In 1986, Kruger and associates proposed a strict criteria for classification of normal and abnormal spermatozoa morphology.⁸ Objective measurements of individual spermatozoa components are included in this system, also known as the Tygerberg strict criteria classification system. According to this system, a spermatozoon is considered normal only if: (1) the head measures 5 to 6 microns in length and 2.5 to 3.5 microns in width; (2) acrosomal staining covers 40% to 70% of the anterior aspect of the sperm head; (3) the midpiece is 1.5 times as long as the head length and less than 1 micron in width; (4) the tail is approximately 45 microns long, uniform, uncoiled, and free from kinks; and (5) cytoplasmic droplets are no more than half the size of the spermatozoa head and occupy the midpiece only. Borderline forms are considered abnormal in this classification scheme.

In men with spermatozoa counts greater than 20 million/mL and motility greater than 30%, fertilization rates during IVF cycles were 91% for men with greater than 14% normal spermatozoa by Tygerberg strict criteria and 37% for men with less than 14% normal forms.⁸ This study established the reference value for normal morphology as 15% or greater by strict criteria.

In 1992, the WHO adopted a grading system based on the Tygerberg strict characterization of distinct spermatozoon components.³ Under these guidelines, a spermatozoon is considered normal if: (1) the head measures 4 to 5.5 microns in length and 2.5 to 3.5 microns in width; (2) acrosomal staining covers 40% to 70% of the anterior aspect of the sperm head; (3) there are no neck, midpiece, or tail defects; and (4) cytoplasmic droplets are less than one third the size of the head. Although these parameters appear quite similar to Tygerberg criteria, slightly looser standards increase the reference value for normal morphology to 30%. Use of both grading schemes varies from laboratory to laboratory.

Spermatozoa Viability

Spermatozoa viability is tested by evaluating the capacity for the spermatozoa membrane to exclude dyes or other substances. Because the percentage of viable spermatozoa is usually slightly greater than that of motile spermatozoa, viability testing is usually performed only when motility is less than 40%.

The eosin–nigrosin test is a common method for assessing viability. Intact cells exclude eosin dye (red); nonviable cells exhibit red staining. Nigrosin functions as a counterstain to visualize unstained live cells. Equal volumes of liquefied semen and eosin–nigrosin stain are mixed on a clean, dry microscope slide. The total number of spermatozoa that do not take up the stain is recorded as percent of viable spermatozoa in the sample.

Another method for assessing spermatozoa viability is the hypo-osmotic swelling test. The test relies on the ability of live spermatozoa to tolerate moderate hypo-osmotic stress as evidenced by controlled swelling. Dead spermatozoa do not show discernible swelling when exposed to hypo-osmotic conditions, whereas viable spermatozoa exhibit a characteristic coiling pattern in the tail.⁹

Functional Spermatozoa Tests

Despite normal semen parameters, spermatozoa may be functionally unable to perform all the requisite steps necessary for fertilization. Several tests have been developed to evaluate potential functional obstructions to fertility. The predictive value of these tests is questionable, however, and their usefulness in therapeutic fertility treatment is debatable, at best.

Acrosome Reaction

Before fertilization, spermatozoa must undergo the acrosome reaction. The acrosome reaction involves the fusion of the acrosomal membrane with the adjacent plasma membrane. This fusion results in the release of acrosomal content. Evaluation of acrosome functional activity may be accomplished by mixing capacitated spermatozoa with inducers of the acrosome reaction. Spermatozoa are then evaluated using fluorescent stain to determine the status of the acrosomal membrane.

Zona Pellucida Binding Assay

The zona pellucida binding assay is another adjunctive test of spermatozoa function. This test assesses the capacity of spermatozoa to bind to the outer surface of the zona pellucida, a prerequisite for binding and penetrating the egg. Microscopically, the zona pellucida is divided and each half is mixed with a sample of the patient’s spermatozoa or a sample of a fertile donor. Spermatozoa bound to the zona pellucida outer surface are counted. A hemizona index is calculated by dividing the total number of patient spermatozoa bound by the total number of donor spermatozoa bound. Successful IVF is associated with an index greater than 60%.¹⁰

Hamster Egg Penetration Assay

The zona pellucida, the acellular glycoprotein layer surrounding the ovum, regulates species-specific fertilization. Removal of this layer allows evaluation of interspecies, spermatozoa–oocyte interaction using the sperm penetration assay (SPA). This procedure assesses the ability of human spermatozoa to penetrate a hamster oocyte as a test of the fertilizing capability of the human spermatozoa.¹¹ Human spermatozoa penetration of zona-free hamster oocytes has been shown to correlate with human spermatozoa penetration of human ova.¹²

Sperm Chromatin Structure Assay/Single-cell Gel Electrophoresis

Spermatozoa DNA integrity tests are useful in the evaluation of the haploid chromatin of the spermatozoa nucleus. Chromosomal quality is determined by both chromatin maturity and chromatin normalcy. Both the sperm chromatin structure assay (SCSA) and single-cell gel electrophoresis test (also known as comet assay) serve as tools for measuring clinically important properties of spermatozoa nuclear chromatin integrity.

Assays utilize direct staining and evaluation of spermatozoa chromatin to assess the integrity of the genetic material. The tests provide useful information on the presence of genetic damage to spermatozoa DNA. Briefly, both procedures evaluate shifts from intact, double-stranded DNA to denatured, single-stranded DNA.¹³ These assays can be used to assess male spermatozoa DNA integrity as related to fertility potential and embryo development as well as effects of reproductive toxins.¹⁴

Collection-Specific Spermatozoa Evaluation and Processing

For most healthy men, masturbation is a straightforward method of semen collection. For men with ejaculatory dysfunction or azoospermia, however, other methods of spermatozoa recovery are necessary for fertility evaluation or therapeutic maneuvers. Men who are unable to ejaculate may require clinical spermatozoa collection; men with azoospermia may be aided by surgical intervention.

