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. 2016 Oct 3;22(12):842–851. doi: 10.1093/molehr/gaw064

A grafted ovarian fragment rescues host fertility after chemotherapy

Iordan Stefanov Batchvarov 1, Rachel Williamson Taylor 1,2, Ximena Bustamante-Marín 1,3, Michael Czerwinski 1, Erika Segear Johnson 4, Sally Kornbluth 4, Blanche Capel 1,*
PMCID: PMC6459082  PMID: 27698028

Abstract

STUDY QUESTION

Can host fertility be rescued by grafting of a fragment of a healthy ovary soon after chemotherapy?

SUMMARY ANSWER

We found that grafting a green fluorescent protein (GFP)-positive fragment from a healthy isogenic ovary to the left ovary of a chemo-treated host rescued function and fertility of the grafted host ovary, and resulted in the production of host-derived offspring as late as the sixth litter after chemotherapy (CTx) treatment, whereas none of the ungrafted controls produced a second litter.

WHAT IS KNOWN ALREADY

In women and girls undergoing chemotherapy, infertility and premature ovarian failure are frequent outcomes. There are accumulating reports of improved endocrine function after autotransplantation of an ovarian fragment, raising the possibility that the transplant is beneficial to the endogenous ovary.

STUDY DESIGN, SIZE, DURATION

We first established a CTx treatment regimen that resulted in the permanent loss of fertility in 100% of female mice of the FVB inbred strain. We grafted an isogenic ovary fragment from a healthy female homozygous for a GFP transgene to the left ovary of 100 CTx-treated hosts, and compared fertility to 39 ungrafted controls in 6 months of continuous matings, using GFP to distinguish offspring derived from the graft, and those derived from the host.

PARTICIPANTS/MATERIALS, SETTING, METHODS

Immunofluoresece and western blot analysis of 39 treated ovaries during and 15 days after CTx treatment revealed elevated apoptosis, rapid loss of granulosa cells and an increased recruitment of growing follicles. Using immunofluorescence and confocal imaging, we tracked the outcome of the grafted tissue over 4 months and its effect on the adjacent and contralateral ovary of the host.

MAIN RESULTS AND THE ROLE OF CHANCE

Fifty-three percent of grafted females produced a second litter whereas none of the ungrafted females produced a second litter. The likelihood that this could occur by chance is very low (P < 0.0001).

LIMITATIONS, REASONS FOR CAUTION

These results are shown only in mice, and whether or how they might apply to chemotherapy patients subjected to different CTx regimens is not yet clear.

WIDER IMPLICATIONS OF THE FINDINGS

Our experiments prove that rescue of a chemo-treated ovary is possible, and establish a system to investigate the mechanism of rescue and to identify the factors responsible with the long-term goal of developing therapies for preservation of ovarian endocrine function and fertility in women undergoing chemotherapy.

LARGE SCALE DATA

No large datasets were produced.

STUDY FUNDING/COMPETING INTEREST(S)

Duke University Medical Center Chancellor's Discovery Grant to BC; ESJ was supported by an NRSA 5F31CA165545; SK was supported by NIH RO1 GM08033; RWT was supported by the Duke University School of Medicine Ovarian Cancer Research Fellowship; XBM was supported by CONICYT. The authors have no conflicts of interest to declare.

Keywords: fertility, chemotherapy, transplantation, ovary, granulosa cells, oocytes

Introduction

Advances in chemotherapy (CTx) have had a dramatic impact on cancer survival, and have led to treatments for autoimmune disease and for conditioning before hematopoietic stem cell transplant. However, the cytotoxicity of CTx causes a wide range of side effects (Jensen et al., 2011; Cox and Liu, 2014). In premenopausal females, damage to the reproductive system resulting from chemotherapeutic treatment often leads to ovarian failure, including infertility and loss of ovarian endocrine function. Primary ovarian insufficiency contributes to an increase in all-cause mortality, cardiovascular disease, osteoporosis and neurocognitive deficits, as well as to the vasomotor and sexual dysfunction associated with menopause (Bath et al., 2002; Marder et al., 2012; Cox and Liu, 2014).

Oocyte or embryo cryopreservation in advance of CTx is currently the most successful means of preserving fertility, but hyperstimulation protocols cannot be used for girls before puberty or for patients with estrogen-sensitive cancers (Anderson and Cameron, 2011; Jensen et al., 2011; Gonzalez et al., 2012; Siristatidis et al., 2012; Roque et al., 2013). Other fertility preservation methods with recent advancements include in vitro culture of ovarian tissue, follicles and oocytes (Tingen et al., 2011), and oocyte or ovarian tissue cryopreservation followed by autotransplantation (Waimey et al., 2013; Shapira et al., 2014). More than 40 live births have now been reported after human ovarian tissue cryopreservation and autotransplantation. Investigators are working to establish consistent success rates, and the community has begun the debate to design best clinical practices for this procedure (Donnez et al., 2013, 2015a,b; Salama et al., 2013; Macklon et al., 2014; Dittrich et al., 2015; Donnez and Dolmans, 2015).

