Macroenvironment

In order to minimize exposures to contaminants, IVF laboratories must rigorously control air quality. To date, there is no standard practice for the optimization of IVF outcomes. However, IVF laboratories generally have their own central air handling unit (AHU) and/or mobile filter units where the outside (plus recirculating) air is forced through high-efficiency particulate air (HEPA) filters, along with activated charcoal, potassium permanganate, and/or activated carbon filters. The HEPA filters remove particulate matter (i.e., ≥0.3 µm in size) and absorb mold spores and bacteria. The carbon or charcoal filters remove VOCs. In addition, many laboratories have filters, housed within the HVAC ducts, that use chemical/absorbent filtration and/or UV light irradiation to further clean the air and remove microbes and particulates (≥0.01 µm in size), bacteria, and viruses. To maintain the clean air environment, the laboratory should have a high tightness factor (walls and ceilings should have no penetrations), have multiple air exchanges per hour (i.e., 15 to 25 air exchanges/hour), and be under positive pressure to prevent contaminants from adjacent rooms entering the laboratory.

To minimize VOC emissions, all furniture within the laboratory should be stainless steel. The laboratory should have smooth, nonporous walls and lighting should be sealed within special units in order to reduce airborne particles. Inline HEPA and activated carbon filters should be present to remove potential contaminants, such as benzene, isopropanol, and pentane from compressed gas (N ₂ and CO ₂ ).

A major task for the laboratory is to ensure that a uniform and optimal environment is maintained. It is therefore necessary to use laboratory equipment and media that provide ideal conditions for embryo development: stable temperatures (37°C) and a pH range of 7.2 to 7.4. Fluctuations in temperature have been shown to be detrimental to the oocyte’s microtubular system, causing reductions in spindle size, disorganization of microtubules within the spindle, and, in some cases, complete absence of microtubules. Interference with spindle organization can prevent chromosome disassociation, resulting in aneuploidy. Changes in pH could perturb metabolic homeostasis, stimulate or prevent induction of specific molecular pathways, alter gene expression profiles, and influence embryo development.

IVF laboratories have many pieces of equipment, including incubators, microscopes, lasers, heating stages, water baths, and storage Dewars. There are a variety of options for each piece of equipment; the choice may depend on cost, specific procedures, flow, and size of the laboratory. Incubators may be typical box incubators or bench-top incubators. The incubators may be humidified or dry and/or capable of maintaining low oxygen tension. Stereoscopes may be used for stripping the oocytes, while inverted microscopes with micromanipulators are used to perform intracytoplasmic sperm injection (ICSI), “assisted hatching,” or an embryo biopsy. Regardless of selection, appropriate equipment quality control measures should be performed. An example of a schedule is shown in Table 32.1 .

Additionally, quality control testing must be performed on all consumables (e.g., media and plastic ware) that are either newly introduced into the laboratory or are a recurring supply (i.e., a new lot). The purpose of the testing is to make sure that no embryotoxic substances are introduced into the system. This testing is performed using a bioassay (e.g., mouse embryo development, human sperm survival assay) that compares the test results of new material with those of control material that was previously used successfully in the laboratory.

Microenvironment

The media system is a vital component in the process of handling gametes and embryos. The goal is to emulate the in vivo environment as closely as possible. However, the in vivo milieu is dynamic while the in vitro environment is static. Therefore a single ideal culture medium is elusive. The media used in IVF laboratories today consider these and other physiologic issues and provide the most appropriate components at various stages of the IVF process.

Media can be classified based on nutrients (simple or complex) or culture process (one-step, sequential). Numerous studies on embryo metabolism as well as examination of tubal and uterine fluid composition have led to advances in media. There are several complex formulations, each composed of various amounts of substrates including ions, carbohydrates, amino acids, protein, antioxidants, antibiotics, buffers, and chelators. The media system used by the laboratory is dependent on the overall culture system and clinical outcomes.

Media can also be classified based on the buffering system in use. As mentioned previously, the pH of the media is critical to embryo development. The pH of the surrounding environment influences sperm function, embryo metabolism, and organelle localization. Safety concerns and outcomes with respect to human embryo development have narrowed the buffering systems in use within the IVF laboratory. The buffering systems most utilized depend on the particular purpose and the equipment available ( Table 32.2 ). Typically, a sodium bicarbonate buffered based medium is used for culture purposes because detailed studies have shown superior embryo development in the presence of carbon dioxide (CO ₂ ) versus room atmosphere. The pH is maintained by way of gas equilibrium; therefore, the equipment must have CO ₂ sensors and adjust the amount of CO ₂ pressure to maintain the desired pH unique to the medium. A medium that is sodium bicarbonate–controlled and remains in room atmosphere (outside of a CO ₂ -controlled environment) even for a short time is susceptible to significant pH changes. Therefore, for procedures performed in room atmosphere, the inclusion of a zwitterionic buffer such as MOPS (pH = 6.5 to 7.9; pK _a 7.0 to 7.3) or HEPES (pH = 6.8 to 8.2; pK _a 7.2 to 7.5) is required. Both are effective and have been shown to work well clinically. This nonbicarbonate medium is used mainly for procedures such as oocyte collection, oocyte stripping, ICSI, and biopsy. Alternatively, laboratories may have isolettes where CO ₂ can be supplied and controlled and a bicarbonate buffer system can be used.

In addition to the environment itself, laboratories must be staffed adequately with properly trained personnel. The work in the laboratory is extremely labor-intensive. Laboratory personnel must have discipline, be detail-oriented, and function as a team. The number of staff depends on the number and types of procedures performed. An IVF lab that is performing all procedures requires a minimum of two embryologists, with an additional embryologist for every 100 to 150 cycles. Each laboratory procedure is standardized. Embryologists follow the standard operating procedures (SOPs) created for every procedure performed in the laboratory. Adherence to the SOPs ensures consistency and reduces systematic errors. As part of the quality assurance/control system, analytics are periodically performed and each embryologist participates in internal and/or external quality assurance programs for each procedure.

Procedures

The procedures performed in the laboratory are specific to the developmental stage of gametes and embryos ( Fig. 32.1 ). Additionally, embryologists must coordinate efforts so the timing of events is performed precisely. The following sections cover the concepts underlying and a detailed description of common procedures.

Timing of embryology procedures.

Daily embryology tasks are dictated by the current day in culture. Culture media are prepared the day prior to any procedure to allow for pH equilibration. Tasks that need to be performed are represented by bars. AM/PM represent time of day.

Oocyte Collection

The egg harvest entails aspirating follicular contents from antral follicles. The FF is transferred to the laboratory for evaluation. Approximately 75% to 80% of the follicular aspirates will contain an oocyte. The size of the follicle dictates the likelihood of oocyte recovery and the likelihood that it is mature ( Fig. 32.2 ). Nuclear and cytoplasmic maturation of the oocyte is critical for normal fertilization and embryo development.

Odds of aspiration of a nuclear mature oocyte (Metaphase II, MII) relative to lead group greater than 18 mm.

(Data from Rosen MP, Shen S, Dobson AT, et al: A quantitative assessment of follicle size on oocyte developmental competence. Fertil Steril 90:684–690, 2008.)

In order to prepare for an oocyte collection, each individual case must be prepared for in advance. The day before retrieval, culture dishes are made and equilibrated under controlled conditions with a constant pH and stable temperature at 37°C (see previous text).

