ORIGINAL RESEARCH REPORT

Ultrastructural Characterization of MouseEmbryonic Stem Cell-Derived Oocytes and Granulosa Cells

Olympia E. Psathaki,1,* Karin Hubner,1,* Davood Sabour,1 Vittorio Sebastiano,2 Guangming Wu,1

Fumihiro Sugawa,1 Peter Wieacker,3 Petra Pennekamp,3 and Hans R. Scholer1,4

Germ cells are a unique population of cells responsible for transmitting genetic information from one generationto the next. Our understanding of the key mechanisms underlying germ cell development in vivo remains scarcebecause of insufficient amounts of cell materials available for conducting biological studies. The establishment ofin vitro differentiation models that support the generation of germ cells from mouse pluripotent stem cellsprovides an alternative means for studying reproductive development. The detection and analysis of stem cell-derived germ cells, however, present technical challenges. Methods for determining the developmental stage ofgerm cells ex vivo, such as gene expression and/or immunochemical analyses are inadequate, frequently ne-cessitating the use of alternative, elaborate methods to prove germ cell identity. We have generated putativeoocytes and granulosa cells in vitro from mouse embryonic stem cells and utilized electron microscopy tocharacterize these cells. Here, we report on the striking ultrastructural similarity of in vitro-generated oocytesand granulosa cells to in vivo oocytes developing within follicles.

Introduction

Afundamental concept in developmental biology is thedefinition of a robust and reproducible in vitro culture

system that supports the induction and differentiation offunctional and specific cells from totipotent or pluripotentstem cells. Commitment of mouse embryonic stem cells(ESCs) to differentiate into germ cells under defined cultureconditions represents an important milestone in develop-mental biology [1]. Although the functionality of mouseESC-derived sperm has been demonstrated [2], that of oo-cytes has not. Human ESCs can also be used to form pri-mordial germ cells (PGCs), an indication of their potential tocontribute to germline development [3–8]. The potentialgeneration of an unlimited source of germ cells and gametesfrom human ESCs and other pluripotent cells, for example,induced pluripotent stem cells (iPs), would facilitate thestudy of reproductive development and provide a new ap-proach to the development of assisted reproduction thera-pies that do not rely on human samples. A prerequisite tothis fascinating approach, however, is the step-by-stepcharacterization of the generated cells of all developmentalstages to ensure appropriate recapitulation of germ cell de-velopment in vitro. The growth and maturation of oocytes

are complex processes of folliculogenesis. A bidirectionalmode of communication between the oocyte and the com-panion granulosa cells (GCs) directs development of theoocyte [9,10]. This communication relies upon the structuralintegrity of the granulosa–oocyte cell interface [11]. The oo-cyte and surrounding GCs are structurally and functionallyassociated via specialized cytoplasmic processes calledtranszonal projections (TZPs), representing a highly dy-namic interaction regulating folliculogenesis in vivo andthereby ensuring appropriate oocyte maturation and ovu-lation [12,13]. Significant information on the adaptive mor-phological changes taking place in GCs during thereproductive cycle has been obtained by electron microscopy(EM) in situ as well as in vitro [14–17]. We used EM to de-termine the morphological characteristics and cellular in-terface ultrastructure of in vitro-generated GCs and oocytes.EM techniques can not only provide information on thefunctionality of the cells under study, but also be performedwith minute quantities of a sample. Such techniques aretherefore not subject to the experimental restrictions associ-ated with the use of fetal tissues. Further, information gar-nered using EM techniques in the mouse model would beinstrumental toward the establishment of differentiationsystems with human pluripotent stem cells.

1Department of Cell and Developmental Biology, Max Planck Institute for Molecular Biomedicine, Munster, Germany.2Institute for Stem Cell Biology and Regenerative Medicine, Stanford University School of Medicine, Stanford, California.3Institute for Human Genetics, Westfalische Wilhelms-University, Munster, Germany.4Medical Faculty, University of Munster, Munster, Germany.*These authors contributed equally to this work.

STEM CELLS AND DEVELOPMENT

Volume XX, Number XX, 2011

� Mary Ann Liebert, Inc.

DOI: 10.1089/scd.2010.0575

1

Materials and Methods

Electron microscopy

For transmission electron microscopy (TEM), cells werefixed with 2.5% glutaraldehyde (Merck) in 0.1 M sodiumcacodylate buffer (pH 7.4), postfixed in 1% aqueous osmiumtetroxide, dehydrated stepwise in a graded ethanol series,and embedded in Epon 812 (Fluka, Buchs, Switzerland).Ultrathin (50 nm) sections were prepared with an ultrami-crotome (EM UC6; Leica), stained first with 1% uranyl ace-tate and then with 3% lead citrate, and subsequentlyexamined using a Zeiss EM 109 electron microscope (Zeiss).Images were taken on 70 mm films (Maco ORT 25c ortho-chromatic; Hans O. Mahn & Co., Photo Division).