Processing of Semen Collected by Masturbation

Exposure to seminal plasma results in decreased spermatozoa motility and viability. Prolonged exposure can irrevocably diminish the fertilizing potential of spermatozoa. To obtain the heartiest sample for both laboratory tests of spermatozoa function and clinical application, spermatozoa must be separated from the seminal plasma after ejaculation. There are three fundamental approaches to the separation of spermatozoa from seminal plasma: spermatozoa washing; spermatozoa migration, or swim-up; and spermatozoa density gradient separation. These general spermatozoa processing techniques are applicable to both intrauterine insemination (IUI) and IVF procedures.

Spermatozoa Washing

Spermatozoa washing is the most common method for the separation of spermatozoa from seminal plasma. Washing not only quickly and efficiently removes detrimental factors present within the seminal plasma, but also concentrates spermatozoa within the ejaculate. Spermatozoa washing is accomplished by mixing the ejaculate with an appropriate, protein-supplemented medium. Mixtures are then centrifuged and spermatozoa pellets form at the bottom of the container. The supernatant is carefully aspirated and the pellet is resuspended. This procedure can be recommended for samples with good concentrations of highly motile spermatozoa and very few nonspermatozoa contaminating cells.

Spermatozoa Migration Techniques (Swim-up Method)

In the female reproductive tract, migration into the periovulatory cervical mucus isolates a population of potentially functional spermatozoa. Although simple washing removes seminal plasma from a semen sample, it does nothing to accomplish this goal. Techniques that enrich the population of motile or morphologically normal spermatozoa from a sample may be utilized for IUI or IVF, and include the classic swim-up procedure. In the spermatozoa swim-up procedure, the innate ability of motile spermatozoa to migrate against gravity is used to enrich for motile spermatozoa.

Two different methods of swim-up separation are commonly used. In the first (swim-up from semen), freshly liquefied semen is split into several fractions and placed in tubes beneath a layer of low-viscosity culture medium. In the second method (swim-up from pellet), fresh semen is first washed and pelleted. The supernatant is removed and medium is gently layered over the pellet. In both methods, motile spermatozoa naturally migrate from the lower layer (fresh semen or pelleted spermatozoa) into the overlying culture medium during incubation at 37°C.

Spermatozoa washing of a sample before enrichment for motile spermatozoa (swim-up from pellet) has been criticized because dead and abnormal spermatozoa present in the original sample remain in the pellet. Aitken and Clarkson showed that the functional fertilizing capacity of motile spermatozoa isolated from such a pellet can be impaired when compared to that of motile spermatozoa isolated before spermatozoa washing and pelleting.¹⁵ The authors propose that the production of oxygen free radicals by abnormal spermatozoa in the pellet may result in cellular damage in motile spermatozoa, leading to abnormal spermatozoa function. Additionally, the centrifuged pellet has a relatively small interface with the overlying layer, making it difficult for motile spermatozoa in the bottom of the pellet to migrate into the media.

Density Gradient Separation

Density gradient separation relies on a density gradient coupled with centrifugation to separate fractions of spermatozoa based on motility, size, and density. Various techniques have been described using continuous or discontinuous (discrete layers interrupted by defined interfaces) gradients. A variety of commercially available gradient materials are in use. An aliquot of the spermatozoa sample is gently layered over gradients of such a material and centrifuged. The resulting pellet is washed in culture medium before resuspension in medium at the desired concentration. Additionally, one can perform density gradient separation and washing followed by spermatozoa swim-up in bicarbonate-buffered media at 37°C in a CO₂ incubator to select for a highly motile population of spermatozoa for insemination in IVF.

Processing of Spermatozoa Collected Following Retrograde Ejaculation

Retrograde ejaculation involves the flow of semen into the bladder during orgasm. During normal ejaculation, the urinary sphincter remains closed to guard against retrograde expulsion of semen into the bladder. If the sphincter is incompetent or otherwise nonfunctioning, semen may preferentially travel back into the bladder rather than through the urethra. Retrograde ejaculation is a rare cause of male infertility. It should be suspected in patients with oligospermia or aspermia. Any condition that interferes with the sympathetic innervation of the bladder neck or vas deferens may result in retrograde flow. The condition is diagnosed by examining postejaculate urine for spermatozoa. The presence of 10 to 15 spermatozoa/HPF confirms the diagnosis and differentiates retrograde ejaculation from anejaculation.

Viable spermatozoa may be retrieved from the urine of patients failing medical therapy.¹⁶ Because urine’s acidity makes it naturally toxic to spermatozoa, it is first suggested to alter the chemical milieu of the bladder to minimize its deleterious effects on spermatozoa quality. Alkalinization of the urine above a pH of 7.5 can be achieved by instructing the patient to ingest sodium bicarbonate for several days prior to collection. The patient empties the bladder before masturbation. Immediately after orgasm, the patient urinates into a sterile container, which is taken to the laboratory for spermatozoa extraction. Alternatively, for men unable to volitionally void, the bladder may be catheterized and washed with fresh, pH-adjusted, spermatozoa processing medium. After irrigation, a small volume of medium is left in the bladder. The patient masturbates and ejaculates, and the bladder is catheterized again to obtain fluid containing retrograde ejaculate, which is collected and transported to the laboratory for processing. Laboratory procedures begin with centrifugation and removal of urine from the spermatozoa. The resuspended pellet can then be processed as previously described for samples produced from masturbation.