An advantage of transplanting cryopreserved ovarian tissue is that it is associated with improved endocrine function, at least partially ameliorating the patient's risk for the complications of ovarian insufficiency (Donnez et al., 2013; Imbert et al., 2014; Stoop et al., 2014; Andersen and Kristensen, 2015; Anderson et al., 2015; Stoop et al., 2015; Kim et al., 2016). In earlier studies, resumption of ovarian function was also reported after autotransplantation of an ovarian fragment to the abdominal wall, and, in one case, a spontaneous implantation occurred in the uterus with full term pregnancy (Oktay and Tilly, 2004; Oktay, 2006). These results suggested that the transplant might be beneficial to the endogenous ovary. However, because resumption of ovarian function after CTx occurs spontaneously at some frequency (Bines et al., 1996), and because offspring from the graft could not be distinguished from offspring from the host, no conclusions could be drawn as to whether the developing embryo came from the endogenous ovary or the graft. This is also the case when cryopreserved ovarian tissue is transplanted to the ovary as an autologous graft. In this situation, it is likely that fertility is recovered based on oocytes in the cryopreserved tissue. However, the possibility that oocytes in the endogenous ovary are partially rescued by the graft cannot be ruled out.

Cell-based therapies have also been reported to improve ovarian function in CTx-treated mice after transplantation of bone marrow (Santiquet et al., 2012), mesenchymal tissue (Takehara et al., 2013) and cells from amniotic fluid (Lai et al., 2013; Xiao et al., 2014). In addition to these surgical interventions, the preconditioning and continual administration of GnRH analogues during CTx has also been investigated as a means to preserve ovarian function (Blumenfeld et al., 2015; Del Mastro and Lambertini, 2015). These results are important because they support the idea that ovarian recovery is possible.

We designed an experimental plan to investigate whether transplantation of a fragment of a healthy ovary could rescue host fertility after chemotherapy. We first established a CTx-treatment regimen that resulted in infertility of treated female mice, and investigated the effects of CTx on the ovary. We then grafted some of the CTx-treated females with a fragment of a genetically marked ovary soon after the end of CTx. We mated grafted and ungrafted females, and compared fertility between them for 8 months.

Materials and Methods

Experimental design overview

One hundred ninety-seven 8-week old FVB females were included in these experiments (Fig. 1). Of these, nine were sacrificed (S) at the outset to serve as untreated controls. The remaining 188 were treated with three doses of CTx at 48-hour intervals. Nine females were sacrificed 12 hours after CTx1 and CTx2 treatments, and 12 were sacrificed after CTx3. Ovaries from these sacrificed females were used for the fluorescent immunocytochemistry and western blot experiments. Six females did not survive the CTx regimen (D).

Figure 1.

Figure 1

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Experimental flow chart. One hundred ninety-seven FVB females participated in this study. The top table shows the number of females that were sacrificed (S) for experimental purposes, or died (D) after each CTx treatment (CTx1, CTx2 and CTx3). One hundred fifty-two surviving females were divided into two groups, 105 females received a graft (grafted - right table), while 47 females did not receive a graft (ungrafted – left table). Surviving grafted and ungrafted animals were placed into continuous matings, and the number that produced a first, second, third, fourth, fifth and sixth litter/total surviving females at that stage is reported. The number of animals that were sacrificed (S) for experimental purposes, or died (D) between grafting and production of each subsequent litter is shown.

Of the 152 animals surviving CTx treatments, 105 received an ovarian graft from a healthy GFP+ 8-week-old isogenic donor on Day 11 after the beginning of CTx (Fig. 1). The graft was placed inside the bursa of the left ovary. The right ovary was not operated, but served as an internal control to determine whether a graft on the contralateral side could rescue both ovaries. If the graft rescued primarily by stimulating the hypothamic/pituitary axis, then both the grafted and ungrafted ovary in the host should be rescued. If the graft had a local paracrine effect, only the left ovary should be rescued. The graft came from an FVB donor, homozygous for a GFP transgene, therefore all offspring derived from oocytes within the graft would be GFP-positive, whereas those derived from the host would be GFP-negative. The GFP transgene was expressed from a ubiquitous promoter, marking any cells emanating from the graft.

Forty-seven of the CTx-treated females did not receive a graft. To investigate whether the surgery alone had an effect on fertility, four of the ungrafted cohort underwent sham surgery. Four ungrafted and four grafted animals were sacrificed for experimental purposes prior to Day 15. One grafted and four ungrafted animals died during this period. On Day 15 after the beginning of CTx, all surviving females (39 ungrafted and 100 grafted) were placed in continuous matings (Figs. 1, 2A, 4A). Figure 1 reports the number of females that produced a first, second, third, fourth, fifth and sixth litter/total surviving females at that stage, for both the ungrafted and grafted cohorts. The number of animals that died (D) or were sacrificed (S) for experimental samples during the 250 day mating period is recorded in Fig. 1. Collected samples were used for fluorescent immunostaining to investigate the state of the graft at various times after grafting, and to histologically compare ovaries from ungrafted animals to the left and right ovaries from grafted animals.

Figure 2.

Figure 2

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CTx-treated females produce only a single litter. (A) Timeline for CTx treatment and mating. Eight-week-old females were CTx-treated by intraperitoneal injection on the first, third and fifth days (see text). On Day 15, treated animals were set up in matings with males. (B) 27% of CTx-treated females did not produce any offspring, while 73% produced one litter averaging eight pups between Day 21–49 after matings began. No subsequent litters were produced.