For scanning electron microscopy (SEM), cells were fixedwith 2.5% glutaraldehyde in 0.1 M phosphate buffered saline(pH 7.4) and dehydrated stepwise in a graded ethanol series.Samples were dried with 100% ethanol via CO2 in a critical-point apparatus (Balzers). Dried samples were mounted ontoaluminum stubs with leit-tabs and coated with gold film to athickness of 40–50 nm in a sputter coater (Leitz). Cells wereviewed under a Hitachi S-530 scanning electron microscopeoperated in a secondary mode at 20 kV.

Cell culture

Mouse ESC lines gcOct4-GFP (XY) [1] and OG2 (DPE-Oct4-GFP, XY; XX) [18] were differentiated into oocytesaccording to 3 protocols: a previously described serum-containing protocol [1], a serum-containing/serum-freedifferentiation combination protocol, and a serum-free dif-ferentiation protocol. Briefly, for the serum-containing dif-ferentiation procedure, cells were grown in tissue cultureplates in Dulbecco’s modified Eagle’s medium containing4.5 g/L glucose and supplemented with 15% fetal calf serum(Hyclone), 2 mM l-glutamine (Invitrogen), 100mM nones-sential amino acids (Invitrogen), 1 mM b-mercaptoethanol(Sigma), 50 mg/mL penicillin and streptomycin (Invitrogen)in the absence of mouse embryonic fibroblast (MEF) feedercells and leukemia inhibitory factor (LIF) for up to 30 days.The medium was changed every 3 days for 9 days and everyother day thereafter. Floating cell aggregates after day 12 ofdifferentiation were collected and processed as published [1].For the serum-containing/serum-free differentiation combi-nation technique, ESCs were differentiated under serum-containing conditions for 7–12 days. The adherent cell layerwas digested with accutase (Invitrogen) and the single-cellsuspension was then plated as a suspension culture at a 1:1ratio in in vitro growth (IVG) media [modified a-MEMsupplemented with 3 mg/mL bovine serum albumin (Milli-pore), 50mg/mL each penicillin and streptomycin (Invitro-gen), 2 mM glutamine (Invitrogen), 0.23 mM pyruvate(Invitrogen), 10 mg/mL insulin, 5 mg/mL transferrin, 5 ng/mL selenium (Invitrogen), 1 ng/mL human recombinantepidermal growth factor (Invitrogen), and 50 ng/mL re-combinant mouse stem cell factor (Millipore)] in a humidi-fied incubator at 37�C in 5% O2 and 5% CO2. Cells werecultured for 7–15 days, and putative oocytes were then in-dividually picked with a drawn-out glass capillary. For theserum-free differentiation protocol, feeder-free ESCs wereplated on gelatin-coated tissue culture plates in N2B27 me-dium according to the method of Ying et al. [19] and cultured

in 5% CO2 for 48 h. The adherent cell layer was then digestedwith accutase and the single-cell suspension was replated at a1:2 ratio in IVG media supplemented with stem cell factor (SCF)and leukemia inhibitory factor (LIF) in suspension plates. After3 days of culture in 5% CO2, cell aggregates were transferred togelatinized 24-well tissue culture plates at a density of 5–6 ag-gregates/well and further cultivated for 5 days in mediumlacking LIF. Cultures were then partially digested in trypsin/ethylenediaminetetraacetic acid 0.05% supplemented with0.02% DNAse I and replated in oocyte growth media (modifieda-MEM supplemented with 3 mg/mL bovine serum albumin[Millipore], 50mg/mL each penicillin and streptomycin [In-vitrogen], 0.23 mM pyruvate [Invitrogen], 5mg/mL insulin,5mg/mL transferrin, 5 ng/mL selenium [Sigma], 1 ng/mLhuman recombinant epidermal growth factor [Invitrogen],100 ng/mL recombinant mouse stem cell factor [Millipore], and5 ng/mL rhFSH [Sigma]) and cultured in a humidified incu-bator at 37�C in 5% O2 and 5% CO2 for additional 12–15 days.Putative oocytes were cultivated in maturation media as pre-viously described to assess their maturation status [20].

RNA isolation

RNA was isolated using the RNeasy Micro Kit (QIAGEN)with on-column DNase digestion as per the manufacturer’sinstructions. RNA quality was determined using a Nanodrop(ND-1000) spectrophotometer and an Agilent RNA 2100Bioanalyzer.

For single-cell analysis of oocytes, the Applied Biosystemskit was used for preamplification according to the manu-facturer’s recommendations.

Reverse transcription–polymerase chainreaction and real-time quantitative reversetranscription–polymerase chain reaction

Two hundred fifty nanograms of RNA was reverse-tran-scribed to cDNA using the High-Capacity cDNA ReverseTranscription Kit (Applied Biosystems). Real-time quantita-tive TaqMan reverse transcription–polymerase chain reac-tion (q-RT-PCR) was performed on 7900ht and ht fastdevices (Applied Biosystems) with milliQ H2O and TaqManUniversal PCR Master Mix (Applied Biosystems). Raw datawere analyzed using SDS 2.3 software (Applied Biosystems).Gene expression was normalized to the mouse housekeepinggene Hprt. Quantification of relative gene expression wascalculated using the DDCt method. Three technical replicateswere used for each real-time q-RT-PCR reaction; a re-verse transcriptase blank and a no-template blank servedas negative controls. The following primers wereused: Oct4: Mm00658129_gH; Blimp1: Mm00476128_m1;Ddx4: Mm00802445_m1; Figla: Mm00488823_m1; Nobox:Mm00453743_m1; Prdm14: Mm01237813_m1; Sohlh1:Mm01338424_g1; Stella: Mm00836373_g1; Sycp3:Mm00488519_m1.

