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Ovary |
Department of Biological Regulation, The Weizmann Institute of Science, Rehovot 76100, Israel
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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follicle, follicular development, oocyte development, ovary, ovulation, reproductive technology
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Among the possibilities to preserve fertility of patients at risk of premature ovarian failure are cryopreservation of embryos, oocytes, or ovarian tissue [3]. Efficient embryo or oocyte cryopreservation techniques require hormone stimulation to increase the number of oocytes. Such stimulation delays initiation of the anticancer therapy and might directly induce progression of hormone-dependent tumors. Embryo cryopreservation provides superior viability relative to oocyte cryopreservation; however, it is only appropriate for patients who have a partner or are willing to accept donor sperm for fertilization. In cryopreservation of both embryos and oocytes, the number of oocytes that can be retrieved is relatively low.
The procedure of ovarian tissue cryopreservation permits retention of hundreds of immature oocytes kept within the protective environment of the original ovarian tissue. An important advantage of this technique is the lack of unnecessary delays in providing anticancer treatment. Additionally, because the primordial follicles are small and structurally simple, they are much more tolerant to manipulation and to the freeze-thaw procedure compared with the large growing follicles that readily undergo atretic degeneration and granulosa cell apoptosis [4–7]. In most cases, ovarian tissue can be collected using laparoscopic ovariectomy or multiple ovarian biopsy [8]. Even though autotransplantation (transplanting the ovarian tissue back into the patient) seems to be the natural choice for preservation of fertility, the potential risk of retransferring cancer cells along with the grafts is a major drawback of this option. Shaw et al. reported that ovarian tissue from a mouse donor with lymphoma transferred the malignancy to recipient mice [9]. However, none of the mice that were grafted with ovarian tissue taken from human lymphoma patients developed the disease [10]. Ovarian tissue xenotransplantation (transplantation between different species) can serve both as a model/research system and as a safer clinical option by which the risk of malignant cancer cell transmission is reduced. In addition to the benefit to human patients who suffer from premature ovarian failure, this approach can also serve for preservation of endangered animal species [11, 12].
One of the most important factors for successful ovarian graft transplantation is the rapid establishment of a rich blood supply, which is crucial for survival of the ovarian follicles [13]. Without vascular anastomosis, grafts are solely dependent on posttransplantation vascularization. Dissen et al. showed that transplanted immature rat ovaries become profusely revascularized within 48 h after autotransplantation. Vascular growth was accompanied by increased expression of genes that encode the angiogenic factors, vascular endothelial growth factor (VEGF), and transforming growth factor-ß1 (TGFß-1) [14]. An important issue in ovarian transplantation is follicle survival. Transplantation of fresh mouse ovaries resulted in 50% reduction in follicle numbers [15]. In another study, only 35% of the oocytes survived fresh sheep tissue grafting into SCID mice [16]. In both studies, the grafts were transplanted into the well-vascularized kidney capsule.
Prior to revascularization, implants are vulnerable to ischemia-reperfusion injury, which is the main obstacle to the survival of a tissue after transplantation. Kim et al. showed a 30%–70% reduction in graft size and significant fibrotic changes within most grafts [17]. There is a direct correlation between the size of the follicles and their susceptibility to insufficient blood supply, namely, larger antral follicles invariably undergo degeneration while smaller ones survive. Most of the damage occurs mainly during the first few days after transplantation [15, 18].
The aim of this study was to investigate the angiogenic processes that occur after ovary xenotransplantation in order to find a way to protect ovarian grafts from posttransplantation damage. The experimental model of rat ovaries transplanted into immunodeficient mice was used for monitoring vascular remodeling by magnetic resonance imaging (MRI) and histology. The rat ovarian cycle is well defined and has been studied extensively, including some MRI studies on ovarian angiogenesis [19], while the nude mice provide a good model for MRI studies of angiogenesis of tumor xenografts [20, 21]. The ovaries were transplanted into two sites that represent tissues abundantly (intramuscular) or poorly (subcutaneous) vascularized. Significant ovarian damage was observed in both cases within the first 24 h after transplantation. However, while the intramuscular grafts recovered, most of the subcutaneous grafts remained impaired and necrotic. Regions with high pericyte distribution showed improved follicular integrity relative to regions devoid of pericytes. As previously shown in other systems, these perivascular cells are important for vascular stabilization and maintenance [22].
