Hum. Reprod. Advance Access originally published online on January 23, 2008
Human Reproduction 2008 23(3):619-626; doi:10.1093/humrep/dem405
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Back muscle as a promising site for ovarian tissue transplantation, an animal model
1 Centre for Reproductive Medicine, Ghent University Hospital, De Pintelaan 185, Oost Vlanderen, 9000 Ghent, Belgium 2 Gynecological Oncology, Department of Obstetrics and Gynecology, Ghent University Hospital, De Pintelaan 185, 9000 Ghent, Belgium 3 Department of Pathology, Ghent University Hospital, De Pintelaan 185, 9000 Ghent, Belgium 4 Present address: Centre for Reproductive Medicine, Universitair Ziekenhuis Brussel, Laarbeeklaan 101, 1090 Brussels, Belgium
5 Correspondence address. E-mail: reza.soleimani{at}ugent.be, rezasoleimani{at}yahoo.com
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Abstract |
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BACKGROUND: The aim of this study was to evaluate the optimal transplantation site for ovarian tissue fragments in murine hosts. We compared the transplantation to the back muscle (B) versus the kidney capsule (K) in a mouse allograft model.
METHODS: Hemi-ovaries from 12-day-old mice were allografted into B and K of bilaterally ovariectomized same strain recipients which had undergone gonadotrophin stimulation (n = 15). Graft survival after 27 days, angiogenesis and follicle development were scored and compared to age-matched control ovaries (38-day old, n = 5). The ability of oocytes to be fertilized was studied after IVF, ICSI and embryos were transferred to recipient mothers. Anti-mouse CD 31+ antibody was used to evaluate neo-vascularization in grafts.
RESULTS: Primordial follicle survival was higher (P < 0.01) and vascular support was better (P < 0.01) in B- than in K-grafts. From 34 oocytes retrieved from B-grafts (15 metaphase I, of which 14 matured in vitro, and 19 collected at metaphase II), 18 morulae were obtained. Transfer of 12 embryos obtained by ICSI led to three live offspring, and transfer of six IVF embryos to another recipient mother yielded four offspring, one of which was born dead and one showed placental anomalies.
CONCLUSIONS: The back muscle is a promising site for ovarian allografts in mice. This is the first report of live offspring obtained after back muscle grafting using both IVF and ICSI.
Key words: back muscle grafting/follicle survival/live offspring/ovarian tissue transplantation/graft neo-vascularization
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Introduction |
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Improvements in cancer therapy in the last decade led to a significant increase in survival rate of young female patients. Unfortunately, long-term side effects of anti-cancer treatments are sub-fertility and premature ovarian failure (Lushbaugh and Casarett, 1976
; Chapman et al., 1979a, b; Wallace et al., 1989; Apperley and Reddy, 1995; Meirow and Nugent, 2001; Critchley et al., 2002; Larsen et al., 2003; Falcone and Bedaiwy, 2005). One of the only options for fertility preservation in these patients is cryopreservation of ovarian tissue, containing primordial follicles, followed by grafting of frozen–thawed ovarian tissue (Yang et al., 2007).
Despite the fact that human ovarian tissue banking is being offered worldwide, to date there are only a few reports of successful clinical use of frozen ovarian tissue leading to pregnancy or live birth (Donnez et al., 2004; Meirow et al., 2005).
Optimal grafting conditions are difficult to study in clinical trials both practically and ethically. Therefore, animal models have the advantage of exploring the adequacy of clinically used sites, such as the s.c. site, and also more non-conventional transplantation sites. The strategy of studying follicle development in animal allografting and human–mouse xenografting models has been reported for different transplantation sites such as the kidney capsule (K) (Gosden et al., 1994; Newton et al., 1996; Oktay et al., 1998; Gook et al., 2001, 2003, 2005; Hernandez-Fonseca et al., 2004; Waterhouse et al., 2004), a s.c. site (Weissman et al., 1999; Van den Broecke et al., 2001; Kim et al., 2002, 2005; Schmidt et al., 2003; Hernandez-Fonseca et al., 2004) and an i.m. site (Revel et al., 2000).
