Ovarian Graft Retrieval

Ovaries were collected from donor mice at the beginning of their sexual maturity, at 5–6 wk of age. The following three mouse strains were used in this study: 1) Mice with transgenic tetracycline-repressible endothelial-specific Akt1 activation under the VE-cadherin promoter (VE-cadherin-tTA;TET-myrAkt1) were received from Professor Laura E. Benjamin (Department of Pathology, Beth Israel Deaconess Medical Center, and Harvard Medical School) [8]. Ovarian grafts were retrieved 4 days after induction of expression of myrAkt1 in the donor animals by removal of tetracycline from the drinking water. Ovarian grafts from TET-myrAkt1-negative mice were used as matching controls; 2) Mice that express tdTomato red fluorescent protein in endothelial cells under the VE-cadherin promoter, that is, B6.Cg-Gt (ROSA)26Sor < tm14 (CAG-tdTomato)Hze > /J (Jackson Laboratory); and 3) C57BL/6 black wild-type mice (Harlan Laboratories).

Mice were euthanized using CO₂ inhalation, and ovaries were collected and cleaned of fat and other tissues in a PBS solution maintained at 37°C. This model focuses on angiogenesis and vascular remodeling during the critical ischemic interval posttransplantation, and thus, we used a fresh ovarian graft that was transplanted within 15 min from retrieval to minimize preimplantation ischemic damage to the tissue.

Ovarian Graft Transplantation

Ovarian grafts were transplanted in 6- to 9-wk-old CD-1 nude female mice (Harlan Laboratories). At 2–3 wk prior to ovarian tissue transplantation, recipient mice underwent ovariectomy under anesthesia with 100 mg/kg ketamine and 20 mg/kg xylazine, injected intraperitoneally. Two different transplantation sites were used in this study: 1) Intramuscular ovarian graft transplantation was applied for in vivo dynamic contrast-enhanced MRI. A 5 mm incision was made in the biceps femoris muscle of the thigh, 5 days prior to transplantation, to form granulation tissue at the site of implantation. It was previously reported that transplantation into granulation tissue improves graft revascularization [10]. On the day of ovarian graft transplantation, the surgical incision in the biceps femoris muscle was cleaned of blood clots and debris and rinsed with PBS. Then the ovary was transplanted intramuscularly at the site of the granulation tissue created by the incision 5 days earlier. The edges of the muscle were closed over the graft with 7/0 Prolene sutures (Ethicon), taking care not to traverse and damage the ovarian tissue. Skin incision was closed with 5/0 Mersilk sutures (Ethicon) and covered with Super Glue Gel (ethyl cyanoacrylate; Loctite) to seal the wound. 2) Intraperitoneal transplantation was applied for fluorescent imaging. The peritoneal surface was exposed through a midline laparotomy incision. Ovarian grafts were anchored to the parietal peritoneum with 7/0 Prolene sutures. The peritoneal cavity was closed with continuous sutures of 5/0 Mersilk. The skin was closed with interrupted 5/0 Mersilk sutures and sealed with Super Glue Gel.

Drug Interventions

Superovulation was induced 30 days after transplantation. Mice were subcutaneously injected with 5 units of equine chorionic gonadotropin (eCG) (National Hormone & Peptide Program, Harbor-UCLA Medical Center) dissolved in 100 ml of PBS. Five units of human chorionic gonadotropin (hCG) (Sigma Aldrich) was intraperitoneally injected to the animals 48 h after the eCG [11]. Ovarian grafts were monitored using MRI.

Activation of Akt by simvastatin was tested in 0.1 and 1mg/kg doses by ex vivo immunoblotting (see below). To activate the simvastatin prodrug (Sigma), 4 mg of simvastatin was dissolved in 100 μl of ethanol. Then, 150 μl of 0.1 N NaOH was added to the solution that was subsequently incubated at 50°C for 2 h. The pH was brought to 7.0 by HCl, and the final concentration of the stock solution was adjusted to 4 mg/ml. The stock solution was kept at 4°C [12]. Recipient mice received daily intraperitoneal (IP) injections of low-dose simvastatin (0.1 mg/kg/day) or saline, starting 1 day before grafting until 7 days thereafter, when complete reperfusion is expected [2]. Ovarian grafts were monitored using MRI.