Isolation of germinal vesicle-stage oocytesfrom ovaries of adult mice

Eight- to 10-week-old CD1 female mice were super-ovulated by injection of 7.5 IU pregnant mare serum gona-dotropin (PMSG) and, 48 h later, by injection of 7.5 IU human

2 PSATHAKI ET AL.

chorionic gonadotropin (hCG). Females were sacrificed bycervical dislocation at 17–18 h post-hCG stimulation. Ovarieswere dissected and digested for 10 min at 37�C in 0.5 mg/mLcollagenase type 1A (Sigma), washed in PBS, and placed inHepes-buffered CZB medium containing 1 g/L glucose and0.1% (w/v) polyvinylpyrolidone (ICN). Follicles were releasedinto the medium by mechanical dissociation of the tissue. Cu-mulus cells were removed mechanically and oocytes with aclearly visible germinal vesicle were collected, washed 3 timesin Hepes-buffered CZB medium, and lysed in RNeasy lysesbuffer (RLT) to extract RNA for gene expression analysis orfurther processed for EM. Mice were maintained and used forexperiments according to institutional guidelines.

Results

Granulosa cells and the GC–oocyte interfaceof ESC-derived follicles

The entry of PGCs into meiosis in vivo and in vitro ap-pears to be a cell autonomous process. Subsequent devel-opment of PGCs into competent oocytes and ultimatemeiotic arrest during folliculogenesis is dependent on 2-waysignaling interactions between the oocyte and surroundingGCs [21,22]. Figure 1A–D shows ESC-derived follicle-likestructures. As aggregation of PGCs and granulosa-like cellstakes place randomly in this system, we obtained sphericalaggregates with centrally located single as well as multipleputative germ cells surrounded by various layers of attachedpresumptive GCs and extracellular matrix (ECM). Lightmicroscopy revealed germ cell clusters surrounded byloosely attached somatic granulosa-like cells (Fig. 1A). SEManalysis demonstrated follicle-like structures, that is, germcells surrounded by a dense layer of cuboidal cells (Fig. 1B).Figure 1C shows ESC-derived putative GCs in direct prox-imity to a zona pellucida-like matrix. The granulosa-like cells

exhibit a polygonal shape and a smooth surface with fewmicrovilli (Fig. 1C). SEM and TEM analysis revealed astructure specific for GCs, that is, TZPs (Fig. 1C, 1D, arrow)that extend toward the putative oocyte.

Figure 2A shows a light microscopy image of anESC-derived follicular structure, with presumptive GCssurrounding the green fluorescent protein (GFP)-positiveoocyte-like cell. To analyze the GC–oocyte interface by SEM,in vitro-derived follicles were cracked. Figure 2C and Dshows both parts of a cracked aggregate, revealing theirmorphological similarity to natural follicles. Even though thepresumptive oocyte was destroyed during the crackingprocedure, a round cavity of 50–60mm in diameter with re-sidual oocyte cell material is evident (Fig. 2C, D). Multiplelayers of GCs surround the cavity. The most central onesexhibiting the typical cuboidal-elongated phenotype of nat-ural polarized cumulus cells, that is, those cells in directcontact to the oocyte (Fig. 2C), whose TZPs orientate towardthe oocyte (Fig. 2B). TZPs have been well characterized inmany mammals by EM [23,24], and their number and formhave been demonstrated to change dynamically during fol-licular development [12]. To investigate the GC–oocyte in-terface in more detail, we performed TEM analyses andcompared GCs of natural follicles to ESC-derived presump-tive GCs. The ultrastructure of TZPs of in vitro-derived GCsappeared to be indistinguishable from the TZPs of theirnatural counterparts, indicating generation of the interfacebetween the oocyte and GCs, the major control structure forhealthy follicle development [12]. ESC-derived GCs of highelectron density exhibited long cellular processes containingthe same dark fine granulation as the cytoplasm (Fig. 2E).These processes penetrated the matrix, with some reaching adiameter of *1mm, very similar to the branched processes ofpolarized cumulus cell in natural follicles. The detection oforganelles, for example, mitochondria [12,25], within the

FIG. 1. ESC-derived follicle-likestructures. (A) Light microscopy imageof ESC-derived follicle-like structureswith clusters of granulosa cells looselysurrounding the oocyte. (B) SEMimage of a defined follicle structure.Note the layer of densely attachedcuboidal cells (asterisk). (C) SEM imageof polygonal-shaped granulosa cellsaround a smooth zona pellucida-likesurface (asterisk). The pole of a granulosacell is facing toward the oocyte (arrow).(D) TEM image of a granulosa cell withan extension stretching towards the oo-cyte (arrow) ( · 4,890). ESC, embryonicstem cell; SEM, scanning electron mi-croscopy; TEM, transmission electronmicroscopy.

