Developmental potential of murine germinal vesicle stage cumulus–oocyte complexes following exposure to dimethylsulphoxide or cryopreservation: loss of membrane integrity of cumulus cells after thawing

C.J. Ruppert-Lingham1,3, S.J. Paynter1, J. Godfrey1, B.J. Fuller1 and R.W. Shaw1,2

1 Department of Obstetrics and Gynaecology, University of Wales College of Medicine, Heath Park, Cardiff CF14 4XN, UK

2 Present address: Academic Division of Obstetrics and Gynaecology, Derby City General Hospital, Uttoxeter Road, Derby DE22 3NE, UK

3 To whom correspondence should be addressed. e-mail: Ruppert-LinghamCJ{at}cardiff.ac.uk


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
BACKGROUND: Cumulus cells of the cumulus–oocyte complex (COC) are important in oocyte maturation. Thus, in preserving immature oocytes it is prudent to also preserve their associated cumulus cells. The survival and function of oocytes and their associated cumulus cells was assessed following cryopreservation or exposure to cryoprotectant without freezing. METHODS: Immature COCs were collected from mice primed with pregnant mare’s serum. COCs were either slow-cooled or exposed to 1.5 mol/l dimethylsulphoxide without freezing. Treated and fresh COCs were stained for membrane integrity or, after in-vitro maturation and IVF, were assessed for developmental capability. Development of cumulus-denuded fresh oocytes, as well as denuded and frozen–thawed oocytes co-cultured with fresh cumulus cells, was assessed. RESULTS: Slow-cooled oocytes had significantly reduced coverage by intact cumulus cells compared with fresh COCs. Cumulus cell association and developmental capability were not substantially affected by exposure to cryoprotectant without freezing. Denuded fresh oocytes and cryopreserved COCs had decreased developmental potential that was not overcome by co-culture with fresh cumulus cells. CONCLUSIONS: Loss of association between oocyte and cumulus cells was induced by cryopreservation, but not by treatment with cryoprotectant alone. The data indicate that direct physical contact between cumulus cells and the oocyte, throughout maturation, improves subsequent embryo development.

Key words: cryopreservation/cumulus–oocyte complex/dimethylsulphoxide/GV-stage oocytes/maturation


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The cryopreservation of mature oocytes is problematic (Paynter, 2000Go), partly due to the presence of the temperature-sensitive microtubular spindle which is required for normal fertilization and embryo development. Germinal vesicle (GV)-stage oocytes have not yet formed this spindle. At this stage, the chromosomes are decondensed and enclosed within the nuclear membrane, potentially making them more resilient to cooling.

Regimes for the collection of fully-grown GV-stage human oocytes have been devised that involve modification of mature oocyte collection techniques (Trounson et al., 1994Go) and reduction in hormonal stimulation. However, at present only a limited number of successful pregnancies have arisen from immature human oocytes (Cha et al., 1991Go; Tucker et al., 1998Go; Wu et al., 2001Go). The clinical use of GV-stage oocytes to be matured for IVF may be preferable to traditional IVF treatment as the risk of ovarian hyperstimulation syndrome is reduced. The cost and duration of hormonal treatment could also be reduced. The development of a reliable method for long-term storage would allow these immature oocytes to be banked for future use. One major application would be the storage of oocytes from cancer patients prior to their receiving high-dose chemotherapy and total body irradiation, which can impair ovarian function. As well as allowing storage of oocytes for a patient’s own use, oocyte banking would simplify the process of oocyte donation by avoiding the need to synchronize donor and recipient cycles.

Spindle abnormalities have been reported following cryopreservation of GV-stage human oocytes (Park et al., 1997Go). However, other cryopreservation studies have emphasized the high levels of spindle and chromosome normality (~80%) observed following cryopreservation and in-vitro maturation (IVM) of immature human oocytes (Baka et al., 1995Go). Cryopreserved GV-stage human oocytes have been shown to be capable of completing nuclear maturation and becoming fertilized (Toth et al., 1994Go; Wu et al., 2001Go). However, the development of embryos resulting from cryopreserved immature human oocytes was impaired (Toth et al., 1994Go; Son et al., 1996Go; Wu et al., 2001Go). The proportion of immature human oocytes that reach maturity in vitro remains low, even among non-cryopreserved GV-stage human oocytes.