In Vivo Dynamic Contrast-Enhanced MRI of Ovarian Grafts

MRI experiments were performed at 9.4 T on a horizontal Biospec spectrometer (Bruker), using a linear coil for excitation and a 2 cm surface coil for detection (Bruker). Animals were anesthetized with isoflurane (3% for induction, 1%–2% for maintenance); (Abbott Laboratories Ltd.) in oxygen (1 L/min) delivered through a muzzle mask and kept under respiratory monitoring. Body temperature was maintained using a heated bed.

Recipient mice were serially scanned at predefined time intervals: Day 2 to obtain a baseline during the ischemic period of the graft; Day 7 at the end of the ischemic period when graft reperfusion is expected; Day 30 before superovulation (pre-eCG/hCG); and Day 33 after superovulation was completed (post-eCG/hCG). During MRI experiments, the macromolecular contrast agent biotin-bovine serum albumin-gadolinium diethylenetriaminepentaacetic acid (biotin-BSA-GdDTPA) (10 mg/mouse in 0.2 ml of PBS; SyMO-Chem BV) was injected through a tail vein silicone catheter. Three-dimensional (3D)-gradient echo images of the graft were acquired before and sequentially for 40 min after intravenous administration of the contrast agent. Precontrast, T1-weighted, 3D-gradient echo images were acquired with variable flip angle (repetition time: 10 msec; echo time: 2.8 msec; flip angles: 5°, 15°, 30°, 50°, 70°; two averages; matrix: 256 × 256 × 64; field of view: 35 × 35 × 35 mm). Postcontrast images were carried with a single flip angle (15°).

Analysis of Dynamic MRI Data

Pixel by pixel analysis was performed using Matlab software (Math Works Inc.) to generate R1 maps and consequently concentration maps of biotin-BSA-GdDTPA in the grafts and surrounding tissues for selected slices. Vascular properties were derived from the dynamics of contrast agent accumulation [13].

Mapping the Origin of Ovarian Graft Blood Vessels with Fluorescence Microscopy

At 1 wk after transplantation, VE-cadherin-tdTomato ovarian grafts that were grafted on the parietal peritoneum were imaged using a Zoom Stereo Microscope SZX-RFL-2 (Olympus), equipped with a fluorescence illuminator and a Pixelfly camera (PCO), and analyzed using ImageJ software ( http://rsbweb.nih.gov/ij/). To map both host (peritoneal) and graft endothelial cells in functional blood vessels, dextran-fluorescein isothiocyanate (FITC) (500 kDa; Sigma Aldrich) was injected (3 mg/mouse in 0.1 ml PBS) via a tail-vein catheter. The host peritoneal blood vessels were depicted after exposure of the peritoneal surface by the signal of FITC in their lumen (green). Graft-derived vascular endothelial cells were identified by the red signal of the tdTomato protein.

To confirm that the tdTomato signal originated from the graft's endothelial cells, the flow inside the blood vessels was imaged noninvasively in vivo with Laser Speckles Imaging [14]. The endothelial cells were imaged in vivo using the tdTomato expression under the VE-cadherin promoter in transgenic mice. In vivo imaging of the ovary was performed in the ovarian imaging window, a tissue preparation enabling intravital longitudinal imaging of the mouse ovary [15].

To image the endothelial cells together with contrast material inside the lumen of blood vessels, 2 mg of 500  dextran-FITC (Sigma-Aldrich) was injected intravenously into a tail vein through a catheter. The ovarian vasculature was imaged shortly after the injection ex vivo with two-photon microscope (LSM 880 Zeiss Laser Scanning Microscope; Carl Zeiss). The imaging revealed tdTomato fluorescence in endothelial cells and FITC signal inside the lumen.