Once the FF has been collected, it is passed to the IVF lab where the oocytes are identified using a stereomicroscope and heated stage, usually at 8 to 60× magnification. The embryologist pours the contents of each tube into a petri dish, forming a thin layer of fluid, which can quickly be scanned for the presence of an oocyte. The oocytes are then washed in bicarbonate-based culture medium, transferred into preequilibrated medium, and stored in a CO ₂ environment (i.e., a CO ₂ incubator) for incubation until further processing.

Assessment of the Oocyte

Techniques that assess the nuclear and cytoplasmic maturation of the oocyte fall into two major categories: invasive and noninvasive. The common practice in embryology laboratories includes a noninvasive morphologic assessment using light microscopy. Each oocyte is assessed for nuclear maturation and morphologic characteristics; this includes evaluating each of the anatomic components of the oocyte (cumulus, ZP, perivitelline space, polar body, and cytoplasm) ( Fig. 32.3 ). The morphologic features of the oocyte have been studied as factors that may be associated with pregnancy outcomes. However, a consensus on whether each morphologic characteristic is associated with outcome has been difficult because many investigators have used different terminology to define similar observations. Although no consensus exists, it is common practice to record these observations. An oocyte scoring system has been developed that takes into account all morphologic characteristics.

The cumulus-oocyte complex.

(A) Upon denudation, egg structures become visible: (z) zona pellucida, (pv) perivitelline space, (pb) polar body if oocyte is mature (MII). (B) Compacted dark cumulus mass (c) surrounding an oocyte (o) . (C) Expanded cumulus mass (c) surrounding the oocyte (o) .

Oocytes collected at retrieval are difficult to visualize because they are surrounded by a cumulus mass composed of follicular cells in a polymerized matrix of hyaluronic acid ( Fig. 32.3B and C ). Therefore, in the case of conventional insemination (CI; see further on), the morphologic assessment is limited to the cumulus-oocyte complex (COC). The volume, density, and expansion of the CCs can suggest the stage of oocyte nuclear maturity. The grading of cumulus expansion is a visual inspection of whether the cells are compact or expanded. Compact CCs take on a darker appearance due to the dense association of these cells around the oocyte (see Fig. 32.3B ); they are usually associated with immature oocytes, while expanding or fully expanded CCs are usually associated with mature oocytes (see Fig. 32.3C ).

Unlike CI, in which intact mature COC are inseminated, fertilization with micromanipulation (ICSI) requires denudation of oocytes (i.e., removal of the surrounding cumulus and corona cells). The denudation allows for an additional detailed evaluation of each component of the oocyte ( Fig. 32.4 ). Starting from the outermost layer, the oocyte is encased in a thick glycoprotein shell called the zona pellucida (ZP). The ZP normally averages 15 to 20 µm in width and is intact and translucent in a mature oocyte (see Fig. 32.3A ). Studies have evaluated whether the darkness and/or thickness of the ZP is associated with clinical outcome. Although darkness has shown little correlation with clinical outcome, the thinner ZP has been associated with higher fertilization rates. Other reported alterations of ZP morphology, including fractured or broken ZP, have not been demonstrated to be associated with clinical outcomes. The etiology of these findings may result from postmaturity of the oocyte or excess pressure during oocyte aspiration.

Denudation of oocytes.

Cumulus-oocyte complexes (COC) can be separated with a combination of chemical and mechanical agitation using hyaluronidase in a process known as denudation. Briefly, COCs are submerged into a drop of hyaluronidase and gently pipetted for no more than 40 seconds to minimize toxicity and prevent detachment of the polar body from its position. The enzymatic reaction is stopped by transferring the oocytes into a clean microdrop without hyaluronidase. Additional mechanical pipetting may be necessary with a narrower bore to remove remnant coronal cells attached to oocytes. The oocytes are then graded for the presence of a polar body to indicate maturity. GV , Germinal vesicle.

The space between the ZP and oolemma is called the perivitelline space. Although several studies have suggested that an enlarged perivitelline space ( Fig. 32.5F ) is associated with poorer fertilization and embryo development, there is no clear consensus. An enlarged perivitelline space has been hypothesized to relate to oocyte postmaturity. Sometimes the perivitelline space may contain granules, which do not appear to influence oocyte or embryo development. The first polar body appears in the perivitelline space. Unusually large polar bodies may indicate disturbances in the position of the meiotic spindle, which may occur during oocyte aging. Accordingly, a large polar body has been associated with poor embryo quality and low pregnancy rates. Fragmentation of the first polar body has not been consistently identified as an abnormality.

Oocyte morphologies.

Atypical morphologies and their appearance. (A) Cytoplasmic granularity, (B) centralized granular area, (C) smooth endoplasmic reticulum clusters (*) , (D) highly vacuolated cytoplasm, (E) abnormal zona pellucida, (F) enlarged perivitelline space with fragments, (G) comparison of normal (left) and giant oocyte (right) diameters, (H) multinucleated oocyte.

(Data from Rienzi L, Vajta G, Ubaldi F: Predictive value of oocyte morphology in human IVF: a systematic review of the literature. Hum Reprod Update 17:34–45, 2011; Lehner A, Kaszas Z, Murber A, et al: Giant oocytes in human in vitro fertilization treatments. Arch Gynecol Obstet 292:697–703, 2015; Munne S, Cohen J: Chromosome abnormalities in human embryos. Hum Reprod Update 4:842–855, 1998.)

The cytoplasm of a normal egg is typically uniform, translucent, and free of inclusions. A relatively common abnormality that can be visualized by light microscopy is central granularity (see Fig. 32.5B ). This granular area likely represents a polarized distribution of mitochondria and may have implications for cytoplasmic maturity. Studies evaluating granularity, either generalized or central, have failed to demonstrate a consistent association with clinical outcome. Although some studies have shown that cytoplasmic darkness is associated with a poorer fertilization rate or embryo development, others have not. It has also been hypothesized that localized dark and granular cytoplasm may be related to atresia.

The presence of vacuoles, an aggregation of smooth endoplasmic reticulum (SER) within the cytoplasm, has been associated with decreased fertilization, embryo quality, and ongoing pregnancy rates (see Fig. 32.5C ). Vacuoles may be associated with oocyte aging and degenerative processes in oocytes. Case reports have suggested that the presence of SER is associated with an increased risk of Beckwith–Wiedemann syndrome, diaphragmatic hernia, multiple malformations, and ventricular septal defect. A recent study reported healthy live births from embryos derived from such oocytes and recommended a careful follow-up of the children born. As a result, it is recommended that this cytoplasmic abnormality be noted. Inclusions, or refractile bodies, are inconsistently associated with a compromise in embryo quality and pregnancy outcomes. Although their cellular basis is unknown, the presence of inclusions within an oocyte may be associated with atresia.

Various shapes and sizes of the oocyte have been observed. A normal human oocyte is roughly 120 µm in diameter. Oocytes with a diameter of 200 µm or more are commonly called giant oocytes (see Fig. 32.5G ). These oocytes are thought to be giant due to errors in mitosis during proliferation of the oogonia. As a result, these oocytes are likely diploid and the resultant embryos are at increased risk for dygynic triploidy, mosaicism, or polyploidy. The presence of a giant egg in a cohort of oocytes is not associated with overall decreased pregnancy outcomes, but some data have shown that embryos derived from the cohort may show an increased incidence of abnormality in the cleavage rate.

