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Biology of Reproduction 63, 1014-1023 (2000)
© 2000 Society for the Study of Reproduction, Inc.

ARTICLES

Development of Mouse and Rat Oocytes in Chimeric Reaggregated Ovaries after Interspecific Exchange of Somatic and Germ Cell Components1

John J. Eppig2,a, and Karen Wiggleswortha

a The Jackson Laboratory, Bar Harbor, Maine 04609


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The germ cell and somatic cell compartments of newborn rat and mouse ovaries, which contain only primordial stage follicles, were completely exchanged and reaggregated to produce xenogeneic chimeric ovaries. The reaggregated ovaries were grafted beneath the renal capsules of ovariectomized SCID mice to develop for periods up to 21 days. Xenogeneic follicles developed with essentially normal morphological characteristics. Both rat and mouse oocytes with species-specific characteristics grew within follicles that were composed of somatic cells exclusively of the alternative species. Rat oocytes grown in mouse follicles became competent to resume meiosis, and progressed to metaphase II when they were removed from follicles and cultured. In addition, mouse oocytes grown in rat follicles underwent fertilization and preimplantation development in vitro, and developed to term after embryos were transferred to pseudopregnant mouse foster mothers. Therefore, despite an estimated 11 million years of divergent evolution, oocytes and somatic cells of rat and mouse ovaries can be exchanged and can produce functional oocytes. It is concluded that factors involved in oocyte-somatic cell interactions necessary to support oocyte development and appropriate differentiation of the oocyte-associated granulosa cells are conserved between rats and mice. Moreover, although granulosa cells play important roles in oocyte development, the development of species-specific characteristics of oocytes occurs without apparent modification by a xenogeneic follicular environment.

developmental biology, follicle, follicular development, gametogenesis, meiosis, oocyte development, ovary, ovum


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Oocyte development requires a complex interaction of intrinsic programming and coordinated communication of the germ cell with companion somatic cells. Oocytes undergo extensive growth, increasing in diameter from about 20 µm to 80 µm, or more in some species. During their growth, oocytes accumulate essential structural, informational, and regulatory molecules to complete meiosis, fertilization, and early embryogenesis. They also acquire species-specific morphological characteristics and synthesize the principal components of the zona pellucida, which limits the number and species of sperm that can enter the egg. Oocyte growth depends upon interactions with companion somatic cells. These interactions are mediated by gap junctions, which metabolically couple the two cell types [1–3] and by paracrine factors such as KIT ligand (KL) [4–6] and other uncharacterized molecules [7].

Like oocyte growth, the progression of meiosis is also regulated by interactions between granulosa cells and oocytes. Fully grown oocytes are maintained at prophase of meiosis by arresting factors transmitted from granulosa cells to oocytes via gap junctions [8–10]. In addition, the resumption of meiosis is believed to be induced by signals transmitted by this same pathway [11, 12].

Communication between oocytes and companion granulosa cells is bidirectional. Oocytes profoundly influence the development and function of the granulosa cells associated with them. Factors secreted by oocytes are critical for the stimulation of cumulus expansion [13, 14], the promotion of granulosa cell proliferation [15, 16], suppression of progesterone production [17–20], and regulation of the expression of luteinizing hormone receptor and KL mRNAs [21–23]. The oocyte-produced growth differentiation factor-9 is probably one of the important paracrine factors in oocyte control of granulosa cell differentiation [24–26], but oocytes secrete many proteins throughout their development, the functions of which are unknown at this time [27].

Differences among mammalian species in the factors that mediate oocyte-granulosa cell communication have not been well-resolved; however, some differences are believed to exist. For example, although oocyte factors are critical for the stimulation of cumulus expansion in rats and mice, expansion of the cumulus oophorus in cattle and pigs seems to occur independently of such factors; yet, one or more factors secreted by oocytes of these domestic species can promote expansion of the murine cumulus oophorus [28–30]. Nevertheless, we hypothesize that fundamental mechanisms governing oocyte development and the interactions of oocytes with their companion somatic cells are evolutionarily conserved.