Histological Assessment of Microvascular Density, Permeability, and Follicle Count

Ovarian graft and surrounding muscle were retrieved for histology at the end of the last serial MRI session, approximately 40 min postinjection of biotin-BSA-GdDTPA. Prior to tissue retrieval, additional contrast material, bovine serum albumin (BSA), labeled with a rhodamine derivative (ROX; Invitrogen) [16], was injected through the tail vein catheter (2.5 mg/mouse in 0.2 ml PBS). Animals were sacrificed 2 min after the administration of BSA-ROX such that it remained inside the lumen of blood vessels with minimal leakage.

To preserve the biotin and fluorescent tags, retrieved tissue was placed in Carnoy fixative solution (6:3:1 ethanol:chloroform:acetic acid) for 24 h at 4°C and then transferred to 70% ethanol. The fixed samples were embedded in paraffin blocks and sectioned serially at 4 μm thickness. Morphometric data was obtained by multilevel sampling of the graft at 120 μm intervals.

Density of functional blood vessels was assessed directly from the fluorescent signal of the intravascular BSA-ROX. Vascular density was derived from five representative histological midsections and normalized by dividing the density of pixels with fluorescent signal above a predefined threshold in the graft to that in the adjacent muscle. Permeability was assessed from the distribution of the extravasated biotin-BSA-GdDTPA, visualized by staining the histological sections with avidin-CY2 conjugate (Jackson Laboratory). Images were obtained with a Zeiss Axio observer microscope equipped with a fluorescence illuminator and an Olympus DP72 camera and analyzed using ImageJ software.

Immunohistochemical staining was used for assessment of maturation of blood vessels according to the degree of coverage with α-smooth muscle actin (αSMA)-expressing perivascular pericytes or vascular smooth muscle cells. After deparaffinization, slides were incubated overnight at 4°C with αSMA antibodies (1:500; Novus Biologicals) and were detected using a second antibody conjugated with alkaline phosphatase (Jackson Laboratory) and its substrate, Fast Red (Sigma). Slides were counterstained with Mayer hematoxylin solution. Slides were examined with a Nikon Eclipse E800 microscope.

Immunoblot Analysis of Akt1 Activation by Simvastatin

C57BL/6 female mice received IP injection of 0.1 or 1 mg/kg simvastatin or saline once daily for 3 days (n = 5 for each treatment group). Protein was extracted on the fourth day from the thigh muscle using meticulous tissue dissociation and phosphatase inhibitors in order to preserve the active phosphorylated form of Akt. Cell lysates were resolved by SDS-PAGE (10%) followed by Western immunoblot analysis, using rabbit polyclonal anti-phosphorylated Ser473 residue of Akt antibody (1:1000; Cell Signaling Technologies). To verify the amount of loaded proteins and inactivated fraction, blots were reprobed with rabbit polyclonal anti-Akt1 antibody (1:1000; Santa Cruz). To explore the effect on eNOS, blots were probed with rabbit monoclonal anti-eNOS (Ser1177) antibody (1:1000; Cell Signaling Technologies).

Statistical Analysis

PS and fBV data were analyzed using repeated-measure ANOVA. The effect of genotype or simvastatin administration was analyzed with simvastatin treatment as a between-subject factor and with the effect of individual mice nested within the treatment factor. Where comparisons were made within a single day, a nested ANOVA model was used with the effect of individual mice nested within the treatment factor. Follicle counts were compared using an independent-samples t-test. Western blot data was analyzed using a one-way ANOVA followed by a Tukey post hoc test. Differences were considered significant at P < 0.05. Results are presented as mean ± SEM. All statistical analyses were performed using Statistica, version 12 (Statsoft, Inc.).