Figure 4.

Figure 4

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CTx-treated and grafted females produced multiple litters containing host- and graft-derived pups. (A) Timeline for CTx treatment, grafting and mating. On Day 11, host animals underwent surgical grafting of a small ovarian fragment from a GFP-positive syngeneic donor to the host ovary. Animals in this cohort recovered from surgery and were set up in matings with males on Day 15. (B) Grafted animals recovered fertility after CTx treatment and continued to produce litters containing pups derived from both host and graft tissue. Host-derived pups declined over time, but were born up to 220 days after matings were initiated. (C) Litters typically consisted of both GFP-negative host-derived pups and GFP-positive donor-derived pups.

Mice

Mice were maintained in a barrier facility, using an individually ventilated cage system (Allentown PNC) with polysulfonate 75 square-inch cages, and Enrich O'Cob bedding (The Andersons). The light cycle was maintained as 12 hours on and 12 hours off. Water was delivered ad libitum from an automatic watering system (Edstrom) with reverse osmosis water (<1% (by volume) chlorine bleach added). The feed was Purina Lab Diet #5053, irradiated and fed ad libitum. The colony was maintained at a temperature of 22°C ± 0.1°C, with humidity at 30–70%.

FVB mice were acquired from Jackson Laboratory in breeding pairs and expanded in our colony. Host mice were female 8-week-old FVB/NJ. GFP-positive (GFP+/+) donor mice were isogenic female 7- to 8-week-old FVB.Cg-Tg(GFPU)5Nagy/J(EGFP). Donors were obtained and maintained as a homozygous breeding line, validated by test matings with wild type males. As a second GFP donor for cell mixing analysis, we used B6.Cg-Tg(Gt(ROSA)26Sor-EGFP)I1Able/J mice grafted to C57BL6/J hosts. These donor mice were used in two sets of grafting experiments and are not included in the flow chart (Fig. 1). As they cannot be maintained as homozygotes, they were not used in breeding experiments.

Ethical approval

All procedures were carried out according to protocols approved by the Duke University Institutional Animal Care and Use Committee (protocol number A148-14-06) and in accord with NIH guidelines.

Treatment with busulfan and cyclophosphamide (CTx)

We identified a CTx regimen that preserved viability in most treated females, but led to permanent infertility in 100% of survivors (Table S1). The final mixture of the chemotherapeutic agents busulfan (butane-1,4-diyl dimethanesulfonate) and cyclophosphamide (RS)-N,N-bis(2-chloroethyl)-1,3,2-oxazaphosphinan-2-amine 2-oxide) was made by dissolving 8.75 mg busulfan (Sigma USA Cat. No B2635) in 2 ml N,N-Dimethylacetamide (EMD USA Cat. No DX 1544-6) at 45°C. This solution was stable stored at 4°C. Just prior to administration, 25 mg of Cyclophosphamide (Sigma USA Cat#C0768-1G) were dissolved in 1 ml of 0.9% Saline solution at 37°C. One ml of this solution was mixed with 1.5 ml of the busulfan solution. Of this final solution, 200 ul/20 g mouse was injected IP three times at 48-hour intervals, resulting in a dose of 100 mg/kg of cyclophosphamide and 8.75 mg/kg of busulfan per injection (Supplementary data, Table S1, No. 10).

Transplantation of ovary fragments and matings

On Day 11 after initiation of CTx, surgery was performed on host mice to transplant a healthy (not CTx-treated) fragment of an ovary from a GFP+ donor mouse beneath the left bursa. Ovaries were harvested from the GFP+ donor mice and each ovary was divided into four equal fragments (~0.5–1 mm3). A small incision was made in the left ovarian bursa of the host on the side radial to the opening of the oviduct, and a 0.5–1.0 mm3 fragment of the host ovary was removed. For transplantation surgeries, a GFP+ donor ovary fragment was inserted into the bursal sac and rotated away from the opening. For sham surgeries, a small incision was made in the left ovarian bursa of the host on the side radial to the opening of the oviduct, and a 0.5–1.0 mm3 fragment of the host ovary was removed, but no graft was inserted, and the cut edge of the ovary was rotated away from the opening in the bursal sac. The right ovary was not operated. The body wall was closed, and 4 days after surgery (Day 15), all experimental and control females were set up in continuous matings with male mice, independent of estrous cycle (see diagram of experimental flow, Fig. 1).

Fluorescent immunocytochemistry and western blot analysis

Nine females were sacrificed (column ‘S’ in Fig. 1) prior to the initiation of CTx (controls), 12 hours after CTx1, 12 hours after CTx2, 12 hours after CTx3 and 15 days after initiation of CTx. Right and left ovaries were dissected and either fixed and used for immunohistochemistry or pooled and homogenized for western blot analysis.