In now classic experiments, spermatozoa that developed from rat spermatogonia within mouse seminiferous tubules acquired a morphological phenotype that was typical of rat sperm. Moreover, the pace of xenogeneic spermatogenesis after transplantation was intrinsic to the genotype of the germ cell [31–33]. Thus, the complex intercellular communication between the male germ cell and its supporting somatic cells was sufficiently similar between these species, which diverged an estimated 11 million years ago [34], to promote apparently normal xenogeneic spermatogenesis. The species-specific characteristics of development, however, were governed by intrinsic germ cell programs.

To address similar questions regarding the role of germ cell-somatic cell interactions in oocyte development, it was necessary to completely exchange the oocyte and somatic cell components of ovaries. To achieve this, we adapted and improved a method that was originally described by O and Baker [35] in studies in which germ cell and somatic cell components were exchanged between male and female fetal gonads. We used this method to produce reaggregated chimeric ovaries and completely exchanged the germ cell and somatic cell compartments of newborn rat and mouse ovaries, which contain only primordial follicles. The chimeric reaggregated ovaries were grafted beneath the renal capsules of immuno-compromised SCID mice for development. We then addressed the following questions: Will morphologically normal-appearing rat/mouse xenogeneic follicles form and develop? If so, will oocytes grow and acquire species-specific characteristics in xenogeneic follicles? Will meiotic arrest be sustained in oocytes in a xenogeneic follicular environment? Does oocyte development in a xenogeneic follicular environment affect the species specificity of sperm penetration? Do oocytes developing in a xenogeneic environment acquire competence to undergo fertilization and embryo development?


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals

B6SJLF1 and B6,129S-Gtrosa26 (ROSA26) mice were reared in the research colonies of the investigators. ROSA26 mice ubiquitously express a bacterial lacZ gene [36]. Neonatal Sprague-Dawley rats were provided by the specific pathogen-free facility of The Jackson Laboratory's Importation Unit. The parental rats were originally obtained from Taconic (Germantown, NY).

Preparation of Chimeric Ovaries

The method to exchange the oocyte and somatic cell components of ovaries takes advantage of differences between germ and somatic cells in their adherence to a culture dish after dissociation into single cells. Living somatic cells adhere tightly, whereas germ cells do not [37]. Ovaries from 6–8 newborn mice were pooled in plastic Petri dishes containing Dulbeccos PBS (GIBCO-BRL, Grand Island, NY) with 1 mg/ml BSA, crystallized and lyophilized (ICN, Costa Mesa, CA), and then removed from their bursae. The solution used for ovarian dissociation was calcium- and magnesium-free Hanks buffered salt solution with 0.05% trypsin and 0.53 mM EDTA (GIBCO-BRL), supplemented with 0.02% DNase-I (Sigma Chemical Co., St. Louis, MO) and maintained at 37°C. Dissociation into a single cell suspension was aided by frequent gentle agitation by repeatedly drawing the tissue in and out of a Pasteur pipette. After complete dissociation, the cell suspension was transferred to a 15-ml centrifuge tube containing an equal volume of warmed Medium 199 (M199; GIBCO-BRL), prepared as described previously [38] and supplemented with 10% fetal bovine serum (FBS). Tubes were centrifuged for 5 min at 2000 x g to gently pellet the cells. After centrifugation, the supernatant medium was removed and the pellet was gently resuspended in 3 ml M199/FBS. The resuspended cells were transferred to a tissue culture dish and cultured in modular incubation chambers with an atmosphere of 5% CO2, 5% O2, and 90% N2, and incubated at 37°C.