These results suggest that interventions aimed at improving ovarian transplantation should be focused on the establishment of perivascular support as early as possible after the implantation.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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All animal experiments were approved by the Institutional Animal Care and Use Committee. Immature 15-day-old Wistar rats were killed using CO2 and subsequent cervical dislocation, and ovaries were collected. The ovaries were cleaned from fat in L-15 Leibovitz medium (GibcoBRL, Invitrogen Corporation, Paisley, Scotland, UK) supplemented with 0.1% BSA and antibiotics (penicillin/streptomycin; 100 IU/ml) at room temperature. Several cleaned ovaries were fixed for morphological evaluation and served as controls. Ovaries were transplanted into CD-1 female nude mice, 6–10 wk old, within 20–30 min after collection. Briefly, mice were anesthetized by intraperitoneal administration of 75 mg/kg Ketaset (ketamine; Fort Dodge Laboratories, Fort Dodge, IA) and 3 mg/kg XYL-M (xylazine; VMD, Arendonk, Belgium) followed by a subcutaneous addition of approximately half of the initial dose in order to prolong the duration of anesthesia. Ovarian fragments were transplanted either subcutaneously at the hind limb above the gluteus superficialis muscle or in the muscle itself. In the subcutaneous transplantations, the graft was inserted through a 1-cm full-thickness dermal incision and placed about 3 cm away from the incision using tweezers. The incision was sealed with Super Glue Gel (ethyl-2-cyanoacrylate; Loctite, Cleveland, OH). Intramuscular grafts were inserted through a cut along the muscle fibers. The muscle was sutured using silk thread in order to allow detection of transplantation site after the animal was killed (anesthesia overdose).
In all transplantations, one ovary piece was transplanted into each mouse. In the subcutaneous transplantation, half ovaries were transplanted into 13 mice (Table 1). The ovaries were retrieved between Days 1 and 24 after transplantation. In the intramuscular ovarian transplantation, half ovaries were transplanted into 31 mice (Table 1). The grafts were retrieved between Days 1 and 31 after transplantation. In addition, different sizes of intramuscular ovarian grafts, obtained by cutting the ovary by fine tweezers under a binocular microscope, were studied by MRI (see below) and retrieved after 5–35 days. The grafts ranged from intact (6 mm3) to 1/8 ovary (0.75 mm3; Table 1).
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In Vivo MRI Studies
MRI experiments were performed on a horizontal 4.7 T Bruker (Karlsruhe, Germany) Biospec spectrometer using an actively decoupled 1.5-cm surface coil embedded in a Perspex board and a birdcage transmission coil.
Mice, 28 day (n = 3) and 35 day (n = 4) after intramuscular ovarian transplantation, were anesthetized as described above. Anesthetized mice were placed supine with the graft located above the center of the surface coil. The mice were immobilized using adhesive tape and covered with a paper blanket in order to reduce temperature drop during the measurement.
BSA-based macromolecular contrast material, biotin2-BSA-gadolinium-DTPA23 (biotin-BSA-GdDTPA; about 82 kDa) was prepared as reported previously [23] and bolus injected through a tail vein catheter (12.4 mg/mouse in 0.2 ml).
A series of precontrast spin echo images, with repetition time (TR) values of 1000, 500, 200, and 100 msec, were acquired to determine the precontrast R1. For dynamic postcontrast imaging, T1 weighted spin echo images were obtained 4–32 min after the contrast agent was administered (TR 200 msec, echo time 10.6 msec, two averages, spectral width 50 000 Hz, field of view 35 mm, slice thickness 1 mm, matrix 256 x 256, in plane resolution 137 µm, acquisition time 105 sec).
At the end of the MRI measurements, mice were killed by anesthesia overdose, and the grafts were retrieved for histological analysis.
Analysis of the Dynamic MR Data
MR data were analyzed on an Indigo-2 O2 work station (Silicon Graphics, Mountain View, CA) using Matlab (MathWorks Inc., Natick, MA). Pixel-by-pixel analysis was used to generate concentration maps of biotin-BSA-GdDTPA from spin-echo data sets as reported [23]. The concentration maps were used for derivation of vascular permeability (apparent permeability surface area product, APSMRI), namely the rate of contrast accumulation, derived by linear regression of the first 10 min of postcontrast agent administration. Only pixels with significant regression (r > 0.9) were considered. Data are presented using a color scale for APSMRI values between 0 and 0.8 µM/min, overlaid on a gray-scale anatomical image. Mean changes in APSMRI were calculated from selected regions of interest, mainly the graft and the adjacent muscle.
