maturation. Hum Reprod 1995;10(9):2343–2349 Liu L, Oldenbourg R, Trimarchi JR, Keefe DL. A reliable, noninvasive technique for spindle imaging and enucleation of mammalian oocytes. Nat Biotechnol 2000;18(2):223–225 Waterman-Storer CM. Microtubules and microscopes: how the development of light microscopic imaging technologies has contributed to discoveries about microtubule dynamics in living cells. Mol Biol Cell 1998;9(12):3263–3271 Pickering SJ, Johnson MH. The influence of cooling on the organization of the meiotic spindle of the mouse oocyte. Hum Reprod 1987;2(3):207–216 Baka SG, Toth TL, Veeck LL, Jones HW Jr, Muasher SJ, Lanzendorf SE. Evaluation of the spindle apparatus of in-vitro matured human oocytes following cryopreservation. Hum Reprod 1995; 10(7):1816–1820 Eichenlaub-Ritter U, Stahl A, Luciani JM. The microtubular cytoskeleton and chromosomes of unfertilized human oocytes aged in vitro. Hum Genet 1988;80(3):259–264 Wang WH, Meng L, Hackett RJ, Odenbourg R, Keefe DL. The spindle observation and its relationship with fertilization after intracytoplasmic sperm injection in living human oocytes. Fertil Steril 2001;75(2):348–353 Wang WH, Meng L, Hackett RJ, Keefe DL. Developmental ability of human oocytes with or without birefringent spindles imaged by Polscope before insemination. Hum Reprod 2001; 16(7):1464–1468 Eichenlaub-Ritter U, Shen Y, Tinneberg HR. Manipulation of the oocyte: possible damage to the spindle apparatus. Reprod Biomed Online 2002;5(2):117–124 Moon JH, Hyun CS, Lee SW, Son WY, Yoon SH, Lim JH. Visualization of the metaphase II meiotic spindle in living human oocytes using the Polscope enables the prediction of embryonic developmental competence after ICSI. Hum Reprod 2003;18(4):817–820 Cooke S, Tyler JPP, Driscoll GL. Meiotic spindle location and identification and its effect on embryonic cleavage plane and early development. Hum Reprod 2003;18(11):2397–2405 Keefe D, Liu L, Wang W, Silva C. Imaging meiotic spindles by polarization light microscopy: principles and applications to IVF. Reprod Biomed Online 2003;7(1):24–29 Held E, Mertens EM, Mohammadi-Sangcheshmeh A, et al. Zona pellucida birefringence correlates with developmental capacity of bovine oocytes classified by maturational environment, COC morphology and G6PDH activity. Reprod Fertil Dev 2012;24(4): 568–579 Payne D, Flaherty SP, Barry MF, Matthews CD. Preliminary observations on polar body extrusion and pronuclear formation in human oocytes using time-lapse video cinematography. Hum Reprod 1997;12(3):532–541 Tesarik J, Greco E. The probability of abnormal preimplantation development can be predicted by a single static observation on pronuclear stage morphology. Hum Reprod 1999;14(5): 1318–1323 Nagy ZP, Dozortsev D, Diamond M, et al. Pronuclear morphology evaluation with subsequent evaluation of embryo morphology significantly increases implantation rates. Fertil Steril 2003; 80(1):67–74 Scott LA, Smith S. The successful use of pronuclear embryo transfers the day following oocyte retrieval. Hum Reprod 1998; 13(4):1003–1013

