When the Y chromosome of Mus musculus domesticas (YDOM) was introduced onto the C57BL/6 (B6) mouse background, half of the XY progeny (B6.YDOM) developed bilateral ovaries and female internal and external genitalia. We examined the fertility of the B6.YDOM sex-reversed female mouse. The chromosomal sex of the individual mouse was identified by dot hybridization with mouse Y chromosome-specific DNA probes. The results indicated that all XY females lacked regular estrous cyclicity although most were able to mate and ovulate after treatment with gonadotropins. When they had been ovariectomized and grafted with ovaries from the XX female litter mate, they initiated estrous cycli-city. Reciprocally, the XX female that had received XY ovarian grafts did not resume estrous cyclicity.

Development of the XY ovary was morphologically comparable to the XX ovary until 16 day of gestation (d.g.), when most germ cells had reached the zygotene or pachytene stage of meiotic prophase. However, by the day of delivery (19 or 20 d.g.), no oocyte remained in the medullary cords of the XY ovary. In the control XX ovary, the first generation of follicles developed in the medullary region, and 5Δ-3β-hydroxysteroid dehydrogenase (3β-HSDH) activity appeared first in the stromal cells around growing follicles by 10 days after birth. In contrast, in the XY ovary, follicles were not formed in the medullary region, and 3β-HSDH activity appeared in epithelial cells of the oocyte-free medullary cords. Primordial follicles in the cortex region continued development in both the XX and XY ovaries.

These results suggest that the XY female is infertile due to a defect inside the XY ovary. The prenatal loss of oocytes in the medullary cords may be a key event leading to abnormal endocrine function, and thereby, the absence of estrous cyclicity.

It has been generally accepted that the presence of the Y chromosome determines the development of testes, and its absence results in the development of ovaries in mammals. Page and others have recently isolated a DNA fragment of human Y chromosome (ZFY), which is conservatively present on the Y chromosome of most mammalian species examined (Page et al. 1987). They have shown that the ZFY sequence is translocated onto either the X chromosome or autosomes of most XX male patients, translocated onto the X chromosome of the XX sex-reversed (Sxr) male mouse, and deleted from the Y chromosome of some XY female patients. These results suggest that the ZFY region contains the gene responsible for primary testis-determination (called TDF in human or Tdy in the mouse).

On the other hand, Eicher and others have reported that, when the Y chromosome from Mus musculus domesticus (or poschiavinus) is placed onto the C57BL/6J (B6) mouse background, all XY progeny (B6.YDOM) fail to develop normal testes during fetal life, and some develop bilateral ovaries and the female phenotype (Eicher et al. 1982; Eicher & Washburn, 1983). These findings suggest that the mere presence of the Y chromosome is not sufficient to induce testicular differentiation. Eicher and others have postulated that the Tdy gene from M. m. domesticus (TdyDOM) needs to interact with another dominant autosomal gene (Tda-1) to impose testicular development, and that the TdyDOM gene cannot properly interact with recessive autosomal genes (tda-V) from the B6 mouse strain. We have previously reported that the (tda-1) -related sexreversal can be transferred to other mouse strains, but the effect is less than that seen when the YDOM chromosome is placed onto the B6 background (Naga-mine et al. 19876). The mechanism of the B6.YDOM sex-reversal remains to be investigated.

It was of interest to determine if the B6.YDOM XY sex-reversed female was fertile. Eicher and others have reported that the XY female is infertile with rare exceptions; all the females that had been placed with normal males mated; however, only one female produced a litter (Eicher et al. 1982). They explained that sterility is caused by rapid depletion of oocytes in the XY female between 4 and 8 weeks of age. The cause of oocyte loss in the XY female was not discussed. In the present study, we investigated the infertility of the B6.YDOM XY female mouse.

Preparation of B6. YDOM mice

Male mice carrying the B6 genetic background and the Y chromosome from M. m. domesticus (N9-N13 backcrossing generations) were prepared as previously described (Nagamine et al. 1987a). For matings, each B6.YDOM male mouse (50 – 180 days old) was housed with three B6 female mice (50 – 100 days old) overnight, and the presence or absence of copulation plugs was examined next morning. The day of copulation was defined as 0 day of gestation (d.g.). The day of delivery was defined as 0 day postpartum (pp.). The B6 male and female mice were puchased from Charle’s River (Toronto).