Processing of Spermatozoa Collected Following Vibratory Stimulation/Electroejaculation

Failure of emission is termed anejaculation. Causes of anejaculation are similar to those of retrograde ejaculation, although spinal cord injury is a much more common cause. Because most of these patients do not respond to sympathomimetic therapy, they may require the use of external stimulatory devices to achieve emission. The most common methods of stimulation are penile vibratory stimulation and rectal electroejaculation. Penile vibratory stimulation entails the application of a penile vibrator to the glans penis, leading to a reflexive ejaculation. This technique induces ejaculation in approximately 70% of anejaculatory men with spinal cord injuries.¹⁷

Men with lesions above the level of T10 are more likely to respond than those with lesions below that level. Electroejaculation may be used in men who do not respond to vibratory stimulation. This procedure involves carefully inserting a probe into the rectum, followed by the intermittent application of electrical current to the periprostatic nerve plexus. The current stimulates the muscular tissue of the prostate, seminal vesicle, vas deferens, and epididymis to contract and propel semen into the urethra. It is successful in approximately 75% of patients.^18,¹⁹ If antegrade ejaculation ensues, it may be collected from the urethra. If retrograde ejaculation occurs, then the urine is alkalinized beforehand and the bladder irrigated with medium and catheterized to remove any spermatozoa-bearing fluid.

Risks of electroejaculation include rectal thermal injury or perforation. The rectum should be examined with proctoscopy before and after the procedure. Life-threatening autonomic dysreflexia, or massive, uncoordinated sympathetic release, may occur in patients with spinal cord injuries above the level of T6. Blood pressure monitoring is imperative in this patient population. Semen obtained by vibratory stimulation and electroejaculation frequently has low volume, high spermatozoa concentration, and poor motility, likely owing to stasis within the genital tract.²⁰ Again, processing of samples produced with vibratory stimulation or electroejaculation continues as in samples produced with masturbation.

Processing of Spermatozoa Collected Following Surgical Spermatozoa Collection

The management of severe male factor infertility has been dramatically changed with the introduction of intracytoplasmic sperm injection (ICSI). Because only one spermatozoa is needed to inject into each oocyte, few spermatozoa are needed to ensure fertilization of retrieved oocytes with ICSI. Advances in surgical techniques of spermatozoa retrieval combined with ICSI have allowed men with certain male infertility factors previously considered to be incompatible with fertility to father biologic progeny. Spermatozoa may be retrieved in men with both nonobstructive and obstructive azoospermia. Chapter 53 discusses the technical details of the specific surgical procedures.

Various surgical techniques have been refined to collect viable spermatozoa from the epididymis and testes. Because low quantity and quality spermatozoa are usually recovered, specialized processing is required when ICSI is indicated.

Microsurgical Epididymal Sperm Aspiration

Microsurgical epididymal sperm aspiration (MESA) is indicated for men with reconstructible or nonreconstructible obstructive azoospermia. It is a relatively simple procedure requiring regional or general anesthesia. MESA involves direct exposure of the epididymis with microscopic retrieval of spermatozoa from that structure. An incision is made through the scrotal skin, dartos, and tunica vaginalis to expose the testis and epididymis. A small incision is made in the wall of an epididymal tubule, and a needle or hollow glass rod is used to aspirate fluid from the lumen.

Although pregnancy rates are similar, MESA is preferred to the other, less-invasive method of epididymal spermatozoa extraction, percutaneous epididymal sperm aspiration (PESA), because it provides the setting to correct reconstructible etiologies of azoospermia identified at the time of exploration. Additionally more spermatozoa are recovered with MESA than PESA.²¹

Testicular Sperm Extraction/Testicular Sperm Aspiration

Testicular sperm extraction (TESE) is an effective procedure for the retrieval of spermatozoa for ICSI. It is indicated in men with nonobstructive azoospermia as well as men with obstructive azoospermia in whom spermatozoa could not be retrieved from the epididymis.²² TESE entails testicular biopsy with retrieval of spermatozoa. An incision is made through the scrotal skin, dartos, and tunica vaginalis to expose the testis. The tunica albuginea is then opened. The testicle is gently squeezed to deliver seminiferous tubules through the incision. Several tubules are cut and immediately placed in medium for processing.

Testicular sperm aspiration (TESA) is a simple, percutaneous procedure that utilizes a small-gauge needle and syringe to retrieve testicular tissue. The procedure is the least invasive of the testicular spermatozoa retrieval methods and can be performed in the office setting under local anesthesia. It is indicated in patients with obstructive azoospermia. In this procedure, a fine-gauge, beveled needle attached to a 10-mL syringe is inserted through the scrotal skin into the anterior testis. Back-pressure is applied to the syringe while the needle is gently moved back and forth until tissue is aspirated into the syringe. The needle is withdrawn and the tissue is transferred into sterile medium, which can then be sent for processing.

Microscopic Testicular Sperm Extraction

Microscopic testicular sperm extraction (microTESE) is a relatively new procedure designed to maximize the yield of testicular spermatozoa from men with nonobstructive azoospermia while minimizing the amount of tissue excised.²³ Testicular microdissection has several advantages over the standard, open procedure. Use of the operating microscope allows for visualization of small blood vessels and minimizes the risk of injury to blood supply. Additionally, spermatozoa production in patients with nonobstructive azoospermia is not constant throughout the testis, with isolated areas showing spermatogenesis even while the majority of the tissue is quiescent.

Schlegel’s discovery that tubules with active spermatogenesis appear larger and fuller than nonactive tubules with microscopy make microTESE a much more sensitive procedure than TESE for identifying active areas. Much smaller volumes of tissue are needed for spermatozoa retrieval, avoiding the impairment in testosterone production that may occur after removal of larger amounts of tissue with TESE.²⁴

As in TESE, the testis is exposed and the tunica vaginalis is opened. An operating microscope is used to identify a relatively avascular area on the anterior surface of the tunica albuginea. A “microknife” is used to make a small incision in this area. The seminiferous tubules are expressed through the incision and careful dissection is used to identify large, full tubules. If such a tubule is encountered, it is excised with sharp, curved scissors. The sample is placed in medium and examined under microscopy for the presence of spermatozoa after microdissection. If spermatozoa are not found other areas or, if necessary, the contralateral testis is examined.