After overnight culture, most of the viable somatic cells were attached to the culture dish, whereas the unattached cell population consisted of oocytes, nonviable somatic cells, red blood cells, and a few rounded mitotic somatic cells. To remove the unattached cells, including oocytes, the dish was gently swirled so that unattached cells moved to the center, where the cells were then carefully removed (without dislodging the monolayer underneath) and transferred to a new tissue culture dish. Sufficient fresh M199/FBS was added to adjust the volume to about 3 ml. The monolayer of somatic cells remaining in the original dish was then rinsed free of any remaining unattached cells by vigorously washing the layer with PBS/BSA three times, and the rinse medium was discarded. The monolayered cells were removed from the tissue culture dish by treatment with 1.7 ml trypsin/EDTA without DNase. The cells were then transferred to a centrifuge tube, an equal volume of fresh M199/FBS was added, and the tube was centrifuged at 2000 x g for 5 min to pellet the cells. After pelleting the cells, the supernatant medium was removed and cells were resuspended in 3 ml of M199/FBS. This cell suspension was then transferred to a new tissue culture dish and both groups of cells were cultured for an additional 6 h. This second round of differential adhesion was found to be essential for complete separation of cell types.

A similar procedure was used to dissociate 1- and 2-day-old rat ovaries and separate the germ cells and somatic cells. However, it was found that the rat somatic cells did not adhere as firmly as mouse somatic cells; therefore, the procedure was slightly modified for the rat cells by coating the culture dish with extracellular matrix before the cells were added. Dishes were coated with components of extracellular matrix (i.e., entactin, collagen IV, and laminin—ECL). These coated dishes were prepared by adding a 20 µg/ml aqueous solution containing these components purified from the Englebreth Holm-Swarm mouse tumor to the dishes and incubating them for 1 h at 37°C as described by the supplier (Upstate Biotechnology, Inc., Lake Placid, NY). The dishes were then washed thoroughly with M199/FBS.

After the 6 h culture for the second differential adhesion step, both the oocytes and somatic cells were collected and washed as described earlier. The supernatant was removed and discarded and the pellets were each resuspended in 100 µl of M199/FBS. The different cell types, either from the same or different species, were then mixed in microcentrifuge tubes and the volume was adjusted to 1 ml. Phytohemagglutinin (PHA; Sigma Chemical Company) was added to a final concentration of 35 µg/ml. PHA facilitates the removal of the pellet with a Pasteur pipette after centrifugation (10 000 x g for 1 min) by promoting the adhesion of the cells.

Pellets were then cultured overnight as described previously for intact newborn mouse ovaries [39]. Briefly, the reaggregated ovaries were transferred with a Pasteur pipette and a drop of medium to a Millicell-PC membrane (3-µm pore size, 24-mm diameter; Millipore Corp., Bedford, MA). Approximately 1.5 ml of medium had been placed in the compartment below the membrane insert so that when the drop of medium was added to the surface of the membrane, excess medium was drawn into the compartment below the membrane, leaving the reaggregated ovary pellet covered by only a thin film of medium. The medium for pellet culture was Waymouths MB752/1 (GIBCO-BRL) supplemented with 0.23 mM pyruvic acid, 50 mg/L streptomycin sulfate, 75 mg/L penicillin-G (Sigma) and 10% FBS. Samples were cultured at 37°C in modular incubation chambers (Billups Rothenberg, Del Mar, CA) with an atmosphere of 5% CO2 and 95% air. After overnight culture, the reaggregated ovary pellets were surgically implanted beneath the renal capsule of bilaterally ovariectomized SCID mice.

Assessing Oocyte and Follicle Development

At 7, 14, and 21 days after grafting beneath the renal bursa, reaggregated ovaries were removed and fixed for 3–5 h in 2.5% glutaraldehyde and 2.5% paraformaldehyde in 0.083 M sodium cacodylate buffer, pH 7.2, at 4°C. They were then washed in 0.1 M sodium cacodylate buffer, pH 7.2, for 24 h before embedding in JB-4 (glycol methacrylate) plastic (Polysciences, Inc., Warrington, PA). Sections of 2-µm thickness were stained with Periodic Acid-Schiff (PAS) reagent and hematoxylin. The method of Gossler et al. [40] was used for localization of lacZ expression by ROSA26-derived cells. For electron microscopy, the aldehyde-fixed samples were postfixed in 1% osmium tetroxide and stained en bloc with uranyl acetate using standard procedures.