The refinement of IVM protocols for fully-grown murine GV-stage oocytes has resulted in oocytes with a similar level of developmental competence to that observed among in-vivo-matured oocytes when the donors were pre-treated with gonadotrophins (Schroeder and Eppig, 1984Go, 1989). High rates of nuclear maturation have been reported following cryopreservation and IVM of murine GV-stage oocytes using a variety of cryopreservation techniques (Schroeder et al., 1990Go; Van der Elst et al., 1993Go; Candy et al., 1994Go; Van Blerkom and Davis, 1994Go). In some cases, the rate of fertilization was similar for thawed and fresh control oocytes (Candy et al., 1994Go). However, there have also been reports of decreased rates of fertilization following cryopreservation (Schroeder et al., 1990Go; Van der Elst et al., 1993Go; Van Blerkom and Davis, 1994Go). In general, the developmental capacity of cryopreserved GV-stage murine oocytes is poor (Schroeder et al., 1990Go; Van der Elst et al., 1992Go, 1993; Candy et al., 1994Go; Van Blerkom and Davis, 1994Go).

It has been established that coupling of somatic cumulus granulosa cells with the GV-stage oocyte is vital to the progression of oocyte maturation and subsequent embryo development (Fagbohun and Downs, 1991Go). Studies have shown that GV-stage oocytes which are stripped of cumulus cells have a reduced developmental capacity compared with that of cumulus-enclosed GV-stage oocytes (Schroeder and Eppig, 1984Go). Cryopreservation has been reported to cause cumulus cell loss (Van der Elst et al., 1993Go; Cooper et al., 1998Go; Goud et al., 2000Go). The three-dimensional COC is likely to be particularly prone to physical disruption caused by ice crystal formation. Even when ice crystal formation is avoided in the process of vitrification, the vast difference in size between the oocyte and its associated cumulus cells means that they are likely to react very differently to the stresses applied during cryopreservation. The high levels of survival, morphological normality and maturation combined with the apparently low levels of developmental competence observed in freeze–thawed GV-stage oocytes could be an indication of damage and/or disruption to the somatic cumulus cells and their association with the oocyte.

The present study aimed to investigate the extent of damage inflicted on the cumulus cells of the COC by cryoprotectant loading/unloading either alone or with the additional stress of ice formation during slow-cooling and transfer to –196°C using dimethylsulphoxide (DMSO) as a cryoprotectant. The effect of this damage on the survival and subsequent development of the oocytes was examined. The effect of co-culture of fresh cumulus cells with fresh denuded GV-stage oocytes and frozen–thawed COCs was also investigated.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
For each experiment, a minimum of 12 ovaries was used and each experiment was repeated at least in quadruplicate.

Source of oocytes
Immature/GV-stage oocytes
Female CBAxC57 (6- to 8-week-old) mice bred from stock (obtained from Harlan, Bicester, UK) were kept under controlled conditions (14 h light, 10 h dark) and fed water and pellets ad libitum. The ovaries of the animals were stimulated by i.p. injection of 0.1 ml pregnant mare’s serum (PMS), 50 IU/ml (Folligon; Intervet UK Ltd, Milton Keynes, UK). After 46 h, the animals were killed by cervical dislocation and the ovaries removed immediately into modified phosphate-buffered saline (PBS) (Invitrogen, Paisley, UK) supplemented with 4 mg/ml bovine serum albumin (BSA) (Albumin fraction V powder; Sigma, Poole, Dorset, UK). The contents of the ovarian follicles were released by repeated puncturing with a 28 G micro-inject needle into 1 ml of standard maturation medium (SMM) which consisted of MEM Earles (Invitrogen, Paisley, UK) containing 10% fetal bovine serum (FBS) heat-inactivated (Invitrogen), 27.5 mg/l sodium pyruvate (Invitrogen), 50 mg/l streptomycin (streptomycin-sulphate BP; Evans, UK), 60 mg/l penicillin (crystapen benzyl penicillin sodium BP; Britannia, Redhill, Surrey, UK), 1 µg/l epidermal growth factor (Sigma), 1 mol/l L-glutamine (Invitrogen) and 0.1 mg/ml dcAMP (Sigma).