MOUSE STEM CELL-DERIVED OOCYTES 3

TZPs of in vitro-derived GCs (Fig. 2F, G) is indicative ofrecapitulation of folliculogenesis in vitro. Further, the ultra-structural texture of the pale amorphous matrix in directproximity to the GCs in vitro (Fig. 2E) resembles the zonapellucida of the GC–oocyte interface in vivo (Fig. 2H).

Ultrastructural characterization of in vitro-generated GC-like cells revealed additional analogies to characteristics ofnatural GCs described in the literature, for example, thepresence of both dark- and light-appearing GCs. Dark GCspossess a dark background nucleoplasm and backgroundcytoplasm (Fig. 3A, C). The electron density is due to the finegranulation of the cytoplasm, which does not correspond tofree ribosomes, as free ribosomes would be much larger and

stand distinctly out from the granular background (Fig. 3E,F). The cytoplasm of the GCs appears to be partly vacuolatedand is populated by mitochondria, ribosomes, and the mostprominent organelle of ESC-derived GCs, the rough endo-plasmic reticulum. Cisternae of swollen rough endoplasmicreticulum containing granular material wind through thecytoplasm of dark and light in vitro-derived GCs (Fig. 3A, E)as well as pale in vivo GCs (Fig. 3F). The ribosomes of nat-ural dark and light GCs and ESC-derived GCs are either freeor associated with rough endoplasmic reticulum (Fig. 3A, B,E, F). The presence of free ribosomes is a characteristic fea-ture of protein-secreting cells, such as the GCs of grow-ing follicles. The mitochondria of both dark in vivo and

FIG. 2. Granulosa cell–oocyte interfaceof ESC-derived follicles. (A) Light mi-croscopy image of an ESC-derived folliclestructure. Granulosa cells surround thegcOct4-GFP oocyte. (B) SEM analysis ofan ESC-derived follicle-like structure. Along process originates from a somaticgranulosa cell toward the oocyte (arrow).(C, D) SEM analysis of both halves of acracked ESC-derived follicle-like struc-ture. The position of the oocyte is clearlyapparent by the 50–60-mm-large, roundcavity (asterisk in C) and residual oocytecell material in D (asterisk). The granulosacells surround the oocyte in multiple lay-ers, with the cell layer in direct contactbeing cuboidal-elongated in shape (C, ar-row). (E) TEM analysis of ESC-derivedgranulosa cells. Cells exert long processes(TZPs) toward the oocyte (arrow). Notethe texture of the pale amorphous matrix(asterisk), which is penetrated by the TZPs( · 4,890). (F) TZPs contain organelles suchas mitochondria (arrow) and have thesame ultrastructural texture as the cellcytoplasm ( · 4890). (G) TZPs of ESC-derived granulosa cells partly exhibit ex-treme branching processes (arrow) andreach diameters of almost 1mm ( · 12,210).(H) Cellular processes of cumulus cellswith branching ends (arrow) of in vivofollicles closely resemble those of in vitro-derived granulosa cells. The ultrastruc-ture of the zona pellucida (asterisk)resembles that of in vitro-derived folliclesin E ( · 4,890). TZPs, transzonal projec-tions.

4 PSATHAKI ET AL.

in vitro-derived GCs appear irregular and elongated, con-taining pale cristae and an electron-dense matrix (Fig. 3A, B).The mitochondria of light GCs exhibit an oval shape andhave tubular–vesicular cristae (Fig. 3E, F). Lipid dropletsfilled with a transparent gray substance and empty vacuolesfrequently surrounded by a membrane (Fig. 3B, D) are dis-tributed throughout the cytoplasm of in vivo GCs. Vacuoli-zation appears to be a prominent feature of ESC-derived GCs(Fig. 3C), manifested mainly by empty white vacuoles andvacuoles containing a residual gray substance (Fig. 3C).Empty vacuoles appear to be an artifact of sample manipu-lation, created when the gray substance of lipid droplets iswashed out during the dehydration step. The nuclei of both

in vitro-derived and in vivo GCs are large and have anirregular contour (Fig. 3A–D). The nucleoplasm of ESC-derived GCs, enclosed by the inner nuclear membrane,contains finely dispersed chromatin and clumps of hetero-chromatin scattered throughout (Fig. 3A, C, E); the nucleo-plasm of in vivo GCs is arranged similarly (Fig. 3B, D, F).Taking these data together, the ultrastructural features of invitro-derived GCs are indistinguishable from natural GCs.Most importantly, to the best of our knowledge, there is noother cell in the body capable of forming unique and typicalTZPs. The observed activated rough endoplasmatic reticulum,indicative for metabolically active and steroid-producing cells,further supports their GC-like identity.

