The effect of osmotic stress on the metaphase II spindle of human oocytes, and the relevance to cryopreservation.

S.F. Mullen1, Y. Agca1, D.C. Broermann2, C.L. Jenkins2, C.A. Johnson2 and J.K. Critser1,3

1 Comparative Medicine Center and Department of Veterinary Pathobiology, University of Missouri at Columbia, Columbia MO 65211, and 2 The Atlanta Center for Reproductive Medicine, Woodstock GA 30189, USA

3 To whom correspondence should be addressed. e-mail: critserj{at}missouri.edu


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
BACKGROUND: Knowing osmotic tolerance limits is important in the design of optimal cryopreservation procedures for cells. METHODS: Mature human oocytes were exposed to anisosmotic sucrose solutions at concentrations of 35, 75, 150, 600, 1200, or 2400 (±5) milliosmolal (mOsm) at 37°C. A control treatment at 290 mOsm was also utilized. Oocytes were randomly allocated to each experimental treatment. After the treatment, the oocytes were cultured for 1 h, then fixed in cold methanol. Immunocytochemical staining and fluorescence microscopy were used to assess the morphology of the metaphase II (MII) spindle. Logistic regression was used to determine if media osmolality had a significant effect on spindle structure. RESULTS: Osmolality was a significant predictor of spindle morphology. Hyposmotic effects at 35, 75, and 150 mOsm resulted in 100, 67, and 56% of oocytes having abnormal spindles, respectively. Hyperosmotic effects at 600, 1200, and 2400 mOsm resulted in 44, 44, and 100% of the spindles with abnormal structure, respectively. CONCLUSIONS: Anisosmotic conditions lead to disruption of the MII spindle in human oocytes. Applying this fundamental knowledge to human oocyte cryopreservation should result in increased numbers of cells maintaining viability.

Key words: cryopreservation/human/MII spindle/oocytes/osmotic tolerance


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Since the birth of the first human through assisted reproduction in 1978 (Steptoe and Edwards, 1978Go), the number of couples seeking infertility treatment has increased dramatically; over 80 000 cycles of clinical infertility treatment occur annually in the United States alone (http://www.cdc.gov/nccdphp/drh/art.htm). This is due in part to the increase in patient awareness and acceptability of fertility treatments. Additionally, technological improvements have led to a higher likelihood of a successful outcome. During routine clinical treatment, ovarian stimulation occurs to increase oocyte production. With increasing success rates have come increasing concerns regarding multiple gestational pregnancies (Collins and Graves, 2000Go; Derom and Bryan, 2000Go). This has led many clinicians to limit the number of embryos transferred during a cycle. It is common to cryopreserve the supernumerary embryos.

Cryopreservation of human spermatozoa and embryos has been conducted for several decades and contributes to the increased success of reproductive medicine (Bunge and Sherman, 1953Go; Trounson and Mohr, 1983Go). Unfortunately, human oocyte cryopreservation has proven much more challenging (Paynter, 2000Go). Having effective measures to cryopreserve oocytes would provide therapy for several patient groups. Patients facing potential iatrogenic sterility would have the option of cryopreserving oocytes, just as male patients can cryopreserve sperm prior to other medical treatments (Gosden, 2001Go; Posada et al., 2001Go). Women who wish to delay childbearing may consider cryopreserving their oocytes for future clinical use. The establishment of oocyte banks could improve the safety of fertility treatments for women using oocyte donors by allowing improved screening of donors for potential transmittable diseases. Finally, patients who find gamete cryopreservation more acceptable than embryo cryopreservation could cryopreserve their oocytes, reducing the number of supernumerary embryos generated (Karow, 1997Go).