Fluorescent immunocytochemistry was performed on sections from ovaries collected from at least three independent animals at each time point, for analysis of apoptosis and cell-type-specific markers. Ovary samples were fixed in 4% paraformaldehyde overnight at 4°C and washed in TBS (0.25 M Tris Base, 4.5% NaCl, pH 7.4). For immunofluorescent staining on sections, fixed and washed samples were passed through a 5–20% sucrose gradient, then incubated in a 1:1 mixture of 20% sucrose and OCT overnight at 4°C. Samples were frozen in blocks in liquid nitrogen and stored at −80°C. Samples were cryosectioned, washed and permeabilized with TBS containing 0.1% Triton-X 100 or Tween 20 (TBST) for cleaved-Caspase-3 staining. Slides were blocked in TBST with 10% goat serum for 1 hour at room temperature and incubated with primary antibodies in TBST with 5% goat serum overnight at 4°C. The next day, slides were washed, incubated with secondary antibodies in TBST + 1% goat serum, and 1 ug/ml Hoechst 33342, and mounted in DABCO. Antibodies used were rabbit polyclonal anti-Cleaved Caspase-3 (cCaspase-3) (Asp175 antibody) (9661 S, 1:500, Cell Signaling, USA), goat polyclonal anti-FOXL2 (ovary somatic cells) (NB100-1277, 1:250, Novus Biological, USA), chicken anti-GFP (ab13970, 1:5000, Abcam, USA), rabbit anti-MSY2 (oocytes at all stages) (1:4000 dilution, kindly provided by Richard Schultz) and rat anti-mouse F4/80 (macrophages) (MCA497RT, 1:1000, AbD Serotec, USA).

For western blot analysis, three ovaries from independent females were pooled for each lane, and western blots were performed four times. Ovaries were homogenized in 100 uL RIPA buffer (50 mM Tris Base, 1 mM EDTA, 1% NP-40, 0.25% sodium deoxycholate, 0.1% SDS, pH 7.4) with 100 mM PMSF and 5 ug/mL aprotinin and leupeptin. 50 ug of protein was separated on a 12% SDS PAGE gel, transferred to PVDF membrane, blocked, incubated with primary antibodies (1:1000 dilution in TBST) overnight at 4°C, and incubated with infrared fluorescent secondary antibodies (1:10 000 dilution in TBST) for 1 hour at room temperature. Membranes were washed and analyzed using the Li-Cor Odyssey Infrared Imaging System (Lincoln, NE, USA). Antibodies used were rabbit polyclonal anti-cCaspase-3 (Asp175 antibody; 1:1000) (9661 S, Cell Signaling, USA), rabbit polyclonal anti-PARP (1:1000; 9544 P, Cell Signaling, USA) and mouse monoclonal β-Actin (sc-47778, 1:1000; Santa Cruz Biotechnology, USA). Secondary antibodies used at 1:10 000 were IRDye® 800CW goat (polyclonal) anti-mouse (Odyssey, USA) and Alexa Fluor® 680 goat anti-rabbit (Invitrogen, USA).

To follow the status of the graft, three ovaries were collected from grafted females at 15, 30 and 120 days after the initiation of CTx (Fig. 1). Whole mount imaging of ovaries grafted with GFP+ ovarian fragments was performed on live, unfixed tissue. Immediately after dissection, whole ovaries were incubated for 15 minutes in PBS with 0.1% Triton-X 100 and 1 ug/ml Hoechst 33342 at RT. The staining solution was removed and replaced with fresh PBS for imaging. After whole mount imaging, grafted ovaries were processed through a sucrose gradient, cryosectioned (as described above) and stained with anti-cCaspase-3, chicken anti-GFP and anti-MSY2.

Oocyte size measurements

Ovaries from three independent females/stage were sectioned at a thickness of 10 µm, and every 10th section from each sample was stained with rabbit anti-MSY2 to facilitate collection of data across the entire mass of the ovary while avoiding regional bias or double counting of oocytes observed in adjacent serial sections. Images of entire ovary sections, ranging in resolution from 2000 to 4000 pixels, were collected through automated stitching of 20× magnification image grids overlapping by 10% across each section. Unsupervised oocyte size measurements were collected using Fiji (ImageJ distribution) by applying an auto threshold v1.15 (Yen method) followed by the ‘analyze particles’ command to find oocyte area in pixels. The analyze particles command identified coherent circular patches of high pixel density, calculated the total number of pixels in that two dimensional space, and used this measurement as a proxy for the diameter of an oocyte in cross section. To reduce noise, patches with area measurements below a cutoff value determined by manual measurement of oocytes in primordial follicles were removed from the data set. This unbiased machine approach likely underestimated the size of some oocytes that were not captured in cross section, and some oocytes with weak diffuse staining were missed. However, these cases would tend to underestimate the significance of our results. The oocyte area measurements were taken from samples collected from untreated control mice and treated mice 12 hours after CTx1, CTx2, CTx3 and 15 days after CTx1. Measurements were used to perform a one-way analysis of variance of oocyte area by treatment stage. Analysis to determine if any samples differed significantly from the control was done by comparison of sample means using the Tukey test.