human pronuclear embryos is positively related to blastocyst development and implantation. Hum Reprod 2000;15(11): 2394–2403 Tesarik J, Kopecny V. Assembly of the nucleolar precursor bodies in human male pronuclei is correlated with an early RNA synthetic activity. Exp Cell Res 1990;191(1):153–156 Balaban B, Yakin K, Urman B, Isiklar A, Tesarik J. Pronuclear morphology predicts embryo development and chromosome constitution. Reprod Biomed Online 2004;8(6):695–700 Zollner U, Zollner KP, Hartl G, Dietl J, Steck T. The use of a detailed zygote score after IVF/ICSI to obtain good quality blastocysts: the German experience. Hum Reprod 2002; 17(5):1327–1333 Salumets A, Hydén-Granskog C, Suikkari AM, Tiitinen A, Tuuri T. The predictive value of pronuclear morphology of zygotes in the assessment of human embryo quality. Hum Reprod 2001;16(10): 2177–2181 James AN, Hennessy S, Reggio B, Wiemer K, Larsen F, Cohen J. The limited importance of pronuclear scoring of human zygotes. Hum Reprod 2006;21(6):1599–1604 Nicoli A, Capodanno F, Moscato L, et al. Analysis of pronuclear zygote configurations in 459 clinical pregnancies obtained with assisted reproductive technique procedures. Reprod Biol Endocrinol 2010;8:77 Jaroudi K, Al-Hassan S, Sieck U, Al-Sufyan H, Al-Kabra M, Coskun S. Zygote transfer on day 1 versus cleavage stage embryo transfer on day 3: a prospective randomized trial. Hum Reprod 2004; 19(3):645–648 Ludwig M, Schöpper B, Al-Hasani S, Diedrich K. Clinical use of a pronuclear stage score following intracytoplasmic sperm injection: impact on pregnancy rates under the conditions of the German embryo protection law. Hum Reprod 2000;15(2): 325–329 Montag M, van der Ven H; German Pronuclear Morphology Study Group. Evaluation of pronuclear morphology as the only selection criterion for further embryo culture and transfer: results of a prospective multicentre study. Hum Reprod 2001;16(11): 2384–2389 Gardner RL, Davies TJ. An investigation of the origin and significance of bilateral symmetry of the pronuclear zygote in the mouse. Hum Reprod 2006;21(2):492–502 Edwards RG, Beard HK. Oocyte polarity and cell determination in early mammalian embryos. Mol Hum Reprod 1997;3(10): 863–905 Antczak M, Van Blerkom J. Temporal and spatial aspects of fragmentation in early human embryos: possible effects on developmental competence and association with the differential elimination of regulatory proteins from polarized domains. Hum Reprod 1999;14(2):429–447 Boiso I, Veiga A, Edwards RG. Fundamentals of human embryonic growth in vitro and the selection of high-quality embryos for transfer. Reprod Biomed Online 2002;5(3):328–350 Shoukir Y, Campana A, Farley T, Sakkas D. Early cleavage of invitro fertilized human embryos to the 2-cell stage: a novel indicator of embryo quality and viability. Hum Reprod 1997; 12(7):1531–1536 Sakkas D, Shoukir Y, Chardonnens D, Bianchi PG, Campana A. Early cleavage of human embryos to the two-cell stage after intracytoplasmic sperm injection as an indicator of embryo viability. Hum Reprod 1998;13(1):182–187 Trounson AO, Mohr LR, Wood C, Leeton JF. Effect of delayed insemination on in-vitro fertilization, culture and transfer of human embryos. J Reprod Fertil 1982;64(2):285–294 Balakier H, MacLusky NJ, Casper RF. Characterization of the first cell cycle in human zygotes: implications for cryopreservation. Fertil Steril 1993;59(2):359–365