Similar to other cells, oocytes are susceptible to injury during cryopreservation (Van Blerkom and Davis, 1994Go). Mature mammalian oocytes have unique properties which make them difficult to cryopreserve. Several studies have documented the sensitivity of the MII spindle to various exposures experienced during cryopreservation. Included among these are subphysiologic temperatures: mouse (Magistrini and Szollosi, 1980Go); bovine (Aman and Parks, 1994Go); human (Sathananthan et al., 1988Go; Pickering et al., 1990Go) and exposure to cryoprotective agents (CPAs): mouse (Johnson and Pickering, 1987Go; Van der Elst et al., 1988Go); bovine (Saunders and Parks, 1999Go); human (Sathananthan et al., 1988Go; Gook et al., 1993Go). Of particular importance is how quickly spindle damage occurs. Two recent studies showed that within a matter of minutes, exposure to temperatures below 37°C caused disruption of the spindle in human oocytes, highlighting the challenge of preserving the integrity of this structure (Wang et al., 2001Go; Zenzes et al., 2001Go). It is noteworthy that previous studies (Pickering and Johnson, 1987Go; Eroglu et al., 1998Go) have documented the ability of the MII spindle in mouse oocytes to undergo repair after disruption. This ability is diminished in human oocytes (Pickering et al., 1990Go; Wang et al., 2001Go). This difference likely contributes to the higher success of oocyte cryopreservation in the mouse compared to other species (Schroeder et al., 1990Go; Stachecki et al., 1998Go). Disruption of the MII spindle will result in failure of the final meiotic reduction division and the potential for the creation of an aneuploid embryo; a condition that is fatal in nearly all instances in human development (Hassold et al., 1996Go; Hassold and Hunt, 2001Go). Previous studies demonstrated the susceptibility of mature mammalian oocytes to failure in chromosome reduction after cryopreservation (Kola et al., 1988Go; Carroll et al., 1989Go). It is our thesis that understanding the effects of the various non-physiological conditions associated with cryopreservation on the MII spindle will facilitate oocyte cryopreservation.

While most reports on human oocyte cryopreservation have been based upon slow-cooling procedures, and some very recent reports have shown improvements in success compared to those from the past two decades (Quintans et al., 2002Go; Boldt et al., 2003Go; Fosas et al., 2003Go), the success rates remain sub-optimal, with highly variable fertilization rates [e.g. from 17 to 100% (Quintans et al., 2002Go) and 0 to 100% (Boldt et al., 2003Go)] as well as in vitro development rates (non-frozen cohort embryo data was not presented in these reports). Vitrification methods are a viable alternative to slow-cooling procedures (Kuleshova and Lopata, 2002Go), and improvements in human oocyte cryopreservation have been reported using such procedures (Kuleshova et al., 1999Go; Katayama et al., 2003Go; Yoon et al., 2003Go). While vitrification may afford protection against the chilling sensitivity of human oocytes, it has its associated challenges. To achieve vitrification, concentrations of cryoprotective agents need to be much higher than those used in slow cooling methods (~5–6 M vs 1.5 M). The difficulties associated with these concentrations result from the potential toxic effects as well as the changes in osmotic pressure associated with their addition and removal (Fahy et al., 1984Go). During the addition and removal of CPAs, as well as during freezing and thawing, cell volume changes occur as CPAs and water enter and exit cells in response to osmotic and chemical gradients. These changes in cell volume associated with changes in osmotic pressure can impair a cell’s viability (Lovelock, 1953Go; Lovelock, 1957Go; Williams and Shaw, 1980Go; Mazur and Schneider, 1986Go).

Often, the methods used for CPA addition and removal are empirically derived, and not based upon the cell’s tolerance to osmotic stress. Determining the osmotic tolerance limits for cells is a common approach to optimizing cryopreservation protocols, and has been performed on numerous cell types including mouse and bovine embryos (Mazur and Schneider, 1986Go); fertilized mouse ova (Oda et al., 1992Go; McWilliams et al., 1995Go) and unfertilized rhesus monkey (Songsasen et al., 2002Go), mouse (Pedro et al., 1997Go), and bovine oocytes (Agca et al., 2000Go); together with human (Gao et al., 1995Go), mouse (Willoughby et al., 1996Go), porcine (Gilmore et al., 1996Go), and bovine (Guthrie et al., 2002Go) spermatozoa, CD34+ cells derived from human placental/umbilical cord blood (Woods et al., 2000Go), canine pancreatic islets (Zieger et al., 1999Go), and rabbit corneal epithelial cells (Pegg et al., 1987Go). Knowing the osmotic sensitivity of the cell will allow the CPA addition and removal techniques to be adapted to keep a cell within its osmotic tolerance limits (Gilmore et al., 1997Go). Control of such volume changes can be achieved by stepwise addition and removal of CPAs and inclusion of non-permeating compounds in the freezing and thawing media (Leibo and Mazur, 1978Go; Gao et al., 1995Go). To determine an optimal method for CPA addition and removal, biophysical properties such as the cell’s permeability to water (LP) and CPAs (PCPA), the osmotically inactive cell volume (Vb), and the activation energies (Ea) of LP and PCPA, must be known. These values have been determined in mature human oocytes for dimethylsulfoxide and 1,2-propanediol (Paynter et al., 1999Go, 2001Go).