Histology and immunohistochemistry

Experimental grafted (left) ovary and the control (right) ovary of individual grafted mice were compared to age-matched samples of ungrafted mice treated with CTx. Samples from ungrafted animals were collected in parallel with samples from grafted animals on Day 15 and between each litter. Samples were fixed in Bouin's and embedded in paraffin. Serial sections of 8 µm were subjected to antigen retrieval by submerging in sodium citrate buffer (10 mM sodium citrate, 0.05% Tween, pH 6.0) at 90°C for 20 minutes. After antigen retrieval sections were stained for MSY2 similar to our immunofluorescence protocol, and positive cells were detected using a peroxidase conjugated goat-anti rabbit IgG (Jackson ImmunoResearch, USA: 111-036-003; 1:500). Staining was achieved using the standard protocol for the Vector DAB Peroxidase Substrate Kit (Vector Labs, USA: SK-4100). The reaction was stopped with water and the slides were counterstained lightly with hematoxylin. The left grafted ovary and right control ovary were compared qualitatively for the presence/absence of follicles 8 weeks after the initiation of CTx.

Detection of cell mixing between the graft and host ovary

Cells derived from the grafted GFP+ ovarian fragment could be identified by ubiquitous expression of the GFP transgene. Cell mixing between the graft and the host ovary was investigated on Day 15 (4 days after grafting), Day 30, and Day 120. Three FVB females grafted with tissue from an FVB.Cg-Tg(GFPU)5Nagy/J (EGFP) donor were sacrificed at each time point, and fluorescent immunocytochemistry for native GFP was used to detect cells from the graft, combined with antibodies against cCASPASE3, to detect cells undergoing apoptosis, and MSY2 to detect oocytes at all stages of development. FVB.Cg-Tg(GFPU)5Nagy/J (EGFP) mice express GFP ubiquitously in all cells. However, they do not express GFP at uniformly high levels in all cells as is evident from Fig. 5. To verify that we were not missing a significant migratory population because GFP was not expressed in cells from the graft at high enough levels for detection, we used a second GFP+ donor graft/host combination (B6.Cg-Tg(Gt(ROSA)26Sor-EGFP)I1Able/J mice grafted to C57BL6/J hosts). This transgenic expressed GFP at lower levels (requiring the use of an antibody against GFP for detection), but expression was uniform in all cell types in the ovary.

Figure 5.

Figure 5

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Grafts occupied a larger proportion of the ovary over time. (A) A representative sample at 15 days (4 days after surgery) showed a small GFP+ fragment (green) weakly attached to the left grafted host ovary, and a right ungrafted host ovary similar in size to the left. (B) By 30 days after CTx, the graft was firmly connected to the left grafted host ovary, while the right ungrafted host ovary was slightly smaller. C. By 120 days after CTx, much of the left grafted ovary was taken over by graft tissue, while the right ungrafted ovary was much smaller. Fluorescent images are at equal magnification. (D–F) Representative samples at 15 days (D), 30 days (E) and 120 days (F) after the beginning of CTx sectioned and stained with DAPI (blue) to detect nuclei, antibodies against GFP (green) to detect graft tissue, and cCaspase-3 (red, D,E) to detect apoptosis, or MSY2 (red, F) to detect oocytes. Higher magnification images of boxed regions in D, E and F (bottom row) show the junction between the graft and host tissue. A few cells from the graft are found in proximate host tissue, although graft and host tissue do not intermingle extensively. Note that MSY2+ oocytes are still present in host tissue (GFP-negative).

Statistical analysis

JMP Pro v12.0.1 software was used to compare the frequency of appearance of a second litter between the grafted and ungrafted groups. JMP Pro v12.0.1 software was also used to determine statistical significance for the changes in oocyte size across the CTx treatment period and 15 days post-treatment initiation. We ran a Tukey test on the comparison of means between all treatment stages and the control. Sample means were considered significantly different if the P-value was ≤0.05.

Results

Establishment of a viable experimental cytotoxic treatment that invariably results in infertility

We established a treatment regimen that resulted in infertility in all treated females while preserving viability in most animals. We originally tested a scheme based on a published report (Shiromizu and Mattison, 1984), but FVB females treated at 6–8 weeks of age spontaneously recovered fertility after this treatment (see Supplementary data, Table S1 for details of other treatments tested). Using a dosage of 100 mg cyclophosphamide and 8.75 mg busulfan per kg of body weight administered intraperitoneally three times at 48-hour intervals, 152/158 (96%) females that were not intentionally sacrificed survived to Day 11 (Figs. 1, 2A). Only six CTx-treated females died (Fig. 1, top, column D) using this dosing regimen. Of survivors, 27/37 (73%) produced a first litter and invariably produced no further offspring (Fig. 1, left table). These single litters averaged eight pups delivered between Day 32 and Day 78 after the initiation of treatment (Fig. 2B). Treated females were continuously mated for as long as 10 months after CTx, and none (0) spontaneously recovered fertility. Survival of these females was recorded at the time of each subsequent litter in parallel with grafted females (Fig. 1).

CTx treatment leads to apoptosis of granulosa cells and stimulation of oocyte growth

Using fluorescent antibodies to detect apoptosis, including TUNEL (data not shown), cleaved PARP (cPARP) (data not shown) and cCaspase-3 on cryosections from ovaries of untreated and CTx-treated females, we rarely detected apoptosis in oocytes. Instead we observed an increase in apoptotic staining in granulosa cells (Fig. 3A).

Figure 3.