Oocyte development was assessed 20–22 days after grafting. Oocytes were isolated by puncturing the antral follicles with 25-gauge needles, and all oocytes were collected, whether or not they were enclosed by cumulus cells. The cumulus-enclosed oocytes were then denuded to make it possible to accurately score whether the oocytes were in germinal vesicle (GV) stage or had undergone maturation before isolation. GV-stage oocytes were matured in Whittens medium supplemented with 20% FBS as described in detail elsewhere [41–43]. All groups of oocytes were matured 16–17 h at 37°C in modular incubation chambers infused with 5% O2, 5% CO2, and 90% N2. After maturation, the progression of meiosis as indicated by the presence or absence of either the GV or a polar body was assessed. Mature oocytes with a polar body were designated as metaphase II-arrested oocytes, those that had undergone germinal vesicle breakdown (GVB) without forming a polar body were designated as metaphase I-arrested oocytes. Metaphase II-arrested mouse oocytes were inseminated, and those that had cleaved to the 2-cell stage 24 h later are referred to as fertilized. The 2-cell stage embryos were cultured exactly as described [43–45] to determine percentage of embryos developing to the expanded blastocyst stage. Fertilization of mature rat oocytes was not attempted. Morula- and blastocyst-stage mouse embryos were transferred to the uterus of pseudopregnant foster mothers, offspring were cesarean delivered 17 days after embryo transfer, and uteri examined for macroscopic fetal resorption sites.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Morphological Aspects of Xenogeneic Follicle and Oocyte Development

The ovaries of newborn rats and mice do not contain follicles that have developed beyond the primordial stage. After reaggregation, the ovaries were disorganized, but by 7 days after grafting, the morphology of primary and secondary follicles beneath the renal capsule appeared essentially normal (Fig. 1). An advantage of PAS staining of sections is that it allows visualization of an obvious difference between mouse and rat oocytes, one that has been described as polar cytoplasm or "nuage" in rat oocytes [46]. In primary follicles, red PAS staining usually appears uniformly throughout the cytoplasm of rat oocytes, but in secondary follicles, PAS-positive granules become eccentrically localized (Fig. 1A). This characteristic was also observed in rat oocytes that developed within mouse follicles (Fig. 1B). PAS staining also enabled visualization of the formation of the zona pellucida, regardless of syngeneic or xenogeneic follicular development of oocytes (Fig. 1).



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  		FIG. 1. Photomicrographs of reaggregated ovaries 7 days after grafting beneath the renal capsules of SCID mice. A) Mouse oocytes and somatic cells. B) Mouse oocytes and rat somatic cells. C) Rat oocytes and somatic cells. D) Rat oocytes and mouse somatic cells. Bar = 100 µm. FIG. 2. Photomicrographs of reaggregated ovaries 14 days after grafting beneath the renal capsules of SCID mice. A) Rat oocytes and mouse somatic cells. B) Mouse oocytes and rat somatic cells. Arrows in A point out polarized nuage granules. Bar = 100 µm

Secondary and early antral follicles were present 14 days after grafting beneath the renal capsule (Fig. 2). When rat oocytes developed in mouse follicles, there was a tight apposition of oocytes with their companion granulosa cells (Fig. 2A). However, the contact of rat granulosa cells with mouse oocytes appeared to be less intimate because regions of the oocyte were devoid of contact with granulosa cells (Fig. 2B). This condition was also observed when rat oocytes developed in rat follicles (not shown) and may have been related to the development of rat ovarian tissue at the extragonadal site of SCID mice. Nevertheless, it is obvious that oocyte growth, zona pellucida formation, and eccentric deposition of nuage occurred despite the apparently limited association of rat granulosa cells with the oocyte.

Large antral follicles were present 21 days after grafting of reaggregated ovaries (Fig. 3). Examination of oocytes with an electron microscope revealed the development of apparently normal fine structure in xenogeneic oocytes (Fig. 4). All oocytes were surrounded by a zona pellucida that was traversed by cytoplasmic processes that originated from the cumulus cells and terminated on the surface of the oocyte. The overall fine structural characteristics of rat and mouse oocytes appeared essentially as has been described elsewhere in detail [47–49]. Structures that have been described as "ribosomal lattices" in mouse oocytes [49, 50] and "membrane packets" in rat oocytes [48] established their species-specific characteristics despite xenogeneic oocyte development (Fig. 4).