Although exposure to changes in osmotic pressure has been shown to decrease the viability of mouse and bovine oocytes as discussed above, to our knowledge, the effect of anisosmotic exposure on the MII spindle has not been investigated in any species. The purpose of the present experiment was to test the hypothesis that the probability of morphological disruption of the MII spindle in a human oocyte is related to the level of anisosmotic exposure. By understanding this relationship, improved CPA addition and removal methods can be designed which will prevent detrimental volume excursion as well as minimize the time to which the cells are exposed to the CPAs.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Chemicals
All chemicals were purchased from Sigma chemical company (St. Louis, MO, USA) unless otherwise indicated.

Source of oocytes
The present study was approved by the Gwinnett Medical Institutional Review Board and the informed, written consent of patients was obtained. Patients underwent ovulation induction primarily using luteal phase leuprolide for down regulation followed by two ampoules of FSH combined with two ampoules of HMG daily in divided doses. A step down approach was utilized as the lead follicles reached the 14 mm level and hCG, 10 000 units, was given when two or more follicles reached 18 mm. Oocyte retrieval was carried out 35 h after hCG administration. All manipulations were conducted in a manner to ensure the temperature of the oocytes remained near 37°C, preventing any detrimental effects of cooling on the spindle. Only oocytes having extruded the first polar body were used in these experiments.

Anisosmotic solutions
Anisosmotic solutions consisted of sucrose as the extracellular, impermeable osmolyte supplemented with 0.23 mM sodium pyruvate and 3 mg/ml bovine serum albumin, dissolved in Milli-Q water (18 M{Omega}; Millipore, Bedford MA, USA). Solution osmolality was determined using a vapor pressure osmometer (VAPRO 5520, Wescor Inc., Logan, UT, USA). Treatment levels were 35, 75, 150, 600, 1200, and 2400 (±5) mOsm. A control treatment at 290 mOsm was also included (HEPES–HTF + 5% HSA: HEPES–HTF, Irvine Scientific, Santa Ana, CA, USA; HSA, InVitroCare, Inc., San Diego, CA, USA). An additional control treatment was utilized, using an isosmotic solution similar in composition to the treatment solutions, to test for effects of the non-physiological solution independent of osmolality.

Oocyte fixation and immunocytochemistry
Oocyte fixation was performed by transferring the cells to cold methanol (~–10°C) containing 5 mM EGTA for 5 min (Simerly and Schatten, 1993Go). They were then transferred to an immunocytochemical blocking buffer [PBS + 0.2% Triton X-100, 0.1% saponin, 130 mM glycine, 3 mM sodium azide, 2% bovine serum albumin, and 5% horse serum (horse serum was obtained from Gibco BRL, Grand Island, NY, USA)] at 4°C and incubated overnight. The remaining procedures were carried out at room temperature. The following morning the cells were transferred to a solution of the blocking buffer containing an anti-{alpha}-tubulin mouse monoclonal antibody (clone DM1A) diluted 1:4500, and incubated for 1 h. They were subsequently washed in ~2 ml of blocking buffer without antibody for 1 h, and then transferred to ~2 ml of blocking buffer which contained the secondary antibody (donkey anti-mouse IgG conjugated to Texas-Red®, Jackson Immunoresearch, West Grove, PA, USA) diluted 1:750 and incubated for 1 h. Finally, the cells were transferred to ~2 ml of blocking buffer without antibody and washed for 2–4 h. The oocytes were mounted on microscope slides using ProLong® antifade reagent containing either DAPI (1 µg/ml final concentration), or SYTOX® green (1 µM final concentration) to counterstain the chromosomes. These products were obtained from Molecular Probes (Eugene, OR, USA).

