Oocyte maturation, follicle rupture and luteinization in human cryopreserved ovarian tissue following xenografting

Debra A. Gook1,2,4, D.H. Edgar1,2, J. Borg1,2, J. Archer1,2, P.J. Lutjen3 and J.C. McBain1,2

1 Reproductive Services, Royal Women’s Hospital, Carlton, Victoria, 2 Melbourne IVF, East Melbourne, Victoria and 3 Department of Obstetrics and Gynaecology, Sandringham Hospital, Sandringham, Australia

4 To whom correspondence should be addressed at: Reproductive Services, Royal Women’s Hospital, 132 Grattan Street, Carlton, Victoria 3053, Australia. e-mail: debra.gook{at}rwh.org.au


    Abstract
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 Abstract
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 Materials and methods
 Results
 Discussion
 References
 
BACKGROUND: Previous studies have demonstrated development of antral follicles in cryopreserved human ovarian tissue after autografting and xenografting, thus indicating successful preservation of follicular function. The study aim was to assess whether these follicles could also undergo periovulatory changes in response to hCG. METHODS: Ovarian tissue from three patients were dehydrated in propanediol (PROH)/sucrose and cryopreserved using the slow cooling/rapid thaw procedure. Thawed tissue was placed under the kidney capsule in immunodeficient mice. Following growth (>20 weeks) in the presence of gonadotrophin, hCG was administered and ovarian tissue examined histologically. RESULTS: Thirty-two antral follicles (diameter range 0.6 to 5 mm) were examined. Histological evidence of a response to hCG was evident in all follicles. Disruption of the concentric layers of mural granulosa and theca cells was apparent in all antral cavities. In 17 (53%) follicles the exterior follicular wall had reduced to a few cells thick, and in eight (25%) the wall had ruptured. Mucified oocyte–cumulus cell complexes were present in 32 follicles, 17 of which had begun to detach from the pedicle. Resumption of meiosis had occurred in over half the oocytes (five metaphase II and seven metaphase I oocytes, eight germinal vesicle breakdown). Two corpora lutea were also detected. CONCLUSIONS: Follicles cryopreserved within human ovarian tissue using the PROH procedure, can develop to the antral stage and undergo periovulatory changes following xenografting and exposure to a luteinizing stimulus.

Key words: cryopreservation/human/luteinized follicle/mature oocyte/xenotransplantation


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 Abstract
 Introduction
 Materials and methods
 Results
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Sterility and/or premature menopause are significant complications associated with the aggressive cytotoxic treatments commonly used to treat a variety of diseases (Arnon et al., 2001Go; Meirow and Nugent, 2001Go). The concept of circumventing these problems by cryopreserving female gametes remains an attractive possibility (Gook and Edgar, 1999Go; Oktay, 2001Go; Gosden et al., 2002Go) in the absence of realistic alternatives other than possible transposition of the ovary in cases where it may be protected from radiotherapy (Morice et al., 2000Go). In cases where cytotoxic therapy is less urgent, ovarian stimulation and mature oocyte cryopreservation—a clinical approach which has resulted in numerous live births (Porcu et al., 1997Go; 1998; 2001; Antinori et al., 1998Go; Borini et al., 1998Go; Polak de Fried et al., 1998Go; Winslow et al., 2001Go)—would seem to offer the most realistic hope for these young women. In the face of imminent therapy, however, many clinics have turned to cryopreservation of ovarian tissue (Bahadur and Steele, 1996Go; Meirow, 2000Go; Oktay and Karlikaya, 2000Go; Gelety et al., 2001Go; Poirot et al., 2002Go). Although this approach is now well established, the reality remains that little is known of the future potential of much of this tissue.