31

32

33

34

35

36

37

38

39

40

41

42

43

44

45

46

47

49

50

51

52

53

54

55

56

57

58

59

60

61

62

63

64

65


Vol. 32

No. 2/2014

123




Abeyta, Behr

66 Capmany G, Taylor A, Braude PR, Bolton VN. The timing of

67

68

69

70

71

72

73

74

75

76

77

78

79

80

81

82

83

84

85

pronuclear formation, DNA synthesis and cleavage in the human 1-cell embryo. Mol Hum Reprod 1996;2(5):299–306 Van Blerkom J. Occurrence and developmental consequences of aberrant cellular organization in meiotically mature human oocytes after exogenous ovarian hyperstimulation. J Electron Microsc Tech 1990;16(4):324–346 Meseguer M, Herrero J, Tejera A, Hilligsøe KM, Ramsing NB, Remohí J. The use of morphokinetics as a predictor of embryo implantation. Hum Reprod 2011;26(10):2658–2671 Montag M, Liebenthron J, Köster M. Which morphological scoring system is relevant in human embryo development? Placenta 2011;32(Suppl 3):S252–S256 Edwards RG, Purdy JM, Steptoe PC, Walters DE. The growth of human preimplantation embryos in vitro. Am J Obstet Gynecol 1981;141(4):408–416 Van Royen E, Mangelschots K, Vercruyssen M, et al. Multinucleation in cleavage stage embryos. Hum Reprod 2003;18(5): 1062–1069 Munné S, Cohen J. Unsuitability of multinucleated human blastomeres for preimplantation genetic diagnosis. Hum Reprod 1993;8(7):1120–1125 Lopata A, Kohlman D, Johnston I. The fine structure of normal and abnormal human embryos developed in culture. In: Beier HM, Lindner HR, eds. Fertilization of the Human Egg In Vitro. Heidelberg: Springer; 1983:189 Tesarík J, Kopecný V, Plachot M, Mandelbaum J. Ultrastructural and autoradiographic observations on multinucleated blastomeres of human cleaving embryos obtained by in-vitro fertilization. Hum Reprod 1987;2(2):127–136 Hardy K, Winston RML, Handyside AH. Binucleate blastomeres in preimplantation human embryos in vitro: failure of cytokinesis during early cleavage. J Reprod Fertil 1993;98(2):549–558 Winston NJ, Braude PR, Pickering SJ, et al. The incidence of abnormal morphology and nucleocytoplasmic ratios in 2-, 3and 5-day human pre-embryos. Hum Reprod 1991;6(1):17–24 Hertig AT, Rock J, Adams EC, Mulligan WJ. On the preimplantation stages of the human ovum: a description of four normal and four abnormal specimens ranging from the second to the fifth day of development. Contrib Embryol 1954;35:199–220 Plachot M, Mandelbaum J, Junca AM, Salat-Baroux J, Cohen J. Impairment of human embryo development after abnormal in vitro fertilization. Ann N Y Acad Sci 1985;442:336–341 Sadowy S, Tomkin G, Munné S, Ferrara-Congedo T, Cohen J. Impaired development of zygotes with uneven pronuclear size. Zygote 1998;6(2):137–141 Pickering SJ, Taylor A, Johnson MH, Braude PR. An analysis of multinucleated blastomere formation in human embryos. Hum Reprod 1995;10(7):1912–1922 Kligman I, Benadiva C, Alikani M, Munne S. The presence of multinucleated blastomeres in human embryos is correlated with chromosomal abnormalities. Hum Reprod 1996;11(7): 1492–1498 Ambroggio J, Gindoff PR, Dayal MB, et al. Multinucleation of a sibling blastomere on day 2 suggests unsuitability for embryo transfer in IVF-preimplantation genetic screening cycles. Fertil Steril 2011;96(4):856–859 Pelinck MJ, De Vos M, Dekens M, Van der Elst J, De Sutter P, Dhont M. Embryos cultured in vitro with multinucleated blastomeres have poor implantation potential in human in-vitro fertilization and intracytoplasmic sperm injection. Hum Reprod 1998;13(4): 960–963 Saldeen P, Sundström P. Nuclear status of four-cell preembryos predicts implantation potential in in vitro fertilization treatment cycles. Fertil Steril 2005;84(3):584–589 Guerif F, Le Gouge A, Giraudeau B, et al. Limited value of morphological assessment at days 1 and 2 to predict blastocyst