Spermatozoa found in the epididymis and testis are not fully mature. Those isolated from these structures may exhibit some movement. However, specimens retrieved from the testis are frequently immotile. Motility of epididymal- and testicular-derived spermatozoa, reflecting viability, is a critical factor in predicting the success of ICSI. Additionally, testicular spermatozoa are enclosed within the surgical specimen and must first be released before subsequent processing. Methods for recovery and maturation of surgically retrieved spermatozoa have been described.²⁵ In MESA, the aspirate is flushed in a small volume of medium. If spermatozoa are present, an aliquot should be used for concentration and motility determination. If present, fresh epididymal spermatozoa may be used for injection or cryopreservation.

For testicular-derived samples, the collected tissue is flushed into a small volume of medium and minced with sterile scissors. Using microscopy, each seminiferous segment is slit with a small-gauge needle and “milked” to tease any spermatozoa present into the medium. The seminiferous tubule contents are then pipetted through a finely pulled glass pipette to produce a single-cell suspension. A portion of this is assessed for spermatozoa. If spermatozoa are present, care is taken to minimally dilute the sample yet to suspend the sample in media before 24- to 48-hour culture to enrich for a population of motile spermatozoa.

Spermatozoa Preservation

Cryopreservation of human spermatozoa became a feasible method of storage in 1949, when Polge discovered that fowl spermatozoa could be protected from the effects of freezing by the addition of glycerol. Soon after, Sherman described successful cryopreservation and thawing of human spermatozoa with subsequent pregnancy.²⁶ Since that time, a large body of research has focused on understanding the physical and chemical changes that occur during freezing and in evaluating methods and agents to optimize freezing. Although many advanced aspects of cryobiology are beyond the scope of this text, several basic principles are important to discuss.

Cryopreservation has become an important component of ART. Patients with ejaculatory dysfunction requiring vibratory stimulation or electroejaculation may freeze spermatozoa and use aliquots to attempt fertilization over several cycles without the need for repeat procedures. It has been recommended that surgically retrieved spermatozoa be cryopreserved for future cycles of IVF with ICSI to obviate the need for repeat procedures.²⁷ Donor insemination relies on the reliable, prolonged storage of large numbers of samples. Patients facing potentially fertility-damaging medical therapies may choose to store spermatozoa for future use.

OOCYTES: PREPARATION AND EVALUATION

Advances in culture medium composition have significantly influenced embryo quality and pregnancy rates over the years. All media for ART can now be purchased commercially. Before use they must be tested for toxins and their ability to support the growth and development of embryos. Media should only be opened in a laminar flow hood, with attention to maintaining sterility during the addition of protein. After any media preparation or addition, it should be filtered for assurance of sterility.

Oocyte Processing Area

A great deal of consideration must be made to the setup of the laboratory to ensure the best results for all procedures. The laboratory should be as close as possible to the procedure room where oocyte retrievals and embryo transfers are taking place. It needs to be a controlled environment devoid of environmental toxins. It may be recommended that the laboratory have positive-pressure airflow; the temperature and lighting need to be controlled for optimal results. A laminar flow hood is indispensable for any preparation of media and manipulation of gametes and embryos.

All equipment in the IVF laboratory must be monitored daily for proper performance. A stringent quality control program is essential for the assurance of consistent results in an embryology laboratory. All media, equipment, and other materials such as dishes and pipettes that will come in contact with gametes and embryos should be biocompatible with mammalian embryos. The mouse embryo assay can be used to detect toxins, suboptimal quality, or suboptimal culture conditions. This assay consists of the culture of mouse embryos from the zygote stage until the blastocyst stage in laboratory test conditions.

Laboratory and Media Preparation

There are two basic kinds of media used for ART procedures, one used during the handling of gametes and embryos out of the CO₂ incubator, called processing medium, and another used for culture while in CO₂ incubators, called culture medium. Both consist of a combination of nutrients necessary to maintain early embryo metabolism and proper pH and osmolarity. A source of protein must be added, such as albumin or synthetic serum, in a percentage that can vary from 2 to 15 mg/L. This addition of protein is not only important for the stability of osmolarity and pH, but it also helps with the manipulation of oocytes and embryos by reducing cell attachment to any surface, as is their natural tendency.

Processing media are modified culture media, with the addition of a buffer called HEPES, capable of maintaining medium pH at a stable state out of the CO₂ incubation. HEPES, however, is known to alter ion channel activity in the plasma membrane and may have a degree of toxicity relative to exposure concentration, temperature, and time. For this reason it is recommended to minimize exposure time of cells to HEPES-buffered media. The addition of an anticoagulation factor, such as heparin, in low concentrations, may be recommended for the processing medium used at oocyte retrieval to help avoid formation of clumps of coagulated blood. These coagulants can make it difficult to visualize the oocyte–cumulus complex.

Media used for the culture of embryos are usually bicarbonate-buffered and kept in an incubation chamber. It is important that this medium be maintained at a stable pH and temperature, because embryos are extremely sensitive to variations of these two factors. Culture media, used for embryo development inside incubation chambers, should always be equilibrated inside the CO₂ environment before use. Overlaying of the medium with mineral oil is recommended to help avoid evaporation and increase stability of pH while the dish is temporarily out of the CO₂ environment. Oil overlay provides an effective barrier to atmospheric volatile organic compounds, which can be embryotoxic if exposed directly to the culture medium.

Oocyte Collection and Evaluation

Before oocyte retrieval, it is necessary to set up the laboratory to allow optimal conditions for rapid, efficient, and safe oocyte collection and placement within an incubator. Proper setup minimizes variations in oocyte environment outside of the incubator.

Room Conditions

It is important to ensure that all room conditions in the laboratory are stable before oocyte collection. All surfaces should be clean and warming blocks should be maintained at 37°C to maintain the temperature of the tubes used for the follicular fluid collection. It is recommended that all pipettes and dishes be rinsed with culture medium before use.