Weissmann et al. (1999) studied different grafting conditions for s.c. human ovarian tissue xenotransplantation in the non-obese diabetic/severely compromised immunodeficient (NOD–SCID) mouse. They showed that using male instead of female recipient mice and using warm instead of ice cold transportation conditions yield better results. They did not find any significant difference in follicular survival and development between intact and pituitary down-regulated host mice after hormonal stimulation.
Israely et al. (2003, 2006) studied the reduction of ischemic damage in rat ovarian fragments xenografted into a s.c. position, the gluteus superficialis muscle of the hind limb and also an angiogenic granulation tissue created during wound healing in the hind limb muscle. Ovarian grafts from an angiogenic granulation tissue showed significantly improved graft vascularization and follicular survival.
Other interesting studies showed that retransplantation of ovarian tissue, back to the original animal, leads to hormonal function restoration and to live offspring after grafting of fresh and cryopreserved ovarian tissue in a number of different species, such as mice (Cox et al., 1996; Gunasena et al., 1997; Candy et al., 2000; Shaw et al., 2000; Snow et al., 2002), sheep (Gosden et al., 1994; Baird et al., 1999; Salle et al., 2002), hamsters (Parrott, 1959) and rabbits (Petroianu et al., 2002). Clinical trials also have led to endocrine function after grafting cryopreserved ovarian tissue fragments heterotopically to the forearm (Oktay and Karlikaya, 2000; Oktay et al., 2001) or in an orthotopical transplantation to ovarian and peritoneal sites combined with a heterotopical transplantation to the abdominal s.c. site (Demeestere et al., 2006).
Liu et al. (2002a, b) could obtain life offspring after IVF of oocytes retrieved from cryopreserved primordial mouse follicles by sequential in vivo transplantation into K-site and in vitro maturation (IVM). Snow et al. (2002) also reported generation of live young from xenografted mouse ovarian tissue into rat.
From all those trials it became obvious that the site of grafting is very important, since it influences tissue survival and follicle development as well as the quality of the obtained oocytes after transplantation. Yang et al. (2006) showed that fertilization and two-cell development rates after IVF are significantly lower in oocytes retrieved from transplanted ovarian tissues compared to the in situ group. They could also obtain live offspring from mouse ovary fragments grafted into the bursal cavity, s.c. and K-site.
On the basis of previous reports of i.m. xenografting sites (Revel et al., 2000; Israely et al., 2003, 2006), we allografted mouse hemi-ovaries to the back muscle (B) and compared this site to K-site. In addition to histological observations, we studied graft functionality by recovering oocytes for IVF and subsequently analysed in vivo developmental competence. We report live offspring from oocytes retrieved from B-site ovarian grafts and fertilized by IVF or ICSI in the mouse.
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The study was approved by our institutional review board. Animal handling was performed with an approval of the Ghent University Hospital Ethical Committee on the use of experimental animals.
Experimental animals
Twelve-day-old (C57BL/6j x CBA/Ca) F1 female hybrid mice (B&K Universal, Hull, England) were used as ovarian tissue donors. Mice of the same strain were used at 8-week-old (F1 female hybrids) to act as ovarian tissue recipients and at 38 days for collection of age-matched control ovaries. For the IVF experiments 8-week-old F1 female hybrids were used for the collection of control ovulated oocytes and as pseudo-pregnant recipients for embryos resulting from B-site graft derived oocytes.
Twelve-week-old F1 male hybrids from the same strain were used for sperm donation.
Transplantation procedure
Tissue donor animals were sacrificed by cervical neck dislocation. Left ovaries were recovered, and cut approximately to half, each in separate 35 mm tissue culture dishes (Falcon; BD 35-3001, VWR, Leuven, Belgium) containing 2 ml of pre-warmed MCDB 105 (Sigma, M-6395, Bornem, Belgium)xM199 medium (Sigma, M-2154), ratio 1:1. Recipient animals were anesthetized by i.p. injection of Natrium pentobarbital 25 mg/kg (Nembutal, CEVA Sante Animale, France) at room temperature.