Contrast Agent Distribution in Intact Rat Ovaries
Rats were anesthetized by intraperitoneal administration of assival (Biogal; Teva Pharmaceutical Industries, Debrecen, Hungary), 125 mg/kg and ketamine (Fort Dodge Laboratories), 6.25 mg/kg. The contrast agent was applied to 15–70-day-old Wistar rats (via the tail vein or the heart). The animals were killed after 45 min in order to mimic the MRI procedure. Ovaries were retrieved for histological analysis after fixation.
Histology
Samples from the subcutaneous transplantations included the skin with the attached graft, while the grafts from the intramuscular transplantations were retrieved with the surrounding muscle as marked by the suture. The samples were placed overnight in Carnoy fixative solution (6:3:1 ethanol:chloroform:acetic acid), transferred into 70% ethanol, and stored at room temperature until processing. Fixed tissues were embedded in paraffin blocks and sectioned serially at 4-µm thickness.
Two out of each three slides of the subcutaneous transplants and every third slide in the intramuscular transplants were stained by eosin-hematoxylin, while the other slides were stained for endothelial cells, for smooth muscle cells, and for the biotinylated MRI contrast material. The paraffin-embedded unstained sections were deparaffinized with xylene for 5 min, followed by sequential ethanol hydration and double distilled water. Endogenous peroxidase was inactivated with 3% H2O2 in PBS for 5 min at room temperature. Sections were then washed with PBS for 5 min and were blocked by overnight incubation in 1% BSA in PBS at 4°C.
Endothelial staining Endothelial cells, which serve as a major component of the vascular inner layer, were stained by horseradish peroxidase conjugated Bandeiraea simplicifolia BS-1 Isolectin (Sigma, St. Louis, MO) and visualized with 3-amino-9-ethylcarbazole (AEC; Sigma) [24]. Specimens were counterstained with Mayer hematoxylin solution (Sigma).
Pericyte and smooth muscle cell staining
Mature blood vessels are coated with pericytes and smooth muscle cells (SMC), which express 
-smooth muscle actin (SMA). In order to detect the mature vasculature, the sections were stained with monoclonal anti--smooth muscle actin antibodies (SMA; Sigma), conjugated to alkaline phosphatase, and visualized with Fast red (Sigma). The slides were counterstained with Mayer hematoxylin solution.
Contrast agent staining The biotin-BSA-Gd-DTPA contrast agent was stained by avidin-FITC conjugate (Sigma) and sealed with a cover slip using Vectashield (antifade mounting with DAPI) (Vector Laboratories, Burlingame, CA).
The slides were examined with an Optiphot2 microscope (Nikon, Kanagawa, Japan) and photographed by a CCD camera (DVC Company, Austin, TX).
Follicle Examination
The eosin-hematoxylin-stained slides were used for follicle counts. The follicles were counted in all the detected grafts (Table 1). Both viable and atretic follicles were counted. Follicles were considered atretic if more than 1% pyknotic nuclei were found in the granulosa cells or if the oocyte began to degenerate [25]. In order to avoid double counting of the same follicle and to assure inclusion of the largest cross-sections of follicles, the counts were performed in the section in which the nucleolus was within the germinal vesicle (oocyte nucleus). Primordial follicles were counted if the oocyte appeared to have a definite nuclear membrane [7]. The follicles were classified into seven types based on the classification of Pedersen and Peters [26]: a) primordial, containing an oocyte surrounded by a single layer of flattened cells; b) primary, characterized by a single layer of cuboidal pregranulosa cells; c) secondary: containing two complete layers of granulosa cells; d) follicles containing three complete layers of granulosa cells; e) follicles containing four to five complete layers of granulosa cells; f) early antral follicle, containing multiple layers of granulosa cells; g) antral follicle, with the cavity occupying most of the total follicular volume. All sections were examined at 400x magnification under light microscopy. The examined sections were photographed at 20x or 100x magnification, and the area of the ovaries was measured by NIH Image. The data are presented as number of follicles per square millimeter of ovarian section.