Figure 3

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Large numbers of granulosa cells are lost by apoptosis during CTx, and many follicles/oocytes initiate growth. (A) Immunofluorescence on sections of control (Ctrl) and CTx-treated ovaries 12 hours after CTx1, CTx2, CTx3 and on d15 after the beginning of treatment. Samples were stained with anti-FOXL2 (green), a marker of granulosa cells, cCaspase-3 (red), a marker of cell death and DAPI (blue) to detect nuclei. Images are representative from three biological replicates. (B) Elevated levels of cCaspase-3 (cC3), full length (FL), and cleaved (CL) PARP were detected by western blot at 12 hours after CTx1 and CTx2, relative to controls (Ctrl). (C) The number of growing follicles increased during CTx treatments. Samples are stained with anti-MSY2 (red), a marker of oocytes and DAPI (blue) to detect nuclei. An increase in growing follicles was observed by 12 hours after CTx2. After CTx3, many follicles with only 1–2 layers of cuboidal granulosa cells (inset, bottom, yellow arrowhead) contained an oocyte comparable in size to that of an antral follicle (inset, top, outlined oocyte indicated by white arrowhead). Images are representative from three biological replicates. (D) Average oocyte size (see Methods) increased from control (C) to a maximum 12 hours after CTx3. *Oocyte sizes in CTx3 and d15 samples are significantly different than controls, P < 0.001 and P = 0.0067, respectively.

Consistent with immunocytochemical detection of cCaspase-3, we found elevated levels of cCaspase-3 and cPARP by Western analysis 12 hours after CTx1 and 12 hours after CTx2 (Fig. 3B). Staining with the macrophage marker F4/80 on Day 15 after the initiation of CTx revealed a high incidence of macrophage invasion inside and outside follicles (Supplementary Fig. S1).

Analysis of MSY2 immunostaining in cryosections from ovaries 12 hours after each CTx treatment (CTx1, CTx2, CTx3) and on Day 15 after initiation of treatment revealed a disproportionally large number of growing oocytes in atypical follicles (Fig. 3C). Because of their atypical structure, we could not classify follicle stages based on traditional histological methods. Instead, we measured average oocyte size 12 hours after each of the three CTx treatments and on Day 15 (see Methods) as an indicator of oocyte activation. Twelve hours after the third treatment, many oocytes had reached a size typical of oocytes in antral follicles (Fig. 3C,D), yet most resided in abnormal follicles comprised of only 1–2 layers of granulosa cells (see example in Fig. 3C, inset).

Transplantation of a healthy ovarian fragment rescued fertility of the CTx-treated host

Having established a robust and reproducible protocol for CTx treatment that invariably resulted in infertility, we investigated whether transplantation of a fragment from a normal ovary into the ovarian bursa at the end of CTx-treatment could rescue the host ovary. In preliminary experiments, we empirically determined that Day 11 was the earliest day after CTx that mice were strong enough to survive surgery. Because we observed a negative correlation between the time to surgery and rescue of fertility in a limited number of trials, we performed all surgeries on Day 11.

Four days after surgery, all ungrafted and grafted females were placed in continuous matings with virile males to test long-term fertility. In contrast to ungrafted controls (Fig. 2B), CTx-treated females that received a fragment of a GFP-positive ovary produced multiple litters (Fig. 4B). Using a Fisher's Exact Test, the frequency of second litter appearance between groups was significantly different (P < 0.0001) and the likelihood of the occurrence of a second litter was significantly higher in the grafted group (P < 0.0001). We found no differences between CTx-treated mice that received sham surgery but no graft, and mice that were not operated. Consequently, these mice were all considered part of the ungrafted cohort.

The second and subsequent litters were comprised of pups that were both GFP-negative, therefore derived from the host ovary, and GFP-positive, therefore derived from the donor ovary fragment (Fig. 4C). CTx-treated and grafted females produced up to six litters containing both host-derived and graft-derived pups (Fig. 4B,C; Supplementary Fig. S2). Of 100 CTx-treated and grafted females set up in matings, 82 (95%) produced a first litter averaging slightly fewer than seven pups, of which an average of six were host-derived and one was donor-derived (Supplementary Fig. S2). The time of delivery of the first litter averaged 32 days after the initiation of matings (Fig. 4). Of the 66 CTx-treated and grafted females set up in second matings, 39 (59%) produced a second litter an average of 36 days later, averaging six animals, of which approximately half were derived from the CTx-treated host ovary (Fig. 4, Supplementary Fig. S2). The average time between litters was 36 days between litters 2 and 3, 33 days between litters 3 and 4, 23 days between litters 4 and 5 and 36 days between litters 5 and 6. The average size of the third, fourth, fifth and sixth litters progressively declined as did the proportion of host derived offspring. However, 7/10 (70%) of the grafted animals set up in fifth matings produced a fifth litter. In this group, one female produced 3/4 host derived pups, another produced 4/4 host derived pups, and the remaining five females produced litters entirely derived from the graft (Figs. 1, 4B; Supplementary Fig. S2). In addition, 1/5 surviving grafted animals produced a sixth litter of one host-derived and one donor-derived pup nearly 6 months after initiation of matings. There was no phenotypic evidence of mutations in the offspring, which proved capable of breeding and producing healthy progeny.