FIG. 3. TEM analyses of ESC-deriveddark- and light-colored granulosa cells.(A) Cuboidal-elongated in vitro-derivedgranulosa cells with electron dense cyto-plasm and swollen rough endoplasmaticreticulum. Mitochondria (insert) exhibitpale tubular cristae ( · 4,890). (B) In vivo-derived dark granulosa cell. Note lipiddroplets (asterisk) ( · 4,890). (C) Dark ESC-derived granulosa cells with numerousvacuoles and droplets containing residualgray translucent substance indicative oflipids (asterisk). Note prominent nucleolus( · 4,890). (D) In vivo dark granulosa cellwith numerous pseudopodia, branchingprocesses and protrusions of cytoplasmicinvaginations, vacuoles (arrowhead), andlipid droplets (asterisk) as detected in invitro-derived granulosa cell shown in C( · 6,950). (E) ESC-derived light granulosacell with microvilli at the cell surface. Thecell exhibits the typical swollen roughendoplasmatic reticulum (asterisk) andoval-shaped tubular mitochondria withdark cristae (insert) ( · 4,890). (F) In vivo-derived light granulosa cell with swollenrough endoplasmatic reticulum (asterisk)and vesicular–tubular mitochondria (ar-row) ( · 4,890).

MOUSE STEM CELL-DERIVED OOCYTES 5

ESC-derived oocytes

We have developed a suspension culture system thatsupports the generation of ESC-derived oocytes of differentfollicular stages. Independent of the differentiation mediaused to initiate differentiation of ESCs, we obtained putativeoocytes of various developmental stages at different yields.Serum-free conditions appeared best to promote germ celldifferentiation, and XY-ESCs yielded a higher percentage ofoocytes than XX-ESCs. Expression of specific oocyte markergenes was assessed by qRT-PCR in groups of 3 in vitro-derived oocytes and compared with superovulated germinalvesicle-stage oocytes and ESCs. As expected, the generatedoocytes (Fig. 4A) exhibited a gene expression pattern verysimilar to that of control samples (Fig. 4B). Morphologically,in vitro-generated oocytes (Figs. 4A and 5A) and in vitro-matured oocytes generated from 18.5 days postcoitum (dpc)germ cells under the same culture conditions (Fig. 5C) ex-hibited a very similar shape, size, and microvillous cell sur-face (Fig. 5C, D). Interestingly, the surface topography ofsome in vitro-derived oocytes analyzed by SEM resembledexactly that of unfertilized oocytes in vivo, with the typicalsparse and relatively uniform microvillous area and a clearmosaic surface containing a polar microvillous-free region(Fig. 5B–D). TEM analyses of generated oocytes revealedlarge, slightly eccentric nuclei with a characteristic palebackground nucleoplasm (paler than the ooplasm) (Fig. 5E).Free ribosomes appeared individually or arranged in clustersof 5–10 (Fig. 5E, insert). Elongated staples of fibrils not as-sociated with ribosomes were found throughout the cyto-plasm (Fig. 5E, insert, F). Oocyte-specific cortical granulescontaining a characteristic electron-dense center and a paleouter ring (Fig. 5F, arrow) were mostly distributedthroughout the cytoplasm and appeared only sporadically at

the cell periphery (Fig. 5F and inserts), a distinct feature ofimmature oocytes. Further, we detected round as well aselongated mitochondria with a dense matrix and pale, ir-regularly arranged cristae and intermitochondrial vacuoles,all features of natural oocytes. Noteworthy is that the ma-jority of ESC-derived oocytes showed signs of apoptosis andautophagy, which might be indicative of atresia (Fig. 5E, H).The oocyte shown in Fig. 5E lost its spherical shape. Nu-merous lysosomes, clear vesicles, and autophagosomes aswell as dark cytoplasmic structures were detected. The ESC-derived oocyte shown in Fig. 5H contains numerous apo-ptotic bodies and secondary lysosomes within the darkcondensed cytoplasm. Mitochondria and other organellescannot be identified. The zona pellucida and most of themicrovilli at the oocyte surface appear to have been lost. Incomparison, oocytes derived from natural 18.5-dpc PGCsgrown for 13 days in the same culture medium as in vitro-derived oocytes (Fig. 5G) also exhibit dark-stained mito-chondria without cristae, numerous vacuoles, and secondarylysosomes; other organelles cannot be identified. Microvillieither have been lost or have retracted from the zona pellu-cida. These observations indicate that our culture conditionssupport the maturation of PGCs into oogonia, but furtheroptimization of these conditions is required for the genera-tion of functional oocytes.

Discussion

The first systematic study of the culture and growth ofmouse oocytes in vitro was published by John Eppig in 1977[26]. Eppig and coworkers demonstrated that mouse oocytesfrom primordial follicles could be grown and matured invitro to produce live offspring after fertilization [27,28].These data marked a major breakthrough in the field of

FIG. 4. Gene expression of oocytemarkers in a pool of 3 in vitro-derivedoocytes. (A) Light microscopy imageof one of the analyzed oocytes. (B)Real-time quantitative reversetranscription–polymerase chain reac-tion of oocyte markers in a pool of 3ESC-derived oocytes compared withXX-ESCs.