GV-stage oocytes, surrounded by at least two layers of cumulus cells, were selected and placed into 30 µl droplets of SMM (as detailed previously) containing 0.1 mg/ml dcAMP, using a pulled Pasteur pipette. COCs were held in this medium, under mineral oil (Sigma) at 37°C in an atmosphere of 5% CO2 in air until all COCs had been isolated from the ovaries (a maximum of 120 min).

In-vivo matured oocytes
Female CBAxC57 (6- to 8-week-old) mice were stimulated by i.p. injection of 0.1 ml PMS as detailed above, followed by 0.1 ml hCG (100 IU/ml) (Chorulon; Intervet UK Ltd) administered 53 h later. After a further 13 h, these animals were killed by cervical dislocation and the oviducts removed immediately into modified PBS supplemented with 4 mg/ml BSA.

Removal of cumulus cells
Freshly collected GV-stage COCs, held in 30 µl droplets of SMM + 10% FBS + 0.1 mg/ml dcAMP, were denuded by being drawn up and down a pulled Pasteur pipette with an internal diameter of 80–90 µm. Oocytes and cumulus cells were transferred into separate 30 µl droplets of SMM + 10% FBS + 0.1 mg/ml dcAMP. dcAMP was included in all media and cryopreservation solutions with the exception of the ‘hormone-supplemented’ maturation media. This was to ensure that the COCs remained at the GV stage throughout the manipulations. Morphologically normal oocytes were selected for IVM either in isolation or in the presence of pooled cumulus cells; that is, cumulus cells derived from twice the number of COCs to be included in the maturation group (to allow for cell loss during isolation). Freshly removed cumulus cells were also added, in similar proportions, to thawed COCs prior to entry into the IVM protocol.

In-vitro maturation (IVM)
Intact COCs, denuded oocytes, denuded oocytes with cumulus cells, thawed oocytes or thawed oocytes with cumulus cells were held in 30 µl droplets of SMM + 10% FBS + 0.1 mg/ml dcAMP at 37°C in an atmosphere of 5% CO2 in air. Each group was then placed into a 30 µl droplet of SMM + 10% FBS containing 0.75 IU/ml Gonal-F® (rFSH, Serono, London, UK), under mineral oil for 4 h at 37°C in an atmosphere of 5% CO2 in air. After this time, the medium was replaced, using a Gilson pipette, with 30 µl of SMM + 10% FBS containing 7.5 IU/ml Humegon® (FSH/LH at a 1:1 ratio; Organon, Cambridge, UK) at 37°C in an atmosphere of 5% CO2 in air for 18 h.

Cryopreservation of immature oocytes
Slow-freezing
Freshly collected COCs were transferred, by pulled Pasteur pipette, from 30 µl droplets of SMM supplemented with dcAMP into 1 ml of cryoprotectant solution containing 1.5 mol/l DMSO (Sigma) diluted in modified PBS +10% FBS + 0.1 mg/ml dcAMP. After 5 min of exposure to the solution at 0°C, the oocytes were pipetted into a small column (15–20 µl) of the same cryoprotectant solution within freezing straws, that had been partially filled with the diluent, 0.1 mol/l sucrose (Aristar; BDH, Poole, UK; sucrose was added to minimize the toxic effects of DMSO) made up in modified PBS +10% FBS + 0.1 mg/ml dcAMP. Straws had been prepared previously, and stored on ice. Approximately 20 COCs were placed into each straw.