Microscopic observation of denuded oocytes allows for assessment of the presence or absence of a GV. Fig. 32.6 shows three stages of oocyte maturation. On average, 3.9% of the intact oocytes are at the metaphase I (MI) stage, having undergone breakdown of the GV but not extrusion of the first polar body. Approximately 10.3% of the intact oocytes are at the GV stage and approximately 85.8% are in the MII stage, defined by the presence of the first polar body. ICSI is carried out only on MII oocytes, because only such oocytes have reached the haploid state and thus can be fertilized normally. It has been reported that 74% of the MI oocytes complete meiosis within 20 hours after retrieval. Another study reported that 27% of MI oocytes extruded their polar bodies within 4 hours of egg retrieval. Oocytes from this study were injected on the day of the egg retrieval in parallel with the oocytes retrieved at MII. This study demonstrated that MI oocytes that completed their maturation in vitro had lower fertilization rates compared with those aspirated at MII. However, no differences were observed in embryo quality between the two groups of oocytes. Additional studies support these observations, showing that oocytes matured in vitro can be normally fertilized; however, embryos derived from these oocytes rarely result in pregnancies. These results suggest that rescue of MI oocytes of patients with few MII oocytes may increase the number of embryos for transfer; but the chance of improving pregnancy rates by this procedure is minimal. GV-stage oocytes require overnight (24 to 48 hours) incubation in culture media supplemented with gonadotropins in order to reach the MII stage. Few pregnancies have been reported from oocytes that were retrieved at the GV stage, although standard IVF treatment with controlled ovarian hyperstimulation was performed. Because of the poor results, GV oocytes are usually discarded. Denuded and rinsed oocytes are incubated until the time of microinjection.

Stages of oocyte maturation.

(A) Immature germinal vesicle (GV) present in the cytoplasm. (B) Oocyte following GV breakdown currently at meiosis I. (C) Extrusion of the first polar body signifies a mature oocyte at meiosis II, awaiting fertilization.

Polscope imaging provides valuable information on the structure and architecture of the meiotic spindle. The integrity of the meiotic spindle in MII oocytes is crucial for normal fertilization and subsequent development. Overall, meiotic spindles can be detected in up to 91% of human MII oocytes at the time of ICSI. Studies have shown the presence of a birefringent spindle in a human oocyte can predict not only a higher fertilization rate but also greater embryo developmental competence. High degrees of misalignment of the meiotic spindle and the first polar body have also been documented and are thought to occur as a result of polar body displacement during cumulus corona removal. The relative position of the spindle within the oocyte, however, has little influence on the developmental potential of the resulting embryos.

Because most oocytes possess spindles, the mere presence of a spindle is likely of limited value. Whether quantitative spindle analysis (estimating the density of tubules within the spindle) offers added value remains to be seen. So far, visualization of the spindle and the appearance of the first polar body are accurate indicators of oocyte maturity and may help to determine the timing of ICSI (see ICSI ).

The developmental competence of the oocyte has also been assessed through a biomarker approach. Several strategies have been applied, including a comparison of transcriptomes, proteins (proteomics), or metabolites (metabolomics) of granulosa cells (GC) and/or CCs or FF. Evidence suggests that CCs from follicles of embryos that result in a pregnancy may have differential expression. Others have shown that gene-expression profiling in either GC or CC has no association with fertilization outcome but may be associated with advanced maternal age. Estes et al. found that the FF proteome in women 32 years of age or younger predicted ovarian response and live birth. Another study reported that oocytes that resulted in pregnancy presented high amounts of proteins, and those oocytes that resulted in no pregnancy presented high amounts of ubiquitinated peptides. Several investigators have demonstrated associations between FF metabolome and the developmental competence of the oocytes. Although intriguing, the available testing is not ready for routine clinical use. However, given that they represent cellular regulatory processes, biomarkers may be more informative in the future than morphology.

Sperm Collection

Sperm are obtained either through ejaculation (i.e., masturbation, electroejaculation) or by surgical retrieval. The evaluation and assessment of semen is very important for the diagnosis of infertility and for treatment decisions. The collection method for IVF is dependent on the clinical scenario (see Chapter 31 ). The collection processing procedures and assessment of the sperm depend on the method of collection and are discussed in the following paragraphs. The choice of sperm preparation technique depends on the nature of the semen sample, particularly with respect to motile count, ratio between motile and immotile count, volume and presence of antibodies, agglutination, and pus cells or debris.

The majority of sperm collections are obtained through masturbation into a collection cup. The container to collect the sample has to be wide necked and plastic or sterile glass, in addition to being cytotoxically tested. It must be labeled with the patient’s full name, identification number, date of birth, date and time of collection, and number of days of abstinence. Semen samples are collected after 2 to 10 days of abstinence. A shorter period of time may be associated with lower volume but higher motility, while longer abstinence can be lead to reduced sperm motility. In cases of male infertility, shorter durations of abstinence (1 day) may be associated with higher motility and better morphology. The use of lubricant (other than mineral oil or selected lubricants) is discouraged because they may be spermicidal. Other collection methods include the use of Silastic condoms during intercourse.

Most of the samples should be collected on site in a private room set aside for this purpose. The room can be adjacent to the lab and have a pass-through for dropoff; alternatively, the sample can be given to a staff member with proper chain of custody (for specimen verification) protocol. If for personal reasons the patient cannot produce on site, he can produce at home or at a place nearby that will allow sample delivery to the laboratory within an hour. Samples should be kept at room temperature during transport (20 to 37°C) and can stay at room temperature in the laboratory for up to 1 hour in order for liquefaction of the semen to occur.

Assessment of Ejaculated Sperm

Prior to processing the sperm for IVF, a semen assessment is performed, which entails a sperm count and motility evaluation ( Table 32.3 ). The assessment dictates how the sperm is processed and the procedure type for insemination. Sperm counting and motility assessment are obtained through standard procedures. The goal of preparation of sperm is to concentrate the motile spermatozoa in a fraction that is free of seminal plasma and debris. The sperm preparation methods attempt to mimic the natural process whereby viable sperm are separated from the seminal plasma and other constituents as they migrate through the cervix. The seminal plasma contains substances that inhibit capacitation and prevent fertilization. Sperm processing should be performed within 30 minutes if liquefaction is completed or no later than 1 hour after production. If delayed, the uncontrolled production of reactive oxygen species (ROS) that exceeds the antioxidant activity of seminal plasma will lead to oxidative stress and result in decreased sperm capacity. The ROS are produced from leukocytes and other debris present in the seminal plasma. After sperm processing, the insemination process ideally should be performed within 4 to 6 hours. After 4 to 6 hours, sperm DNA fragmentation may be increased. Several sperm preparation techniques are described in the following paragraphs.

Table 32.3

Semen Parameter Reference Value According to World Health Organization Manuals

Semen prameters	WHO, 2010 ‡	Strict morphology analysis: Observed morphologies in sample include (a) large acrosome, (d) tail defect, (l) leukocyte, (m) macrocephalic head, (n) normal morphology, (r) round head, (t) tapered head.
Volume	1.5 mL
Concentration	15 × 10 6 /mL
Total sperm concentration	39 × 10 6
% Motile	40%
Progressive motility *	32% (a+b)
Vitality †	58%
Morphology (Kruger strict) normal forms, %	4%
Leukocyte count	<1 × 10 6 /mL

 

Values obtained from lower fifth centile value.