6 PSATHAKI ET AL.

reproductive biology. Based on this success, various in vitroculture systems for female gametes of different mammalianspecies were established, such as rat [29], hamster [30], cat[31], pig [32], sheep [33], goat [34], cow [35,36], and human[37]. All these maturation systems, including the mousemodel, are still under investigation in an effort to broadenour understanding of the complex mechanisms involved ingerm cell development and maturation. In this study, weperformed comparative ultrastructural analyses of follicle-like structures and bona fide oocytes from mouse ESCs withtheir natural counterparts. With our data, we supplement

the genetic and immunological data obtained by standardanalyses with a description of the unique ultrastructuralproperties of oocytes and their supporting GCs during ga-metogenesis in vivo. Specifically, we performed SEM anal-ysis on in vitro-derived follicle-like structures and comparedtheir morphology with that of natural mammalian folliclesdescribed by Makabe and coworkers in 2006 [13]. Strikingmorphological similarities indicate that our culture systemsupports the derivation of germ cells of different develop-mental follicular stages. During the early growth of follicles,the shape of GCs progressively changes from flattened to

FIG. 5. Ultrastructural analysis ofin vitro-derived oocytes. (A) Light mi-croscopy image of an in vitro-derivedoocyte. (B) SEM image of an in vitro-derived oocyte with a polar microvillous-free surface (asterisk). (C) SEM image offetal in vivo oocytes cultivated under thesame culture conditions as the in vitro-derived oocytes. Note the polar microvil-lous-free region (asterisk). (D) SEM imageof an in vitro-derived oocyte with a polarmicrovillous-free surface (asterisk). Notethe similarity of the shape and surfacestructure to in vivo oocytes shown in C.(E) TEM image showing an overview ofan in vitro-derived oocyte. Numerous ly-sosomes, clear vesicles, lipid droplets (ar-row), and autophagosomes (arrowhead) aredistributed throughout the cytoplasm. n:nucleus ( · 1,100). Insert: Elongated mito-chondria with tubular–vesicular cristaeand vacuoles are arranged betweenstaples of fibrils ( · 6,950). (F) TEM imageof oocyte-specific cortical granules (arrow)with the typical electron-dense centerand pale outer ring of in vitro-derivedoocytes. Cortical granules appear onlysporadically in the cell periphery (inserts).Note vacuolated mitochondria (arrowhead)( · 6,950). (G) TEM image of oocytes de-rived from natural 18.5-day postcoitumprimordial germ cells grown for 13 daysin the same culture medium as in vitro-derived oocytes. The dark-stained mi-tochondria contain only few cristae,numerous vacuoles, autophagosomes (ar-rowhead), and secondary lysosomes. Otherorganelles are not identifiable. Microvilliare lost or retracted from the zona pellu-cida (arrow) ( · 6,950). (H) TEM image ofin vitro-derived oocyte showing numer-ous apoptotic bodies (arrow) and second-ary lysosomes in the dark, condensedcytoplasm. Mitochondria or other organ-elles are not identifiable. The zona pellu-cida and most of the microvilli at theoocyte surface are lost ( · 6,950).

MOUSE STEM CELL-DERIVED OOCYTES 7

cuboidal and to columnar [38] [39]. Well-defined in vitro-derived follicles (Fig. 1B), covered with a layer of denselyattached polyhedral cells, form the stratum granulosum, in-dicating that the follicles are in the late phase of growth.Cracked in vitro-derived follicles, such as the one shown inFig. 2C and D, exhibit the same morphological features ascracked mammalian follicles, with a residual macerated oo-plasm as previously described [13]. We utilized TEM ana-lyses to assess differences in the dynamic structure of TZPs,which form the morphological basis of the oocyte–GC in-terface and play a crucial role in the structural integrity ofdeveloping follicles and maturing oocytes [12]. Many of ourin vitro-derived GCs developed microvilli, cytoplasmic in-vaginations, pseudopodia, and cytoplasmic protrusions (notshown here), which have been associated with cell expansionand are considered indirect signs of GC luteinization both invitro and in vivo [40]. Further, we observed morphologicallydifferent subpopulations of ESC-derived GCs, that is, thoseexhibiting differences in the darkness of background cyto-plasm and nucleoplasm as well as in content and distributionof lipid droplets and rough endoplasmic reticulum. Thisheterogeneity of a given GC population has also been re-ported by Nottola and coworkers [16] for in vivo follicularaspirates cultured in vitro. Further, cuboidal GCs with a darkbackground cytoplasm and nucleoplasm have been found inprimordial polyovular and unilaminar follicles of the goldenhamster in vivo [41]. We also detected these light and darkGCs in natural mouse follicles (Fig. 3B, D, F). Both dark andlight GCs are frequently found in developing endocrine tis-sues and reflect a change in the complex secretory cycle [41].The fine dispersion of chromatin and the peripheral patchesof heterochromatin, which we found in the nuclei of ESC-derived GCs (Fig. 3A, C, E), are features typical of meta-bolically active cells [40,42]. Mitochondria of ESC-derivedGCs vary in size and shape and have mostly tubular–vesic-ular cristae (Fig. 3A, E). These types of mitochondria arefrequently found in luteinizing GCs in vivo [43] and in hu-man GCs of follicles containing a fertilizable oocyte [16]. Invitro-derived dark GCs contain mitochondria with a densematrix and pale cristae (Fig. 3A, insert), whereas light GCscontain mitochondria with a pale matrix and dark cristae(Fig. 3E, insert). Makino et al. [44] found an association be-tween mitochondria of human GCs exhibiting a pale matrixwith poor fertilization outcome in vitro. Mitochondria withlongitudinally oriented cristae have been found in humanovarian carcinoma and in the lungs, liver, and ureter ofdifferent mammals [16]. Researchers have hypothesized thatthis morphological transformation of mitochondria could bedue to negative environmental conditions, such as high ox-ygen concentration, presence of cytostatic drugs or antibi-otics and aging [45–47]. We did not observe suchmitochondria in our culture-derived GCs, indicating that wesuccessfully generated functional GCs in vitro. A most pro-minent feature of in vitro-derived GCs is a rich network ofswollen rough endoplasmic reticulum and a well-developedGolgi apparatus. These features indicate that there is ongoingluteinization [40,43,48,49]. Isolated rough endoplasmic re-ticulum and ribosomes of luteinizing GCs are responsible forsynthesizing the enzymes involved in steroid production[50]. The presence of numerous lipid droplets within thecytoplasm of in vitro-derived GCs (Fig. 3C), an early sign ofluteinization [50], correlates with active steroidogenesis [16].