The cryopreservation protocol used slow-cooling and warming rates. The straws were sealed with wet plastic plugs and placed on ice. When the COCs had been in the presence of DMSO at 0°C for a total of 15 min, the straws were cooled at –2°C/min to –6°C in a controlled rate freezer (Planer Kryo10, Series III) which had been precooled to 4°C. The straws were held at this temperature (–6°C) for 10 min, after which ice nucleation was instigated by touching each straw with cooled forceps. The straws were held at –6°C for a further 10 min. Cooling was then continued at a rate of 0.3°C/min to –60°C. When the samples reached a temperature of –60°C the straws were removed from the controlled-rate freezing machine and plunged into liquid nitrogen. The straws were stored at –196°C for between 1 and 12 weeks.

Thawing
The straws were removed from liquid nitrogen storage and placed in a controlled-rate freezing machine precooled to –70°C. They were then warmed to 4°C at a rate of 8°C/min. The contents of the straws were flushed with 1 ml of 0.1 mol/l sucrose made up in modified PBS +10% FBS + 0.1 mg/ml dcAMP into a culture dish. After being held in 0.1 mol/l sucrose solution at room temperature for 5 min, the COCs were placed into modified PBS + 10% FBS + 0.1 mg/ml dcAMP at room temperature for 5 min. They were finally placed into a fresh droplet of the same solution on a hot plate at 37°C for 5 min. After thawing, the oocytes were either stained for membrane integrity or were placed into the IVM protocol.

Exposure of COCs to cryoprotectant without freezing
COCs were transferred, by pulled Pasteur pipette, from SMM supplemented with dcAMP into 1 ml of cryoprotectant solution (1.5 mol/l DMSO +10% FBS + 0.1 mg/ml dcAMP) and held on ice for 15 min. After this time, the COCs were transferred into a dish containing 1 ml of 0.1 mol/l sucrose at room temperature. After 5 min the COCs were treated in a manner identical to thawed COCs. Following exposure to and dilution of the cryoprotectant, the COCs were either stained for membrane integrity or were placed into the IVM protocol.

Assessment of COCs and oocytes
Membrane integrity staining
Freshly collected GV-stage COCs, COCs which had been exposed to the cryoprotective agent without freezing, and thawed COCs were incubated in the dark (immediately after the 5 min incubation in modified PBS) at 37°C in modified PBS +10% FBS + 0.1 mg/ml dcAMP, containing 0.1 mg/ml carboxy fluorescein diacetate (Sigma) and 0.1 mg/ml propidium iodide (Sigma) for 10 min. The COCs were then washed twice in modified PBS +10% FBS + 0.1 mg/ml dcAMP. The COCs were then placed into a droplet of the same solution on a cavity slide and viewed using an Optiphot-2 microscope (Nikon, Tokyo, Japan) fitted with a filter capable of detecting fluorescence in the range 450–490 nm. Cells with an intact cell membrane fluoresce green, whereas those with a damaged cell membrane fluoresce red. Each COC was double-blind scored for membrane integrity of the cumulus cells using the following scoring system. COCs were scored according to the proportion of the oocyte surface covered by cumulus cells with intact membranes: score 1 = 76–100% coverage; score 2 = 51–75% coverage; score 3 = 26–50% coverage; and score 4 = 0–25% coverage.

IVF and assessment of development
In-vivo- or in-vitro-matured oocytes were released or placed (respectively) into 0.9 ml of Tyrode’s medium (Invitrogen) supplemented with 16 mg/ml BSA, and then incubated for 10 min at 37°C in an atmosphere of 5% CO2 in air. After this time, 0.1 ml of capacitated sperm (sperm incubated for 1–1.5 h in Tyrode’s medium supplemented with 16 mg/ml BSA at 37°C) was added to the oocytes, and the mixture was further incubated at 37°C in an atmosphere of 5% CO2 in air for 5 h. Oocytes were then transferred through three droplets of Tyrode’s medium supplemented with 4 mg/ml BSA under mineral oil and incubated in the final droplet for a further 15 h at 37°C, in an atmosphere of 5% CO2 in air. At this point, the cells were assessed for normality and progression to the 2-cell stage. The number of blastocysts and hatching blastocysts was counted 4 days later.