Depicting the Dynamics of Ovarian Graft Reperfusion Using In Vivo Imaging

Graft-induced vascular remodeling was followed longitudinally using a macromolecular blood-pool contrast agent (biotin-BSA-GdDTPA), which extravasates specifically from newly formed hyperpermeable blood vessels. Ovarian grafts showed severe ischemia 2 days after grafting followed by recovery of perfusion 5 days later (Fig. 2, A and B). MRI-derived blood volume (fBV) gradually improved from baseline measurement on Day 2 until 30 days posttransplantation, with an additional increase after hormonal induction of ovulation using eCG/hCG (Fig. 2C). As the graft's newly formed blood vessels matured, mean vessel permeability (PS) declined from Day 2 until Day 30. It modestly increased after eCG/hCG administration (Fig. 2D).

FIG. 2

The dynamics of ovarian graft reperfusion. A–D) Ovarian graft harvested from C57/B wild-type mice were transplanted in the thigh muscle of recipient mouse. Three-dimensional contrast-enhanced MRI revealed the transition of ovarian graft from the ischemic period 2 days after grafting (A), characterized by minimal filling of the graft's vasculature and extravasation of contrast agent from leaky peripheral blood vessel (arrowheads), to reperfusion 7 days thereafter (B). Reperfusion is marked by contrast enhancement in the graft vasculature with minimal leakage to the surrounding muscle. Maximal intensity projections presented here were acquired 40 min after intravenous injection of biotin-BSA-GdDTPA. G = ovarian graft; B = urinary bladder. Bar = 5 mm. MRI-derived vascular parameters reflects the changes in blood volume fraction (mean fBV; C) and vessel permeability (mean PS; D) over time. The graft's new vasculature is characterized by gradual increase in fBV (C) and gradual decrease in PS (D) over time. Pre-MRI on Day 30 was performed prior to eCG/hCG-induced superovulation. Post-Day 33, 14–17 h after hCG administration. Data presented as mean ± SEM (n = 7). E–I) Ovarian grafts with red fluorescent endothelial cells (from a VE-cadherin-tdTomato donor) were transplanted on the recipient peritoneum. Stereomicroscope images acquired 7 days after grafting show that ovarian graft vasculature originates predominantly from the graft's endothelial cells, depicted by the tdTomato fluorescence signal (red; E). Perfused blood vessels in the graft and surrounding peritoneum are highlighted by intravenous injection of dextran-FITC (green; F). Overlay images (G–I) show that most of the grafts vasculature, mainly in the center of the graft, originated predominantly from the graft's own vasculature (red). Some of the vessels were perfused as evident by the colocalization of FITC and tdTomato signals (yellow). A fine network of peritoneal blood vessels (green) invaded the periphery of the graft. H, I) Two representative slides showing ovarian graft blood vessel outgrew (red) toward the peritoneal vasculature (green) to create anastomosis (arrows). Original magnification ×40 (E–I).

To learn more about the origin of the graft blood vessels, ovarian grafts that express tdTomato red fluorescent protein in their endothelial cells (VE-cadherin-tdTomato) (Supplemental Fig. S1; Supplemental Data are available online at  www.biolreprod.org) were grafted intraperitoneally. Most of the graft's vasculature, especially in the center of the graft, originated from the graft. A fine network of recipient-peritoneal blood vessels invaded the graft's periphery (up to one-third of the cross-sectional length of the graft) (Fig. 2, E–G). Most importantly, in all grafts, we observed graft blood vessels that sprouted radially, from the grafts toward the recipient peritoneal vasculature to create functional anastomosis (Fig. 2, H and I).

Selective Activation of Endothelial Akt1 Expression Induces Enhanced Posttransplantation Angiogenic Response in Ovarian Grafts

To explore the role of endothelial Akt1 in graft reception, ovarian grafts with tetracycline-regulated expression of activated Akt1 selectively in endothelial cells (VE-cadherin-myrAkt1) were transplanted into the thigh muscle. Mean fBV was significantly higher from 7 up to 30 days after grafting. Mean fBV increased by almost 400%, compared to the baseline level, on Day 2 and remained relatively constant on Days 7, 14, and 30. Mean fBV in the controls increased by 46%–87% on Days 7–30 (Fig. 3, A–D). Mean PS of VE-cadherin-myrAkt1 grafts was gradually reduced over time, albeit at a slower rate compared to controls and was significantly higher 14 days after transplantation (Fig. 3E). The development of this new vasculature in myrAkt1 was accompanied by a significant increase in graft volume compared to controls (Fig. 3F).