Vol. 32

No. 2/2014

86

87

88

89

90

91

92

93

94

95

96

97

98

99

100

101

102

103

development potential: a prospective study based on 4042 embryos. Hum Reprod 2007;22(7):1973–1981 Guerif F, Lemseffer M, Leger J, et al. Does early morphology provide additional selection power to blastocyst selection for transfer? Reprod Biomed Online 2010;21(4):510–519 Ziebe S, Petersen K, Lindenberg S, Andersen AG, Gabrielsen A, Andersen AN. Embryo morphology or cleavage stage: how to select the best embryos for transfer after in-vitro fertilization. Hum Reprod 1997;12(7):1545–1549 Holte J, Berglund L, Milton K, et al. Construction of an evidencebased integrated morphology cleavage embryo score for implantation potential of embryos scored and transferred on day 2 after oocyte retrieval. Hum Reprod 2007;22(2):548–557 Scott L, Finn A, O’Leary T, McLellan S, Hill J. Morphologic parameters of early cleavage-stage embryos that correlate with fetal development and delivery: prospective and applied data for increased pregnancy rates. Hum Reprod 2007;22(1):230–240 Cummins JM, Breen TM, Harrison KL, Shaw JM, Wilson LM, Hennessey JF. A formula for scoring human embryo growth rates in in vitro fertilization: its value in predicting pregnancy and in comparison with visual estimates of embryo quality. J In Vitro Fert Embryo Transf 1986;3(5):284–295 Claman P, Armant DR, Seibel MM, Wang TA, Oskowitz SP, Taymor ML. The impact of embryo quality and quantity on implantation and the establishment of viable pregnancies. J In Vitro Fert Embryo Transf 1987;4(4):218–222 Staessen C, Camus M, Bollen N, Devroey P, Van Steirteghem AC. The relationship between embryo quality and the occurrence of multiple pregnancies. Fertil Steril 1992;57(3):626–630 Giorgetti C, Terriou P, Auquier P, et al. Embryo score to predict implantation after in-vitro fertilization: based on 957 single embryo transfers. Hum Reprod 1995;10(9):2427–2431 Van Royen E, Mangelschots K, De Neubourg D, et al. Characterization of a top quality embryo, a step towards single-embryo transfer. Hum Reprod 1999;14(9):2345–2349 Wong CC, Loewke KE, Bossert NL, et al. Non-invasive imaging of human embryos before embryonic genome activation predicts development to the blastocyst stage. Nat Biotechnol 2010;28(10): 1115–1121 Cruz M, Garrido N, Herrero J, Pérez-Cano I, Muñoz M, Meseguer M. Timing of cell division in human cleavage-stage embryos is linked with blastocyst formation and quality. Reprod Biomed Online 2012;25(4):371–381 Lemmen JG, Agerholm I, Ziebe S. Kinetic markers of human embryo quality using time-lapse recordings of IVF/ICSI-fertilized oocytes. Reprod Biomed Online 2008;17(3):385–391 Lassalle B, Testart J, Renard JP. Human embryo features that influence the success of cryopreservation with the use of 1,2 propanediol. Fertil Steril 1985;44(5):645–651 Alikani M, Calderon G, Tomkin G, Garrisi J, Kokot M, Cohen J. Cleavage anomalies in early human embryos and survival after prolonged culture in-vitro. Hum Reprod 2000;15(12):2634–2643 Racowsky C, Combelles CM, Nureddin A, et al. Day 3 and day 5 morphological predictors of embryo viability. Reprod Biomed Online 2003;6(3):323–331 Steer CV, Mills CL, Tan SL, Campbell S, Edwards RG. The cumulative embryo score: a predictive embryo scoring technique to select the optimal number of embryos to transfer in an in-vitro fertilization and embryo transfer programme. Hum Reprod 1992; 7(1):117–119 Desai NN, Goldstein J, Rowland DY, Goldfarb JM. Morphological evaluation of human embryos and derivation of an embryo quality scoring system specific for day 3 embryos: a preliminary study. Hum Reprod 2000;15(10):2190–2196 Hnida C, Ziebe S. Total cytoplasmic volume as biomarker of fragmentation in human embryos. J Assist Reprod Genet 2004; 21(9):335–340