Statistical analysis
For membrane integrity data, comparisons were made between freshly collected COCs, COCs which had been exposed to cryoprotectant without freezing and cryopreserved COCs using the {chi}2-test. With data obtained following IVF, three data sets were compared using the Kruskal–Wallis test: either two treatment groups and in-vitro-matured controls for maturation experiments; or one treatment group (either exposure to cryoprotectant or cryopreservation) and in-vivo- and in-vitro-matured control groups. Where a significant difference was found using the Kruskal–Wallis test, pair-wise comparisons were made for this data set using the Mann–Whitney U-test.


    Results
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 Abstract
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 Materials and methods
 Results
 Discussion
 References
 
Exposure of GV-stage COCs to 1.5 mol/l DMSO without cooling
Normality was assessed following IVM and IVF. Exposure to DMSO did not cause a decrease in normality of the oocytes when compared with in-vitro- and in-vivo-matured control oocytes (Table I). Rates of fertilization were similar for COCs exposed to cryoprotectant and the in-vitro-matured control group. These rates were not lower than those observed in the in-vivo-matured control group but differences were not statistically significant. Exposure to DMSO did not cause a decrease in developmental potential when compared with in-vitro-matured control oocytes.


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Table I. Fertilization and development of GV-stage murine COCs exposed to 1.5 mol/l DMSO compared with in-vivo- and in-vitro-matured controls
 
Slow-cooling of GV-stage COCs
The normality of GV-stage COCs that were slow-cooled in the presence of 1.5 mol/l DMSO was significantly (P < 0.05) reduced compared with in-vitro- and in-vivo-matured controls (Table II). Oocytes that were assessed as normal became fertilized in similar proportions for all groups. Thawed COCs were found to have low developmental potential compared with controls when development was calculated from the number of fertilized oocytes or from the total number of oocytes in that group.


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Table II. Fertilization and development of freeze–thawed GV-stage murine COCs compared with in-vivo- and in-vitro-matured controls
 
Cumulus cell membrane integrity staining
Staining of fresh, treated and slow-cooled COCs is shown in Figure 1. The majority of the fresh COCs had a high proportion of cumulus cell membrane integrity. Following exposure to 1.5 mol/l DMSO, the percentage of the surface of the oocyte covered with cumulus cells that had intact membranes was found to be similar to that observed in fresh intact COCs (Figure 2). However, few of the cumulus cells of the frozen–thawed COCs retained an intact plasma membrane, and there was found to be significantly (P < 0.001) less direct contact between the surface of the oocyte and cumulus cells with intact membranes when compared with the fresh and DMSO treated COCs (Figure 2).



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Figure 1. Fluorescent micrographs depicting carboxy fluorescein/propidium iodide membrane integrity staining of: (A) freshly collected-untreated control COCs; (B) COCs exposed to 1.5 mol/l DMSO at 0°C for 15 min; and (C) COCs slow-cooled and thawed in 1.5 mol/l DMSO. Cells with an intact cell membrane fluoresce green whereas those with a damaged cell membrane fluoresce red.

 


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Figure 2. Distribution of scores assigned following membrane integrity staining of freshly collected COCs, cryoprotectant treated COCs and frozen–thawed COCs. Scoring system used was percentage of the oocyte surface in contact with cumulus cells with intact membranes: 100–76% = score 1; 75–51% = score 2; 50–26% = score 3; 25–0% = score 4.