FIG. 3

Changes in fBV, PS, and graft volume over time in myrAkt1 and control grafts. A–C) Maximal intensity projections of dynamic contrast-enhanced MRI in myrAkt1 and control grafts 7 days after transplantation. Images show early vascular filling at the first scan, 5 min after the injection of biotin-BSA-GdDTPA (A), contrast agent extravasation (permeability), and accumulation 40 min postinjection (B), and overlay image of the 5 min (red) and 40 min (green) time points (C). Bar = 5 mm. D) Mean blood volume fraction (fBV) was significantly higher in myrAkt1 grafts compared to controls from Day 7 onward. E) Mean vessel permeability (PS) decreases gradually in both groups. However, PS was significantly higher in myrAkt1 grafts on Day 14. F) Mean volume of myrAkt1 ovarian grafts (measured from MRI) was significantly higher from Day 7 onward (n = 9 for myrAkt1; n = 5 for control). *P < 0.05 by nested ANOVA (D, E) and by independent t-test (F). Data presented as mean ± SEM.

The morphological changes that resulted in these very high fBV values were clearly evident in histological sections. Ovarian tissue, including follicles and stroma, was enclosed by numerous, abnormally appearing blood vessels characterized by large diameter blood lakes and tortuous vessels (Fig. 4A). All myrAkt1 grafts contained antral follicles, and 66% of the myrAkt1 grafts that were assessed were at least partially functional, as was evident by the presence of corpora lutea. The transplantation site consisted of two morphologically distinct blood vessels: 1) numerous normal blood vessels in the muscle adjacent to the graft and 2) large aberrant blood vessels in the graft, congested with erythrocytes. Compared to controls, (Fig. 4, E and F), some of the large stromal blood vessels in myrAkt1 grafts were mature and expressed αSMA, while this marker was not found in some of the immature ones (Fig. 4, C and D). Thirty days after grafting of myrAkt1 grafts, although PS values were similar to controls, some of the newly formed abnormal vessels lacked mural cell support. Using fluorescence microscopy, we recorded high signal intensity of BSA-ROX (indicating perfused vasculature) in abnormally enlarged blood vessels inside myrAkt1 grafts. Furthermore, the area of the grafts that was covered by patent blood vessels was, on average, 72.86% ± 8.50% in myrAkt1 compared to 39.14% ± 6.74% in controls (P < 0.05 by t-test; myrAkt1 n = 3 vs. control n = 6; Fig. 4, G–J).

FIG. 4

Increased vascularity and abnormal growth of irregular blood vessels in myrAkt1 ovarian grafts. A, B) H&E staining 30 days after grafting. Ovarian tissue of myrAkt1 graft, including follicles and stroma, was enclosed in numerous, abnormally appearing blood vessels characterized by large diameter and tortuous vessels (A). Controls showed normal morphology of ovarian graft (B). Arrow = antral follicles, m = muscle. C–F) Alpha smooth muscle actin (α-SMA) staining (red). C, D) Two different areas in myrAkt1 graft showing maturation of large diameter blood vessels in myrAkt1 graft (*) with a thickened wall. Arrowheads = normal blood vessels in the border between the muscle and graft. E, F) Controls, normal blood vessels distribution in the graft and surrounding muscle. Arrowheads = normal blood vessels surrounding growing follicles and inside a corpus luteum. Arrows = antral follicles. G–J) High blood volume in abnormally enlarged vessels inside myrAkt1 graft. H&E staining (G, I). Fluorescence microscopy (H, J): red = BSA-ROX injected 1 min prior to animal sacrifice; green = extravasation of avidin-CY2-biotin-BSA-GdDTPA from permeable blood vessels injected 40 min prior to animal sacrifice and visualized using avidin-CY2. Bars = 500 μm (A, C, D), 0.5 mm (B, G, I), 100 μm (E, F), and 0.2 mm (H, J).