Cryopreservation of animal ovarian tissue was initially reported during the 1950s (Deanesly, 1954Go; Green et al., 1956Go; Parkes, 1958Go; Parrott, 1960Go), and live births were subsequently documented following the autografting of such tissue (Parrott, 1960Go; Gosden et al., 1994aGo; Gunasena et al., 1997Go; Sztein et al., 1998Go; Candy et al., 2000Go; Shaw et al., 2000Go). In particular, the birth of a lamb (Gosden et al., 1994aGo) and resumption of cycling for a period of 2 years in ewes (Baird et al., 1999Go) following grafting of cryopreserved ovine ovarian tissue suggested that similar success may be possible in the human given the similarity in follicle distribution and density of stromal tissue. Initial attempts to freeze human ovarian strips using the same procedure as that used in the sheep studies (Gosden et al., 1994aGo) indicated that some primordial follicles had survived the cryopreservation (Hovatta et al., 1996Go; Newton et al., 1996Go; Oktay et al., 1997Go). Further studies with human tissue have demonstrated initiation of mitosis in some primary follicles following xenografting (Oktay et al., 2000Go). However, the reduction in follicle numbers, high rate of fibrosis in the tissue (Kim et al., 2000Go; Nisolle et al., 2000Go) and histological evidence of follicular damage associated with manipulation of the cryopreservation procedure (Gook et al., 1999Go), highlight some fundamental concerns associated with the cryopreservation of human ovarian tissue. Until recently, growth of follicles following xenografting had only been established in non-frozen human ovarian tissue (Oktay et al., 1998Go; Weissman et al., 1999Go). More recent reports of growth to the antral stage within cryopreserved ovarian tissue following xenografting (Gook et al., 2001Go; Van den Broecke et al., 2001aGo; Kim et al., 2002Go) are encouraging, and indicate preservation of developmental potential in the follicles. Reports of albeit transient resumption of cycling in two patients following auto-transplantation of cryopreserved ovarian tissue (Oktay and Karlikaya, 2000Go; Radford et al., 2001Go) further support the potential of the technology. In contrast to these encouraging results, there is a lack of fundamental evidence that the primordial follicles cryopreserved within human ovarian tissue can fulfil their full developmental potential and progress to ovulation of mature oocytes.

The aim of the present study was to examine whether periovulatory changes could be induced by administration of hCG in human antral follicles which had developed within xenografted cryopreserved ovarian tissue.


    Materials and methods
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 Introduction
 Materials and methods
 Results
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Ovarian tissue was donated from three women who were at risk of loss of fertility: a 20-year-old diagnosed with osteosarcoma; an 18-year-old with acute myeloid leukaemia; and an 18-year-old with a paraspinal tumour. This study was approved by the Hospital Research and Ethics Committee. Collection, preparation and cryopreservation procedures for ovarian tissue have been described previously (Gook et al., 2001Go). Briefly, thin slices of ovarian cortex (~4x2x1 mm) were dehydrated for 90 min in phosphate-buffered saline containing 1.5 mol/l propanediol (PROH; BDH, Kilsyth, Victoria, Australia), 0.1 mol/l sucrose (BDH) and 10 mg/ml human serum albumin (HSA; Albumex; CSL, Camperfield, Victoria, Australia) at room temperature. Slices were loaded into cryo-vials containing 1 ml dehydration solution and frozen using a slow rate of freezing with manual seeding (Gook et al., 1999Go). The tissue was finally stored in liquid nitrogen vapour for a minimum of 6 months before thawing using a rapid procedure described previously (Gook et al., 1999Go).