Hemi-ovaries from the left ovary of each donor animal were selected randomly and co-allotransplanted into the K-site and the B-site of 15 bilateral ovariectomized recipient mice.
Under the kidney capsule
The kidney was exteriorized through a dorso-horizontal incision in skin and body wall at the left side. A hemi-ovary was selected randomly and inserted under the capsule of the left kidney through a small hole made using fine watchmakers’ forceps. At the same time, both ovaries of recipients were removed by cauterization at the top of the uterine horns.
Back muscle site
The other hemi-ovary was inserted into the back muscle about 10 mm away from the edge of the incision by using fine watchmakers’ forceps to make a hole 3–5-mm deep.
Finally, body wall and skin incisions were closed. All procedures were performed under aseptic conditions.
Gonadotropin treatment in transplanted animals
In a preliminary study, different stimulation schedules were compared with a non-stimulated control group and the best was chosen to be used in present study. There was no significant difference whether starting FSH stimulation from the day after grafting or one week after grafting in the number of antral follicles and ovulated oocytes in the grafts, but both methods yielded higher oocyte numbers compared to the non-stimulated group (35 ± 20 and 31 ± 22 versus 18 ± 11, antral follicle numbers respectively, mean ± SD; P < 0.05).
We also observed that ending the stimulation with two doses of 5 IU FSH gives a slightly but significantly higher developing follicle number (data not shown).
Therefore 1 week after grafting, ovarian stimulation was started by i.p. injection of one IU FSH (Puregon, NV Organon, Oss, The Netherlands) given every second day for 2 weeks. Finally, two doses of 5 IU FSH were given i.p. followed by one dose of 5 IU HCG (Chorulon, Intervet, Boxmeer, The Netherlands) i.p., 48 h later. Animals were autopsied 14 h after HCG injection and grafts from K and B sites were recovered.
Follicular assessment, histology and quantitative study of angiogenesis
Histological assessment was carried out on grafted hemi-ovaries and age-matched non-grafted controls. Age-matching was done at the start of the experiment (12-day-old control ovaries, code O12, n = 5) and at the time of collection of the grafts (38-day-old control ovaries, code O38, n = 5).
All recovered grafted and control tissues were fixed in 4% buffered para-formaldehyde (Klinipath, Geel, Belgium. Ref No: 4078.9020) at 4°C for 2 h and stored in 0.1% pre-cooled para-formaldehyde/phosphate-buffered solution (PBS) at 4°C and processed manually for paraffin embedding. Histomorphological examination was carried out after serial sectioning to 4 µm thickness and staining with hematoxylin–eosin. To prevent recounting of the same follicle, follicles were only counted when the dark staining nucleolus was seen within the nucleus of the follicles. Follicles were classified as primordial (oocytes surrounded by one layer of flattened pregranulosa cells), primary (surrounded by one layer of cuboidal granulosa cells), preantral (with two or more layers of granulosa cells without antrum), antral (with an antral cavity), ovulated metaphase II (MII) (oocytes found in a cavity formed in grafting site) and corpora lutea (Jones and Krohn, 1961). To prevent any mistake in counting, slides were counted by two independent individuals and results were compared.