So far, we showed that long-term constitutively active Akt1 improved the vascularity of the graft. Physiologically, pro-angiogenic factors are required during the first week after transplantation. After that period, the graft should be adequately perfused. To assess if transient expression of myrAkt1 can be used to improve ovarian graft reception, expression of myrAkt1 was suppressed 7 days after transplantation (myrAkt1^transient). As expected, 7 days after transplantation myrAkt1^transient grafts had significantly higher fBV and also showed a trend toward higher PS values. At this point in time, myrAkt1 was suppressed by addition of tetracycline to the drinking water; consequently, on the following scans, fBV and PS values returned to a level similar to the controls. The myrAkt1^transient and control grafts responded similarly to eCG/hCG, by increasing their fBV, while PS showed a small decrease and increase in myrAkt1^transient and controls, respectively (Fig. 5, A–D).

FIG. 5

Angiogenic response to transient induction of myrAkt1 in ovarian grafts. A, B) Representative contrast-enhanced MRI of myrAkt1 graft taken 40 min after injection of contrast agent. The high signal intensity and accumulation observed in ovarian graft 7 days after grafting (A) diminished in volume and extent by Day 14, which is 7 days after the suppression of myrAkt1 (B; same graft as in A). Bar = 5 mm. Elevated mean blood volume fraction (fBV; C) and mean vessel permeability (PS; D) in myrAkt1 ovarian grafts 7 days after grafting gradually return to control values after myrAkt1 suppression. Red dashed line indicates time of myrAkt1 suppression. MRI on Day 30 was performed prior to eCG/hCG-induced superovulation and repeated on Day 33, 14–17 h after hCG administration. E, F) Ovarian graft follicle count and follicle density. No differences were found in follicle counts and density 33 days after grafting; n = 5 (transient induction of myrAkt1, myrAkt1trans), n = 9 (controls). CL = corpora lutea. Early-immature follicles up to the secondary stage. *P < 0.05, ⁺P = 0.06 (trend) by nested ANOVA (C, D) and by independent t-test (E). Data presented as mean ± SEM.

Follicular reserve was estimated by multilevel histological sections. We found no differences in primordial, primary, secondary antral corpora lutea and atretic follicle counts and follicle density per cubic millimeter (Fig. 5, E and F). One month after transplantation (3 wk after myrAkt1 suppression), myrAkt1^transient ovarian grafts were adequately perfused and developed extensive vascularity within the graft's stroma (Supplemental Fig. S2).

Improved Ovarian Graft Angiogenesis by Pharmacological Induction of Akt1

In the next step, we tested whether the administration of a 3-hydroxyl-3-methylglutaryl coenzyme A reductase inhibitor, that is, simvastatin, to the recipient can activate Akt1 at the transplantation site and thus improve ovarian graft reperfusion. First, we tested whether Akt1 can be activated in the muscle by the administration of 0.1 and 1 mg/kg simvastatin for 3 days. This treatment caused a 3-fold increase in Akt phosphorylation on Ser473 (P < 0.05). Raised phosphorylated Akt level was correlated with a downstream increase in phosphorylated eNOS (Ser1177). Simvastatin administration caused an approximate 3- and 7-fold increase in phosphorylated eNOS at 0.1 and 1 mg/kg dosage, respectively (P < 0.05) (Fig. 6).

FIG. 6

Induction of activated Akt1 and eNOS in the muscle after 3 days of simvastatin therapy. A) Western immunoblot of p-Akt (Ser473), p-eNOS (Ser1177), and total Akt1 in muscle lysate from c57/B mice treated with daily intraperitoneal injections of 0.1 and 1 mg/kg simvastatin or saline for 3 days. B) Quantitative Western blot densitometry results showing that simvastatin administration caused a 3-fold increase in p-Akt (Ser473) relative to control. C) Simvastatin administration caused approximately 3- and 7-fold increase in phosphorylated eNOS at 0.1 mg/kg and 1 mg/kg respectively (n = 5 animals in each group). *P < 0.05 by one-way ANOVA followed by a Tukey post hoc test. Data presented as mean ± SEM.