Cryopreserved ovarian tissue was xenografted into 12 mice, using methods reported previously (Gook et al., 2001Go). Following thawing, slices were transferred to Hams F12 medium (Trace Scientific Ltd, Noble Park, Australia) with 10 mg HSA/ml, and cut into smaller pieces (~0.5x0.5x1 mm). Female, 16-week-old severely compromised immunodeficient (SCID) mice were anaesthetized with an i.p. injection of 2,2,2-tribromoethanol (0.4 mg/g body weight; Avertin; Sigma-Aldrich). A single piece of ovarian cortex was inserted under the capsule of each kidney, and the mice were oophorectomized bilaterally. An i.p. injection of antibiotic (4 µg/g body weight; Gentamycin; Pharmacia-Upjohn) was given while the animal was anaesthetized. At 7 days after surgery, i.p. injections of gonadotrophin (1 IU recombinant FSH; Gonal F; a gift from Serono, Australia, and Puregon; a gift from Organon, Australia) were commenced and continued every second day until completion of the study. At >=27 weeks following grafting, mice received an i.p. injection (20 IU) of hCG (Profasi; Serono). The kidneys were removed at 30–36 h after hCG administration, examined initially for gross morphological changes in the appearance of antral follicles, and subsequently fixed in 4% paraformaldehyde (ProSciTech, Thuringowa, Queensland, Australia) for histology. Follicular diameters were measured using an ocular micrometer both prior to fixation and at maximum diameter on histological section. Fixed tissue was processed followed by embedding in paraffin wax, and 3 µm serial sections were cut and stained with haematoxylin and eosin. Due to the possibility of folds occurring within the follicle wall, only cavities containing an identifiable oocyte were classified as antral follicles.


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 Materials and methods
 Results
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 References
 
Initial observations, made under the dissecting microscope, identified at least one antral follicle (ranging in diameter from 0.6 to 5mm) in all mice. In contrast to a previous study (Gook et al., 2001Go) in which antral follicles were not exposed to hCG, most antral cavities in the present study were discoloured and occasionally bloody in appearance. In five mice, antral follicles were observed on only one site within the animal. Histological sectioning revealed the presence of multiple antral cavities on some xenograft sites. A summary of the characteristics of individual follicle histology is given in Table I.


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Table I. Individual follicle characteristics following administration of hCG
 
Granulosa and theca cell layers
Following hCG administration, clear signs of disorganization and degradation were observed in the layers of mural granulosa cells. The cellular arrangement was more random, suggesting loss of cell–cell interactions, and varied in thickness throughout the follicle. Degradation of the mural granulosa cells manifested as two common forms: (i) a separation between the theca layer and the granulosa or stromal layer (peeling apart; Figure 1A); and (ii) swelling and lysis of cells in the surface layer and sloughing into the cavity, followed by degradation of subsequent layers (Figure 1A and B; arrow). Large droplets of follicular fluid, which stain relatively heavily with eosin, were often associated with the granulosa cells. All follicles examined showed degradation of the granulosa cell layers, with both forms often occurring in the same follicle. Although theca layers were apparent in some parts of the follicular wall, disorganization was also evident, with a lack of definition between granulosa and theca layers (Figure 1B). Generally, the exterior follicle wall was reduced to only a few cells in thickness (Figure 1C). Examination of serial sections demonstrated that this phenomenon was not artefactual, being consistently observed in a number of follicles as preferential degradation of the follicle wall (Figure 1D, E and F; Figure 1G, H and I). The follicle wall in these regions showed gaps between the putative cell layers, which often consisted merely of cell remnants with an occasional nucleus (Figure 1D, E and G). These features were consistent with the point of follicle rupture. These regions of the follicle wall were usually observed diametrically opposed to the pedicle (Figure 1C; arrow). Complete breakage of the follicle wall (represented as + in Table I) was observed in some of the large follicles (Figure 1G, H and I), and in many of the other follicles the wall was reduced to remnants with gaps but had not completely broken (~ + in Table I). In other follicles the wall was deteriorating but there was no dramatic preferential thinning (– in Table I).