124

Abeyta, Behr

104 Scott L. The biological basis of non-invasive strategies for selec-

124 Lin DP, Huang CC, Wu HM, Cheng TC, Chen CI, Lee MS. Comparison

tion of human oocytes and embryos. Hum Reprod Update 2003; 9(3):237–249 Sela R, Samuelov L, Almog B, et al. An embryo cleavage pattern based on the relative blastomere size as a function of cell number for predicting implantation outcome. Fertil Steril 2012;98(3): 650–656, e4 Roux C, Joanne C, Agnani G, Fromm M, Clavequin MC, Bresson JL. Morphometric parameters of living human in-vitro fertilization embryos; importance of the asynchronous division process. Hum Reprod 1995;10(5):1201–1207 Van Royen E, Mangelschots K, De Neubourg D, Laureys I, Ryckaert G, Gerris J. Calculating the implantation potential of day 3 embryos in women younger than 38 years of age: a new model. Hum Reprod 2001;16(2):326–332 Rienzi L, Ubaldi F, Iacobelli M, et al. Significance of morphological attributes of the early embryo. Reprod Biomed Online 2005; 10(5):669–681 Hardarson T, Hanson C, Sjögren A, Lundin K. Human embryos with unevenly sized blastomeres have lower pregnancy and implantation rates: indications for aneuploidy and multinucleation. Hum Reprod 2001;16(2):313–318 Magli MC, Gianaroli L, Ferraretti AP, Lappi M, Ruberti A, Farfalli V. Embryo morphology and development are dependent on the chromosomal complement. Fertil Steril 2007;87(3):534–541 Ziebe S, Lundin K, Loft A, et al; CEMAS II and Study Group. FISH analysis for chromosomes 13, 16, 18, 21, 22, X and Y in all blastomeres of IVF pre-embryos from 144 randomly selected donated human oocytes and impact on pre-embryo morphology. Hum Reprod 2003;18(12):2575–2581 Munné S. Chromosome abnormalities and their relationship to morphology and development of human embryos. Reprod Biomed Online 2006;12(2):234–253 Puissant F, Van Rysselberge M, Barlow P, Deweze J, Leroy F. Embryo scoring as a prognostic tool in IVF treatment. Hum Reprod 1987;2(8):705–708 Alikani M, Cohen J, Tomkin G, Garrisi GJ, Mack C, Scott RT. Human embryo fragmentation in vitro and its implications for pregnancy and implantation. Fertil Steril 1999;71(5):836–842 Plachot M, Junca AM, Mandelbaum J, de Grouchy J, Salat-Baroux J, Cohen J. Chromosome investigations in early life. II. Human preimplantation embryos. Hum Reprod 1987;2(1):29–35 Pellestor F, Sèle B. Assessment of aneuploidy in the human female by using cytogenetics of IVF failures. Am J Hum Genet 1988;42(2): 274–283 Munné S, Cohen J. Chromosome abnormalities in human embryos. Hum Reprod Update 1998;4(6):842–855 Magli MC, Gianaroli L, Ferraretti AP. Chromosomal abnormalities in embryos. Mol Cell Endocrinol 2001;183:29–34 Racowsky C, Ohno-Machado L, Kim J, Biggers JD. Is there an advantage in scoring early embryos on more than one day? Hum Reprod 2009;24(9):2104–2113 Kashyap S, Cedars M, Shen S, Rosen M, Grady D, Wells G. Real data mathematical modeling to predict successful embryo implantation. Fertil Steril 2008;90:S224 Pelinck MJ, Hoek A, Simons AH, Heineman MJ, van Echten-Arends J, Arts EG. Embryo quality and impact of specific embryo characteristics on ongoing implantation in unselected embryos derived from modified natural cycle in vitro fertilization. Fertil Steril 2010;94(2):527–534 Fisch JD, Sher G, Adamowicz M, Keskintepe L. The graduated embryo score predicts the outcome of assisted reproductive technologies better than a single day 3 evaluation and achieves results associated with blastocyst transfer from day 3 embryo transfer. Fertil Steril 2003;80(6):1352–1358 Wilding M, Dale B, Marino M, et al. Mitochondrial aggregation patterns and activity in human oocytes and preimplantation embryos. Hum Reprod 2001;16(5):909–917