Taken together, in vitro-derived GCs exhibit ultrastructuralcharacteristics typical for metabolically active and steroid-producing cells.

The ovarian follicle represents a morphological and func-tional unit wherein both somatic and germ cells play a piv-otal role in follicle maturation and formation of fullycompetent, fertilizable oocytes [51]. Under appropriate cul-ture conditions, in vitro-matured cumulus-enclosed oocytesof mouse, rat, cow, and sheep have been shown to possess asimilar developmental capacity to mature oocytes in vivo[52–54]. SEM analysis clearly demonstrated that ESC-derivedcuboidal-shaped GCs closely surround the oocyte (Fig. 2C,D) and extend long TZPs toward the oocyte (Fig. 1D). Invivo, TZPs are most abundant in preantral follicles. By TEManalysis, we demonstrated that ESC-derived cumulus cellsextend typical TZPs, some of which contain mitochondria, anobservation consistent with literature [12,25]. Albertini et al.[12] demonstrated the active bidirectional movement of mi-tochondria within the TZPs by vital staining of intact mousefollicles with MitoTrackerTM, a cell-permeant probe thatcontains a thiol-reactive chloromethyl moiety for labelingmitochondria. TZPs are an essential requirement for healthyfollicle development [12] and the ultrastructure of TZPs ofour in vitro-derived GCs demonstrates a profound confor-mity to that of natural GCs. This data demonstrate the es-tablishment of an interface, that is, the major control sitebetween the oocyte and GCs in vitro. TEM analysis showedthat TZPs penetrate a mucous substance, which appears as agray homogeneous material (Fig. 2E, F). Cumulus cells syn-thesize and secrete hyaluronic acid as well as produce ahyaluronan-rich ECM matrix [55,56]. The ultrastructure ofthe ECM matrix of the in vivo cumulus–oocyte complex (Fig.2H) closely resembles that of the matrix penetrated by theTZPs of the in vitro-derived GCs (Fig. 2E).

Comparative morphological analysis of ESC-derived oo-cytes and in vivo oocytes revealed remarkable similarities(Fig. 5A–H). The oocyte surface varies greatly from one stageof the cell cycle to the other and even within a given stage,suggesting a close association between the surface charac-teristics and the maturation status of the oocyte [57]. A ma-ture, unfertilized mouse oocyte is characterized by smallblebs and sparse, relatively uniform microvilli [58] as wellas a clear mosaic surface, that is, a microvillous membranewith a smooth, microvillous-free polar region. This region isrelatively free of organelles [59] and has fewer CGs [60]. SEManalysis of in vitro-derived oocytes from late cultures re-vealed a mosaic surface topography typical for mature un-fertilized oocytes (Fig. 5D).

TEM analysis demonstrated a background nucleoplasmthat was paler than the background cytoplasm (Fig. 5E).ESC-derived oocytes exhibited slight vacuolization of oo-plasm, a feature typical for immature and aged oocytes [61].Mature oocytes showed spontaneous or vesicle-fused va-cuolization, with the vacuoles usually filled with fluid [62].Culture-derived oocytes had numerous mitochondria ofvaried shapes (Fig. 5F). These mitochondria exhibited anelectron-dense matrix and pale, irregularly arranged cristae,a typical feature of fetal mouse oocytes [63]. Some mito-chondria showed a vacuolated matrix, which has been de-scribed for oocytes of polyovular and unilaminar follicles inthe golden hamster [41]. Round, oval, and elongated mito-chondria are found in mouse growing oocytes [63] and

8 PSATHAKI ET AL.

contain large intermitochondrial vacuoles, as shown in Fig.5F (arrowhead). Numerous fibrillar arrays were detectedamong the mitochondria of our in vitro-derived oocytes (Fig.5E, insert). This arrangement is seen in mature oocytes andcan only be detected in glutaraldehyde-fixed samples [25]. Inthe mouse, these fibrils form polysomes and a hexagonalcrystalline pattern. The fibrillar bundle seen in Fig. 5E (insert)appeared long, without any pattern, and was not associatedwith ribosomes as described by Zamboni [25]. Future workshould address whether this nonassociation with ribosomescan be attributed to a smaller size, different metabolic ac-tivity, or a thinner and atypical zona pellucida of the oocyte.