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Figure 1. Morphology of the follicular wall following hCG administration. (A) Separation of the follicle wall from the stromal cell layers (S). Individual mural granulosa cells can be seen sloughing into the cavity. Within the cavity is a clump of swollen mural granulosa cells (arrow), some of which no longer contain a nucleus. Original magnification, x5. (B) The initiation of loss of contact between the theca cell layer (T) and stromal tissue. The follicle wall on the right-hand side still consists of theca and mural granulosa cell layers, whereas on the other wall (#) there has been a loss of cellular demarcation. Clumps of degrading mural granulosa cells (arrow) are present in the cavity. Original magnification, x5. (C) The exterior follicle wall is extremely thin. This thinning has occurred in the wall adjacent to the pedicle (arrow). Original magnification, x5; scale bar = 100 µm. (D,E,F) Consecutive sections through the same region of the exterior follicular wall, showing progressive deterioration of the wall which has almost ruptured (classified in Table I as ~ +). Gaps form between cells (D), a single cell or cell remnants on both the inner and outer side of the wall (E) maintain the follicular wall. Finally, the outer connections rupture, resulting in the follicular wall being held together by one or two cells (F). Note the densely stained follicular fluid engulfing the inner layer of granulosa cells and droplets in the cavity. Original magnification, x20. (G,H,I) Consecutive sections through the same region of another exterior follicular wall which has ruptured (classified in Table I as +). Gaps form between the cells (G) and on the outer side only a thin piece of cell remnant maintains the connection. This connection breaks (H) and the follicular wall has completely ruptured (I). Original magnifications: G, x20; H and I, x10.

 
Antral cavities
The antral cavities contained densely stained follicular fluid, swollen granulosa cells, cell remnants and, occasionally, a few luteinized cells (Figure 2A). The cell remnants had not been completely destroyed in some follicles and remained as a fibrous mesh. Variable amounts of blood were also present in the cavities of some follicles; estimates of the relative blood content are detailed in Table I. Erythrocytes were the main blood component and were used to estimate the amount of blood, although macrophages and polymorphs were also occasionally present. Erythrocytes within the stromal tissue were seen streaming through the tissue and the follicle wall into the cavities of the follicles which contained more significant amounts of blood (++ and +++ in Table I). The distribution of blood was not random within the cavity but appeared to be concentrated in a certain region of the cavity (Figure 2B). Irrespective of the amount of blood within the cavity, there was a close association between the blood and what remained of the granulosa and theca cells at a specific region of the follicle wall. This was often on the exterior side of the wall. In follicles with large amounts of blood (++ and +++), blood had infiltrated the exterior wall forming a blood-filled pocket (Figure 2C). At this point in subsequent sections, the wall had finally ruptured (Figure 2D).



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Figure 2. Morphology of the antral cavity following hCG administration. (A) Luteinized cells (L) within the cavity. Only a few cell nuclei remain in the follicle wall, mainly on the inner side (arrow) of the exterior wall. Original magnification, x20. (B) A follicle abutting the renal (r) surface. Within the follicular cavity is an oocyte (O) and a large amount (Table I, ++) of blood (b) which has aggregated in a specific area of the cavity. In contrast to the follicular wall in other areas, the follicular wall adjacent to this bloody area (arrow) has a different structure; it contains less cellular material and is more fibrous. Original magnification, x5; scale bar = 100 µm. (C,D) Consecutive sections of the follicular wall at the position of the arrow in (B), showing blood infiltration and formation of a pocket in the follicular wall (C), resulting in the loss of cell–cell connections in the wall, and almost complete rupture of the follicular wall (D). Again, the outer wall is only held together by a thin amount of cell remnants. Original magnification, x20.