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  		FIG. 3. Photomicrograph of a reaggregated ovary, rat oocyte, and mouse somatic cells 21 days after grafting beneath the renal capsule of an SCID mouse. Arrow indicates polarized nuage granules in rat oocyte. Bar = 100 µm



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  		FIG. 4. Electron micrographs. A) A mouse oocyte in a rat follicle. B) A rat oocyte in a mouse follicle 21 days after grafting the reaggregated ovary beneath the renal capsules of SCID mice. Arrowheads indicate cytoplasmic processes from cumulus cells traversing the zona pellucida. Arrows in A indicate crystalline lattices characteristic of mouse oocytes and in B the arrows delineate membrane packets that are distinctive of rat oocytes. Magnification: A, x7590; B, x9480

To assess the effectiveness of the oocyte-somatic cell separation and exchange protocols, the two cell types were exchanged between rats and ROSA26 mice. Cells derived from ROSA26 mice were identified by blue staining after incubation with X-gal. As demonstrated in Figure 5, the exchange of oocytes and somatic cells was complete; no cross-contamination of the preparations was observed. This established xenogeneic oocyte and follicle development with certainty.



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  		FIG. 5. Photomicrographs demonstrating the complete exchange of somatic cells and oocytes between the ovaries of rats and ROSA26 mice. Panels A and B show chimeric reaggregated ovaries containing ROSA26 oocytes (blue) and rat somatic cells. Panels C and D show chimeric reaggregated ovaries containing ROSA26 somatic cells (blue) and rat oocytes. Ovaries were grafted beneath the renal capsule of ovariectomized mice for 9 days. A, C) Bar = 100 µm. B, D) Bar = 50 µm

Functional Aspects of Xenogeneic Oocyte Development

To assess functional aspects of xenogeneic oocyte development, fully grown oocytes were isolated from antral follicles 21 days after grafting beneath the renal capsule. Cumulus cells were removed from oocytes that were still cumulus cell-enclosed after isolation, and the oocytes were examined for the presence of a GV. Seventy-eight percent of the mouse oocytes that developed in mouse follicles and 87% of the mouse oocytes that developed in rat follicles were at the GV stage (Table 1). After culturing the GV-stage mouse oocytes for 15 h, the percentage of oocytes progressing to metaphase II when developed in syngeneic or xenogeneic follicles was 80% and 81%, respectively (Table 1). When metaphase II oocytes were inseminated, 45% and 47% of the oocytes developed in syngeneic or xenogeneic follicles, respectively, cleaved to the 2-cell stage. Moreover, 17% of the oocytes from each group developed to the blastocyst stage (Table 1; Fig. 6). Finally, one live offspring derived from both groups was produced after transfer of embryos to pseudopregnant foster mothers. Similar numbers of resorbed fetuses were observed in both groups (Table 1). The female mouse, named Romula, derived from an oocyte of a xenogeneic follicle, has given birth to 83 apparently normal offspring.


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  		TABLE 1. Maturation and developmental competence of mouse oocytes after isolation from reassembled ovaries composed of mouse or rat somatic cells



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  		FIG. 6. Photomicrographs of morula/blastocyst stage mouse embryos derived from oocytes of reaggregated ovaries. A) Morula/blastocyst stage mouse embryos derived from oocytes that developed in mouse (syngeneic) follicles. B) Morula/blastocyst stage mouse embryos derived from oocytes that developed in rat (xenogeneic) follicles. Nomarksi interference optics. Bars = 100 µm. FIG. 7. Photomicrographs of rat eggs derived from oocytes of reaggregated ovaries and matured in vitro from GV-stage oocytes. A) Rat oocytes grown in rat follicles. B) Rat oocytes grown in mouse follicles. Nomarski interference optics. Bars = 100 µm

Fifty-two percent of the rat oocytes that developed in rat follicles and 68% of the rat oocytes that developed in mouse follicles were at the GV stage upon isolation (Table 2). When GV-stage rat oocytes were cultured for 15 h, 55% and 62% matured to metaphase II after growth and development in syngeneic and xenogeneic follicles, respectively (Table 2; Fig. 7). No attempt was made to inseminate rat oocytes.