Determination of chromosomal sex

The chromosomal sex of individual mice was determined by dot hybridization with mouse Y chromosome-specific DNA probes according to the methods described previously (Nishioka & Lamothe, 1986; Nishioka, 1988). Briefly, tissue from individual fetuses or the tail tip of neonatal mice (50 – 100 μ g wet weight) was homogenized and incubated with proteinase K (Boehringer Mannheim, W. Germany) and SDS at 60°C for 30 min. The total DNA was extracted with phenol mixture, followed by chloroform and isoamylalcohol. Then the DNA was precipitated with ethanol in the presence of NaCl. This procedure yielded 50-500 μ g DNA/sample. After treatment with NaOH, 2 and 5 μ g DNA of each sample were applied to hybridization transfer membranes (NEN Res., Boston) and air-dried. The membrane sheet was hybridized with 32P-labelled mouse Y-specific DNA probe AC11 or 145SC5, 1x106ctsmin-1 in 10ml hybridization solution overnight at 42°C. The DNA probe was labelled with 32P using a nick translation kit from BRL (Bethesda, ML). After washing four times in 0·l × SSC at 45°C, the membrane sheet was exposed to an X-ray film. The loading of DNA in representative samples was examined by hybridization with non-specific DNA probe R17 (Nishioka, 1989).

Fertility

Each female offspring (50 – 120 days pp.) was housed with a B6 male mouse (50 – 150days pp.) which was known to be fertile. The presence or absence of copulation plugs was examined between 8:00 and 11:00a.m.every morning up to 7 days. If copulation plugs were not found, mating was repeated two more times. The female mouse that had copulated was housed singly until the expected delivery day (19 or 20 d.g.). If pregnancy was not confirmed after 12d.g., this process was’ repeated two more times.

Estrous cyclicity

Female offspring (70 – 90 or 110 – 150 days pp.) were housed singly at least one week prior to examinations. Smears were obtained from the vagina daily between 8:00 and 11:00a.m. for 3 – 4 weeks. The dried smears on microscope slides were fixed in absolute methanol, and stained in 4% Giemsa solution. Smears were then classified as to the stage of estrous cyclicity according to the criteria described by Nelson et al. (1982).

Transplantation of ovaries

Bilateral ovaries were removed from two female mice (XX or XY) (30 – 60 days pp.) at a time, and transplanted beneath the kidney capsule of the reciprocal female mouse (e.g. the right ovary from one mouse into the right kidney of the other mouse). After 2 – 3 weeks post-transplantation, smears were obtained to determine the estrous cyclicity as described above. The female mice that did not show cyclicity were sacrificed and the ovarian grafts examined with light microscopy. The female mice that showed clear or ambiguous estrous cyclicity were kept until 4 months pp., when smears were taken again. After these experiments, females were sacrificed, and the condition of ovariectomy examined. The data were discarded when any residue of the original ovary was found.

Induction of ovulation

Female offspring (25 – 65 days pp.) were injected intraperitoneally with 5i.u. pregnant mare’s serum gonadotropin (PMSG) between 2:00 and 4:00p.m. on the first day, 5i.u. human chorionic gonadotropin (hCG) between 6:00 and 8:00 p.m. on the third day, and sacrificed between 8:00 and 10:00a.m. on the fourth day. (Gonadotropins were purchased from SIGMA Chemicals.) The ovaries and oviducts were removed, and examined under a stereo microscope with transillumination. Ova were flushed out from the oviduct and counted.

Gross anatomy, light and electron microscopy

Bilateral gonads were dissected from offspring between 12 d.g. and 3 months pp., and examined for gonadal structures under a stereomicroscope with transillumination. Specimens were fixed in Karnovsky’s solution (Karnovsky, 1965), postfixed in OSO4, and embedded in Epon. Semithin sections (l μ m thickness) were stained with toluidine blue and examined with a light microscope. Serial sections from, at least, 5 ovaries of each stage were examined. Ultrathin sections were stained with uranyl acetate and lead citrate, and examined under a Zeiss EM9 electron microscope.