 
Oocyte–cumulus complexes
In follicular cavities, part of the granulosa cell layer had differentiated forming a pedicle of cumulus cells which encapsulated the oocyte. An oocyte was detected in the cavity of all follicles, except the putative corpus luteum. Pedicle formation was commonly observed on the interior (renal side) of the cavity wall. A rough correlation was observed between cavity size and expanse of the pedicle. However, irrespective of the cavity size, the cumulus cells of the pedicle were interspersed by an amorphous matrix (Figure 3A). This mucification of the cumulus cells extended throughout the pedicle separating the cumulus cells and, in some of the larger follicles, had completely dissociated the oocyte–cumulus complex from the pedicle (Figure 3B and + in Table I). Partial separation (~ +) and no obvious evidence of detachment (–) from the pedicle were also observed, the latter occurring in the smaller follicles. Erythrocytes were occasionally observed in association with the matrix (Figure 3A; arrow). Matrix was also observed interspersed between the corona cells which abutted the zona pellucida of the oocyte. No cellular processes extending from the corona cells down through the zona pellucida were evident in any of the oocyte cumulus complexes. With one exception (* in Table I), in which the oocyte was completely naked, all other oocytes were encapsulated by corona cells and matrix which were in turn surrounded by mucified cumulus cells.



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Figure 3. Stages of oocyte nuclear maturation observed following hCG administration. (A) A mucified cumulus complex showing the cumulus cells interspersed with an amphorous matrix (m). Disorganization in the cellular arrangement of both the cumulus and corona cells is apparent, and there are no corona cell processes traversing the zona pellucida. Erythrocytes (arrow) are occasionally observed within the cumulus complex. The absence of a germinal vesicle, remnants of the germinal vesicle or a polar body suggest the oocyte is at metaphase I. Original magnification, x20; scale bar = 100 µm. (B, C) Two sections through the same metaphase II oocyte–cumulus complex. Polar body chromatin (arrow) can be seen (B) and the alignment of the metaphase chromosomes (C; arrow). The oocyte–cumulus complex has broken away from the pedicle (referred to in Table I as +). Original magnification: B and C, x20. (D) Germinal vesicle breakdown (GVBD) has commenced (arrow), the membrane is absent, but the chromatin is still in a circular arrangement with an irregular-shaped nucleolus (left) and other more condensed areas (right). Separation between the oocyte–cumulus complex and the pedicle has commenced (referred to in Table I, detached pedicle as ~ +). Original magnification, x20. (E) Although staining has revealed a circular area, the germinal vesicle membrane and nucleolus are both absent, suggesting more advanced GVBD. Around the edge of the circular region are condensed clumps of chromatin (arrow). Original magnification, x10. (F) In this oocyte no evidence of a polar body or germinal vesicle could be detected. The oocyte chromatin (arrow) was in a linear arrangement (metaphase I). Original magnification, x20. (G) This oocyte was similar to the previous, in which GVBD had been completed, but in this case the chromatin was in two separated linear alignments (arrows; late telophase I). Original magnification, x40.

 
Oocytes
All stages of nuclear maturation were observed within the oocytes and are detailed in Table I. Polar bodies within the perivitelline space (Figure 3B) and condensed chromatin in a linear formation indicating metaphase chromosomes (Figure 3C) were observed in five oocytes. Figure 3B and C are from two sections through the same metaphase II (MII) oocyte. These MII oocytes were detected in large follicles (>=2.7 mm) and had a mean diameter (including the zona) of 116 µm. In contrast, in the smaller follicles (<1.5 mm), the oocyte chromatin was identified at the germinal vesicle stage. In these oocytes, the chromatin was completely decondensed apart from the nucleolus, and enclosed by a distinct membrane. Germinal vesicle breakdown (GVBD) was also observed in some oocytes. This was characterized by the absence of a membrane enclosing the germinal vesicle chromatin. The nucleolus was generally larger, slightly misshapen and less condensed, and the chromatin had started to condense within the circular region (Figure 3D). In two oocytes (Table I; MIa), GVBD was complete but there was no evidence of a nucleolus and the chromatin had completely condensed (Figure 3E). In seven oocytes there was no evidence of GVBD and/or MII, indicating that these oocytes were at metaphase I stage (Figure 3A). Aligned chromatin was observed in one of these oocytes (Figure 3F). In another oocyte, the well-condensed chromatin was separated into two populations, one close to the oolemma in a clump and the other towards the centre of the oocyte, in a linear formation (late telophase I; Figure 3G; Donahue, 1970Go).

