Generation of Miniaturized Ovaries by In Vitro Culture from Mouse Gonads

Pregnant female ICR mice at 12.5, 13.5, and 14.5 dpc were employed to isolate gonad for dissociation experiments. The following reagents and chemicals were procured from respective suppliers: Dulbecco’s modified Eagle’s medium (1×) (DMEM; 11965-092, Gibco, Grand Island, NY, USA), phosphate-buffered saline (PBS; Gibco, cat. no. 10010-023), Minimum Essential Medium Eagle, alpha modification (1×) (α-MEM; cat. no. 12571-063 and 32571-036, Gibco), 2-mercaptoethanol (55 mM; cat. no. 21985-023, Gibco), fetal bovine serum (FBS; cat. no. 16000-044, Gibco), Pen Strep Glutamine (100×) (cat. no. 10378-016, Gibco), ascorbic acid (150 mM; cat. no. G0394, TCI), ICI182 and 780 (cat. no. 1047, Tocris), follicle-stimulating hormone (FSH; cat. no. F4021, Sigma-Aldrich, St. Louis, MO, USA), epidermal growth factor (EGF; cat. no. 2028-EG-200, R&D Systems, Minneapolis, MN, USA), polyvinylpyrrolidone (PVP; cat. no. PVP360100G , Sigma-Aldrich), human chorionic gonadotropin (1,200 IU mL–1) (hCG; cat. no. C8554, Sigma-Aldrich), collagenase type IV (1 g mL–1; cat. no. 17104-019, Gibco), trypsin-EDTA, 0.25% (cat. no. 25200-072, Gibco), StemPro-34 SFM (1×), liquid (cat. no. 10639-011, Gibco), and hyaluronidase (cat. no. D8418, TCI).

The following list of equipment was employed to perform the experiments: Clean bench (cat. no. JSCB-1500SB, JSR), water bath (cat. no. JSWB-22T, JSR, Gongju, Korea), CO₂ incubator ( JP/SMA-165D , Astec, Seoul, Korea), stereoscopic microscope (cat. no. CKX53, Olympus, Tokyo, Japan), a suction machine (cat. no. TFS-30, Unim, Gunpo, Korea), centrifuge (cat. no. KLG122618050210, Labogene, Allerød, Denmark), a hemocytometer (cat. no. 2960408,Paul Marienfeld, Lauda-Königshofen, Germany), micropipettes (cat. nos. EP3120000011, EP3120000020, EP3120000038, EP3120000046, EP3120000054, EP3120000062, Eppendorf, Hamburg, Germany), sterile plastic tips (10 μL, 200 μL, 1,000 μL; Thermo Fisher Scientific, Waltham, MA, USA), pipettors, sterile plastic pipettes (5, 10, 25, and 50 mL), 15-mL conical SW Jang, HW Choi centrifuge tubes (cat. no. 338652 or equivalent, Thermo Fisher Scientific), 50-mL conical centrifuge tubes (cat. no. 338652 or equivalent, Thermo Fisher Scientific), four-well culture plates (cat. no. 176740, Thermo Fisher Scientific), Petri dish 35 mm (cat. no. 353001, BD-Falcon), Petri dish 100 mm (cat. no. 353003, BD-Falcon), 24 well-tissue culture plates (cat. no. 3526, Costar, NY, USA), Transwell-COL membrane inserts (cat. no. 3496, Costar), sterile syringes (0.5 mL; cat. no. 328821, BD, New Jersey, USA), tungsten wire (cat. no. W-461387, Nilaco, Tokyo, Japan), cell strainer (40, 70, 100, 150 μm ; cat. nos. 93070, 93100, SPL, Pocheon, Korea), and 35 mm hexagon U-bottom plate (cat. no. H2951000, StemFIT 3D, Hanam, Korea).

No statistical methods were used to predetermine sample size. The experiments were not randomized. The investigators were not blinded to allocation during experiments and outcome assessment.

All methods were performed in accordance with relevant guidelines and regulations of Jeonbuk National University. All experimental protocols were approved by the Institutional Animal Care and Use Committee of Jeonbuk National University (approval numbers CBNU-2019-00395). Pregnant female outbred albino ICR mice were purchased from Company of Samtako (Osan, Korea). Pregnant females were killed by cervical dislocation to obtain E12.5, E13.5 and E14.5 embryos. Only from female embryos, the gonads were isolated and then dissociated for generation of miniaturized ovaries. This study was carried out in compliance with the ARRIVE guidelines (http://www.nc3rs.org.uk/page.asp?id=1357).