We have applied a relative scale in order to estimate the condition of the ovary on the basis of follicle growth and viability. The seven follicle types were scored as follows: the viable ones received a positive grade from 1 to 7; the smallest one received a score of 1 and the largest one a score of 7. The atretic follicles were negatively scored, the smallest one received a score of -7 and the largest one received a score of -1. The rationale for this scoring system is based on the probability that small follicles being atretic is normally low, while as the follicles grow, their probability of being atretic is increased [27]. Therefore, we assume that excessive atresia of small follicles in the transplants is the result of ovarian stress due to insufficient blood supply. Thus, atretic primordial follicles received the most negative score (-7). Viable antral follicles received the highest positive score (+7). Each follicle type was graded as follows: follicle grade x (total number of follicles from the specified type/total area of the examined ovary [mm2]). The total score of an ovary was obtained by summing up the scores of each follicle type.
Histological Quantification of Vascular Permeability
The distribution of the biotinylated MR contrast agent in blood vessels and its extravasation in the ovarian graft and the adjacent muscle were measured by the fluorescence intensity of avidin-FITC. The samples were photographed at 100x magnification at 1 sec of exposure and a gain of 10 dB. Pictures were converted to 256 gray scale by Adobe Photoshop 6.0 ME (San Jose, CA) and the intensity of the different regions was measured using NIH Image. Selected regions of interest of the graft and the muscle were of the same size, whereas the blood vessel was analyzed using a smaller region confined to a well-defined blood vessel. Intensity data were normalized to a scale of 0–1, where 1 represents intensity within blood vessels. The intensity of the contrast agent driven from the histology data is referred to as APShistology.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Ovaries of 15-day-old Wistar rats contain primordial, primary, and small antral follicles (Fig. 1A). The GSL-1 staining represents staining of endothelial cells (e.g., Fig. 1B) and SMA staining represents SMC and pericytes (e.g., Fig. 1C).
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Subcutaneous half ovarian grafts were examined at various time points during the first days after transplantation (Fig. 1, D–L). Out of the 13 grafts, only 6 provided identifiable follicles within the ovarian sections (Table 1). A gradual decrease in the overall preservation (follicle state, vascular integrity, and necrosis) of the ovarian grafts was observed. Damage at the center of the graft was detected 2 days after transplantation (Fig. 1D). Damage was also observed in blood vessels, where pericytes were the first to be affected, as is indicated by reduction in the SMA staining (Fig. 1F). After 6 days, the damage expanded and only follicles at the graft periphery, located away from the necrotic center, survived (Fig. 1, G–I). Spatial correlation was observed between surviving follicles and maintenance of SMA-positive cells (Fig. 1I). After 11 days, most of the ovary was necrotic (Fig. 1, J–L). The condition of the ovary remained poor 24 days after transplantation (even though a few follicles could be observed, most of them were necrotic). It seems that, during the first days after subcutaneous transplantation, extensive irreversible damage to the graft occurred.
Intramuscular Ovary Transplantation: Optimization of Graft Size
Because the subcutaneous region is heterogeneous and relatively poor in its blood vessel support, we chose the muscle, which is more homogeneous and rich with vasculature, as a transplantation site. While the graft (in muscle as well as in the subcutaneous transplantations) should be large enough to contain the maximum pool of oocytes, it should also be small enough to minimize ischemia-reperfusion injury. In order to optimize the graft size in the intramuscular transplantation, we studied various sizes of ovarian grafts, ranging from intact ovaries (6 mm3) to 1/8 ovary (0.75 mm3; Fig. 2). The grafts were evaluated 23–35 days after the transplantation. We detected 23 out of the 25 grafts (Table 1), and all of the detected grafts showed relatively good survival (low necrosis) and follicle maintenance. Normal appearance of blood vessels was observed using endothelial (Fig. 2, middle column) and smooth muscle cell markers (Fig. 2, right column). Follicles in different developmental stages were detected. No significant differences were observed between the average grades of the different graft sizes (data not shown). The follicles developed to the antral stage (stage g) only in the half ovarian grafts (3 mm3; Fig. 2, D–F).