Histochemical staining for 3β -HSDH

Fresh tissues were embedded in OCT medium (Tissue-Tek) in liquid nitrogen, and serial sections of 10 μ m thickness were cut with a cryostat. After air-drying on microscope slides overnight, sections were incubated in Levy’s solution using dehyd-roepiandrosterone as a substrate (Levy et al. 1959).

Organ culture and testosterone determination

Bilateral gonads were dissected from fetal offspring of 14 or 16 d.g. and cultured individually for 3 days as described before (Taketo & Koidc, 1981). The basic culture medium was Eagle’s minimum essential medium containing 10% heat-inactivated horse serum, 50i.u. ml− 1 penicillin G sodium, and 50figml− 1 streptomycin sulfate. (All media and chemicals were purchased from G1BCO, New York.) The culture medium was changed every 24 h, and the spent medium was stored at − 20°C. After culture, the expiants were fixed in Bouin’s solution and embedded in paraffin. All serial sections of 5|Um thickness were stained with hematoxylin and eosin. The fifth section from each end and three more sections, evenly distributed in between, were selected and photographed with a light microscope. The ratio of testicular area to whole gonad was estimated by cutting and weighing the photographs of the gonadal area. The concentration of testosterone in the spent medium during the first day of culture was determined by radioimmunoassay using the kit from Radioassay Systems Lab (Carson, California) without extraction of the sample. The cross-reactivity with dehydrotestosterone and androstenedione was 3 · 4 and 0 · 52%, respectively. The lowest concentration of assay for testosterone was 0 · 1 ng ml− 1.

Statistical analysis

All data were analyzed statistically and evaluated using either Chi-square test or Student’s /-test.

Dot hybridization with mouse Y chromosome-specific DNA probes

An example of dot hybridization is shown in Fig. 1. As a control, total DNA was extracted from B6 male and female fetuses of 14 d.g. The DNA from offspring of the B6 female × B6.YDOM male cross gave all- or-none-type spots at an intensity comparable to B6 male or female controls. The DNA from offspring that possessed mono- or bilateral ovotestes (or testes) gave exclusively positive results.

Fig. 1.

Dot hybridization with a mouse Y-specific DNA probe AC 11. The DNA from individual mice at concentrations of 2·0 μ g (top and third rows) and 5·0 μ g (second and bottom rows) were applied onto the membrane. No. 1, B6 male, no. 2, B6 female, no. 3 – 20, offspring from the B6 female X B6.YDOM male cross. No. 5, 7, & 8 were phenotypical males and others were females.

Fig. 1.

Dot hybridization with a mouse Y-specific DNA probe AC 11. The DNA from individual mice at concentrations of 2·0 μ g (top and third rows) and 5·0 μ g (second and bottom rows) were applied onto the membrane. No. 1, B6 male, no. 2, B6 female, no. 3 – 20, offspring from the B6 female X B6.YDOM male cross. No. 5, 7, & 8 were phenotypical males and others were females.

Sex ratio of offspring at various developmental stages

The chromosomal sex and the gonadal sex of offspring are summarized in Table 1. Testes were characterized by the presence of seminiferous tubules, which were clearly distinguishable from other structures with transillumination or in histological sections. Ovaries were identified by the absence of seminiferous tubules, or the presence of oocytes and follicles in histological sections.

Table 1.

Sex ratio of F1 progeny from the B6 female × B6. YDOM male cross at various developmental stages

Sex ratio of F1 progeny from the B6 female × B6. YDOM male cross at various developmental stages
Sex ratio of F1 progeny from the B6 female × B6. YDOM male cross at various developmental stages

At 12 d.g., 94% of 48 fetal progeny possessed sexually undifferentiated gonadal primordia, which had ovarian appearance under a dissecting microscope (data not included in Table 1). At 14d.g., the largest percentage of progeny possessed monolateral or bilateral ovotestes (Table 1). After 16d.g., the ratio of XX females, XY females (with bilateral ovaries), and XY males (with testicular components) was 1:0-5:0-5 throughout development.