Following this successful induction of activated Akt in the transplantation site, we assessed whether low dosage (0.1 mg/kg/day) simvastatin therapy starting 1 day before grafting until 7 days thereafter can improve ovarian graft vascularization. Variations in fBV and PS values of each voxel within the graft were recorded for each animal at 2, 7, and 30 days after grafting. We found a significant effect of simvastatin treatment on improved blood volume fraction in ovarian transplants. On Day 7, mean fBV was significantly higher in the simvastatin group. Grafts treated with simvastatin improved their mean fBV values by 80% ± 5% compared to baseline levels on Day 2, while mean fBV improved by only 21% ± 3% in the control group (P < 0.05 n = 7 in each group). Mean PS was significantly higher on Day 2 in the simvastatin group, which can be explained by augmented angiogenic response and increased permeability of new blood vessels, in contrast to lower mean PS levels on Day 7, which represents accelerated maturation in the simvastatin group (Fig. 7, A and C).

FIG. 7

Augmented vascularization in response to simvastatin treatment. Recipient mice received daily IP injections of 0.1 mg/kg simvastatin or saline (control) starting 1 day before grafting and until 7 days thereafter. A) Simvastatin significantly improved mean blood volume fraction (fBV) from Day 2 after ovarian grafting to Day 7 and in response to superovulation between Days 30 and 33 (B). C, D) Decline in mean PS was found during these periods. Pre = presuperovulation with eCG/hCG; Post = 14–17 h after hCG administration (n = 7 each). *P < 0.05 by nested ANOVA. Data presented as mean ± SEM.

MRI-derived fBV and PS values were also used to dynamically test angiogenic response in ovarian grafts following superovulation with eCG/hCG. In order to support the massive formation of new blood vessels after ovulation induction, the graft needs an adequate scaffold of blood vessels that will be able to perfuse growing follicles and corpora lutea. Mean fBV significantly increased in the simvastatin group compared to prehormonal levels (60% ± 3.20% vs. 28% ± 2.19% increase in simvastatin and control groups, respectively, P < 0.05, n = 7 in each group). This increase in fBV was coupled with a decrease in PS after eCG/hCG (Fig. 7, B and D).

Next, we wanted to assess vascular density in histological sections retrieved at the end point of the study on Day 33 after eCG/hCG. The ratio of pixels with a BSA-ROX signal above a predefined threshold inside the graft, to those in the surrounding muscle, was calculated in five histological sections representing different areas within the graft. Normalized blood vessel density in the grafts treated with simvastatin was 33 ± 5.6 times higher than the muscle, compared to only 16 ± 0.97 times higher in the controls (P < 0.05) (Fig. 8, A–E). This serves as further proof of improved vascular support in ovarian grafts treated with simvastatin. Follicle counts showed an increase in corpora lutea density, while no differences were found in ovarian graft volume. Furthermore, grafts in the simvastatin group had a lower number of atretic follicles (Fig. 8, F and G, and Supplemental Fig. S3).

FIG. 8

Improved ovarian graft vascular and follicle density after simvastatin treatment. Grafts were treated with simvastatin from 1 day before grafting until 7 days thereafter. A, C) H&E stain. B, D) functional blood vessels depicted by fluorescence signal of BSA-ROX (red) injected 1 min prior to animal sacrifice. Dashed line outlines the graft area. Bar = 0.5 mm (A, C) and 0.2 mm (B, D). E) Mean normalized blood vessel density measured by fluorescence microscopy was about 2-fold higher in the simvastatin group compared to control. F) Mean total normal and atretic follicle counts in histological sections. Follicle atresia was significantly higher in the controls (percentage of atretic follicles from total follicle count). G) Mean ovarian graft volume, measured by MRI, was similar between the groups (n = 5–6 grafts in each group). *P < 0.05 by independent t-test. Data presented as mean ± SEM.