* a, rapid progression (>25 µm/s); b, sluggish (5–25 µm/s).

† % alive.

‡ World Health Organization, Department of Reproductive Health and Research. Who laboratory manual for the examination and processing of human semen, 5th ed. 2010. Available at: http://www.who.int.easyaccess1.lib.cuhk.edu.hk/reproductivehealth/publications/infertility/9789241547789/en/ .

Processing Techniques

Surgical Sperm Retrieval

Several sperm retrieval methods have been developed to retrieve sperm from the epididymis and/or testes in men with azoospermia. The technique employed depends on the clinical scenario (discussed in Chapter 31 ). The evaluation and processing of the sample is dependent on the surgical technique and site of collection. Sperm retrieval should be carried out either on the day of oocyte retrieval or the day before, depending on the laboratory workload. Additionally, the surgically retrieved specimen may be incubated for up to several days if the sample is poor and requires further maturation. All surgical collections require ICSI for optimal fertilization (see further on).

Collection, Processing, and Assessment of Surgical Sperm

Conventional Insemination

Insemination should be done within 2 to 6 hours after the egg retrieval. Evidence has shown that insemination prior to 2 hours postretrieval leads to a compromise in pregnancy rate. Significant delays after retrieval are associated with decreased fertilization and pregnancy outcomes due to spindle instability and subsequent loss or scattering of chromosomes in the oocyte. On average, 55% to 65% of the oocytes will normally be fertilized (2PN). The procedure involves using prepared dishes from the day prior to ensure proper equilibration of medium and oil (37°C and pH). Since incubation is required, insemination dishes are made of droplets composed of sodium bicarbonate–based media under an oil overlay ( Fig. 32.8 ). The droplets typically are 50 to 200 µL in size with one to four COCs in each. Oocyte grouping is determined by the laboratory and is mostly based on efficiency of the procedure. The cumulus is maintained on the oocytes to theoretically ensure maximum fertilization. The processed sperm are then added to the droplets. It is recommended to aseptically dispense 40,000 to 100,000/mL of motile sperm per microdroplet. Fertilization rates have been demonstrated to be significantly higher with 20,000 sperm per egg versus 5000 sperm per egg. This sperm concentration range is optimized for achieving high normal fertilization rates without the increased risk of polyspermy. It is critical that the insemination process be performed as quickly as possible or under an isolette with CO ₂ and on a heated stage to reduce pH changes or cooling. Once insemination has been completed, the gametes are coincubated overnight, and 16 to 20 hours postinsemination, oocytes are observed to check for normal fertilization. In most instances, we like to do the insemination between 1 p . m . and 3 p . m . to optimize the timing for fertilization check the next day (see Fig. 32.1 ). Although traditionally inseminated oocytes are incubated overnight, sperm binding to the zona can take place within 1 to 3 hours after insemination and fertilization occur. Thus a few hours of sperm-oocyte contact can yield the same time course of events that was observed after overnight insemination. One thought is that longer exposure to the high concentrations of sperm may increase exposure of oocytes to high ROS levels and alter the pH in the insemination medium. These derangements may affect embryo viability, cause zona hardening, and lead to a lower pregnancy rate. Therefore some labs may choose to wash the excess sperm free from the oocyte after 3 hours incubation. Regardless of the incubation time, the current standard of care is to evaluate for fertilization 16 to 20 hours postinsemination ( Table 32.4 ).

Conventional insemination.

Two to 6 hours following oocyte retrieval, one to four cumulus-oocyte complexes (COCs) can be placed in a microdrop of approximately 200 µL. Ideal sperm concentration in this environment should be around a total of 40,000 to 100,000 sperm per drop. The following morning, fertilization can be assessed by mechanical stripping of the cumulus cells to visualize the presence of pronuclei in the cytoplasm.

However, with low sperm counts or poor motility, CI is associated with decreased fertilization. In some cases, improved fertilization may be achieved by increasing the number of motile sperm (see earlier discussion of semen preparation). In other instances fertilization can be improved by decreasing the volume of medium, which may increase the interaction between the gametes, such as the insemination technique involving a microdroplet under oil. This technique has been shown to raise the chances of fertilization to over 40% and to provide a clinical pregnancy rate of 24%.

Several methods have been developed to improve fertilization when the CI was poor or unsuccessful. These methods bypass layers of the COC to aid in fertilization. Initially, the cumulus was removed, but fertilization rates were not improved. The techniques subsequently focused on the ZP, given that it was a known barrier to sperm penetration. The various methods were intended to thin the zona so that the inseminated sperm could have direct access to the oolemma. The techniques included zona drilling (ZD), partial zona dissection (PZD), and localized laser cutting. However, the increased incidence of polyspermy and a still relatively low fertilization rate led to subzonal sperm injection (SUZI). This technique involves the direct injection of several motile sperm into the perivitelline space with an injection pipette. But even SUZI had its limitations, such as the inability to overcome acrosomal abnormalities or dysfunction of the sperm–oolemma fusion process. As a result, normal fertilization rates were poor (20%), and pregnancy rates were too low to consider PZD and/or SUZI for routine clinical application.

Intracytoplasmic Sperm Injection

ICSI is a procedure that entails microinjecting a single spermatozoon into the oocyte, bypassing the ZP and the oolemma. These micromanipulated oocytes undergo the normal process of egg activation, including Ca ²⁺ release. This suggests that contact of sperm and egg plasma membranes is not a critical step for egg activation. The first successful use of ICSI was to obtain live offspring in rabbits and cattle, and in 1992, the first human pregnancies and births were reported. ICSI was also found to be superior to SUZI with respect to fertilization rate, embryo yield, and embryo implantation rate.

The average fertilization rates with ICSI range from 70% to 80%, and pregnancy rates are comparable to those of cases with CI and normal sperm counts. Given the success, ICSI is now performed universally when CI cannot be performed because of poor sperm motility or morphology.

ICSI is performed on an inverted microscope equipped with micromanipulators and microinjectors under 200× and 400× magnification while on heated stage. The micromanipulators allow three-dimensional manipulation of coarse and fine movements. One microinjector is used to hold the denuded oocyte and the other to inject the sperm. Movement of the oocyte or sperm is achieved by a micrometer that controls a plunger. The whole setup is placed on a vibration-proof table to avoid possible environmental motions.

ICSI can be described with the following steps: (1) denudation of the oocyte, (2) selection and immobilization of a viable sperm, (3) positioning of the oocyte before injection, and (4) rupture of the oolemma before release of the sperm into the oocyte. Fig. 32.9 shows the injection sequence.

Intracytoplasmic sperm injection (ICSI).

Approximately 1 µL of processed sperm is added to a drop of polyvinylpyrrolidone (PVP) to slow their progression and allow for immobilization via tail break using an injection needle. The approximate path of the needle to break the sperm is shown as a dashed line in the top panel. This sperm will be aspirated and taken to a drop of buffered media holding the oocyte, where it can be directly injected into the cytoplasm. Special attention should be paid to the position of the polar body, and injections should be directed away from this site, where the mitotic spindle apparatus is expected. (A to D) Microscopic images of sperm immobilization (A) to injection (B to D).