Fertility

Six out of nine XY females that had been placed with B6 males mated as evidenced by the presence of copulation plugs (Table 2). None of them were pregnant on 12 day after copulation. In contrast, 95 % of XX females mated with B6 males, and most of them carried the pregnancy to full term.

Table 2.

Fertility of female progeny from the B6 female × B6. YDOM male cross

Fertility of female progeny from the B6 female × B6. YDOM male cross
Fertility of female progeny from the B6 female × B6. YDOM male cross

Estrous cyclicity

All XX females of 70-90 days pp. showed regular estrous cyclicity (3-5 days cycle) (Fig. 2). In contrast, no XY females of the same age showed regular estrous cyclicity, and all stayed at persistent diestrous stage. All of 13 older XY females (110 – 150 days pp.) were in persistent vaginal cornification stage.

Fig. 2.

Estrous cyclicity of the female progeny from the B6 female x B6.YD°M male cross. *, the XX and XY female had been ovariectomized and grafted with ovaries from the XX and XY female (the chromosomal sex of the ovarian graft/that of the host is indicated). Cyclicity +, regular estrous cyclicity; PD, persistent diestrous; PVC, persistent vaginal cornification; PD/PVC, shifted from PD to PVC, and then to PD. The number in parentheses indicates the number of mice examined in each group.

Fig. 2.

Estrous cyclicity of the female progeny from the B6 female x B6.YD°M male cross. *, the XX and XY female had been ovariectomized and grafted with ovaries from the XX and XY female (the chromosomal sex of the ovarian graft/that of the host is indicated). Cyclicity +, regular estrous cyclicity; PD, persistent diestrous; PVC, persistent vaginal cornification; PD/PVC, shifted from PD to PVC, and then to PD. The number in parentheses indicates the number of mice examined in each group.

Estrous cyclicity after transplantation of ovaries

All XY females that had been ovariectomized and grafted with ovaries from the XX female showed regular estrous cyclicity (Fig. 2). They continued estrous cyclicity until, at least, 120 days pp. (data not shown). In contrast, of the seven XX females that had been ovariectomized and grafted with ovaries from the XY female, three stayed in persistent diestrous (PD) stage, three in persistent vaginal cornification (PVC) stage, and one shifted between PVC and PD stages (Fig. 2). No degenerative change was observed in the XY ovarian graft when histologically examined. All control XX females that had been grafted with XX ovaries showed regular estrous cyclicity.

Induction of ovulation

In the XX female, the number of ova released after gonadotropin treatment was the highest at 25 days pp., decreasing to the lowest level at 45 days pp., and increasing again after 55 days pp. (Fig. 3). No significant difference was observed between the right and the left ovaries (data not shown). In the XY female, much smaller numbers of ova were released, reaching the maximum (3 ova/ovary) at 35 days pp. (Fig. 3). At 55 days pp. or later, no ovum was produced by the XY female with only a few exceptions.

Fig. 3.

Number of ova released from each ovary of the female progeny after treatment with gonadotropins. The bar indicates the S.E. of the mean value. The number in parentheses indicates the total number of ovaries examined (including left and right hand sides) in each group. *, significant difference from the control XX female with Student’s t test (P<0·001).

Fig. 3.

Number of ova released from each ovary of the female progeny after treatment with gonadotropins. The bar indicates the S.E. of the mean value. The number in parentheses indicates the total number of ovaries examined (including left and right hand sides) in each group. *, significant difference from the control XX female with Student’s t test (P<0·001).

Fig. 4 shows a representative XY ovotestis at 14 d.g. Sex cords had been dissociated from the surface epithelium by formation of tunica albuginea, and differentiated into testis cords in the mid-portion. Inside the testis cord, germ cells, either in mitotic cell cycle or arrested at the prespermatogonia stage, were enclosed together with fetal Sertoli cells in the basement membrane (Fig. 5). In the cranial and caudal poles, on the other hand, sex cords remained attached to the surface epithelium (Fig. 4). Some germ cells had entered meiotic prophase while others were in mitotic cell cycle (Fig. 6). These structures are characteristics of ovarian differentiation. The location of testicular and ovarian sex cords was consistent in all ovotestes although the ratio of two structures varied among individual gonads.