Media

Oocyte Collection

Tubes containing processing medium plus heparin, to prevent clotting, should be prepared. Using sterile technique, these are placed in a warming block in the room where the collection will be performed by the physician. The aspirated follicular fluid is collected by the physician into processing medium containing protein at 37°C and immediately transported to the laboratory.

In the laboratory, aspirates are examined quickly for the presence of cumulus–oocyte complexes. When an oocyte is identified, it is rinsed in processing medium, quickly graded, and placed in a pre-equilibrated culture dish in the CO₂ incubator. This process must be as fast and efficient as possible. In cases where the laboratory and egg collection rooms are separated by significant distance, the laboratory staff may employ mobile equipment to conduct the egg search in the egg collection room and to provide adequate conditions for oocytes during transit back to the laboratory. A modified pediatric isolette fitted with a microscope is commonly used for this purpose.

Oocyte Assessment

Oocyte–cumulus complexes are scored before incubation. Accurate assessment of oocyte maturity at the time of the retrieval may be informative for timing and success of fertilization and may permit identification of some oocyte abnormalities. Initial oocyte grading relies on direct optical analysis of oocyte– cumulus complex morphology. The grade is based on characteristics of the cumulus oophorus and corona radiata. This first observation should be made quickly while searching through the follicular fluid. In cases of IVF by conventional insemination, the cumulus is not removed before insemination, so observation is somewhat limited.

Immature oocytes are defined as being at a stage of meiosis prior to metaphase of meiosis II. This includes oocytes in prophase of meiosis I, which are identified by the presence of a germinal vesicle or nuclear envelope in the cytoplasm, without any polar body present in the perivitelline space (Fig. 39-1). If present, cumulus and corona are commonly very tightly condensed. As prophase I resumes, the oocyte enters into metaphase of meiosis I. This intermediate stage of maturation is recognized by the disappearance of the germinal vesicle and the absence of the first polar body (Fig. 39-2). For meiosis I oocytes the cumulus may be expanded, but the corona can still be compact. The extrusion of the first polar body marks the transition to a mature oocyte, which is now considered to be at meiosis II (Fig. 39-3). Metaphase II oocytes will usually have a fully expanded cumulus. Under normal circumstances the oocyte will remain at meiosis II until fertilization, when meiosis II resumes, the second polar body is extruded, and the male and female pronuclei form (Fig. 39-4).

Figure 39-1 Germinal vesicle intact or immature human oocyte after cumulus removal.

(Courtesy of Kathleen A. Miller, Reproductive Medicine Associates of New Jersey, Morriston, N.J.)

Figure 39-2 Metaphase I human oocyte without a polar body after cumulus cell removal.

(Courtesy of Kathleen A. Miller, Reproductive Medicine Associates of New Jersey, Morriston, N.J.)

Figure 39-3 Metaphase II (mature) oocyte. A, Oocyte–cumulus cell complex. B, Denuded oocyte showing presence of first polar body.

(Courtesy of Kathleen A. Miller, Reproductive Medicine Associates of New Jersey, Morriston, N.J.)

Figure 39-4 Normal zygote showing presence of two polar bodies and two pronuclei.

(Courtesy of Kathleen A. Miller, Reproductive Medicine Associates of New Jersey, Morriston, N.J.)

Assessment of oocyte morphology after cumulus cell removal may reveal its true morphologic quality. The appearance of cytoplasm, zona pellucida, and polar body should be considered. A homogeneous cytoplasm, with uniform color and granularity and occupying most of the space inside the zona pellucida, represents a good quality oocyte. Vacuoles, a dark color, shrunken shape, or granularity are considered negative signs in the morphological quality evaluation of oocytes. It may be important to observe shape and thickness of the zona pellucida; it should not be cracked and should have an appropriate thickness for its developmental stage.

There are many oocyte morphologic grading systems. The folllowing is an example of one based on visual appearance:

0 = Atretic oocyte, degenerate.

1 = A germinal vesicle (GV) is present. The oocyte is very immature.

2 = The corona is tight/dark, but no GV is present. The cumulus is starting to loosen, but the oocyte is hard to visualize. The oocyte is still immature.

3 = The corona is radiating and loosening, the cumulus is loose and dispersed. The oocyte is visible, but the first polar body is still not present. The oocyte is still immature but likely will extrude a polar body within a few hours.

4 = The optimal level of maturity for insemination. The oocyte has fully expanded cumulus and the first polar body is visible.

IN VITRO MATURATION OF OOCYTES

In vitro maturation of oocytes is the act of culturing and maturing oocytes that have been harvested at an immature state. What is unclear in this definition are the starting material and the endpoint measurements. One must be extremely cautious in extrapolating and comparing data regarding in vitro maturation of oocytes from exogenous hormone-stimulated cycles, human chorionic gonadotropin (hCG)-only-primed cycles, and nonstimulated cycles. Although the starting materials (immature oocytes) from each of these cycle categories may look the same at the light microscope level, their degree of cytoplasmic maturation and thus fertilization and embryonic developmental competence can be very different. In addition, from the definition, the true measure of successful oocyte in vitro maturation is complete nuclear maturation and ability to fertilize and develop into an embryo that is capable of supporting a live birth.

In vitro maturation of human oocytes was first demonstrated by Dr. Edwards in 1965. Although this technique was viewed as having great potential in providing oocytes for early IVF work, the use of exogenous gonadotropin administration proved more efficient and became the gold standard in infertility treatment. Because of the numerous perceived advantages of in vitro maturation, continued interest and investigation have persisted in the area of human oocyte in vitro maturation. Pregnancies have been reported using in vitro maturation of immature oocytes that were retrieved after conventional follicle-stimulating hormone (FSH) stimulation and hCG administration.²⁸^–³⁰ However, such an approach minimizes the advantages that warrant in vitro maturation. In addition, it could be argued that such an approach represents pseudo in vitro maturation of oocytes because the exogenous gonadotropins and resulting follicular development most likely progressed oocytes’ cytoplasmic maturation while nuclear maturation lagged.