Female fetal gonads without mesonephros were cultured in Transwell-COL membranes (3.0-μm pore size, 24-mm diameter, Corning, Corning, NY, USA) soaked in α-MEM-based IVDi medium (α-MEM supplemented with 2% FBS, 150 μM ascorbic acid, 1× penicillin/streptomycin/ L-glutamine, and 55 μM 2-mercaptoethanol). From days 5 to 11 of culture, 10 μM ICI182780 was added to the α-MEM-based IVDi medium (henceforth, referred to as α-MEM). On day 4 of culture, the culture medium was changed to StemPro-based IVDi medium (StemPro-34 SFM supplemented with 10% FBS, 150 μM ascorbic acid, 1× penicillin/streptomycin/L-glutamine, and 55 μM 2-mercaptoethanol). From days 7 to 10 of culture, 500 nM ICI182780 was added to the Stempro-34-based IVDi medium (henceforth, referred to as StemPro). On day 21 of culture, individual secondary follicles (2FLs) were manually dissociated using sharpened tungsten needles. Gonads were cultured for 21 days at 37℃ under 5% CO₂ and 95% air. Approximately half of the medium in each well was replaced with fresh medium every other day.

Electrically sharpened tungsten needles were used to isolate the individual 2FLs. The interstitial cells between 2FLs were carefully removed. Then, 2FLs were separated from the ovaries and placed at regular intervals.

Single 2FLs on Transwell-COL membranes were soaked in IVG-α-MEM medium (α-MEM supplemented with 5% FBS, 2% polyvinylpyrrolidone, 150 μM ascorbic acid, 1× penicillin/ streptomycin/L-glutamine, 100 μM 2-mercaptoethanol, 55 μg mL^–1 sodium pyruvate, 0.1 IU mL^–1 FSH, 15 ng mL^–1 BMP15, and 15 ng mL^–1 GDF9). After 2 days of culture, the medium containing BMP15 and GDF9 was withdrawn, and follicles were then incubated in 0.1% type IV collagenase. After washing several times with α-MEM supplemented with 5% FBS, the follicles were cultured in IVG-α-MEM without BMP15 and GDF9. On day 11 of culture, cumulus-oocyte complexes grown on the membrane were picked using a fine glass capillary, cultured on Transwell-COL or Millicell membranes for another 9–13 days at 37℃ in medium, under 5% CO₂ and 95% air. Then, 1 mL of medium was added to the inside and outside of membrane inserts. Approximately half of the medium in each well was replaced with fresh medium every other day.

Cumulus-oocyte complexes were transferred to IVM medium (α-MEM supplemented with 5% FBS, 25 μg mL^–1 sodium pyruvate, 1× penicillin/streptomycin, 0.1 IU mL^–1 FSH, 4 ng mL^–1 EGF, and 1.2 IU mL^–1 hCG ). After 16 h of culture, swollen cumulus cells were stripped from oocytes by treatment with hyaluronidase, and then MII oocytes were determined by first polar body extrusion.

In brief, oocytes were fixed for 4 h at 20℃ in 4% paraformaldehyde. The cells were then washed with PBS and treated with PBS containing 10% normal goat serum and 0.03% Triton X-100 for 45 min at room temperature. The primary antibodies used were anti-Ddx4 (polyclonal, 1:500; Abcam, Cambridge, MA, USA), anti-Oct3/4 (monoclonal, 1:500; Santa Cruz Biotechnology, Dallas, TX, USA), and GDF9 (polyclonal, 1:500; Invitrogen, Carlsbad, CA, USA). For detection of primary antibodies, fluorescently labeled (Alexa Fluor 488 or 568; Molecular Probes, Eugene, OR, USA) secondary antibodies were used according to the manufacturer’s specifications. All immunofluorescence samples were analyzed using a confocal microscope (LSM700, Carl Zeiss, Oberkochen, Germany).

All experiments were performed in triplicate, and generation of miniaturized ovaries using each method, the mean numbers of intact follicles per ovary was calculated by an arithmetical calculation.