of mitochondrial DNA contents in human embryos with good or poor morphology at the 8-cell stage. Fertil Steril 2004;81(1): 73–79 Zamah AM, Browne RW, Conti G, Jaggavarapu SR, Sridhar V, Fujimoto VY. High density lipoprotein (HDL) particle size and composition predicts embryo fragmentation. Fertil Steril 2009; 92:S96–S97 Browne RW, Bloom MS, Shelly WB, Ocque AJ, Huddleston HG, Fujimoto VY. Follicular fluid high density lipoprotein-associated micronutrient levels are associated with embryo fragmentation during IVF. J Assist Reprod Genet 2009;26(11-12): 557–560 Fujimoto VY, Kane JP, Ishida BY, Bloom MS, Browne RW. Highdensity lipoprotein metabolism and the human embryo. Hum Reprod Update 2010;16(1):20–38 Alikani M. Epithelial cadherin distribution in abnormal human preimplantation embryos. Hum Reprod 2005;20(12):3369–3375 Liu L, Trimarchi JR, Smith PJS, Keefe DL. Mitochondrial dysfunction leads to telomere attrition and genomic instability. Aging Cell 2002;1(1):40–46 Kalmbach KH, Fontes Antunes DM, Dracxler RC, et al. Telomeres and human reproduction. Fertil Steril 2013;99(1):23–29 Liu L, Blasco MA, Keefe DL. Requirement of functional telomeres for metaphase chromosome alignments and integrity of meiotic spindles. EMBO Rep 2002;3(3):230–234 Jurisicova A, Varmuza S, Casper RF. Programmed cell death and human embryo fragmentation. Mol Hum Reprod 1996;2(2): 93–98 Liu L, Blasco M, Trimarchi J, Keefe D. An essential role for functional telomeres in mouse germ cells during fertilization and early development. Dev Biol 2002;249(1):74–84 Keefe DL, Franco S, Liu L, et al. Telomere length predicts embryo fragmentation after in vitro fertilization in women—toward a telomere theory of reproductive aging in women. Am J Obstet Gynecol 2005;192(4):1256–1260, discussion 1260–1261 Alikani M, Schimmel T, Willadsen SM. Cytoplasmic fragmentation in activated eggs occurs in the cytokinetic phase of the cell cycle, in lieu of normal cytokinesis, and in response to cytoskeletal disorder. Mol Hum Reprod 2005;11(5): 335–344 Liu L, Trimarchi JR, Smith PJ, Keefe DL. Checkpoint for DNA integrity at the first mitosis after oocyte activation. Mol Reprod Dev 2002;62(2):277–288 Hardarson T, Löfman C, Coull G, Sjögren A, Hamberger L, Edwards RG. Internalization of cellular fragments in a human embryo: time-lapse recordings. Reprod Biomed Online 2002; 5(1):36–38 Dozortsev D, Ermilov A, El-Mowafi DM, Diamond M. The impact of cellular fragmentation induced experimentally at different stages of mouse preimplantation development. Hum Reprod 1998;13(5):1307–1311 Eftekhari-Yazdi P, Valojerdi MR, Ashtiani SK, Eslaminejad MB, Karimian L. Effect of fragment removal on blastocyst formation and quality of human embryos. Reprod Biomed Online 2006; 13(6):823–832 Buster JE, Bustillo M, Rodi IA, et al. Biologic and morphologic development of donated human ova recovered by nonsurgical uterine lavage. Am J Obstet Gynecol 1985;153(2):211–217 Skiadas CC, Jackson KV, Racowsky C. Early compaction on day 3 may be associated with increased implantation potential. Fertil Steril 2006;86(5):1386–1391 Tao J, Tamis R, Fink K, Williams B, Nelson-White T, Craig R. The neglected morula/compact stage embryo transfer. Hum Reprod 2002;17(6):1513–1518 Feil D, Henshaw RC, Lane M. Day 4 embryo selection is equal to Day 5 using a new embryo scoring system validated in single embryo transfers. Hum Reprod 2008;23(7):1505–1510