The graft rescued only the adjacent grafted ovary

Our experiments were designed to test whether a graft of a normal ovary fragment could rescue fertility by re-establishing signaling between the ovary and the hypothalamus. If this were the case, we would expect both left and right ovaries to be rescued. Ovaries were dissected from host females at sequential times after grafting. On Day 15 (4 days after grafting), four animals were sacrificed from grafted and ungrafted cohorts (Fig. 1) to investigate the effects of CTx on the host at this time point (as described previously), and to evaluate whether the graft adhered to the host ovary. The GFP-positive grafted tissue was evident attached to the left ovary, and both left and right ovaries were similar in size (Fig. 5A).

On Day 30, the left and right ovaries were still similar in size, but the graft tissue comprised a larger proportion of the grafted ovary than on Day 15 (Fig. 5B). By 120 and 130 days after grafting, a dramatic size difference was evident between the ungrafted (right) and grafted (left) ovaries (Fig. 5C). Growing follicles with oocytes at all stages of development were visible throughout the grafted left ovary, but no follicles were evident in the right ovary, indicating that the mechanism of rescue is not systemic. Although the proportion of the total left ovary occupied by the graft continued to increase, and varied from ~50% to 90% in individual animals examined at 120–130 days after CTx, offspring derived from the host ovary were still produced >160 days after CTx (Fig. 4B; Supplementary Fig. S2) indicating that ovulations were still occurring from some host ovaries at these stages.

Differences between the grafted (left) and ungrafted (right) ovary in individual females were analyzed using an antibody against MSY2 to label all oocytes. A representative experiment is shown at 8 weeks after the beginning of CTx (Supplementary Fig. S3). Qualitative assessment showed primordial, primary, secondary and mature follicles throughout the grafted (left) ovary, whereas the ungrafted (right) ovary was dominated by corpora lutea, vacuoles and fibrous stroma.

Intermingling of graft and host tissue was limited

On Day 15, just 4 days after grafting, a tenuous connection between the host and the graft was evident (Fig. 5). On Day 30, the grafted tissue was firmly adhered to the host ovary, and a few GFP+ graft-derived cells were typically detected around host follicles near the border with the graft. At 120 days after the beginning of CTx, graft and host tissue are more integrated, but regions of host and graft cells were still quite distinct. At the two early time points, cCASPASE3+ cells undergoing apoptosis were distributed throughout host tissue, proximate and distant to the graft. Using a second GFP+ transgenic strain to validate these observations, results were very similar: on Day 30, graft-derived cells were typically detected in the theca layer of host follicles near the border with the graft, whereas MSY2+ host oocytes were located proximate and distant to the graft (Supplementary Fig. S4).

The graft may improve lifespan of the host

We compared survival in a subset of the original cohort of CTx-treated hosts continuously mated and followed for a year. In this cohort, 14/42 females were not grafted and 28/42 received grafts. Survival rate was similar until ~5–6 months post treatment. Only a single mouse of the original ungrafted cohort (1/14) survived beyond 190 days, while 9/28 of the original grafted cohort survived beyond this time point. Eight mice of the grafted cohort survived beyond the single remaining control mouse, which died at 204 days, and three of the grafted group survived beyond 270 days post-treatment. Although these results suggest that the graft had a positive effect on longevity, a larger cohort would be required to determine significance of this trend.

Discussion

In the last few decades, significant advances in CTx treatments have led to a continuous increase in the survival rate of patients, many of whom experience premature ovarian failure. We have developed a system of ovarian damage and rescue and collected new data that adds to growing evidence that fertility and function of the ovary can be preserved after CTx. Partial rescue of ovarian function is associated with transplantation of cryopreserved ovarian tissue (Donnez et al., 2013; Imbert et al., 2014; Stoop et al., 2014, 2015; Andersen and Kristensen, 2015; Anderson et al., 2015; Kim et al., 2016), and has been reported after intraperitoneal or direct ovarian injection of various stem cell populations including mesenchymal stem cells from multiple sources (Fu et al., 2008; Takehara et al., 2013), amnionic fluid stem cells (Zhang et al., 2015) and bone marrow stem cells (Fu et al., 2008; Santiquet et al., 2012). The model we have developed provides an opportunity to investigate the mechanisms of ovarian rescue and the factors/signals that mediate this effect.

Among ovo-toxic treatments commonly in use are alkylating agents (cyclophosphamide, busulphan and dacarbazine), platinum complexes (cisplatin, carboplatin) and taxanes (paclitaxel). Most of these lead to DNA breaks and/or trigger apoptosis through effects on microtubules (Roness et al., 2016). It was previously believed that germ cells are the direct targets of cytotoxic agents, leading to the collapse of all follicles (Hutt et al., 2013). However, agents that damage DNA are most toxic to dividing cells. Because oocytes are arrested in meiosis during follicular growth and are not undergoing cell division, we and others (Lopez and Luderer, 2004; Morgan et al., 2012; De Vos et al., 2014) have hypothesized that CTx is most harmful to dividing granulosa cells in growing follicles, and that oocyte loss is secondary to granulosa cell apoptosis. Consistent with previous reports (Freiesleben et al., 2010; Lopes et al., 2014), we find that, rather than acting directly on oocytes, the CTx regimen we tested leads to the apoptosis of granulosa cells in growing follicles, which causes a secondary loss of oocytes. Whether grafting can rescue the ovary after other classes of cytotoxic treatments (e.g. platinum complexes and taxanes) remains to be tested.