Anti-mouse CD31+ immunohistochemical staining (staining epithelial cells of new blood vessels) was performed in 4 µm thick sections to evaluate neo-vascularization of the grafted tissues (no of animals = 5). CD31 (PECAM-1) (Platelet Endothelial Cell Adhesion Molecule-1) is a 130 kDa integral membrane glycoprotein, i.e. expressed on the surface of new blood vessels (DeLisser et al., 1997). Antigen retrieval pretreatment was carried out by boiling the slides twice for 5 min in 10 mM citric acid, pH 6.0, in a microwave oven and cooling down at room temperature, followed by incubation for 10 min using protease 1 solution (0.5 enzyme units/ml) (Ventana, Lillie, France, Ref No: 760-2018). To prevent non-specific antigen binding, sections were incubated for 20 min at room temperature covered by Bovine Serum Albumin 1% (w/v) (BSA, Sigma–Aldrich, Bornem, Belgium, Ref No: a7030). Primary anti-mouse CD31+ antibody (BD-Pharmingen, Bornem, Belgium) dilution 1/200 in 1% BSA/PBS was applied on sections overnight at 4°C. Polyclonal rabbit anti-rat biotinylated immunoglobulin (DakoCytomation, Heverlee, Leuven, Belgium, Ref No: e0468), 1/300 in PBS for 60 min was used as secondary antibody. Streptavidin-horse-radish peroxidase (DakoCytomation, Heverlee, Leuven, Belgium, Ref No: p0397) 1/300 in 1 x PBS was applied for 30 min. Liquid DAB (3,3-diaminobenzidine)+chromogen (DakoCytomation, Heverlee, Leuven, Belgium, Ref No: k3468) was applied for 10 min. The sections were also lightly counterstained with hematoxylin before mounting. Angiogenesis was studied by counting and averaging blood vessels using high power field magnification (x400) in five randomly selected positions in five different sections in each tissue sample.
IVF/ICSI and embryo transfer
In a complementary step, eight mice were used to recover oocytes after allografting of ovarian tissue to the B-site.
On Day 38, recipient animals were autopsied and grafts with surrounding back muscle tissue were removed aseptically and collected in KSOM–HEPES buffered medium (Potassium simplex optimized medium)+4% BSA (w/v) (Calbiochem, #12657, Euro Biochem, Bierges, Belgium). All the components to make KSOM medium were purchased from Sigma–Aldrich Chemie, Bornem, Belgium and HEPES from Gibco, cat. No. 15630-056, Life Technologies, Belgium.
Oocytes were collected mechanically by puncturing the cystic-like cavity formed around the site of transplantation by means of two 24 G needles in a 60 mm Falcon dish (Falcon; BD 35-3002). Antral follicles were also punctured to release oocytes. All retrieved oocytes were placed in pre-warmed KSOM+0.4% BSA.
Oocytes were scored for maturity (Germinal vesicle, metaphase I (MI), MII). Germinal vesicle oocytes were not further included in the study (Liu et al., 2000a, b). MI oocytes were submitted to IVM. In vitro matured oocytes that became MII were fertilized by ICSI. Oocytes that were already MII at collection were fertilized either by IVF or ICSI.
As controls, 35 MII oocytes from super-ovulated mice were collected. Super-ovulation was induced in two female mice from the same strain by i.p. injection of 5 IU FSH (Puregon) followed by one dose of 5 IU HCG (Chorulon) i.p., 48 h later. Animals were autopsied 14 h after HCG injection and oocytes with surrounding cumulus complex were punctured directly from ampula using two 24 G needles in a 60 mm Falcon dish containing KSOM–HEPES medium, fertilized by ICSI and cultured in parallel with the experimental groups. All embryos that reached the morula stage in the experimental and control groups were transferred to pseudo-pregnant recipient mice.
Sperm preparation
Twelve-week-old male (C57BL/6j x CBA/Ca) F1 mice were used to obtain sperm. Caudae epididymidis were collected and placed in 1 ml KSOM with high glucose concentration (5.5 mM) under oil in a sperm dispersion dish (Falcon; BD 35-3037). Contents of the epididymis were squeezed out gently and residual caudal tissue was discarded. The culture dish was incubated for about 20 min to let the sperm disperse under 6% CO2. capacitation was performed by adding 10 µl of sperm suspension to 90 µl of KSOM supplemented with 4.0% BSA crystalline fraction V (Sigma, A-3311) (w/v) with incubation at 37°C in a 6% CO2 atmosphere for 1 h.
IVF
Oocytes were placed in 100 µl fertilization droplets with c. 1 x 106 spermatozoa (10 µl sperm suspension + 90 µl KSOM) for 4 h at 37°C in a 6% CO2 atmosphere. Finally, attached sperm and cumulus cells were removed by means of aspirating the oocytes in and out of a fine-drawn pipette. Inseminated oocytes were incubated in 20 µl droplets of KSOM with low glucose concentration (0.2 mM) culture medium supplemented with 0.4% BSA (w/v) in 60 mm tissue culture dishes at 37°C in a 6% CO2 atmosphere.