 
IVM of denuded GV-stage oocytes
GV-stage COCs were stripped of cumulus cells and matured in vitro, either in the presence of disassociated cumulus cells or in isolation. Normality and fertilization were similar in all groups (Table III). When development was calculated from the total number of oocytes, both groups of denuded oocytes were found to have a low level of developmental competence, compared with intact controls. Co-culture with fresh cumulus cells during maturation did not significantly improve the developmental competence of denuded oocytes. The percentage of blastocysts formed from the original pool of oocytes was significantly greater (P < 0.05) for cumulus intact oocytes matured in vitro.


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Table III. Fertilization and development of denuded GV-stage murine oocytes matured in isolation, denuded GV-stage murine oocytes matured in the presence of fresh cumulus cells, and intact in-vitro-matured controls
 
IVM of freeze–thawed GV-stage COCs in the presence of fresh disassociated cumulus cells
GV-stage COCs were slow-cooled and thawed in the presence of 1.5 mol/l DMSO and then matured in vitro, either in the presence of fresh disassociated cumulus cells or in isolation. Normality was found to be similar in all groups (Table IV). Freeze–thawed oocytes were found to have a lower rate of fertilization and poorer development than in-vitro-matured oocytes (P < 0.05). Co-culture of freeze–thawed COCs with fresh cumulus cells during IVM did not improve any of the parameters compared with similarly treated non co-cultured COCs.


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Table IV. Fertilization and development of freeze–thawed GV-stage murine COCs matured in isolation, freeze–thawed GV-stage murine COCs matured in the presence of fresh cumulus cells, and intact in-vitro-matured controls
 

    Discussion
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
In the current study, denuding of oocytes prior to IVM led to a decrease in developmental potential. Loss of cumulus cells has previously been implicated in the poor development of embryos derived from denuded oocytes (Schroeder and Eppig, 1984Go). Association of the oocyte with somatic cumulus cells promotes competence to undergo fertilization and preimplantation development by supporting the completion of cytoplasmic maturation (Buccione et al., 1990Go). The mechanism by which this benefit is realised is unclear. It has been suggested that a paracrine factor secreted by the cumulus cells is, in part, responsible for the maturation of the oocyte (Byskov et al., 1995Go). Disassociated cumulus cells appear to produce a factor that acts in a positive capacity on denuded oocytes to stimulate meiotic resumption (Downs, 2001Go). Although the presence of a positive paracrine factor has been indicated, evidence fails to support this as a primary mechanism for hormone-induced maturation in isolated mouse oocytes (Downs, 2001Go). In the current study, denuded oocytes that were matured in vitro in the presence of freshly removed cumulus cells showed no increase in fertilization or developmental capacity compared with the denuded oocytes that were matured in vitro in isolation. This does not provide support for the existence of a secreted maturation factor, despite the fact that the concentration of any paracrine factor would be expected to be higher in the present study, where oocytes were cultured in the presence of excess cumulus cells and where culture was performed in a lesser volume (Downs, 2001Go). The failure in the completion of cytoplasmic maturation, as evidenced by a lack of improvement in fertilization and development in the presence of dissociated cumulus cells, emphasizes the importance of the conservation of the original association between the GV-stage oocyte and the cumulus cells during maturation in vitro. A requirement for physical contact with the somatic cells would suggest that intercellular connections such as gap junctions are important at this stage. This therefore supports the view that paracrine factors are overridden by signals transmitted by gap junctions (Downs, 2001Go).