Corpora lutea
On one site (14), a follicle was detected which was morphologically different from all other follicles, and was classified as a recent corpus luteum. The entire cavity was filled with blood (Figure 4A) which precluded the identification of any oocyte. The cavity was surrounded incompletely by a fibrous wall that was thick and contained very few cells, some of which were luteinized (Figure 4B; L). Luteinized cells were also present in the stromal tissue adjacent to the wall. The fibrous wall was completely absent in one region, in which the cavity contents merged with the stromal tissue. Higher resolution examination of the cavity contents (Figure 4B) revealed erythrocytes, polymorphs and macrophages in association with large luteinized cells (some of which contained yellow pigmentation) abutting the wall. Heavily eosin-stained granules were present throughout the remainder of the cavity and also within the large luteinized cells.



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Figure 4. Corpora lutea. (A) The cavity is engorged with blood. The wall (arrow) which is almost entirely fibrous (seen at higher magnification in B) varies in thickness and is absent in some regions (bottom left, adjacent to bar). Original magnification, x5; scale bar = 100 µm. (B) Higher magnification showing the thick almost acellular wall (arrow) and the cellular composition of the cavity [erythrocytes, polymorphs; (p) and large luteinized cells with yellow pigmentation; (L)]. Original magnification, x20. (C) A montage of the ovarian tissue on site 3 showing two antral cavities; one in the top panel and the other on the bottom edge. Traversing the tissue is a corpus luteum (arrow) which has no defined border and consists of large and small heavily luteinized cells. Original magnification, x5.

 
An advanced corpus luteum (Figure 4C), which spanned the width of the tissue, was observed on site 3 between two antral follicles. It had no boundary and consisted of large and small heavily luteinized cells, which frequently contained yellow pigmentation.

Summary of observed periovulatory changes
All the characteristics reported were more prevalent within those follicles (n = 15) with a larger diameter (>=2 mm). Thinning of the exterior follicular wall occurred in over half (17/32) of all follicles, and eight follicles (all of which had a diameter >=2 mm) had completely ruptured. Blood was observed within nine follicles, seven of which were >=2 mm in diameter. Although a mucified cumulus oocytes complex was observed in all 32 follicles, detachment of the complex from the pedicle rarely occurred in the smaller (<2 mm) follicles. In almost all (14/15) of the larger follicles, partial and complete detachment of the complex from the pedicle was observed. Resumption of meiosis had occurred in over half (20/32) of all oocytes. Five were at MII, seven at MI, and GVBD had commenced in eight. Two corpora lutea were also detected.


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The cryopreservation procedure used in this study has been shown previously to preserve the integrity of human primordial and primary follicles (Gook et al., 2000Go) which can develop subsequently into morphologically normal antral follicles following xenografting (Gook et al., 2001Go). The appearance of healthy stromal tissue containing developing follicles and the abundant circulation to antral follicles following xenografting in the present study confirmed these findings. The previous observations of antral follicles in xenografted cryopreserved tissue which had not been exposed to hCG (Gook et al., 2001Go) revealed preovulatory morphological characteristics; that is, antral cavities encapsulated by numerous concentric layers of mural granulosa cells surrounded by theca cell layers. Plump cells and cell–cell interactions were obvious within these cell populations which were segregated into distinct layers. Germinal vesicle oocytes were present in compact cumulus cell pedicles, and processes from the corona cells traversed the zona pellucida. The observations reported in the present study demonstrate a marked contrast in morphology following exposure to hCG; indeed, the morphological consequences of exposure to hCG were restricted to antral follicles and observed in numerous antral cavities.