105

106

107

108

109

110

111

112

113

114

115

116

117 118 119

120

121

122

123

125

126

127

128 129

130 131

132

133

134

135

136

137

138

139

140

141

142

143


Vol. 32

No. 2/2014

125




Abeyta, Behr

144 Pantos K, Makrakis E, Chronopoulou M, Biba M, Perdikaris A,

158 Levens ED, Whitcomb BW, Hennessy S, James AN, Yauger BJ,

Dafereras A. Day 4 versus day 3 embryo transfer: a prospective study of clinical outcomes. Fertil Steril 2008;89(3):573–577 Ebner T, Moser M, Shebl O, Sommergruber M, Gaiswinkler U, Tews G. Morphological analysis at compacting stage is a valuable prognostic tool for ICSI patients. Reprod Biomed Online 2009; 18(1):61–66 Gardner DK, Schoolcraft WB. In vitro culture of human blastocyst. In: Jansen R, Mortimer D, eds. Towards Reproductive Certainty: Infertility and Genetics Beyond 1999. Carnforth: Parthenon Press; 1999:378–388 Ivec M, Kovacic B, Vlaisavljevic V. Prediction of human blastocyst development from morulas with delayed and/or incomplete compaction. Fertil Steril 2011;96(6):1473–1478, e2 Dokras A, Sargent IL, Barlow DH. Human blastocyst grading: an indicator of developmental potential? Hum Reprod 1993;8(12): 2119–2127 Balaban B, Yakin K, Urman B. Randomized comparison of two different blastocyst grading systems. Fertil Steril 2006;85(3): 559–563 Ahlström A, Westin C, Reismer E, Wikland M, Hardarson T. Trophectoderm morphology: an important parameter for predicting live birth after single blastocyst transfer. Hum Reprod 2011;26(12):3289–3296 Guerif F, Lemseffer M, Bidault R, et al. Single Day 2 embryo versus blastocyst-stage transfer: a prospective study integrating fresh and frozen embryo transfers. Hum Reprod 2009;24(5): 1051–1058 Papanikolaou EG, Kolibianakis EM, Tournaye H, et al. Live birth rates after transfer of equal number of blastocysts or cleavagestage embryos in IVF. A systematic review and meta-analysis. Hum Reprod 2008;23(1):91–99 Richter KS, Harris DC, Daneshmand ST, Shapiro BS. Quantitative grading of a human blastocyst: optimal inner cell mass size and shape. Fertil Steril 2001;76(6):1157–1167 Hill MJ, Richter KS, Heitmann RJ, et al. Trophectoderm grade predicts outcomes of single-blastocyst transfers. Fertil Steril 2013;99(5):1283–1289, e1 Dean JH, Chapman MG, Sullivan EA. The effect on human sex ratio at birth by assisted reproductive technology (ART) procedures— an assessment of babies born following single embryo transfers, Australia and New Zealand, 2002-2006. BJOG 2010;117(13): 1628–1634 Alfarawati S, Fragouli E, Colls P, et al. The relationship between blastocyst morphology, chromosomal abnormality, and embryo gender. Fertil Steril 2011;95(2):520–524 Kroener L, Ambartsumyan G, Briton-Jones C, et al. The effect of timing of embryonic progression on chromosomal abnormality. Fertil Steril 2012;98(4):876–880