Assessment of the MII spindle
Assessing the structure of the spindle in fixed and stained cells is challenging using standard immunofluorescence techniques, due to varied spindle orientation in relation to the focal plane. When the spindle poles are parallel to the plane of focus, the classic barrel shape is evident. However, if the orientation is not in this manner, the configuration of the chromosomes and spindle fibers are difficult to assess, and what may appear as a disorganized pattern of chromatin and tubulin may actually represent the 3-dimensional structure of a normal spindle. Confocal microscopy allows the collection of thin optical sections of cellular structures (Figure 1). Using appropriate software, a 3-dimensional interpretation of the overall structure can be made. For the present study, optical sectioning of the spindles was performed using confocal microscopy, and the 3-dimensional structure was interpreted from these images. Spindle morphology was assessed using a Bio-Rad Radiance 2000 laser scanning confocal microscope and LaserSharp 2000TM software (Bio-Rad, Hercules, California, USA). A 60x UPlanApo (numerical aperture = 1.25) objective was used for all experiments, and the zoom setting during image collection was 2.3. Confocal Assistant PC software (version 4.02) was used to perform 3-dimensional analysis of the spindle. Spindles with two anastral poles having the classic barrel shape of a human spindle, and lacking chromosome scattering, were scored as normal. All other spindle morphologies were scored as abnormal.



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Figure 1. A 3-dimensional artistic reconstruction of a metaphsase II spindle from a control human oocyte based on a representative oocyte from the present study. The focal planes approximate the optical sections through the spindle imaged with confocal microscopy. It is evident from the serial optical sections that the microtubule organization (red) is appropriate for a normal meiotic spindle in which the microtubules form from the two opposite poles as seen in sections a and d. Chromosomes (green) are aligned regularly at the metaphase plate located between the two half spindles, (b and c). A 2-dimensional analysis would not have allowed such an accurate interpretation.

 
Experimental design
An incomplete randomized block design was employed for this study. Individual oocytes from each patient were randomly allocated to the experimental treatments for each replicate. Nine replicates were performed at all osmotic levels with the exception of the 35 mOsm level, where only three replicates were performed, and the isosmotic sucrose solution, where six replicates were performed. Oocytes were exposed to the treatment solution for 10 min, and subsequently returned to HEPES–HTF + 5% HSA for 10 min, which allowed the cells to return to their isosmotic volume. Subsequently, the cells were placed in HTF (Irvine Scientific, Santa Ana, CA, USA) + 5% HSA medium in a cell culture incubator (Forma Scientific, Model 3130, Marietta, OH, USA) at 37°C with an atmosphere of 6.5 ± 0.5% CO2 and air. The oocytes remained in the incubator for ~60 min prior to fixation.

Statistical analysis
Spindles scored as normal or abnormal were coded as 0 or 1, respectively. Logistic regression was used to analyze the data (Hosmer and Lemeshow, 2000Go) using SAS (The SAS Institute, Cary, NC, USA). The hypo- and hypertonic treatments were analyzed independently. The probability of a type-1 error ({alpha}) was chosen to be 0.05.


    Results
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Percentage of cells maintaining a morphologically normal spindle
The number of cells with a normal spindle structure at each treatment level are shown in Figure 2. Less than half of the cells exposed to hypotonic levels were able to maintain a normal spindle, with increasing proportions of oocytes with disrupted spindles occurring as the solution concentrations decreased. None of the spindles (n = 3) were normal at the 35 mOsm level. A similar pattern was seen in the cells exposed to hypertonic solutions, with nearly half of the spindles exhibiting abnormal structure at the 600 and 1200 mOsm level; none of the spindles were normal in the cells exposed to 2400 mOsm. In contrast, all of the oocytes exposed to the control isotonic solutions (HTF, n = 9 as well as sucrose, n = 6) maintained a normal spindle structure.



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Figure 2. The number of oocytes in each experimental group which had maintained a normal spindle structure after the osmotic treatment (values measured in milliosmolal, mOsm) and a 60 min culture period is shown. The isosmotic exposure marked with an asterisk contained sucrose as the primary solute, similar to the other anisosmotic treatments. There were six oocytes exposed to this treatment, with all of them maintaining a normal morphology. The proportion of spindles in each treatment that maintained a normal structure decreased as the solution osmolality diverged from isosmotic.