The morphology observed in the present study was consistent with generally accepted periovulatory changes in mammalian ovarian follicles. Difficulties associated with obtaining periovulatory and ovulatory samples have limited reported observations on human tissue. However, in the rodent and rabbit (Burr and Davies, 1951Go; Blandau, 1967Go; Byskov, 1969Go; Motta et al., 1971Go; Bjersing and Cajander, 1974Go) similar observations to those presented in the present study—that is, the destruction of the granulosa and theca cell layers, mainly in the apical region—have been reported. At ~1 h before ovulation in the rabbit, evidence was observed (Blandau, 1967Go) of separation and vacuolation of the granulosa and theca cells at the stigma site. Progressive deterioration was evident, resulting in only a few strands of connective tissue remaining just prior to ovulation. The wall finally ruptured, releasing the mucified cumulus–oocyte complex, which had dislodged from the pedicle. As in the present study, blood has been observed within the cavity (Burr and Davies, 1951Go) or associated with the escaping cumulus oocyte complex (Blandau, 1967Go; Motta et al., 1995Go). The withdrawal of the corona cell processes, mucification of the cumulus–corona cell complex and nuclear maturation of the oocyte seen in the present study are also consistent with ovulated oocytes (Motta et al., 1995Go) and routine observations following follicle aspiration for clinical assisted reproduction (Veeck, 1991Go).

Evaluation of the pre-existing follicle population in cryopreserved human ovarian tissue (Gook et al., 1999Go) and the observed time requirement for antral follicle development following xenografting (Gook et al., 2001Go) indicate that these antral follicles developed to the periovulatory stage, following xenografting, from primordial follicles which were present in the cryopreserved tissue. The full range of ovulatory changes in multiple follicles reported in the present study demonstrates that full functional potential has been preserved—an important prerequisite for clinical application of this technology.

Histological evidence of luteinization in human cryopreserved ovarian tissue following xenografting (Kim et al., 2002Go) has also been observed when dimethylsulphoxide (DMSO) was used as the cryoprotectant. The DMSO cryopreservation regimen, which is frequently used for animal and human ovarian tissue, was initially reported for use with sheep ovarian tissue (Gosden et al., 1994aGo), and numerous live births have been achieved from autografting of ovarian tissue cryopreserved using this DMSO regimen in animals. However, data relating to the extended development of follicles within human ovarian tissue cryopreserved with DMSO are limited, thereby precluding a conclusive comparison of the relative effectiveness of the two cryoprotectants. Recent evidence of temporary resumption of menses in a single patient following autologous transplantation of pieces of a whole ovary, which had been cryopreserved using the DMSO procedure (Radford et al., 2001Go), and the presence of corpora lutea in a subsequent study (Kim et al., 2002Go) in which a modification of this procedure was used, suggest that this method can preserve follicular function within the human ovarian cortex. However, in contrast to the minimal follicle loss (7%) observed as a consequence of cryopreservation damage in sheep (Baird et al., 1999Go), a significant loss of cellular structure has been reported following DMSO cryopreservation and xenografting of human ovarian tissue (Kim et al., 2000Go; 2002; Nisolle et al., 2000Go) and has been estimated to result in a 50% reduction in the number of primordial and primary follicles compared with non-frozen tissue (Nisolle et al., 2000Go). Degenerative changes in oocytes and granulosa cells have also been observed (Nisolle et al., 2000Go; Van den Broecke et al., 2001bGo) in human ovarian tissue cryopreserved using DMSO. The follicular development and luteinization observed in the present study, together with the temporary return to menses in a patient receiving ovarian tissue cryopreserved using the PROH procedure (Oktay and Karlikaya, 2000Go), indicate that this approach, which has previously been optimized for human ovarian cortex (Gook et al., 1999Go; 2000), is consistent with survival and subsequent functional competence. The total number of antral follicles (fewer than 10) previously reported to have developed within human cryopreserved ovarian tissue following autografting or xenografting remains limited (Oktay and Karlikaya, 2000Go; Radford et al., 2001Go; Van den Broecke et al., 2001aGo; Kim et al., 2002Go). The number of follicles which developed in the present study (n = 32) and in a previous study (n = 6; Gook et al., 2001Go) indicate that preservation of cellular function within follicles is highly reproducible using this cryopreservation procedure.