Denudation of the oocyte entails removal of the surrounding cumulus to visualize the oocyte and allow for a precise injection of the oocyte (also called oocyte stripping, Fig. 32.4 ). Since CCs are thought to secrete paracrine growth factors, which play a role in the nuclear and/or cytoplasmic maturation of the oocyte, several investigators have evaluated if early or delayed denudation will impact fertilization rates. The consensus suggests a preincubation time of 2 to 6 hours prior to denudation. However, some evidence suggests that there may be improvement in fertilization rates with longer preincubation times (>3 hours) and shorter times to fertilization once the oocyte has been denuded. Denuded oocytes are more vulnerable to pH changes.

Removal of the surrounding cells is typically achieved by a combination of enzymatic and mechanical procedures (see Fig. 32.4 ). Since hyaluronic acid is a major component of the extracellular matrix of the cumulus, these oocytes are repeatedly transferred to a hyaluronidase solution and HEPES buffered medium for approximately 30 seconds with pipettes of decreasing diameter (150 to 135 μm) to facilitate mechanical denudation. Hyaluronidase-purified preparations of bovine origin are most commonly used, although human recombinant hyaluronidase is available. Regardless of the preparation, hyaluronidase helps to digest the connections between the CCs, freeing the oocyte for grading and ICSI (see previous text). After denudation, oocytes should be thoroughly washed to remove traces of hyaluronidase. It should be noted that both the enzyme concentration and the duration of exposure to the enzyme should be limited because of the possibility of damage or parthenogenic activation of the oocytes.

Once denuded, the oocyte can be clearly visualized, and its morphology and maturation can be assessed (see Fig. 32.5 ). Oocytes that demonstrate a polar body (MII) and are morphologically intact are suitable for ICSI and can be fertilized accordingly. Oocytes that do not show the first polar body (MI) or that are GV can undergo extended culture and subsequently be micromanipulated if the polar body has been extruded. However, as previously mentioned, embryos derived from immature oocytes that mature in vitro after ovarian stimulation rarely result in pregnancies. Therefore GV oocytes are usually discarded. Denuded and rinsed oocytes are incubated until the time of microinjection. Oocytes should be fertilized within 6 hours after collection. However, some evidence suggests that shorter times to insemination once the oocytes have been denuded may improve outcome.

To inject a sperm into the oocyte, the sperm must be selected and immobilized. The process of selection starts with creating a plastic microinjection dish containing microdroplets of HEPES-buffered medium under a layer of mineral oil (see Fig. 32.9 ). Generally there is a central drop (~5 µL) of polyvinylpyrrolidone (PVP) surrounded by more microdroplets (culture medium to support the oocyte during ICSI). Because of the viscous nature of the PVP, the sperm’s progression is greatly slowed, and there is usually a general dispersion aiding in sperm selection and manipulation. The use of PVP also aids in microinjector use, as the viscosity allows for greater control of sperm with the injection needle. Once the semen has been processed, a small amount (~1 µL) of sperm suspension is added to the PVP drop and oocytes are added to the surrounding drops.

The injection pipette is backloaded with PVP, then a viable sperm is located. Viability is evidenced mostly by the sperm’s motility (sometimes only “twitching”) and morphology. Immobilization of the sperm involves rubbing, dragging, or knocking the sperm tail with the injection pipette against the bottom of the dish. This causes a breakage or rupture in the sperm tail’s plasma membrane. Aggressive breakage has been shown to increase fertilization rates. Immobilization of spermatozoa has been proven to be an important factor for oocyte activation, which is achieved by the release of sperm cytosolic factors via the ruptured membrane. Changes in the sperm plasma membrane and acrosome after immobilization have been confirmed by transmission electron microscopy and scanning electron microscopy. Mechanical immobilization with the ICSI needle is the general rule of thumb; however, use of a laser has been described, resulting in identical fertilization rates. After immobilization, the sperm cell is aspirated tail first with the injection pipette and moved to the microdrop containing the oocyte.

The oocyte is picked up and held in position with gentle suction by the holding pipette. The polar body of the oocyte should be at the 12 or 6 o’clock position, which may avoid possible damage to the meiotic spindle, although the first polar body does not always coincide with spindle localization. Nevertheless, there is little evidence demonstrating that spindle monitoring during ICSI reduces spindle damage or increases the chance of fertilization.

If both the holding pipette and the oocyte are in perfect focus, the injection needle, containing the immobilized sperm cell near the tip, can be introduced in the equatorial plane of the oocyte at the 3 o’clock position. Permanent focus of the injection pipette tip ensures that the needle remains in the equatorial plane of the oocyte. Passing through the ZP is fairly easy and is achieved by simply advancing the injection pipette. However, the oolemma is not always immediately pierced by simple injection of the needle. Minimal suction of the membrane into the needle or multiple passes of the needle into the oolemma may help breakage. Even selecting another area of the oocyte may be necessary. A sudden flow of cytoplasm into the needle or “pop” of the oolemma indicates membrane rupture. Once it is determined that the oolemma is broken, the sperm cell can be released into the oocyte with a minimal amount of surrounding media, and the needle then withdrawn.

Once ICSI is completed, all injected oocytes are rinsed and moved to culture microdroplets of bicarbonate-based medium under oil in conditions similar to the CI oocytes. These oocytes are then examined for integrity and fertilization approximately 16 to 20 hours after ICSI ( Table 32.5 ).

Table 32.5

Fertilization outcomes following CI or ICSI.

Outcome	Example	CI (incidence)	CI (ploidy status)	T	ICSI (incidence)	ICSI (ploidy status)	T
0 PN		1% *	>50% diploid	Y	1% *	>50% diploid	Y
1 PN		10%	30–60% diploid	Y	2–5%	Haploid	N
2 PN		55-65%	Diploid	Y	70–80%	Diploid	Y
3 PN		0-6%	Dispermic triploidy †	N	2–4%	Digynic triploidy †	N
>3 PN		Rare	Multiploidy	N	Rare	Multiploidy	N

 

CI , conventional insemination; ICSI, intracytoplasmic sperm injection; PN, pronuclei; T, transfer.

* incidence of 0PN that continue to cleave

† most common chromosomal abnormality

Sperm Selection Techniques

Sperm selection varies based upon processing technique. For ICSI, the technician subjectively evaluates sperm at an optical magnification of approximately 200 to 400× for motility and morphology. However, spermatozoa appearing morphologically normal at this magnification may carry various structural abnormalities at the subcellular level; these remain undetected by the embryologist. Some evidence suggests that the fertilization rate, embryo development, and pregnancy outcomes can be improved with more elaborate selection.

Failed Fertilization Options

TFF usually refers to the failure of all available mature MII oocytes to fertilize, and the etiology may be distinctly different than with low fertilization. The incidence of TFF after CI is 5% to 10% and after ICSI is 2% to 3%. The etiology of TFF after CI is mostly unknown and often is not elucidated, but is thought to be either due to sperm abnormalities (as with low fertilization) or a result of “shock” due to laboratory conditions. Sperm abnormalities are more likely the cause when there is a total absence of sperm binding to the ZP or failure of the sperm to successfully penetrate the ZP. Since the recurrence rate of TFF or low fertilization rate with CI is 30% to 70%, the most common treatment for failed/low fertilization after CI is ICSI the following cycle. Although ICSI may be performed emergently to salvage fertilization, successful pregnancies with this technique are rare (see the discussion of rescue ICSI ). The etiology of TFF after ICSI is more likely due to failure of oocyte activation, which can be attributed to either gamete. After TFF with ICSI, low or moderate fertilization is observed in 30% of repeated ICSI cycles. If there is failed or low fertilization after ICSI, the next step is to try to improve on selection of sperm (see previous text) or use oocyte activation procedures (see subsequent text).