Our results suggest that apoptosis of granulosa cells in growing follicles leads to the recruitment of more follicles into the growth phase, which could explain the sequential loss of the primordial follicle pool after several rounds of CTx, a hypothesis referred to as ‘follicle burn out’ (Kalich-Philosoph et al., 2013). The abrupt activation of many growing follicles may be responsible for the single post-CTx litter born to CTx-treated animals, while others are presumably eliminated through cell death pathways and/or phagocytosis by resident leukocytes. It is not yet clear how grafting a fragment of a healthy ovary rescues the host. We originally anticipated that the graft might re-establish functional signals between the ovary and the hypothalamus. The average time between subsequent litters of CTx-treated, grafted females ranged between 23 and 36 days, as compared to an average of 30 ± 2 days in untreated FVB females. These data suggest a return to relatively normal estrus cycles albeit with more variability. Because we did not rigorously control for the time of removal of pups from the previous litter, these data may be influenced by delayed implantation due to ongoing lactation (McLaren, 1968). However, even if the graft exerts a systemic effect, this alone is insufficient to rescue the right (ungrafted) ovary, which degenerates.

It is possible that cells from the graft intermingle with the host ovary, and afford cellular rescue through signaling or replacement of damaged host cells. Amniotic fluid stem cells transplanted directly into CTx-treated ovaries were proposed to rescue by secretion of factors that mediate follicle survival (Zhang et al., 2015), similar to adipose derived mesenchymal stem cells, hypothesized to promote vascularization and growth factor-mediated repair (Takehara et al., 2013). In the present study, we find some cells from the graft located in theca layers of follicles near the site of the graft. However, we note the growth of apparently healthy follicles distant to the site of the graft, with no evidence of the presence of donor cells. Similar to previous reports of ovarian rescue with genetically labeled donor cells (Santiquet et al., 2012; Takehara et al., 2013), there was no evidence in this study for direct replacement of granulosa cells or oocytes derived from the graft. Our results are consistent with a mechanism of rescue that involves diffusible paracrine substance(s) secreted from the donor ovary tissue.

Previous intraovarian transplantation of primordial follicles failed to rescue CTx-treated ovaries (Park et al., 2013), which could suggest that the source of the rescue substance is the ovarian stroma or growing follicles. It would be very interesting to know whether ovary fragments that have been cryopreserved also rescue the host ovary; however, autologous grafts in patients preclude this analysis. The position or timing of the graft after CTx-treatment could be critical. Grafting as soon as possible after CTx was correlated with an improvement in fertility in our experiments, and results from testing extended CTx regimens suggested that the window for rescue was finite.

In this study, we focused on demonstrating that ovarian rescue can occur. We did not attempt to identify the substance(s) (hormones or signaling molecules) secreted from the donor ovary tissue. Because there are many possibilities, further experiments to determine whether grafting prevents apoptosis of granulosa cells (perhaps by blocking their entry into the cell cycle), promotes the formation of new granulosa cells, or favorably affects the vasculature or neuronal circuits in the ovary could help refine a list of candidates.

Rescue of the ovary during CTx would raise questions about damage to the resident oocytes and the possibility that offspring would carry a heavy mutational load. However, we found no obvious phenotypic mutations in >720 offspring born from CTx-treated females, and were routinely able to breed from these animals, consistent with a previous study (Takehara et al., 2013). Further investigation is necessary to determine whether oocytes in a rescued ovary carry an elevated mutational load, and, if not, how repair is being effected. A recent report showed that oocytes are capable of significant repair after DNA damage (Kerr et al., 2012).

Enhanced survival of a female receiving an ovarian graft has been reported previously (Cargill et al., 2003; Mason et al.,, 2009; Mason et al., 2011). The fact that some grafted females survived longer than ungrafted females in this study may be due to the beneficial effect of ovarian function on female physiology. An understanding of the mechanism by which the healthy grafted tissue prevents the failure of the damaged endogenous ovarian tissue could lead to important therapies for the preservation of ovarian endocrine function and fertility in women undergoing chemotherapy.

Supplementary data

Supplementary data are available at http://molehr.oxfordjournals.org/.

Supplementary Material

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Acknowledgements

We are grateful to many members of the lab for helpful suggestions in the course of this work and to Jennifer McKey for careful reading of the manuscript.

Authors’ roles

B.C. designed the experiments; I.S.B. and R.W.T. conducted CTx and grafting experiments; X.M. and R.W.T. performed histology; X.B.-M. and M.C. performed immunocytochemistry studies; S.K. and E.S.J. performed Western analysis; all authors contributed to the writing of the manuscript.

Funding

Duke University Medical Center Chancellor's Discovery Grant to B.C.; E.S.J. was supported by an NRSA 5F31CA165545; S.K. was supported by NIH RO1 GM08033; R.W.T. was supported by the Duke University School of Medicine Ovarian Cancer Research Fellowship; X.B.-M. was supported by CONICYT.

Conflict of interest

None declared.

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