 
Figure 3 shows representative images of the various spindle morphologies seen in the treated oocytes. In some cases, as in panels A and B, the morphology of the spindle appeared normal. Note the tubulin and chromosome staining associated with the first polar body in panel B to the lower right of the MII spindle. In other cases, such as the spindles shown in panels C–E, significant structural disturbances were evident. The image in panel E shows metaphase II chromosomes where microtubule staining was absent. In our experience, this pattern occurs in oocytes which lost the integrity of the oolemma during the treatment. In one of the cells exposed to 150 mOsm, the microtubule pattern suggested that the cell was activated during the treatment and fixation occurred as the second polar body was being extruded (Inagaki et al., 1996Go). In the oocytes in which the spindle had a normal morphology, the chromosomes were aligned along the metaphase plate.



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Figure 3. Confocal microscopy images showing representative structures of MII spindles in oocytes exposed to different osmolalities. Examples of spindles with normal microtubule and chromosome organization are shown in (A) and (B). Note the tubulin and chromatin staining associated with the first polar body located to the lower right of the spindle in (B). Panels (CE) represent microtubule organization that was abnormal for a spindle. Note the reduced spindle diameter and slight chromosome scattering in (C). The spindle in (D) has tubulin that is more dispersed, lacking discrete spindle poles and chromosome scattering. The absence of tubulin staining in (E) is indicative of a cell that lysed during the treatment.

 
Logistic regression analysis
Osmolality was a significant predictor of loss of normal spindle structure for both the hypo- and hyperosmotic levels as assessed by the –2 Log Likelihood {chi}2 test (P < 0.001). For the hyposmotic range, each 1 milliosmolal decrease in solution concentration increases the odds of spindle disruption by a factor of 1.006 to 1.038, with 95% confidence. For the hyperosmotic range, each 1 milliosmolal increase in solution concentration increases the odds of spindle disruption by a factor of 1.001 to 1.004 with 95% confidence.


    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The current report documents the effects of volume changes associated with anisosmotic exposure on the structure of the MII spindle in human oocytes. Our analysis allowed a determination of the overall structure of the spindle as well as the maintenance of the chromosomes at the metaphase plate. In those spindles classified as normal, the chromosomes were located at the metaphase plate, suggesting the bonds between the microtubules and kinetochores, as well as the sister chromatids, remained intact. We have shown that an increasing proportion of spindles exhibited abnormal morphology as the cell volume diverged from isotonic resulting from exposure to anisosmotic solutions.

During the process of cryopreservation, cells are exposed to numerous non-physiological conditions. During the pre-freeze phase, the cells experience exposure to potentially toxic CPAs and cooling which can cause cold shock and disruption of cellular constituents. Thawing can also be detrimental with rehydration and the potential for the crystallization of previously vitrified cytoplasmic water (known as devitrification). The nature of these exposures is variable, depending on the approach one uses for cryopreservation. It has long been established that the presence of cryoprotective agents is necessary for cells to survive freezing (Polge et al., 1949Go). Exposing cells to CPAs causes transient volume changes due to osmotic pressure differences across the cell membrane, and these osmotically-induced volume changes can also have detrimental effects on cell viability. The hyperosmotic tolerance of human oocytes, as measured using a vital dye assay, was previously assessed (McWilliams et al., 1995Go). While this study showed that human oocytes can tolerate anisosmotic exposure in a similar range of concentrations used in the present report (~600–3000 mOsm), our analysis suggests that the tolerance is more limited. The vital dye assay, while able to determine membrane integrity and enzymatic function, would have failed to detect any changes to the structure of the MII spindle, as shown in the present report.

The present study used a design to isolate the effects of osmolality, where sucrose was the primary extracellular, non-permeating compound, to avoid the potential confounding effect of high ionic concentrations (Gao et al., 1993Go). We included an additional control to test for effects of the brief absence of ionic conditions usually present in culture media. It was determined that there was not an effect of the composition of the treatment solutions independent of osmotic effects with this additional control. It should be noted that the present treatments did not include cryoprotective agents, which may have a transient effect on spindle stability during cryopreservation (George et al., 1996Go).