Rescue Intracytoplasmic Sperm Injection

Rescue ICSI is an option after failed fertilization with CI that involves the microinjection of unfertilized oocytes with a single sperm from the original sample. This can take place the following day (late rescue) or 8 to 10 hours after oocyte retrieval (early rescue). In some countries this procedure is banned (e.g., the United Kingdom) as it cannot be determined if fertilization has occurred and is just delayed. Others reserve this procedure for complete fertilization failure following CI. Fertilization rates from rescue ICSI have been shown to be low, and the generated embryos achieve lower pregnancy rates. Although controversial, the poor outcomes have been attributed to chromosomal abnormalities and/or dysynchronous embryo development with respect to the endometrium. The higher aneuploidy might be the cause of the failed fertilization or the result of the in vitro aging process. Preliminary evidence suggests that pregnancy rates can be improved with the performance of early rescue ICSI or frozen embryo transfers. Some centers do perform rescue ICSI, but because of the low pregnancy rate and the potential genetic risk, more studies are needed before rescue ICSI can be considered a standard of care.

Artificial Oocyte Activation

Artificial oocyte activation has been proposed as a way to overcome TFF after ICSI. Calcium ionophores, such as ionomycin or calcimycin (A23187), are the most popular artificial activating substances for human oocytes. The procedure entails first microinjecting the sperm into the oocyte, followed by exposing the oocyte to an ionophore solution on two separate occasions. However, there is no standardization of the procedure as there are disparities in type, concentration, exposure time, and, mode of application of the ionophore among published reports. The lack of standardization may contribute to the discrepancies in outcomes in the literature. Evidence has shown that the success of ionophore treatment is related to the fertilization rate in a previous cycle. The cutoff value for fertilization rate in the preceding cycle that distinguished between an increase in fertilization rate in the presence of A23187 and no improvement was 30%. Although studies have shown a significant increase in fertilization rates with artificial oocyte activation (up to 70% to 77%) and developmental outcome of children aged 3 to 10 years are within expected ranges, the numbers of live births registered are low and the lack of long-term follow-up after after ionophore treatment does not allow proper calculation of the actual risk. Therefore this treatment remains experimental.

Embryo Development and Selection

As early as the pronuclear stage (16 to 18 hours), morphologic scoring, including alignment and size of pronuclei as well as alignment and number of nucleoli, can be performed. While some studies have shown a prognostic effect of PN scoring for pregnancy and aneuploidy, others have not been able to demonstrate the association. However, TLI has shown that the timing of PNB is associated with aneuploidy and implantation and is a key assessment for embryo selection. Of course, the timing of assessment is critical because pronuclear development is a dynamic process; therefore, zygote scoring and/or the timing of PNB should be used with caution and only in conjunction with other methods of evaluation.

Postfertilization, approximately 90% of 2PN oocytes cleave, resulting in multicellular embryos. Studies have shown early cleavage to be a strong biologic indicator of embryonic potential and may be used as an additional criterion for embryo selection. Cleavage is conventionally assessed approximately 25 to 27 hours after fertilization. TLI has also shown that the duration of first cytokinesis may be associated with development outcomes. The early cleavage may be a sign of molecular health and cytoplasmic maturation. Previous studies have found that embryos with early cleavage have higher blastocyst development rate and are associated with higher pregnancy rates as compared to late cleavage embryos; however, other studies have not confirmed these findings. Interestingly, very early cleavage (<20 hours postfertilization) is associated with poor clinical outcomes. When using early cleavage as a predictor for embryo implantation, it may be necessary to consider the potential confounding factors that influence cleavage, such as culture conditions, stimulation protocols, and fertilization methods.

It is generally accepted that a day-3 embryo represents a key time point for embryo development. At this juncture, it is possible to identify embryos with better developmental potential. To obtain this stage of development, there is a required transition from relying on the maternal genome to the activation of the embryonic genome. Therefore, until recently, most centers cultured embryos for 3 days prior to embryo transfer. The embryo(s) transferred would then be selected based on morphology.

There are several grading schemes for cleavage stage embryos, all designed to determine likelihood of implantation. The Society for Assisted Reproductive Technology (SART) grading system approach was developed to help laboratories establish a common standard for embryo grading ( Table 32.6 ). Regardless of the system, the main parameters include the number of blastomeres, symmetry, and fragmentation. In addition, most laboratories evaluate for compaction and note if multinucleation is present.

The single most important indicator of embryo viability at the cleavage stage is the occurrence of cellular division. The frequency of mitotic divisions is related to developmental potential of the embryo. Numerous studies have shown that the number of blastomeres is strongly associated with chromosomal abnormalities, implantation, and pregnancy outcome. Normally developing, good-quality embryos should reach the four-cell and eight-cell stages, respectively, on day 2 (40 to 44 hours) and on the morning of day 3 (64 to 68 hours) postmicroinjection ( Fig. 32.10 ). Studies have shown that embryos with either lower or higher cell numbers have significantly reduced developmental potential. Day-2 embryos with four cells result in a significantly higher implantation rate than 2- or 3-cell embryos. When transferring on day 3, embryos that comprise seven and eight cells have lower rates of chromosomal abnormalities and higher developmental potential than embryos with fewer or even higher numbers of blastomeres.

Embryo cleavage after intracytoplasmic sperm injection.

Only embryos resulting from normally fertilized oocytes (A) will be transferred to patients. Embryo cleavage is evaluated daily. Two-cell embryos (B), four-cell embryos (C), and eight-cell embryos (D) are usually obtained on day 1 (late afternoon), on day 2, and on the morning of day 3, respectively. The blastomere number is recorded and the embryos are scored according to equality of size of the blastomeres and the presence of anucleate cytoplasmic fragments. On day 4 (sometimes on day 3), a certain degree of compaction can be observed (E). For blastocyst (F) scoring, the classification system introduced by Gardner and Schoolcraft is used. Embryo transfer is usually done on day 3 (eight-cell stage) or day 5 (blastocyst stage).

TLI studies have shown that the duration of cleavage events as well as the time to specific embryonic stages are associated with developmental potential. Although some studies suggest that early kinetics is most predictive of subsequent embryo development, others suggest that later kinetics are most predictive of blastocyst development. These studies suggest that optimal cleavage times fall into the two middle quartiles and embryos that reach the eight-cell stage fastest are associated with higher implant potential. Although morphokinetics are indicators of blastocyst development, there is low sensitivity and a lack of evidence to predict implantation above and beyond static morphologic assessments. It is likely that morphokinetics are affected by ovarian stimulation protocols, culture conditions, fertilization method, and individual differences.

Blastomeres should cleave synchronously and give rise to two cells with half the volume of the parent cell. Several studies have shown that even numbered embryos have higher implantation potential. In addition, a number of TLI studies have shown that abnormal early cleavage events—such as direct cleavage, where embryos cleave from two cell to three cells or exist as two cells for a limited time (<5 hours) or have uneven blastomere cleavage (>25% difference)—are associated with poor implantation potential. Examples of variations in blastomere size and cell generations are shown in Table 32.7 . Because embryos with uneven cell cleavage have a lower developmental capacity compared with evenly cleaved embryos, there is an emphasis on blastomere size within conventional embryo scoring systems. Therefore embryos are typically scored according to equality of size of the blastomeres in addition to number.