Fig. 4.

An XY ovotestis with the adjacent mesonephros of the B6.YDOM progeny at 14d.g. Sex cords have differentiated into testis cords (ts) in the mid area. The tunica albuginea (ta) has developed well beneath the surface epithelium. In the cranial and caudal poles, sex cords remain attached to the surface epithelium, characteristics of ovarian differentiation (ov). Note the distribution of stromal cells (st) in the center of the gonad and the mesonephros, md, mesonephric ducts; mt, mesonephric tubules. The bar indicates 0·1 mm.

Fig. 4.

An XY ovotestis with the adjacent mesonephros of the B6.YDOM progeny at 14d.g. Sex cords have differentiated into testis cords (ts) in the mid area. The tunica albuginea (ta) has developed well beneath the surface epithelium. In the cranial and caudal poles, sex cords remain attached to the surface epithelium, characteristics of ovarian differentiation (ov). Note the distribution of stromal cells (st) in the center of the gonad and the mesonephros, md, mesonephric ducts; mt, mesonephric tubules. The bar indicates 0·1 mm.

Fig. 5.

Testicular region of the ovotestis shown in Fig. 4. The testis cord is composed of fetal Sertoli cells (sc) and germ cells. Some germ cells are arrested at the prespermatogonial stage (ps). Note thick layers of stromal cells (st) around the basement membrane (bm) of the testis cord. Magnification, same as Fig. 6.

Fig. 5.

Testicular region of the ovotestis shown in Fig. 4. The testis cord is composed of fetal Sertoli cells (sc) and germ cells. Some germ cells are arrested at the prespermatogonial stage (ps). Note thick layers of stromal cells (st) around the basement membrane (bm) of the testis cord. Magnification, same as Fig. 6.

Fig. 6.

Ovarian region in the caudal pole of the ovotestis shown in Fig. 4. Germ cells, in either mitotic cell cycle (mt) or meiotic prophase (mi), are packed with epithelial cells (ep) in sex cords surrounded by basement membranes (bm). Stromal cells (st) are scarce in these regions. The bar indicates 0’1 mm.

Fig. 6.

Ovarian region in the caudal pole of the ovotestis shown in Fig. 4. Germ cells, in either mitotic cell cycle (mt) or meiotic prophase (mi), are packed with epithelial cells (ep) in sex cords surrounded by basement membranes (bm). Stromal cells (st) are scarce in these regions. The bar indicates 0’1 mm.

The XY ovary was morphologically indistinguishable from the XX ovary between 12 and 16 d.g. At 16 d.g., both ovaries contained abundant germ cells, most of which had reached the zygotene or pachytene stage of meiotic prophase and distributed all over the ovary (Fig. 7, 8).

Fig. 7.

Part of an XX and an XY ovary, respectively, at 16 d.g. Most germ cells in the sex cord (sec arrowheads) have entered meiotic prophase. The bar indicates 0·1 mm.

Fig. 7.

Part of an XX and an XY ovary, respectively, at 16 d.g. Most germ cells in the sex cord (sec arrowheads) have entered meiotic prophase. The bar indicates 0·1 mm.

Fig. 8.

Part of an XX and an XY ovary, respectively, at 16 d.g. Most germ cells in the sex cord (sec arrowheads) have entered meiotic prophase. The bar indicates 0·1 mm.

Fig. 8.

Part of an XX and an XY ovary, respectively, at 16 d.g. Most germ cells in the sex cord (sec arrowheads) have entered meiotic prophase. The bar indicates 0·1 mm.