The spindle in human oocytes has been shown to be very sensitive to differing aspects of cryopreservation. Sathananthan et al. (1988Go) reported on the consequences of slow cooling oocytes in the presence or absence of DMSO to 0°C and holding them at this temperature for 20 or 60 min. All of the oocytes from these treatments showed significant disruption of the meiotic spindle, while non-cooled oocytes displayed a normal microtubule pattern. Another report confirmed the temperature sensitivity of the spindle in human oocytes when the cells were cooled only to room temperature in the absence of CPAs (Pickering et al., 1990Go). In addition to highlighting the temperature sensitivity, this report demonstrated that the human MII spindle has a limited ability to undergo repair upon return to physiologic temperature. This property differs when compared with mouse oocytes (Pickering and Johnson, 1987Go). In more recent reports, the rate of human spindle disruption was assessed at different temperatures. Zenzes et al. (2001Go) documented this phenomenon at 0°C. They showed that within 2 min, noticeable spindle disruption had occurred. An additional 2 min caused complete loss of the microtubules. Wang et al. (2001Go) reported on the effects of cooling between 37 and 25°C on the spindle in live human oocytes using polarized light microscopy. They showed that spindle disassembly begins around 32°C. Although spindles that were disassembled for a brief period were able to recover upon return to 37°C, this was not the case when the spindles remained disassembled for 10 min. None of five oocytes held at 25°C recovered their spindle, and only two of five held at 28°C were capable of rebuilding their spindle. This is the first study to investigate the effects of osmotically-induced volume changes on the MII spindle in human oocytes. In our design, we allowed a 60 min recovery period prior to fixation to allow for spindle repair. Data from this collection of studies reinforce the need to develop methods capable of protecting the MII spindle from disassembly during cryopreservation.

Understanding the effect on human oocytes of all of the factors associated with cryopreservation is crucial to developing optimized procedures. The results from the present study can be used in a fundamental manner to optimize CPA addition and removal procedures for human MII oocytes, as has been done for human spermatozoa (Gilmore et al., 1997Go). Although spindle assessment was not undertaken, Fabbri et al. (2001Go) demonstrated the beneficial effect of increasing the sucrose concentration to 0.3 M in a freezing and thawing medium on the prevention of intracellular ice formation. Although this improvement is likely due to the reduction of intracellular water volume prior to freezing, increasing the sucrose concentration will impose greater osmotic stress and will increase the likelihood of causing damage to the spindle as shown by the present study. In an effort to reduce the osmotic stress on the oocyte during CPA addition and removal, the CPA addition was performed in two steps, and the CPA removal was performed in four steps. Such a procedure may have detrimental effects for reasons not associated with changes in osmotic pressure. The complexity of this procedure necessitates exposing the cells to suboptimal conditions for extended periods (~30 min for CPA removal was required for this procedure). Considering this was performed at room temperature, the extended cooling of these oocytes may have had deleterious effects on the spindle. In a more recent report (Fosas et al., 2003Go), the investigators used a similar approach for their cryopreservation procedure, but significantly reduced the amount of time for the CPA removal step, and found the outcome to be similar to the report by Fabbri et al. (2001Go). Minimizing the amount of time used to remove CPAs, thereby reducing the exposure to non-physiological conditions, is a logical approach to optimization of cryopreservation procedures. The results from the present study suggest that a single-step CPA removal could be accomplished without causing volume changes that would have deleterious effects on the spindle. Thus, having defined the osmotic tolerance limits for the human MII oocyte, one can engineer CPA addition and removal procedures for any cryopreservation procedure and minimize the exposure of the cells to suboptimal volume conditions.

With increasing numbers of human ART cycles being undertaken worldwide, developing improved cryopreservation procedures for human oocytes is essential. The results from the present study demonstrate that there is a relationship between anisosmotic exposure and maintenance of a morphologically normal MII spindle in human oocytes. Understanding this relationship will allow the development of improved methods for CPA addition and removal procedures during human oocyte cryopreservation.


    Acknowledgements
 
The authors thank the staff at the Atlanta Center for Reproductive Medicine for their assistance with oocyte collection and administrative support. Dr Heide Schatten is recognized for her helpful discussions on cell processing and microtubule biochemistry. Don Connor and Howard Wilson are recognized for their artistic abilities in figure preparation. This study was supported by the Gerald J. and Dorothy R. Friedman New York Foundation for Medical Research, the Gwinnett Women’s Pavillion, The Cryobiology Research Institute, and the University of Missouri at Columbia’s Department of Veterinary Pathobiology.


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 Results
 Discussion
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Submitted on January 12, 2004; accepted on February 13, 2004.