The current study aimed to determine at what stage of the cryopreservation process damage occurred, and whether damage was inflicted on the oocyte or the cumulus cells, or both. Following membrane integrity staining, the scores assigned to fresh control COCs and to COCs exposed to DMSO were similar. Thus, no substantial disassociation of the oocyte and cumulus cells occurred as a result of exposure to 1.5 mol/l DMSO for 15 min at 0°C. Exposure to DMSO also had no effect on the morphological normality of the oocytes or on the developmental capacity of the oocytes compared with fresh control COCs. One group (Schroeder et al., 1990Go) found a decrease in morphological normality of GV-stage oocytes following exposure to DMSO at 0°C for 30 min. The lower rates of normality can be attributed to the longer duration of exposure that, due to the kinetics of chilling injury, could lead to more extensive damage. This has been demonstrated in GV-stage bovine oocytes (Zeron et al., 1999Go), while others (Van der Elst et al., 1992Go) reported significantly fewer oocytes with normal spindle morphology following exposure to DMSO at 0°C at the GV stage than in control groups. However, no chromosomal abnormalities were reported, and in all cases the chromosomes were located at the equatorial plane of the spindle (Van der Elst et al., 1992Go).

Following cryopreservation, normality of oocytes was found to be highly variable. This parameter was assessed at 47 h post-treatment and, therefore reflects the ability of the oocyte to survive the freeze–thaw process, to achieve maturation to metaphase II and to survive in culture for this period. In contrast, high survival at 1–5 h post-thaw has been reported (Schroeder et al., 1990Go; Van der Elst et al., 1992Go). However, fertilization was significantly reduced compared with non-frozen controls. In another study, 93% survival at 16 h post-thaw with 83% maturation in vitro and 70% fertilization were reported (Candy et al., 1994Go). In the current study, oocytes that were morphologically normal following cryopreservation were fertilized in similar proportions to controls.

Membrane integrity staining revealed extensive loss of plasma membrane integrity of the cumulus cells of thawed COCs. Thawed COCs therefore had significantly less direct contact with intact cumulus cells compared with fresh COCs and COCs exposed to DMSO without freezing. Following cryopreservation, embryo development was poor. Developmental impairment as a result of cryopreservation has been demonstrated following post-fertilization culture both in vitro (Van der Elst et al., 1993Go) and in vivo (Candy et al., 1994Go). In the current study, it was observed that the blastocysts derived from fresh COCs were larger than those derived following slow cooling. A similar delay in development has been reported following vitrification of GV-stage COCs (Van Blerkom and Davis, 1994Go).

Some studies have reported a loss of cumulus cells from the COC following cryopreservation and thawing (Cooper et al., 1998Go; Goud et al., 2000Go). In the present study, careful pipetting allowed retention of the majority of cumulus cells and assessment of the membrane integrity of COCs immediately after thawing. However, most of the cumulus cells of the thawed COCs became disassociated from the oocyte following a short period of culture. The reduced developmental capacity of the thawed oocytes could therefore be due to a loss of integrity and/or functionality of cumulus cells.

The addition of fresh cumulus cells to thawed COCs did not improve maturation or development of the oocytes. Therefore, as with denuded oocytes, no benefit of co-culture with fresh cumulus cells during IVM was established. This is evidence against the significance of a secretory maturational factor that is lacking in thawed COCs, but supports the assertion that the metabolic coupling which exists between the cumulus cells and the oocyte has a maturational role that is disrupted by the process of slow-cooling and thawing. This evidence also supports the assertion that direct physical contact between the oocyte and cumulus cells, possibly via intercellular junctions, is a requirement for the completion of cytoplasmic maturation and subsequent embryo development.

In conclusion, at some stage during the process of cryopreservation a loss of association between the oocyte and cumulus cells occurred, and the integrity of cumulus cells was compromised. Disassociation of these two cell types prior to the completion of maturation led to a decrease in the developmental capacity of the oocyte. This developmental deficit was not overcome by co-culture with fresh disassociated cumulus cells. The stages in the cryopreservation profile at which this damage occurs need to be identified so that cryopreservation protocols can be designed which avoid or minimize such disruption. This could be achieved for example, by changing the rates of cooling/warming and employing alternative methods of cryopreservation such as vitrification (Hong et al., 1999Go).


    Acknowledgement
 
The authors thank Dr Frank Dunstan, University of Wales College of Medicine for his assistance with the statistical analysis.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
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Submitted on August 1, 2002; accepted on October 22, 2002.