Interestingly, although the follicles which developed in the present study were approximately one-fifth of the diameter of periovulatory human follicles, they contained mature oocytes of normal size and exhibited the full range of morphological characteristics associated with ovulation. Similar or smaller follicle sizes to those in the present study have been observed following xenografting of human tissue, whether non-frozen (Oktay et al., 1998Go; Weissman et al., 1999Go; Nisolle et al., 2000Go; Gook et al., 2001Go) or cryopreserved (Kim et al., 2000Go; 2002; Nisolle et al., 2000Go; Oktay et al., 2000Go; Gook et al., 2001Go; Van den Broecke et al., 2001aGo), suggesting that the inability to achieve normal size does not relate to the cryopreservation. Smaller than normal diameters have also been observed in sheep follicles (Gosden et al., 1994bGo; Salle et al., 1998Go; 1999; Aubard et al., 1999Go; Jeremias et al., 2001Go) which developed following autologous transplantation of cryopreserved tissue. This phenomenon has also been reported following autologous (Lee et al., 2002Go) and heterologous (Candy et al., 1995Go) transplantation of cryopreserved monkey ovarian tissue. These results suggest that restricted follicle size may be a consequence of the transplantation site.

The present report is the first describing mature (MII) oocyte development from cryopreserved human ovarian cortex. The resumption of nuclear maturation in all oocytes within the larger follicles indicates the reproducible nature of the preservation of not only follicular but also oocyte function using the PROH cryopreservation procedure. Oocytes have been observed in antral cavities in previous studies using human cryopreserved ovarian tissue (Kim et al., 2000Go; Oktay et al., 2000Go; Gook et al., 2001Go; Van den Broecke et al., 2001aGo). In these studies, however, no exogenous stimulus for induction of maturation was administered and oocytes remained at the germinal vesicle stage. The identification of mature oocytes is a crucial step towards the success of human ovarian tissue cryopreservation, and also in the future development of ovarian tissue transplantation. Initial clinical reports using various transplantation techniques have, to date, resulted in the recovery of only one metaphase I oocyte (Oktay et al., 2001Go).

In conclusion, the highly reproducible and extensive evidence presented in the present study illustrates clearly that primordial follicles within human ovarian tissue, when cryopreserved using PROH, can subsequently develop to the antral stage and undergo the full range of morphological changes associated with ovulation, luteinization and oocyte maturation. Not only has this confirmed the clinical potential of this approach, but it has also provided a model which has allowed characterization of the process of human ovulation.


    Acknowledgements
 
The authors thank Angela Nelson for her expert care of the animals, and The Pharmacy College, Monash University for the use of their animal facility.


    References
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Antinori, S., Dani, G., Selman, H.A., Vidali, A., Antinori, M., Cerusico, C. and Versaci, C. (1998) Pregnancies after sperm injection into cryopreserved human oocytes. Hum. Reprod. (Abstract Bk. 1), 13, 157–158.

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Aubard, Y., Piver, P., Cogni, Y., Fermeaux, V., Poulin, N. and Driancourt, M.A. (1999) Orthotopic and heterotopic autografts of frozen-thawed ovarian cortex in sheep. Hum. Reprod., 14, 2149–2154.[Abstract/Free Full Text]

Bahadur, G. and Steele, S.J. (1996) Ovarian tissue cryopreservation for patients. Hum. Reprod., 11, 2215–2216.[Abstract]

Baird, D.T., Webb, R., Campbell, B.K., Harkness, L.M. and Gosden, R.G. (1999) Long-term ovarian function in sheep after ovariectomy and transplantation of autografts stored at –196°C. Endocrinology, 140, 462–471.[Abstract/Free Full Text]

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Submitted on March 21, 2003; accepted on May 20, 2003.