Recent reports have utilized a single dose of hCG (no exogenous FSH administration) on day 10 to 14 of the cycle with oocyte isolation 36 hours after hCG administration.^31,³² These studies were performed in patients with polycystic ovary syndrome (PCOS). In this subpopulation the use of single-dose hCG administration coupled with in vitro maturation and ICSI appears to be an efficient means of treating infertility while reducing the cost and concerns of hyperstimulation syndrome. Although the numbers in the study are small, the resulting clinical pregnancy rate is encouraging, at approximately 39%.³² Whether such a treatment regimen will work in non-PCOS patients remains to be determined.

At the other extreme, true in vitro maturation entails collection of oocytes from ovaries without exogenous gonadotropin stimulation. Such an approach will, when it becomes efficient, allow realization of all the perceived advantages of in vitro maturation. Clinical pregnancies using in vitro maturation without exogenous gonadotropin stimulation have been reported.³³^–³⁵ These results suggest that using an approach of in vitro maturation of human oocytes, coupled with IVF, is a means of obtaining live births. However, this technique requires extensive evaluation because the cumulative success in all of these studies, as determined by live births, is less than 5%. Numerous factors individually, or in combination, may be contributing to the poor success of IVF after in vitro maturation. These include oocyte quality, culture conditions, and endometrial receptivity.

IN VITRO FERTILIZATION

IVF is the procedure in which oocytes and spermatozoa are obtained from a couple, with the combination of gametes taking place in a laboratory. The aim of this procedure is to produce embryos that can ultimately give rise to a viable pregnancy and healthy offspring. Spermatozoa are obtained by above-mentioned methods.

Conventional Insemination

If spermatozoa are of sufficient quality and quantity, IVF can be attempted with conventional insemination. Although the terms of sufficient quality and quantity are indeed subjective, each laboratory will have its own criteria for when a couple uses conventional insemination or ICSI to achieve fertilization. Spermatozoa can be processed in numerous manners or with a combination of techniques. The primary goal is to remove seminal plasma and isolate motile spermatozoa of normal morphology with as little nongamete cellular contamination as possible. The processes used include semen washing, density gradient separation, and swim-up procedures.

Independent of the method of spermatozoa recovery, it is advised to incubate spermatozoa for approximately 4 hours in culture medium containing a protein source in a CO₂ incubator to allow capacitation of spermatozoa. Capacitation is defined as the spermatozoa gaining the ability to bind to the oocyte’s zona pellucida, penetrate the zona pellucida and oolemma, and ultimately fertilize the oocyte. Just before insemination the spermatozoa sample should be assessed for concentration and percent motility. With these values, one can calculate the volume of insemination medium containing spermatozoa that will equate to the desired insemination spermatozoa number.

Oocytes are routinely inseminated 3 to 6 hours after oocyte retrieval is performed, taking into consideration their level of maturity for the proper time of incubation. Oocytes can be inseminated in groups or individually. Groups of oocytes can be incubated and inseminated either in organ culture dishes, four-well dishes, or test tubes containing equilibrated medium with or without oil overlay. Individual oocytes can also be inseminated in 30- to 50-μL drops of equilibrated medium in culture dishes with oil overlay. This reduces the number of spermatozoa necessary for the insemination. The final concentration of motile spermatozoa used for conventional insemination will differ between laboratories based on personal experiences. Concentrations that are too high can result in increased incidence of polyspermic fertilizations (more than one spermatozoon penetrating an oocyte). Concentrations that are too low may compromise fertilization rates. Generally concentrations range from 50,000 to 100,000 motile spermatozoa/mL. These numbers may be adjusted in cases of suboptimal parameters of motility or morphology.

Intracytoplasmatic Sperm Injection

ICSI consists of insertion of a single spermatozoon directly into the oocyte cytoplasm. This technique was first successfully applied to human oocytes in 1994 and has since revolutionized the treatment of severe male factor infertility. By injecting a spermatozoon into the oocyte cytoplasm, many steps of spermatozoa processing and developmental prerequisites are bypassed without compromising fertilization rates. There is currently debate as to when to perform ICSI. Some programs utilize the procedure quite freely, whereas others are more conservative. Evidence strongly supports the following indications for the use of ICSI:

• Couple has had a prior failed fertilization by conventional insemination.

• Total motile spermatozoa concentration is less than 10 million/mL.

• Spermatozoa morphology is less than 4% normal forms based on strict criteria (this may depend on other semen parameters such as concentration, motility, and volume).

Technique

The equipment for ICSI, as in any other micromanipulation procedure, consists of an inverted phase contrast microscope with micromanipulators and injectors, which permit the manipulation and cellular surgery. A warming stage may be recommended on the microscope for maintenance of a consistent temperature, although some groups show quite acceptable results without using a warming stage for micromanipulation.³⁶ The hydraulic system in a micromanipulator should work with maximal precision, and the microscope should be placed in a very stable location.

Removal of the cumulus and corona radiata cells surrounding the oocytes (denuding) is essential for ICSI. The cumulus cells need to be removed so that they do not interfere with visualization of the egg or the injection. Two needles may be used to separate most of the cumulus cells by “cutting” the cumulus off and leaving the gamete with just a few cells attached. The oocytes, now almost free of cumulus cells, can be exposed for up to 30 seconds to a solution of processing medium containing 80 IU/mL of hyaluronidase and manually pipetted to remove remaining cells. The oocytes are then immediately washed in processing medium without hyaluronidase by repeated pipetting with a pulled pipette with an inner diameter just larger than the oocyte.