Table 32.7

Cleavage Embryo Grading Guidelines

Fragmentation		Symmetry
Score	Appearance	Score	Appearance
1: No fragmentation		1: Equal size of all blastomeres
2: <10% fragmentation
3: 10%–25% fragmentation		2: Slightly unequal size of all blastomeres
4: 25%–50% fragmentation
5: >50% fragmentation		3: Significantly unequal size of all blastomeres

 

Numerous studies have shown that the presence of anucleate material or fragmentation is negatively associated with decreased blast development and pregnancy outcomes. Fragmentation grading is shown in Table 32.7 . Fragments form when segments of the cytoplasm protrude from the cell upon cell division and then separate. Blastomere fragmentation is observed in approximately 40% of early human embryos. The cause of fragmentation is not completely known. However, several theories include insufficient adenosine triphosphate (ATP) production, apoptosis with subsequent loss of regulatory proteins, and poor-quality gametes. Although evidence suggests that fragmentation may occur in vivo, in vitro conditions such as a decrease in pH, high oxygen tension, or decreases in temperature may lead to destabilization or loss of integrity of membranes and result in fragmentation. Studies have shown a trend toward increased fragmentation percentage with aneuploidy; however, there is no correlation with maternal age. Although several studies have shown that increased fragmentation percentage is associated with decreased implantation rate, the correlation of fragmentation score with outcomes remains controversial, as even highly fragmented embryos result in live births. This may be due to fragments being associated with mosaicism rather than aneuploidy and embryonic self-correction (see the discussion of preimplantation genetic testing [ PGT ]). In summary, fragmentation is a predictor of implantation but should not be the only morphologic criterion assessed.

Multinucleation is a frequently observed phenomenon in cleavage-stage embryos ( Fig. 32.11 ). Evidence has shown that multinucleated cells occur in cleavage stage embryos at the two- to four-cell range in 17% of cases and up to 65% in the third cleavage division. The etiology of multinucleation within a cell involves errors in cytokinesis. Multinucleation affects morphokinetic parameters. The presence of one or more multinucleated blastomeres (≤50%) has been associated with impaired cleavage and increased fragmentation, a poor prognosis for blastocyst formation, and reductions in the ongoing implantation rate, probably due to chromosomal abnormalities. The proportion of multinucleated blastomeres within one embryo is an important parameter. However, not all multinucleated embryos are chromosomally abnormal. Excluding these embryos for transfer when other mononucleated embryos are available is prudent practice.

Multinucleation in a cleavage stage embryo.

Still-frame of an eight-cell embryo containing a multinucleated blastomere (indicated by * ) as captured by time-lapse cinematography.

(Modified from Iwata K, Yumoto K, Sugishima M, et al: Analysis of compaction initiation in human embryos by using time-lapse cinematography. J Assist Reprod Gen 31:421–426, 2014.)

Assessing the degree of compaction is valuable because early compaction on day 3 may be associated with increased implantation potential. On day 4 (sometimes on day 3), a certain degree of compaction can be observed. Compaction is a morphologic change in a preimplantation embryo that results in a mass of cells held by tight junctions and is a fundamental event that leads to the formation of the TE, the ICM, and the blastocoele. Full compaction (16- to 32-cell stage) is followed by immediate cavitation and blastocoele expansion.

In the past it was suggested that the culturing of embryos to the blastocyst stage could naturally select the most viable embryos, increase implantation, and decrease miscarriage rates following a transfer. However, at that time it was difficult to maintain embryo viability to the blastocyst stage. With advancements in commercially available culture media, the common practice at present is to extend embryo culture for 5, 6, and 7 days to the blastocyst stage. Culture systems for blastocyst development may entail refreshing the media on day 3 and possibly on day 5 of culture with similar media, or sequential media systems may be used where media with different constituents specific to the phase of embryonic development are utilized. Other culture systems use one medium without refreshing. The specific culture system used for blastocyst development is dependent on laboratory conditions, clinic success rates, and preference. On average, the blastocyst rate is 50% but is dependent on culture environment and patient characteristics, such as age. As a result of the success with extended culture, transfers with blastocysts are commonly performed.

With improved selection, a blastocyst transfer would allow the transfer of fewer embryos, thereby decreasing the number of multiple pregnancies. However, although blastocyst culture improves selection and pregnancy rates are increased per transfer, the superiority of blastocyst transfer over early embryo transfer per cycle remains to be proven. It is essential for an embryo to develop into a blastocyst to implant, but not all blastocysts will implant and some viable embryos may fail to develop into a blastocyst in vitro. Extended culture also allows for preimplantation genetic screening (see the discussion of PGT ), although the safety of extended culture is not known. It is possible that prolonged culture to the blastocyst stage may increase the potential of aberrant epigenetic programming events. The decision to culture embryos to the blastocyst stage is based on the clinical scenario (see Chapter 31 ) and the laboratory’s capabilities.

Several scoring systems have been created for the evaluation of blastocysts. The most common classification system is shown in Table 32.8 . A distinction between early and expanded blastocysts is made, and the latter category is further scored according to the quality of the ICM and the TE. Ideally, blastocysts should have a thin ZP, smooth cohesive trophoectoderm, equality and close adhesion of blastomeres, a clearly visible blastocele cavity, and well-developed ICM with many closely aggregated cells. Evidence has shown that the single most important scoring factor in determining the likelihood of implantation is the TE score. Although top-quality blastocysts are associated with increased implantation potential, the correlation between blastocyst score and aneuploidy is modest, as more than 50% of aneuploid embryos are capable of achieving good- or excellent-quality blasts.

Table 32.8

Blastocyst Grading Guidelines

Expansion	Inner Cell Mass	Trophectoderm
EB1/2 : Early blastocyst, cavity ≤ volume of embryo	A : Well formed and tightly packed	A : Many cells forming cohesive epithelium
3 : Full blastocyst, blastocele cavity completely fills embryo	B : Loosely grouped with several cells	B : Few cells forming loose epithelium
4 : Expanded blastocyst, cavity > embryo volume and zona is thinning	C : Very few cells	C : Very few large cells
5 : Hatching blastocyst, trophectoderm begins to herniate out of the zona	Expansion, inner cell mass, and trophectoderm grading are used to describe the blastocyst morphology at the time of observation.

 

Assisted Hatching

Developing embryos show a gradual thinning of the ZP during culture. The mammalian late blastocyst both expands and thins its ZP through alternating expansion and contraction. Intrinsic processes allowing thinning before successful hatching include pressure of the blastocyst as it expands against the zona and a lytic mechanism mediated by zona lysins. Finally, the ZP ruptures, allowing the embryo to hatch and implant into the endometrium.

An important observation leading to the clinical introduction of assisted hatching was the finding that there were higher implantation rates from embryos that were fertilized using microsurgical techniques. In addition, there was some evidence that cleaved embryos with thinner zonae had higher implantation rates than those with thick ZPs. Many embryos may not succeed in hatching from the zona for different reasons. A naturally thick ZP or hardening of the ZP due to prolonged in vitro culture conditions may impair or even prevent the natural hatching process, leading to implantation failure. Suboptimal culture conditions may cause TE lysin production to fall below the threshold level needed to promote zona thinning before successful hatching and subsequent implantation. Alternatively, failure of intrinsic lysin production unrelated to culture conditions may inhibit normal hatching.