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Kinetic Studies of Intramuscular Ovarian Transplantation
The subcutaneous grafts showed extensive damage during the first days after transplantation. Therefore, the intramuscular grafts were assessed at different time periods, from 1 to 31 days posttransplantation (Fig. 3). Half ovaries were transplanted into the gluteus superficialis at the hind limb. Of the 31 grafts, 26 were detected (Table 1). One of the ovaries retrieved 24 h after transplantation was found to be disconnected from the muscle (Fig. 3, A–C). In that ovary, both the follicles and the vasculature were extensively damaged. Similar to the subcutaneous transplants, at this early time point after transplantation, damage was also detected in ovaries that were in close contact with the muscular tissue. Nevertheless, in this case, survival of primordial follicles along with some mature blood vessels was observed (Fig. 3, D–I). The damage 3 days after transplantation was limited to the center of the grafts, while at the periphery, viable follicles and intact blood vessels were observed (Fig. 3, J–L). Beyond 6 days after the transplantation, the grafts seemed to recover and resembled normal ovaries (Fig. 3, M–R). Follicles at progressive developmental stages, ranging from primordial to antral, were detected.
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Ovarian Preservation and Follicle Growth in the Grafts
We determined the state of grafted ovaries by the size and integrity of the follicles. Follicles were divided into seven groups according to their size, and both the viable and the atretic follicles were counted and graded. Grafts of half ovaries were assessed. In control ovaries, fixed immediately after the animal was killed (n = 3), all stages of follicular development were detected. In the controls, small atretic follicles were only rarely found at early stages of follicular development (Fig. 4A). In the subcutaneous ovarian grafts (n = 6; 2–24 days after transplantation), the largest viable follicles that were detected reached stage d (three layers of granulosa cells). A large number of degenerated primordial and primary follicles were detected (Fig. 4B). The follicles in intramuscular grafts (n = 26; 1–31 days after transplantation), as the control intact ovary, ranged from the primordial to the antral stages. The number of degenerating primary follicles (stage b) was significantly lower and the number of viable follicles at more advanced stages was significantly higher in intramuscular relative to subcutaneous grafts (P < 0.05; two-tail, unpaired t-test; Fig. 4C). Distribution of follicles at various stages of development 1–3 days (Fig. 4D; n = 11), 6–7 days (Fig. 4E; n = 8), and 28–31 days (Fig. 4F; n = 7) after transplantation are presented. By grading the follicles according to their growth and viability, a marked difference was observed between the control ovaries (which scored 17.6), the subcutaneous grafts (which scored -35.2, i.e., very poor preservation), and the intramuscular grafts (which scored 6.7; Fig. 5). Dividing the intramuscular grafts into three groups according to the time after transplantation reveals that, during the short time periods of 1–3 days after transplantation, the ovaries scored -7.7 (n = 11), after 6–7 days, the score was 25.5 (n = ;8), and at 28–31 days, the score was 8.0 (n = 7; Fig. 5). This scoring method reflects ovarian graft survival and was the lowest in the subcutaneous grafts throughout the 2–24 days after transplantation. The state of the intramuscular grafts was suboptimal during the first 3 days after transplantation; however, it improved after 6–7 days. After a period of about a month, a decrease in the follicular state was observed, although the grafts remained viable. The number of the small primordial follicles per square millimeter in the intramuscular site remained similar during the entire period examined (Fig. 4, D–F).
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Intramuscular Ovary Transplantation: Angiogenesis
In order to detect and evaluate functional blood vessels in the graft and in the adjacent muscle, a high molecular weight contrast agent, biotin-Gd-DTPA-BSA, was administered through the tail vein. In addition to showing the functionality of the vessels, its leakage is also a marker for vascular permeability. The contrast agent was followed by dynamic contrast-enhanced MRI at 28–35 days after transplantation. After administration, the contrast agent was immediately detected in large blood vessels (Fig. 6B) and subsequently extravasated in the ovarian implant. Changes in signal intensity reflective of vascular leakage were followed for 32 min (Fig. 6C). The accumulation of contrast agent was restricted to the graft and was not observed in the surrounding muscle (Fig. 6D). Histological staining of the biotinylated contrast agent was performed using avidin-FITC. Consistent with the MRI data, FITC staining was restricted to blood vessels within the muscle, whereas it leaked into the graft (Fig. 6E). The contrast agent was also detected in the follicular fluid. This leakage of the contrast agent in the ovarian grafts was similar to that observed in intact rats that were injected with the contrast agent intravenously 45 min prior to retrieval of the ovaries. Accordingly, the vascular properties of the ovarian grafts in the intramuscular transplantation appeared to be similar to those of the intact ovary.