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  		TABLE 2. Maturation competence of rat oocytes after isolation from reassembled ovaries composed of mouse or rat somatic cells


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
This study shows that, despite 11 million years of divergent evolution, oocytes and somatic cells of rat and mouse ovaries can be exchanged and can produce functional oocytes. Rat oocytes can grow in mouse follicles, establish rat-specific morphological characteristics, and become competent to resume meiosis and progress to metaphase II. In addition, mouse oocytes grown in rat follicles can undergo fertilization and preimplantation development in vitro, and develop to term after embryos are transferred to pseudopregnant foster mothers.

To produce a functional egg requires coordinated interactions between developing oocytes and their companion somatic cells as well as autonomous developmental programming. Complex communications between granulosa cells and an oocyte, which involve gap junctions and paracrine factors, are required to promote oocyte growth and regulate meiosis. Clearly, the factors that mediate this communication are sufficiently similar in rats and mice to allow the development of apparently normal, fully grown oocytes that are capable of completing meiosis and undergoing fertilization and embryogenesis.

Since the classic experiments of Pincus and Enzmann [8] and Pincus and Saunders [51], it has been realized that factors derived from follicular somatic cells function to maintain fully grown oocytes in meiotic arrest until the preovulatory surge of gonadotropins. The fundamental observation forming the basis of this conclusion was that removal of the oocyte from the follicle and culture resulted in "spontaneous" gonadotropin-independent resumption of meiosis. Whether the somatic cells of all mammalian species use the same meiosis-arresting mechanisms could be questioned because of apparent differing sensitivity to potential meiosis-arresting substances. For example, KL seems to delay the onset of GVB in rat but not mouse oocytes in vitro [52, 53] (and B. Vanderhyden, personal communication). Nevertheless, membrane permeable analogues of cAMP are effective at sustaining meiotic arrest in both mouse and rat oocytes in vitro [54, 55], suggesting that cAMP is important for maintaining meiotic arrest in both species. Here it has been shown that rat and mouse oocytes isolated from xenogeneic follicles undergo spontaneous maturation in vitro. Thus, meiotic arrest had been sustained in vivo until the oocytes were liberated from the follicles. This supports the idea that similar mechanisms sustain meiotic arrest in vivo in both species; however, because GV-stage oocytes were isolated from follicles and matured in vitro, the mechanisms involved in the induction of oocyte maturation in vivo were not assessed.

Oocytes prepare for embryogenesis by accumulating essential informational, structural, and regulatory molecules during growth. Evidence suggests that interactions between granulosa cells and growing oocytes are critical for the acquisition of competence to complete preimplantation development [21, 56]. Inappropriate differentiation of oocyte-associated granulosa cells can have detrimental effects on the acquisition of competence to develop to the blastocyst stage [21, 56]. Oocytes probably promote the appropriate differentiation of oocyte-associated granulosa cells in ways that benefit their own development [27, 57]. Thus, there appears to be a loop of communication between oocytes and their companion somatic cells that regulates the mutual development and function of both cell types. This regulatory loop appears to be functional in xenogeneic follicles because eggs could complete preimplantation development and produce live offspring. The frequency at which oocytes completed preimplantation development was less than the frequencies that we have reported for in vitro-matured oocytes [58]. This is probably due to a combination of factors, including those associated with the unusual conditions of oocyte development in reaggregated ovaries grafted to an extragonadal site beneath the renal capsule. In addition, oocytes were denuded of cumulus cells before maturation in vitro in order to evaluate their stage of meiosis. Nevertheless, oocytes grown in xenogeneic follicles displayed essentially the same competence to complete meiosis and undergo embryo development as oocytes grown in syngeneic follicles.