Dishes for the actual injection can be made in many ways. The procedure can be performed in 5-μL microdrops of processing medium supplemented with protein, overlayed with mineral oil in a shallow culture dish. A 5-μL drop of a 7% polyvinylpyrrolidone (PVP) solution in processing medium can be placed in the center of the ICSI dish. Just before beginning the injections approximately 1 μL of spermatozoa suspension is added to the PVP drop. The high viscosity of this solution slows down spermatozoa movement and facilitates immobilization and aspiration. Additionally, it helps to prevent spermatozoa from sticking to the inside of the injection pipette. At the time of injection, denuded oocytes are placed one per drop into 5-μL microdrops around the 5-μL microdrop of PVP containing processed spermatozoa. Only a couple of oocytes should be added to the dish at a time to avoid prolonged exposure to the environment outside an incubator.

Spermatozoa are aspirated from the drop using the injection pipette. A morphologically normal-appearing, motile spermatozoon is identified and the tail is broken using the injection pipette with a slashing motion against the bottom of the dish. The spermatozoon is then aspirated, tail first, into the pipette in preparation for ICSI and the pipette is moved into the drop with the oocyte. The holding pipette is used to manipulate the oocyte into position with the polar body in the 12 or 6 o’clock position. Negative pressure is applied to the pipette to hold the oocyte in place for the injection. The injection pipette is positioned in accordance with the outer border of the oolemma on the equatorial plane at 3 o’clock.

The spermatozoon is then brought to the tip of the pipette, which is pushed against the zona pellucida. The needle is pushed forward through the zona pellucida and into the oolemma. Gentle suction is applied when the tip is believed to have penetrated the oolemma and a small amount of ooplasm is aspirated to ensure that the membrane has been breached. The spermatozoa is then released into the ooplasm and the pipette is removed. Injection of any extra medium should be avoided. A timeline schematic of the entire ICSI process is depicted in Figure 39-5.

Figure 39-5 Representative scheme of ICSI procedure. (University of Michigan ART Laboratory Procedures Manual.)

PREPARATION AND EVALUATION OF ZYGOTES AND EMBRYOS

Fertilization

The fertilization check of oocytes is performed 15 to 18 hours after insemination for both IVF and ICSI procedures. It is necessary to examine the oocytes/zygotes within this time period to visualize the presence of pronuclei and polar bodies. Fertilization checks before pronuclear formation or after their dissolution can lead to misclassification of a zygote as an oocyte not fertilized or to an inability to recognize abnormally fertilized oocytes. It is important that fertilization checks be performed rapidly yet efficiently. If oocytes have undergone conventional IVF, the cumulus cells must be removed to clearly see the oocyte/zygote. This is done simply by repeatedly pipetting the oocyte/zygote with a pulled pipette with an inner diameter just larger than the oocyte or zygote.

Normal fertilization is characterized by the presence of two pronuclei, one male and one female, in the ooplasm and two polar bodies in the perivitelline space. Polar bodies may fragment, leading to the appearance of more than two. Abnormal fertilization may be represented by a polypronuclear zygote, which means more than two pronuclei in the ooplasm. Abnormal fertilization can be classified as polyspermic, due to the penetration of more than one spermatozoon, or polygynic, due to disturbance in the second meiotic division, retention of the second polar body, and formation of two female pronuclei. A polypronuclear zygote can develop as preimplantation embryos yet are genetically abnormal. This justifies the recommendation to discard all polypronuclear zygotes observed at fertilization check.

Abnormal fertilization may also be represented by oligopronuclear zygotes. The term applies to zygotes that express a single pronuclei. Only one pronuclei and the presence of two polar bodies may be observed in some cases when the oocyte undergoes parthenogenic activation, or failure of the spermatozoa head to decondense. It is possible that a second pronuclei will develop later than the first one, and a repeat observation about 4 hours after the first check is recommended.

Failed fertilization is represented by absence of pronuclei and the presence of one or two polar bodies that may be in the process of degeneration. Failure of the oocyte to fertilize can have many causes, including poor-quality oocytes, problems in the interaction between the gametes, spermatozoa acrosomal reaction problems, or suboptimal culture conditions.

Embryo Assessment

Embryos can be assessed and graded daily while they are in culture. A standard morphologic method of grading is applied according to observations made on embryo development until their transfer to the uterus on day 3 or day 5 (blastocyst stage). There are numerous scoring systems proposed for embryo development. Criteria for grading include the rate of division as judged by numbers of blastomeres, size, shape, symmetry, appearance of the cytoplasm, and presence of cytoplasmatic fragments. Traditionally, embryo morphology has been used as a means of embryo selection for transfer or freezing. Timing of cleavages and formation of fragmentation can be documented. Below is an example of a cleavage-stage embryo two-step grading system that incorporates cell number and morphology:

A: Minimum of four-cell by 40 hours postinsemination; minimum of eight-cell by 64 hours postinsemination.

B: Minimum of two-cell by 40 hours postinsemination; minimum of four-cell by 64 hours postinsemination.

C: Minimum of two-cell by 64 hours postinsemination.

D: No minimums (lowest possible grade). Do not make subtractions.

Subtract from the grade for irregularities as follows:

• Spherical blastomeres with no fragmentation: No subtractions

• Spherical blastomeres with <20% fragmentation: Subtract 1 grade

• Slightly irregular blastomeres with <50% fragmentation: Subtract 2 grades

• Irregular blastomeres with >50% fragmentation: Subtract 3 grades

Examples of how to apply this classification are:

• A three-cell embryo with 15% fragmentation at 40 hours postinsemination: Reported as 3Cell/C

• A four-cell embryo with 10% fragmentation at 40 hours postinsemination: Reported as 4Cell/B

• A seven-cell embryo with 0% fragmentation at 64 hours postinsemination: Reported as 7Cell/B

• An eight-cell embryo with 0% fragmentation at 64 hours postinsemination: Reported as 8Cell/A

It is important to note that evaluation and scoring by morphology alone can be subjective and may not necessarily reveal embryos with the best developmental potential. Currently, there are numerous groups working on translational research focused on noninvasive means of assessing embryos for biomarkers that are indicative of embryonic developmental competence and implantation potential. Such evaluation, in concert with morphology, and genetic analysis (see discussion of Preimplantation Genetic Diagnosis in this chapter) has enormous potential to reshape the practice of embryo selection in the future.