In our early experiments, female gonads obtained from E12.5–E14.5 mice were cultured in α-MEM or StemPro medium for 5 weeks on Transwell-COL membranes. First, we assessed the efficiency of follicle formation in the two culture media. Two control groups were prepared: one cultured using the α-MEM-based culture medium (Morohaku et al., 2016) and the other cultured in StemPro medium (Hikabe et al., 2016). On employing the two cultures for preserving gonads, the formation of follicles was assessed, and secondary follicle formation was observed (Fig. 1A and B). Follicles generated using two different media gradually matured and produced oocytes (Fig. 1B). The in vitro oocytes expressed Oct4 and Ddx4, as well as the oocyte-specific marker GDF9 (Fig. 1C). Oct4 is expressed in the oocyte core, whereas Ddx4 is expressed on the oocyte (Noce et al., 2001; Saitou et al., 2002; Tanaka et al., 2004; Okamura et al., 2008; Pisarska et al., 2011; Sabour et al., 2011). These data revealed that, in both media, the cultures could form normal functional oocytes. Collectively, the gonads from the period between 12.5, 13.5, and 14.5 dpc could develop into follicles in vitro on Transwell-COL membranes, irrespective of the medium employed. The follicle formation efficiency did not significantly differ between the two types of culture media. However, in vitro ovaries in StemPro medium contained considerably more follicles, demonstrating more rounded granulosa cells in the IVG medium than those in the α-MEM medium. Additionally, we confirmed that the granulosa cells were further expanded during the IVG process using StemPro-based medium (Fig. 1D). Therefore, we used StemPro medium to perform all further experiments.

Fig. 1. Optimized culture condition for oocyte formation. (A) Ovary culture. Representative images of ovaries at two weeks of culture in the α-MEM-based medium and StemPro-based medium (scale bar, 200 μm), (B) Immunostaining results for the follicular expression of Ddx4 in vitro, (C) Oocyte phase and expression of the Ddx4, Oct4, and oocyte-specific marker GDF9 in vitro (immunochemistry staining), (D) Image of the follicular growth between the α-MEM-based medium and StemPro-based medium (scale bar, 200 μm).

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Next, to determine whether a single follicle and oocyte possessing a three-dimensional (3D) structure could be formed, we cultured single cells dissociated from mice gonads at 12.5, 13.5, and 14.5 dpc on a Transwell-COL dish using StemPro medium. Additionally, to confirm enzyme efficiency for gonad dissociation into single cells, two enzymes were assessed, i.e., trypsin-EDTA and collagenase IV. First, we used trypsin-EDTA to dissociate the gonads into single cells, which demonstrated high dissociation efficiency into single cells. We cultured single cells on Transwell-COL after trypsin-EDTA treatment to confirm dissociation of mice gonads at 12.5, 13.5, and 14.5 dpc. Spherical cells, which differed from the germline somatic cell shape, were found to grow on Transwell-COL membranes. We analyzed cultured globular cells, and expression levels of the germline cell marker Ddx4 were confirmed by immunocytochemistry (Fig. 2A). Spherical cells displayed Ddx4 expression on the cell surface; this was demonstrated by 12.5 dpc gonads, as well as 13.5 and 14.5 dpc gonads (Fig. 2A). Reportedly, gonadal PGCs are known to demonstrate oogonia, which can differentiate into meiotic cells through multiple cell divisions during migration to the genital ridge (Hayashi & Saitou, 2013; Saitou & Miyauchi, 2016). These results demonstrated that, when single cells dissociated from gonads and PGCs and gonadal somatic cells were co-cultured in Transwell-COL, co-cultured single cells could differentiate into oogonia-like cells expressing Ddx4, without follicle formation (Lei et al., 2006).

Fig. 2. Single-cell dissociation fails to form follicles. (A) Phase and immunostaining data for expression of Ddx4 in oogonia-like cells developed 12.5, 13.5, and 14.5 days post coitum (dpc) (scale bar, 200 μm), (B) Follicle is not formed in the U-bottom plate (scale bar, 200 μm), (C) Cultivation of gonad cells in Matrigel.

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Then, we aggregated and cultured PGCs and germline somatic cells on a U-bottom plate, as we failed to generate a 3D structure on Transwell-COL membranes. Single cells from gonads were seeded at a density of 2.5 × 10⁵ per well in a hexagon-shaped U-bottom plate. The cells aggregated after 1 day, but no follicle formation was observed (Fig. 2B); these results were similarly observed for 12.5, 13.5, and 14.5 dpc gonads. As another method for 3D culturing, we used Matrigel, which is widely used for organoid culture or 3D tissue culture (Lang et al., 2001; Pierzchalska et al., 2012; Dolega et al., 2015). Similar to previous results, it did not facilitate ovary formation (Fig. 2C). These results indicated that once the gonad is dissociated into a single cell, ovary formation is impossible even if clustered again.