Morphologically normal rat sperm formed after transfer of rat spermatogonia to germ-cell free mouse seminiferous tubules [31]. Although sperm were not functionally assessed, they developed morphological characteristics, and at the same pace, as sperm that develop in rat testes [32, 33]. Likewise, it is shown here that species-specific characteristics of oocytes develop in a xenogeneic environment. The formation and polarity of nuage and membrane packets in rat oocytes and ribosomal lattices in mouse oocytes were obvious species-specific morphological characteristics. In rodents, the zona pellucida is produced exclusively by the oocyte and presents an effective barrier to fertilization by more than one sperm, or by sperm of a different species [59, 60]. The fertilization of mouse eggs with mouse sperm that developed in rat follicles further demonstrates the autonomous formation of a functional, species-specific zona pellucida by mouse oocytes.

Gap junctions are membrane specializations that allow the transfer of low molecular weight molecules from one cell to another. Optimal oocyte growth depends upon gap junction-mediated communication between an oocyte and its companion granulosa cells. In fact, the rate of growth of an oocyte is directly related to the number of granulosa cells coupled to it [3]. Because of gap junctional communication, granulosa cells functionally increase the surface area of an oocyte and thereby increase its surface-to-volume ratio. This increases the rate at which small molecules having nutritional or regulatory functions enter an oocyte. Moreover, mammalian oocytes are apparently deficient in the transport systems required for the entry of some molecules. Gap junction-mediated transport of these molecules, which are initially taken up by granulosa cells, is required for metabolic processes that are essential for oocyte growth and development [2, 61–64]. Oocyte and follicular development are both dramatically affected in mice that are deficient in the gap junction protein, connexin-37 [65], which is encoded by the gene Gja4. In Gja4null mice, oocytes fail to complete their growth phase and follicles undergo premature luteinization [65]. Clearly, then, functional gap junctional communication between oocytes and companion granulosa cells was established in the rat-mouse xenogeneic follicles described here. Gap junctional connexins associate with partners in apposing plasma membranes. Therefore, rat-mouse gap junctions must be functional between oocytes and granulosa cells in xenogeneic follicles. That this can happen was originally demonstrated using rat granulosa cells and mouse myocardial cells in vitro [66]. The results presented here show that complex developmental interactions can utilize these heterologous pathways in vivo.

Although there are some notable morphological differences between mouse and rat oocytes, the follicles and oocytes of these two species exhibit generally similar characteristics in structure and developmental dynamics. Therefore, the relative roles of granulosa cell-oocyte interactions or autonomous programming in the pace of either oocyte or follicle development cannot be determined using rat-mouse xenogeneic ovaries. However, the experimental paradigm for such investigation is established by the studies presented here. It will be exciting to determine, for example, whether the kinetics of bovine oocyte development, which are very different from those of mouse oocytes, are sustained when these oocytes develop in mouse follicles. Perhaps bovine oocyte development will be accelerated in mouse follicles. Similarly, it will be interesting to determine whether the unusual features of the ovarian follicle of species such as the musk shrew [67–69] occur when they are under the influence of a mouse oocyte.


   ACKNOWLEDGMENTS
 
We are grateful for suggestions from Drs. Wes Beamer, Barbara Vanderhyden, Rabin De La Fuente, Ieuan Joyce, Rob Taft, and Maria Viveiros in the preparation of this manuscript. In addition, we thank Leslie Bechtold, Deborah Lane, and Gregory Martin of the Biological Imaging Service, and Jennifer Smith of the Multi Media Services, for their assistance in electron microscopy, histology, and preparation of photomicrographs.


   FOOTNOTES
 
First decision: 3 May 2000.

1 This research was funded by the National Institute of Child Health and Human Development (NICHD) through NIH grant HD23839. The development of culture systems was supported by The National Cooperative Program on Non-Human In Vitro Fertilization and Preimplantation Development, which was funded by NICHD through cooperative agreement HD21970. The scientific services of the Jackson Laboratory receive support from a Cancer Center Core Grant (CA34196) from the National Cancer Institute. Back

2 Correspondence. FAX: 207 288 6073; jje{at}jax.org Back

Accepted: May 17, 2000.

Received: April 11, 2000.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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