The formation of cortical granules begins during oocytematuration [25] and the distribution of cortical granules inthe mouse oocyte cortex changes dynamically during meioticmaturation [60]. Cortical granules are round or elliptical inshape, measure 0.2–0.5mm in diameter, and consist of ahighly dense matrix surrounded by a single smooth mem-brane [25]. As described by Ducibella et al. [60], in immatureoocytes, cortical granules are distributed asymmetricallythroughout the cortex, whereas in mature oocytes, they arelocalized mainly to the cortex periphery and are fewer innumber. TEM analysis of ESC-derived oocytes showed nu-merous cortical granules distributed over the entire cortex(Fig. 5F), with only a few granules at the oocyte periphery(Fig. 5F, insert), which is indicative for immature oocytes.The distribution of cortical granules and their competence toundergo cortical reaction are important indicators of oocytecytoplasmic maturation and provide valuable informationabout the fertilizable lifespan of the oocyte [60]. Overall,these ultrastructural data suggest that the analyzed ESC-derived oocytes resembled immature and growing wild-typeoocytes. We carried out some IVF experiments with oocytesderived under the tested differentiation conditions and atlater culture time points, but were not able to achieve fer-tilization. The immature nature and the low yields of oocytesimpeded the appropriate experimental setup for oocyte ac-tivation and subsequent IVF and ICSI. For the ultimate proofof functionality, our differentiation conditions need to befurther improved to obtain higher yields of oocytes.

In vivo, only few follicles ovulate; the vast majority de-generate and die [64] during atresia, a process that can occurat any stage of follicular development [65]. This process as-sures adequate oocyte quality, prevents the survival of oo-cytes with crossover chromosome defects, and limits thenumber of oocytes within the growing follicle [66]. TEManalysis of in vitro-derived oocytes revealed signs of apo-ptosis and autophagy (Fig. 5E, H), which are indicative ofatresia. As shown in Fig. 5E, ESC-derived oocytes had losttheir spherical shape, a structural feature that was used byEscobar et al. to select for atretic rat oocytes. These oocytesshowed clear vesicles, numerous lysosomes, and autopha-gosomes as well as dark cytoplasmic structures, all signs ofautophagy [67]. Signs of apoptosis, such as clumps of com-pact chromatin, apoptotic bodies, and membrane blebbing,were not observed, consistent with the results of Devine andcoworkers [65]. Ribbon-like fibril bundles were foundaround the nucleus, a finding previously reported by Ortizet al. [68] in atretic oocytes of rat antral follicles. In com-parison, in vitro-derived oocytes, such as the one shown inFig. 5H, exhibited signs of apoptosis. Numerous apoptoticbodies and secondary lysosomes were distributed within the

breaking, condensed cytoplasm. Mitochondria could not beidentified and the zona pellucida, organelles, and most of theoocyte surface microvilli were lost. Escobar et al. [67] pro-posed that oocyte cell death probably starts with autophagicdegradation of cytoplasmic components, such as mitochon-dria. It therefore appears that the in vitro-derived oocyteshown in Fig. 5E was analyzed at an initial phase of celldeath, whereas the oocyte shown in Fig. 5H was isolated atan advanced stage of apoptotic cell death. Natural mouseoocytes derived from 18.5-dpc PGCs (Fig. 5G) exhibitedsimilar features when grown under the same culture condi-tions, indicative of ongoing atresia. All oocytes shown in Fig.5 were freely floating in the culture media at the time ofcollection, that is, had already been extruded from the folli-cle-like structures. At this time, it is not known whether theautophagic and/or apoptotic signs shown by our in vitro-derived oocytes are a consequence of the culture conditionsor are caused by apoptotic-inducing stimuli present in thecellular milieu.

Although the derivation and culture of female germ cellsfrom pluripotent stem cells in vitro may lead to ultrastruc-tural alterations in oocytes, this study demonstrates that asimple in vitro culture system can create the essential com-ponents required for oocyte development: oocytes, an ECM-based interface, and granulosa-like cells. Future efforts tooptimize our in vitro differentiation conditions should bringus closer to our ultimate goal of generating fully functionaloocytes and demonstrate whether all key events of germ celldevelopment in vivo can be truthfully recapitulated in vitro.

Acknowledgments

The authors thank J. Lange, M. Sinn, C. Ortmeier, andG. Verberk for providing technical assistance, L. Gentile andT. Esteves for providing helpful discussions, and the De-partment of Zoology (University of Osnabruck) for grantingour group access to their electron microscope. This work wassupported by the Max Planck Society and the DFG grant forResearch Unit Germ Cell Potential (FOR 1041).

Author Disclosure Statement

The authors declare that no competing financial interestsexist.

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Address correspondence to:Dr. Hans R. Scholer

Department of Cell and Developmental BiologyMax Planck Institute for Molecular Biomedicine

Rontgenstraße 20D-48149 Munster

Germany

E-mail: office@mpi-muenster.mpg.de

Received for publication December 18, 2010Accepted after revision January 17, 2011

Prepublished on Liebert Instant Online Month 00, 0000

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