Embryo Culture

With the improvement of culture systems and extended culture media, it is now possible to culture embryos to the blastocyst stage (day 5 or 6). Typically patients who are considered candidates for blastocyst transfer are young women (less than 36 years old) with good ovarian reserve or older patients with a high number of embryos in a cycle (more than 5). One of the most important factors influencing blastocyst culture and transfer success is the quality and quantity of embryos on day 3. An effective blastocyst grading system using both number and letter assignment based on degree of development and assessment of both the trophectoderm and the inner cell mass has been proposed.³⁷ The program should consider advantages and disadvantages in each individual patient and cycle before considering the option of blastocyst culture.

Assisted Hatching

One of the most common unsolved problems in IVF is the fact that embryos with apparently good developmental potential do not always implant. It has been proposed that this may be due to defects of the zona pellucida, uterine receptivity, extensive fragmentation, modifications after freezing and thawing, or even suboptimal culture conditions. Potential defects in the zona pellucida could be overcome by assisted hatching. In this procedure a controlled incision in the embryo’s zona pellucida can be applied in an attempt to improve embryo implantation. The basis of assisted hatching is the hypothesis that weakening the zona pellucida, either by drilling a hole through it or altering its stability, will promote hatching and implantation of embryos that potentially otherwise would not occur. This has become a widely applied technique, but its true value is still under discussion.

There are two common methods of assisted hatching used clinically for embryos at the six- to eight-cell stage: acid Tyrode’s solution and laser assisted hatching. Acid Tyrode’s solution can be prepared in the laboratory, following a standard protocol, and the pH adjusted to 2.5,³⁸ or it can be commercially purchased. It is important to appreciate that use of this acid can be detrimental to the blastomeres closest to the point where the acid is applied. In cases of erroneous manipulation, it can compromise the viability of the embryo. In this procedure, an assisted hatching micropipette expels the acid close to the zona pellucida of the embryo, which is secured by a holding pipette. As soon as a hole is observed, rapid suction is applied to avoid excess acid in the perivitelline space and in contact with blastomeres. After the procedure the embryos must be washed several times before further culture. Use of a laser for assisted hatching may be considered by some to be less dangerous than use of acid Tyrode’s solution assisted hatching, but its application, if not precise, may cause thermal or mutagenic effects in the embryo. Both techniques have risks and mandate fine proficiency.

Preimplantation Genetic Screening

Preimplantation genetic screening (PGS) was originally seen as a means to detect and avoid congenital diseases by evaluation of individual blastomeres of preimplantation embryos. This has value for couples with probability of having a child affected by some genetic disturbance. Initially, molecular PGS was employed for embryo sexing in couples at risk for X-linked diseases.³⁹ Technically, PGS consists of two steps, including micromanipulation and biopsy, and DNA analysis of gametes or embryos.

Micromanipulation for the biopsy includes the mechanical opening of the zona pellucida and retrieval of one or two polar bodies, in the case of diagnosis performed on oocytes or zygotes, or the retrieval of a blastomere in cases of biopsy of embryos. This procedure is briefly illustrated in Figure 39-6. For DNA analysis, there are two main techniques currently employed, polymerase chain reaction (PCR) for single-gene disease detection and fluorescence in situ hybridization (FISH) for aneuploidy.

Figure 39-6 Biopsy procedures before PGS. A, Polar body biopsy, showing removal of first and second polar bodies. B, Blastomere biopsy, showing removal of two blastomeres.

(Courtesy of Kathleen A. Miller, Reproductive Medicine Associates, New Jersey, Morriston, N.J.)

Single-Gene Defects

PCR allows the amplification of a specific DNA sequence. This is a cyclic process in which the number of DNA fragments doubles at each cycle, allowing the observation of specific genes or gene mutations through either radioactive labeling, ethidium bromide, fluorescent dye, or silver staining. Each PCR cycle is divided into three steps, for multiplication of primer-specific gene sequences.

First, natent double-stranded DNA is denatured into single strands by exposure to high temperatures (>90°C). Second, the temperature is lowered, allowing annealing of sequence-specific primers to their matching sites on the single-stranded DNA. Third, thermostable polymerase adds and extends nucleotides from the site of primer annealing. Collectively, this process provides amplification of a specific gene, or mutation, if present.

Even with the high number of copies generated with PCR cycle amplification, a single PCR procedure may yield insufficient genetic material for gene detection. Efficiency and specificity of the DNA amplification can be improved by using a nested PCR method that consists of a two-step procedure.⁴⁰

It is of extreme importance to realize that many factors can influence the reliability of PCR analysis. It is essential that numerous steps in quality assurance be implemented to protect against erroneous results and to ensure confidence in the final results.

Fluorescent In Situ Hybridization

Another way to analyze chromatin for PGS is FISH. This method utilizes direct-labeled probes, with the specific annealing of fluorescent nucleic acid probes to complementary sequences of DNA in a fixed specimen in which chromatin or chromosomes have been isolated.⁴¹^–⁴⁴ Chromatin from polar bodies or blastomeres can be treated and fixed for FISH. Cells in this procedure are fixed on a slide in a manner that ruptures the cell, removes the cytoplasmic material, and spreads and adheres chromosomes. These chromosomes are then hybridized or annealed to the chromosome-specific fluorescent probes. Using a fluorescent microscope one can count specific chromosomes and thus identify normal or abnormal chromosome numbers (Fig. 39-7).