Partial zona dissection and ICSI
Zona dissection was performed at 37°C by means of a fine glass needle. Sperm was collected from the caudae epididymidis and placed in 1 ml KSOM–HEPES. Morphologically normal, motile spermatozoa were randomly selected for ICSI. Immediately before injection 3 µl of freshly thawed 10% polyvinylpyrrolidone solution (PVP; Vitrolife, Sweden AB, Kungs backa, Sweden) was added to the drop with spermatozoa to immobilize and facilitate handling. Sperms tails were separated from heads by pressuring the injection pipette at the tail of a spermatozoon against the bottom of the dish. Separated sperm heads were injected into the zona pellucida-dissected oocytes at 17°C on a pre-cooled inverted microscope. Injected oocytes were left to rest for 20 min at 17°C followed by 15 min at room temperature.
All oocytes after IVF or ICSI were washed four times in KSOM+0.4% BSA (w/v) medium.
Embryo culture and transfer
Fertilized oocytes were cultured in a 60 mm tissue culture dish in 20 µl of KSOM+0.4% BSA (w/v) medium under mineral oil. Pseudo-pregnant F1 female mice of the same strain were used after mating with vasectomized CD1 males (IFFA Credo). Morulae and compacted morulae were transferred with a glass pipette to the upper part of the left uterus horn of Day 3 pseudo-pregnant mice. Recipient mothers in experimental and control groups were left to deliver and raise pups.
Statistical analysis
Follicle and blood vessel counts are presented as mean ± SD. Levene’s test of homogeneity of variances (
= 0.01) and Kolmogorov–Smirnov test of normality ( = 0.01) were performed to choose the appropriate statistical test. One-way analysis of variance followed by Scheffé multiple comparison tests were used to compare the mean numbers of primordial, primary and total follicles present in B-site or K-site grafted hemi-ovaries and in 12- or 38-day-old control mouse hemi-ovaries. The mean number of preantral, antral, ovulated follicles and corpora lutea were compared using a Kruskal–Wallis rank test followed by Dunn tests. The Mann–Whitney U-test was used to evaluate the data for vascularization. When the P-value was <0.05, the difference was considered significant.
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Site of transplantation and ovarian follicle survival
All 30 grafted hemi-ovaries in the B- and K-sites were recovered.
Even though hemi-ovaries of almost the same size were transplanted, it was observed that the overall total follicle count was significantly higher in B- than in K-grafts (447 ± 224 versus 275 ± 138, P < 0.05). In particular, the count of primordial follicles was significantly higher in B-grafts compared to K-grafts (340 ± 158 versus 140 ± 87, P < 0.01).
There was no significant difference between B-grafts and O38 controls in number of primordial follicles (340 ± 158 and 369 ± 177, respectively, P > 0.05) (Table I).
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Total follicle and primordial follicle count was significantly lower in K-grafts compared with O12 and O38 controls (257 ± 138 versus 676 ± 181 and 491 ± 236, respectively, for total; 140 ± 87 versus 589 ± 179 and 369 ± 177, respectively, for primordial, P < 0.05) (Table I).
Follicular development to the antral stage was observed in all B- and K-grafts and in 38-day-old age-matched controls. There was no antral follicle development seen in 12-day-old age-matched control ovaries. There was no significant difference in the number of primary, preantral and antral follicles between B- and K-grafts and O38 controls. Also, there was no significant difference in corpora lutea development in B- and K-grafts (Table I).
Follicular assessment, histology and quantitative study of angiogenesis
Ovulation sites in the back muscle grafts showed a unique picture of free oocyte–cumulus complexes inside a big cavity with a cystic-like structure formed around the ovulation site (Fig. 1). Histological studies showed that this cavity was partially covered with a few layers of flattened mesothelial cells (Fig. 2). In 12 out of 15 B-grafts, ovulating follicles were observed with at least one oocyte found released free in the cavity (Table I). No ovulating follicle was observed in K-grafts.