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The distribution of the contrast agent in the ovarian grafts was significantly higher than in the adjacent muscle according to both the MRI and the histology intensity quantification (P < 0.05; two-tailed, unpaired t-test; Fig. 7, A and B; n = 7). The correlation between the MRI and the histology was highly significant (Fig. 7C; P = 0.008; r2 = 0.46).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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In both subcutaneous and intramuscular transplantations sites, tissue damage was apparent in the ovarian medulla within 24–48 h after the grafting. These first hours are probably the time period during which substantial damage occurs, mainly due to ischemia-reperfusion injuries. During this stage, the graft must receive adequate blood supply, without which it will suffer extensive irreversible damage. It seems that the muscle environment meets this condition for sufficient blood supply while the subcutaneous site does not.
Ovarian fragments implanted intramuscularly showed much better follicular maintenance. Many follicles, from the primordial through the antral stages of development, could be found in the implant, suggesting recovery of ovarian function. At 6 days after implantation, the vasculature in the implant was very similar to that of control ovaries, with normal morphology of blood vessels. Despite this favorable outcome, necrotic regions in the medulla of the graft were detected at the first days (1–3 days) after transplantation, suggesting that early transitory necrosis may be overcome and the maintenance of the grafts improved at a later time (6–7 days). The medullar damage during the first hours after transplantation was accompanied by regression of pericytes and SMC, which was symptomatic of insufficient blood supply.
The superior pericyte coating in the intramuscular versus the subcutaneous grafts seems to be related to the different blood supply between these two sites. The mechanism for vascular preservation in the two transplantation sites remains to be elucidated in future studies. It is possible, e.g., that the hypoxic stress and the subsequent production of factors that contribute to detachment of SMC and pericytes, such as VEGF and ANG-2, are higher at the subcutaneous site. Supplementation of stabilizing factors such as PDGF and/or ANG-1 to the graft, prior to the transplantation, in order to supply perivascular support may prevent pericyte and SMC loss and improve follicular survival also in this region. Administration of growth factors aimed to promote vascular maturation was reported previously to induce growth of stable and mature blood vessels [28, 29].
Considering the fact that the procedure of ovarian transplantation will require a freezing-thawing step in which the large developing follicles will be damaged, the main goal is preservation of the pool of nongrowing small follicles. An encouraging result was the fact that the number of primordial follicles in the intramuscular grafts remained essentially unchanged during a month after transplantation.
Survival of follicles was spatially related to presence of pericytes rather than to endothelial cells in the area of the graft (Fig. 1, G–I; Fig. 2, A–O; Fig. 3, G–R). The ovarian cortex showed better follicular maintenance, probably due to sufficient blood supply (see Fig. 1I). Interestingly, Dissen et al. showed that the mRNA expression of the two angiogenic factors, VEGF and TGFß-1, is upregulated mainly at the ovarian cortex 48 h after transplantation [14]. In the subcutaneous implants, no recovery of the initial damage was observed, whereas in the intramuscular grafts, improvement was observed within 6–7 days postimplantation, showing healthy follicular morphology and vascular integrity, including endothelial cells, pericytes, and SMC.
Vascular permeability in the implanted ovaries was studied by MRI and by histology. In muscle, the contrast material was restricted to the blood vessel lumen, whereas in the ovary, it was found to extravasate to the surrounding tissue. Shalgi et al. showed that the permeability of the blood follicle barrier to serum proteins is inversely related to their molecular weight and smaller proteins like albumin are found at higher concentration in follicular fluid than in the serum [30]. Such hyperpermeability is a well-known characteristic of the ovarian vasculature [31] and was also demonstrated here for the intact normal ovary as well as the transplanted ovary.
In conclusion, we report here that survival of ovarian implants was improved under conditions that support vascular maintenance. Thus, follicular integrity was associated with preservation of pericyte coating while loss of SMA was reflected by follicular damage. Subcutaneous implantation was followed by pericyte loss associated with tissue damage, whereas intramuscular transplantation showed vascular maintenance and was associated with improved follicular preservation.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Correspondence. FAX: 972 8 9342487; michal.neeman{at}weizmann.ac.il
Received: 26 September 2002.
First decision: 25 October 2002.
Accepted: 3 January 2003.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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