To overcome hatching failure, three different micromanipulation procedures (mechanical, chemical, and laser-induced hatching) have been used to produce holes in the ZP of cleavage-stage embryos or to thin the ZP superficially. The advantages and disadvantages are shown in Table 32.9 and the methods are described in the following paragraphs.

PZD is a form of mechanical hatching that was developed to create an artificial opening of the ZP of early cleavage-stage embryos. This technique is performed by stabilizing the embryo with a holding pipette while the zona is pierced with a microneedle pushed tangentially through the space between the ZP and blastomeres until it pierces the zona again. This spearing leaves a small part of the zona trapped against the microneedle. Rubbing this part of the zona against a holding pipette leaves a narrow incision in the zona. The mechanism of PZD is quick to perform; however, it produces holes of variable size that may not always be optimal. A modification of PZD has been described where a second cut is made in the zona below the first cut at a right angle. This leaves a cross-shaped hole on the surface of the zona pellucida (3D-PZD). This technique allows for the creation of a larger opening while maintaining the protection of the zona flaps during embryo transfer.

Chemically assisted hatching involves the use of acidic Tyrode solution (pH 2.35) to “drill” the ZP. A microneedle filled with acidic Tyrode solution is used to produce an opening at the 3 o’clock position, facing either empty perivitelline space or extracellular anucleate fragments. The acidic solution is expelled gently over a small area (30 µm), with the tip of the needle held very close to the zona until an opening is obtained. The mean diameter of the gap created is 20 ± 6.7 µm (range, 10 to 36 µm). It should be noted that the acidic Tyrode solution is embryotoxic and may alter the viability of the embryos. Therefore, with the advancement of alternative methods, chemically assisted hatching is rarely used today.

Laser-assisted ZP drilling has been shown to be a suitable alternative to the use of acidic Tyrode solution for zona dissection. Several types of laser sources have been investigated. In a convenient setup, the laser beam is guided through an optical lens and focused on the biologic specimen. There is no need for holding or cutting tools. Lasers with different wavelengths have been proposed to operate in this no-touch mode; however, infrared radiation is the most appropriate, especially when mutagenic risks are taken into account. The 1.48-µm diode laser allows rapid, precise, and easily controlled lysis of the ZP. The size of the hole can be chosen by varying the irradiation time; typically, a hole with a diameter of 20 µm is produced within 12 to 30 ms irradiation time. Larger-diameter holes are obtained by increasing the irradiation time.

Piezo technology, an alternative to laser technology, also produces a precise and controllable defect in the zona without embryotoxic chemicals. The vibration produced by the piezoelectric pulse is used for the assisted hatching. The ZP is carved easily in a limited area that is conical and approximately 30 µm in diameter. Five to eight applications in adjacent areas are needed to treat approximately one-third of the visible circumference of the ZP. Thereafter, weaker vibrations are used to create an opening 20 µm in diameter in the thinned zona at the junction between blastomeres.

Assisted hatching techniques designed to facilitate embryonic escape from the zona have been used in IVF centers since 1992. The initial indications for assisted hatching were patient age, zona thickness, high basal FSH value, and repeated IVF failure. Several studies assessing assisted hatching in these cases have given disparate results. For example, there remains conflicting evidence of a benefit of assisted hatching in patients with advanced maternal age. Some of the controversy may be due to the different methods used for assisted hatching and the lack of reproducibility of the hole sizes obtained. Disparate patient selection, retrospective study designs, and the lack of an appropriate control group may also explain conflicting findings in previous research. A recent meta-analysis that included randomized controlled trials of assisted hatching versus no assisted hatching and reported live births, clinical pregnancies, or implantation rates, concluded that there is some evidence that assisted hatching may increase the chance of pregnancy in women for whom IVF has been repeatedly unsuccessful and in older women, although definitive recommendations could not be given. As a result, the clinical relevance of assisted hatching of cleavage-stage embryos remains debated.

Preimplantation Genetic Testing (PGT)

For the past 25 years, PGT has been used to select against embryos carrying various forms of genetic abnormalities ( Fig. 32.12 ). There are two types of PGT: preimplantation genetic diagnosis (PGD) and PGS. Although the usefulness of PGD is definitive, the utility of PGS remains an area of active investigation.

Embryo biopsy for genetic testing.

Biopsies can be done at several stages during preimplantation embryo development. Polar bodies (PB) can be removed from the oocyte before fertilization and the following morning after insemination to ensure the collection of both PBs. One to 2 blastomeres can be removed from a Day 3 embryo. The most common method of sampling uses trophectoderm biopsy from embryos at day 5 to 7. These cells are lysed and the DNA is amplified for preimplantation genetic testing (PGT). Chromosomal and gene abnormalities can be assessed via genetic screening (PGS) or genetic diagnosis (PGD). Embryos that are considered normal and/or euploid are selected for embryo transfer.

In 1989, PGD was introduced as a way of identifying genetic mutations in human embryos during an IVF cycle. Today, PGD is used for single-gene disorders, for inherited chromosome abnormalities, or human leukocyte antigen (HLA) matching. The aim of PGD is to avoid the transmission of genetic conditions in the offspring of a couple with increased risk of transmitting a genetic pathology, allowing the transfer of genetically healthy embryos. In principle, the indications for PGD are similar to those for prenatal diagnosis after chorionic villus sampling or amniocentesis, given that PGD at the single-cell level is technically possible.

In 1993, PGS was introduced based on the hypothesis that selection of euploid oocytes and embryos during assisted reproduction would lead to increased implantation rates and reduce spontaneous miscarriage rates. The hypothesis is based on the findings that show that the majority of implantation failure has been attributed to chromosomal aneuploidies. As previously mentioned, grading of the embryos and extended culture has, at most, a modest correlation with chromosome copy number. With the additional genetic testing, the aim is to increase the chance of delivery by preselecting euploid embryos for transfer. The number of referrals for PGS has only increased since its first clinical application, but the benefits of PGS remain a topic of debate.

An informed discussion of the utilization, successes, and challenges of PGT is predicated on a detailed understanding of how genetic material is obtained from the embryo. In this section, we describe the methods for each type of biopsy and discuss the advantages and disadvantages of each. The embryo biopsy procedure, regardless of stage of development, always involves two steps: opening the ZP and removing the cellular material. The methods for opening the ZP are similar to assisted hatching (see Table 32.9 ). However, the biopsy process for each developmental stage of the embryo is different. Three different sources of genetic material are potentially available for PGT: polar bodies from the initial conception, blastomeres from early cleaving embryos, and TE cells from embryos reaching the blastocyst stage (see Fig. 32.12 ). Each of these three types of biopsies is described in further detail in the following text. Next, we discuss the strengths and limitations of various modalities for genetic testing of biopsy samples. Lastly, we will end this section with a discussion of the clinical utility of PGD and PGS.

Timing of Biopsy

Cleavage-Stage Biopsy Procedure

Cleavage-stage biopsy is performed on the morning of day 3 after oocyte retrieval ( Fig. 32.13 ). Unfortunately not all human embryos reach the seven- or eight-cell stage by that time. Six-cell–stage embryos may, therefore, be included for one- or two-cell removal, although they are suspected of being of inferior quality because of delayed cleavage.

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