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In all grafts re-vascularization was more prominent in B- than in K-grafts (2.56 ± 0.36 versus 1.56 ± 0.26, respectively, P < 0.01, n = 5). In all of the B-grafts, re-vascularization providing a specific blood stream around the grafting site was observed (Fig. 3).
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In vivo competence of oocytes from B-site allografts
In total, 19 MII and 15 MI oocytes were recovered from eight B-site grafts. From the 15 MI oocytes, 14 became MII (93%) after IVM and 7 (50%) developed to the morula stage after ICSI. Of the 19 collected MII oocytes, five out of nine (56%) developed to the morula stage after ICSI and six out of 10 (60%) after IVF (Table II). All 12 ICSI embryos were transferred to one mouse and all six IVF embryos to another. Three live born fetuses were obtained in the ICSI group (25%). In the IVF group, in total four fetuses were born (67%). One fetus was born dead, one showed placental anomalies and the other two were born healthy. In the control group, 30 out of 35 oocytes (85.7%) developed to the two-cell stage after ICSI and 27 (77%) reached the morula stage (Table II). Embryos were transferred to pseudo-pregnant females as two groups of 12 and 15 each. Seven healthy pups were obtained from the first group and no pregnancy was seen in the second group.
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Discussion |
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In the present study, mouse ovaries were grafted into two different locations in donor mice to determine the most suitable site for grafting and to compare the follicular survival and development. We report the first live offspring from mouse oocytes retrieved from B-site allografts and fertilized in vitro. The histological sections clearly showed mature oocyte–cumulus complexes and their ovulation in the grafted tissues.
Furthermore, we confirmed the better survival of follicles in the B-site grafts than in the K-site grafts. We quantified and compared neo-vascularization in the B- and K-site grafts by detecting endothelial cell–cell interactions involved in angiogenesis by using anti-mouse CD31 immunohistochemical staining.
Even though murine ovary re-vascularization occurs within 48 h after transplantation (Dissen et al., 1994; Israely et al., 2003), this is long enough to be considered as a main limiting factor for ovarian tissue transplantation (Jeremias et al., 2002), due to the loss of a large fraction of follicles during the initial ischemia. Follicle cell apoptosis occurs shortly after the transplantation, leading to the loss of the primordial content of the grafts (Liu et al., 2002a, b). This can be the result of ischemic-reperfusion injury during generation of new blood vessels in the grafts (Israely et al., 2006). Survival of follicles at different stages of development in grafts may be dependent on their metabolic rate (Nugent et al., 1997). It is obvious that the transplantation adds an additional metabolic stress on follicles but it seems that primordial follicles can survive grafting better than follicles at other stages (Gosden et al., 1994). Indeed, hypoxic conditions may be the main reason of follicle loss initially after transplantation (Aubard et al., 1999). On the other hand, low oxygen supply to the tissue may induce angiogenesis by up-regulation of angiogenic vascular endothelial growth factor (Laschke et al., 2003). Then rapid re-vascularization after grafting is considered essential for maximum follicle survival (Dissen et al., 1994).
Therefore, we looked for a grafting site which facilitates angiogenesis to protect the ovarian tissue graft from ischemia. In a preliminary attempt, we studied different body locations such as a s.c. site and we were only able to recover two out of six grafts (data not shown). Also after grafting to the muscle in the hind limb a large numbers of follicles were lost, as reported previously by Israely et al. (2003).
A recent report from Israely et al. (2006) showed an improvement of follicular maintenance after transplantation into angiogenic granulation tissue in a wound healing area in the hind limb muscle, again stressing the importance of vascularization. Intact oocytes at different stages of maturation (mostly at the germinal vesicle stage) were retrieved but IVF was not attempted. The hind limb muscle may not to be the perfect site for transplantation because of the high muscular activity at this location, which possibly puts additional physical stress on the grafted tissue. For this reason, we chose to graft ovarian tissue into the back muscle.
Our study showed that the B-site supports murine ovarian graft survival. Higher total follicle counts in B- versus K-grafts, and primordial follicles in particular, showed its unique ability to preserve the vitality and potential of resting follicles to survive, probably due to faster and better support of blood vessels in the grafted tissue compared to the K-site (Table I).