We employed several strategies to generate a minimally-sized ovary that can form a single follicle. Accordingly, the gonad was manually separated into three parts, and all three parts formed the ovary, producing fewer follicles than the control (Fig. 3A). In addition, follicles formed in this manner generated superior oocytes. Furthermore, the oocytes within the ovary were well-developed, similar to the oocytes on the outside. Signals and nutrients were well transmitted to the inner part of the ovary. However, the method failed to generate fewer ovarian follicles. Therefore, loss of follicles can still occur during the isolation process. During this method, we used a thin syringe to mechanically cut the gonad into the smallest fragments possible. We then assessed ovary formation and observed that follicles were produced when gonads were cut to the minimum size without isolation into a single cell state. Accordingly, all gonads adequately formed ovaries; however, numerous follicles were generated. Moreover, the number of follicles appeared to random in the in vitro ovaries obtained (Fig. 3B). This method was unsuitable for generating uniform follicles in all ovaries, and it was difficult to individually isolate follicles. Based on these findings, it was confirmed that ovaries were formed even when the gonad was separated into a gonadal mass without isolation into a single cell. Next, we attempted to confirm whether ovary formation occurred when smaller gonadal masses were achieved using a collagenase IV suspension rather than mechanical separation. Using the collagenase IV enzyme, we observed that gonads were dissociated into an appropriate gonad mass that could form ovaries. In the present study, the efficiency of dissociation into an appropriate gonadal mass was increased by controlling the collagenase IV treatment time and suspension. Following collagenase IV treatment for more than 2 min, gonadal masses were broken into single cells; when treatment was performed for less than 2 min, ovaries producing numerous follicles were formed. After approximately 2 min of collagenase IV treatment in suspension, a few minimally crushed masses could be identified. We cultured the masses and found that ovaries with a fewer number of follicles were formed. The ovaries generated using this method were smaller in size than those generated using the previous methods; however, a random number of follicles were produced (Fig. 3C). This strategy successfully reduced ovary size and follicle number; however, precise standardization remained challenging.

Fig. 3. Method optimization for generation of miniaturized follicles per ovary. (A) Image of the ovaries from gonadal tissues dissociated into three parts (left scale bars, 40 μm; right scale bar, 100 μm), (B) Image of the ovaries from gonadal tissues dissociated by mechanical cutting (scale bar, 200 μm), (C) Image of the ovaries from gonadal tissues dissociated with collagenase Ⅵ (scale bar, 200 μm), (D) A schematic of miniaturized ovary production in vitro, (E) Image of the ovaries from gonadal tissues dissociated using 150, 100, 70, and 40 µm meshes (scale bar, 200 μm), (F) Number of follicles in miniaturized ovaries generated from different gonadal mass sizes (n=16, Experiments were performed in triplicate).

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Finally, we cultured gonad tissues after breaking them down to a uniform size using 150, 100, 70, and 40 μm meshes. Collagenase IV treatment was performed for 2 min, followed by repeated suspension two or three times; in the absence of adequate collagenase treatment, all gonads will be similar in size as control gonads. In order to determine the minimum mass size that can accurately form follicles, the gonad was cultured by dividing it into four groups after passing through 150, 100, 70, and 40 μm meshes. Thus, gonad masses were cultured in four groups of 150–100, 100–70, 70–40, and 40 μm or less (Fig. 3D). When cultured by breaking the gonad using a 40 μm mesh, the gonadal mass formed ovaries but no follicles (Fig. 3E). Additionally, the gonadal mass passed through the 70 μm mesh failed to generate follicles (Fig. 3E). In contrast, the gonadal mass passed through the 100 μm mesh and retained in the 70 μm mesh only formed four ovaries, which produced follicles (Fig. 3F). However, owing to the low ovarian production rate, it was challenging to produce numerous oocytes. Gonadal mass passed through the 150 μm mesh and retained in the 100 μm mesh was equal or slightly larger than 100 μm sized mass, formed miniaturized ovaries, and contained an average of 12 uniform follicles per ovary (Fig. 3F). In gonadal masses ranging between 150 μm and 100 μm in size, ovaries generated normally developed follicles during IVG (Fig. 4A). These results indicate that a minimum gonadal mass of 100 μm is required to form an ovary capable of generating follicles. In other words, when the gonad is crushed to a size less than 100 μm, the cells are damaged, and ovaries cannot be formed. In this experiment, we used collagenase IV treatment, manual cutting, and passage through a mesh. On using a mesh, we were able to identify the most efficient and suitable size. Undoubtedly, the method was less time-consuming, resulted in reduced cell loss during isolation, and produced ovaries with fewer follicles. Although there are technical limitations to generating ovaries that form single follicles, these issues can be surpassed. Ten days after IVG, oocytes were well-developed into germinal vesicle (GV) oocytes (Fig. 4A). GV oocytes were treated with IVM medium which used to resume meiosis. After 16 h, the oocytes extruded the first polar body (Fig. 4B).

Fig. 4. Oocyte maturation from the miniaturized ovary. (A) Representative image of the follicle cultivated on a Transwell-COL membrane on days 26–35 (scale bar, 100 μm), (B) Image of the first polar body in MII oocyte (scale bar, 100 μm).

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