Larsen FW. Blastocyst development rate impacts outcome in cryopreserved blastocyst transfer cycles. Fertil Steril 2008; 90(6):2138–2143 El-Toukhy T, Wharf E, Walavalkar R, et al. Delayed blastocyst development does not influence the outcome of frozen-thawed transfer cycles. BJOG 2011;118(13):1551–1556 Hashimoto S, Amo A, Hama S, Ito K, Nakaoka Y, Morimoto Y. Growth retardation in human blastocysts increases the incidence of abnormal spindles and decreases implantation potential after vitrification. Hum Reprod 2013;28(6):1528–1535 Fisch JD, Rodriguez H, Ross R, Overby G, Sher G. The Graduated Embryo Score (GES) predicts blastocyst formation and pregnancy rate from cleavage-stage embryos. Hum Reprod 2001;16(9): 1970–1975 Neuber E, Rinaudo P, Trimarchi JR, Sakkas D. Sequential assessment of individually cultured human embryos as an indicator of subsequent good quality blastocyst development. Hum Reprod 2003;18(6):1307–1312 Neuber E, Mahutte NG, Arici A, Sakkas D. Sequential embryo assessment outperforms investigator-driven morphological assessment at selecting a good quality blastocyst. Fertil Steril 2006; 85(3):794–796 Rienzi L, Ubaldi F, Iacobelli M, et al. Day 3 embryo transfer with combined evaluation at the pronuclear and cleavage stages compares favourably with day 5 blastocyst transfer. Hum Reprod 2002;17(7):1852–1855 Racowsky C, Stern JE, Gibbons WE, Behr B, Pomeroy KO, Biggers JD. National collection of embryo morphology data into Society for Assisted Reproductive Technology Clinic Outcomes Reporting System: associations among day 3 cell number, fragmentation and blastomere asymmetry, and live birth rate. Fertil Steril 2011; 95(6):1985–1989 Vernon M, Stern JE, Ball GD, Wininger D, Mayer J, Racowsky C. Utility of the national embryo morphology data collection by the Society for Assisted Reproductive Technologies (SART): correlation between day-3 morphology grade and live-birth outcome. Fertil Steril 2011;95(8):2761–2763 Newmark JA, Warger WC II, Chang C, et al. Determination of the number of cells in preimplantation embryos by using noninvasive optical quadrature microscopy in conjunction with differential interference contrast microscopy. Microsc Microanal 2007;13(2): 118–127 Nagy ZP, Janssenswillen C, Janssens R, et al. Timing of oocyte activation, pronucleus formation and cleavage in humans after intracytoplasmic sperm injection (ICSI) with testicular spermatozoa and after ICSI or in-vitro fertilization on sibling oocytes with ejaculated spermatozoa. Hum Reprod 1998;13(6): 1606–1612

145

146

147

148

149

150

151

152

153

154

155

156

157


Vol. 32

No. 2/2014

159

160

161

162

163

164

165

166

167

168


126

Copyright of Seminars in Reproductive Medicine is the property of Thieme Medical Publishing Inc. and its content may not be copied or emailed to multiple sites or posted to a listserv without the copyright holder's express written permission. However, users may print, download, or email articles for individual use.

Metabolomic assessment of embryo viability.

Mitochondrial DNA Assessment to Determine Oocyte and Embryo Viability.

Diagnosis of human preimplantation embryo viability.

Inter-laboratory agreement on embryo classification and clinical decision: Conventional morphological assessment vs. time lapse.

Operative assessment of intestinal viability.

Induction of autophagy improves embryo viability in cloned mouse embryos.

Effects of chromic chloride on chick embryo fibroblast viability.

Assessment of hamster blastocysts derived from eight-cell embryos cultured in hamster embryo culture medium-2 (HECM-2): cell numbers and viability following embryo transfer.

Assessment of a test of renal viability.

A morphological study on the pineal gland of human embryo.

Viability assessment of allograft valves by autoradiography.

Morphological transformation of Syrian hamster embryo cells by mezerein.

Bacteria viability assessment after photocatalytic treatment.

Morphological assessment of the retina in uveitis.

Morphological assessment of ancestry using cranial macromorphoscopics.

Rapid assessment of bacterial viability by flow cytometry.

Topics in the routine assessment of newborn puppy viability.

Prognostic value of viability assessment in chronic ischaemic LV dysfunction.

Assessment of planctomycetes cell viability after pollutants exposure.

Assessment of bone viability by scintiscanning in frostbite injuries.

Nuclear cardiac imaging for the assessment of myocardial viability.

Assessment of cell viability after manipulation with insulator-based dielectrophoresis.

Effects of (R)-deoxycoformycin (pentostatin) on intrauterine nucleoside catabolism and embryo viability in the pregnant mouse.

Freezing behavior of adherent neuron-like cells and morphological change and viability of post-thaw cells.