Even though there were slightly more follicles lost in B-grafts compared to O38 controls, the difference was not statistically significant, confirming the supportive nature of the back muscle compartment after grafting. On the other hand, there was a significantly higher follicle loss rate in K-grafts compared to B-grafts, O38 and O12 controls (Table I).
The primordial follicle pool decreases naturally with time in all species (Gougeon et al., 1994) (Table I) and therefore the difference between O12 versus O38 controls and B-grafts is naturally expected. Histological evaluation of the grafted tissues showed improved graft quality in B- versus K-grafts (Fig. 1).
It is known that FSH stimulates granulosa cell mitosis and follicle maturation and inhibits granulosa cell apoptosis after initial stages (Gougeon, 1996). The hormonal supply depends on good re-vascularization of the grafts. Abir et al. (2003) have compared the K-site to s.c. sites, using fresh or frozen human fetal ovarian tissues after transplantation to immunodeficient mice. The K-site has been shown to be superior to the s.c. site in supporting ovarian grafts, and is probably due to its superior vascularization and rich content of angiogenic factors, such as vascular endothelial growth factor (Abir et al., 2003). From the results obtained by different groups (Newton et al., 1996; Oktar et al., 1998, 2000; Gook et al., 2001; Van den Broecke et al., 2001), we can conclude that adult ovarian fragments survive transplantation and develop better than fetal ovarian fragments. Also, earlier, our group has reported successful development of antral follicles after hormonal stimulation of xenografted human ovarian fragments into K-site or s.c.: we showed that primordial follicles from transplanted fresh or frozen–thawed human ovarian cortex can grow and are responsive to hormonal stimulation (Van den Broecke et al., 2001). In the present study, histological examination of B-grafts showed a unique connection between the graft and the site of grafting (Fig. 1). We showed that the B-site provides better supportive neo-vascularization with specific blood stream generation in the transplantation location (Figs 2 and 3). The results were confirmed by finding significantly higher numbers of blood vessels in B- than in K-grafts.
Even though ovarian tissue can survive freezing well (Liu et al., 2002a, b), follicular apoptosis might be a consequence of the freezing and thawing procedures (Rimon et al., 2005). Therefore, only fresh ovaries were used in this experiment, although the ultimate goal for fertility restoration in human is the successful transplantation of frozen–thawed ovarian tissue.
Apart from advantages in terms of better follicular survival and specific rich neo-vascularization around the site of grafting, the B-site also provides a larger manipulation and transplantation field. The back muscle tissue compartment may allow large follicles to develop. The convenience of insertion and subsequent graft accessibility are additional benefits of this site compared to the under K-site. However, it has to be acknowledged that human and murine ovaries are quite different tissue compartments, in terms of follicular content and developmental capacity.
To date, several clinical reports have been published on surgical approaches for fresh or frozen–thawed human ovarian tissue auto-transplantation to different heterotopic body locations such as the pelvic sidewall (Oktay and Karlikaya, 2000), forearm (Oktay et al., 2001), anterior abdominal wall (Kiran et al., 2004; Schmidt et al., 2005), and also orthotopically (Radford et al., 2001; Donnez et al., 2004; Meirow et al., 2005; Schmidt et al., 2005) but it is not yet clearly established which site yields the best results after ovarian tissue transplantation. In view of the demonstrated advantages of muscle sites for ovarian tissue transplantation, they can be subject of further exploration for human ovarian tissue xenografting model.
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R.S. and E.H. are supported by the Special Research Foundation (BOF) of Ghent University. P.D.S. is holder of a fundamental clinical research mandate by the Flemish Fund for Scientific Research (FWO-Vlaanderen).
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Acknowledgements |
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Our special thanks go to the Organon Co. for providing us with recombinant FSH (Puregon) used in this study. The Authors would also like to thank Sharon Bryan, R.N. for her help in reading and editing this manuscript.
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References |
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Submitted on September 18, 2007; resubmitted on November 19, 2007; accepted on November 30, 2007.
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