Between 17 and 19 d.g., many oocytes progressed to the diplotene stage in the medullary cords of the control XX ovary (Fig. 9). In contrast, all oocytes degenerated in the medullary cords of the XY ovary while many oocytes remained in the cortex region (Fig. 10, 11). Under the electron microscope, synaptonemal complexes were often seen in degenerating oocytes of the XY ovary (Fig. 11). Epithelial cells in the medullary cords were ultrastructurally indistinguishable from those in the cortex region.

Fig. 9.

An XX ovary at 18 d.g. Germ cells in meiotic prophase are abundant in both the medullary (md) and cortex (cr) regions. The bar indicates 0-1 mm.

Fig. 9.

An XX ovary at 18 d.g. Germ cells in meiotic prophase are abundant in both the medullary (md) and cortex (cr) regions. The bar indicates 0-1 mm.

Fig. 10.

An XY ovary at 18d.g. The center of medullary region (md) is devoid of oocytes. Oocytes near the cortex (cr) region are undergoing degeneration (see arrowheads). Many oocytes are seen in the cortex region. Magnification, same as Fig. 9.

Fig. 10.

An XY ovary at 18d.g. The center of medullary region (md) is devoid of oocytes. Oocytes near the cortex (cr) region are undergoing degeneration (see arrowheads). Many oocytes are seen in the cortex region. Magnification, same as Fig. 9.

Fig. 11.

Electron micrograph of the medullar region of the XY ovary shown in Fig. 10. Two oocytes (indicated with *) are undergoing degeneration. Note synaptonemal complexes in one of these oocytes (indicated by arrowheads). Epithelial cells (cp) around oocytes show characteristics of undifferentiated ovarian cells. A thin basal lamina (bl) separates the epithelial sex cord from stromal cells (st). bv, Blood vessels. Magnification, × 45 000.

Fig. 11.

Electron micrograph of the medullar region of the XY ovary shown in Fig. 10. Two oocytes (indicated with *) are undergoing degeneration. Note synaptonemal complexes in one of these oocytes (indicated by arrowheads). Epithelial cells (cp) around oocytes show characteristics of undifferentiated ovarian cells. A thin basal lamina (bl) separates the epithelial sex cord from stromal cells (st). bv, Blood vessels. Magnification, × 45 000.

During the second week after birth, follicles had initiated growth in the medullary region of the control XX ovary and they were often undergoing atresia (Fig. 12, 13). In contrast, the XY ovary contained primordial and growing follicles only in the cortex region (Fig. 14). The medullary region was occupied by remnants of sterile sex cords surrounded by stromal tissue (Fig. 14, 15). Around one month after birth, the medullary cords were no longer distinguishable in the

Fig. 12.

An XX ovary at 12 days pp. The section was cut through the center region including the rete ovary (ro). Follicles have initiated growth in the medullary region (md). Many are undergoing atresia (see arrowheads). The cortex region (cr) is occupied by primordial follicles. The bar indicates 0 · 1 mm.

Fig. 12.

An XX ovary at 12 days pp. The section was cut through the center region including the rete ovary (ro). Follicles have initiated growth in the medullary region (md). Many are undergoing atresia (see arrowheads). The cortex region (cr) is occupied by primordial follicles. The bar indicates 0 · 1 mm.

Fig. 13.

Part of the XX ovary shown in Fig. 12. Follicles in the medullary region, formed with cuboidai follicular cells (fc) and large oocytes (oo), are undergoing atresia. The rete ovary (ro) is formed by compacted epithelial cells (indicated by arrowheads), which occasionally continue to a lumen (indicated with *). The bar indicates 0 · 1mm.

Fig. 13.

Part of the XX ovary shown in Fig. 12. Follicles in the medullary region, formed with cuboidai follicular cells (fc) and large oocytes (oo), are undergoing atresia. The rete ovary (ro) is formed by compacted epithelial cells (indicated by arrowheads), which occasionally continue to a lumen (indicated with *). The bar indicates 0 · 1mm.

Fig. 14.

An XY ovary at 11 days pp. The medullary region (md) is completely devoid of follicles, and occupied by remnants of sex cords. Primordial and growing follicles are seen in the cortex region (cr). Magnification, same as Fig. 12.

Fig. 14.

An XY ovary at 11 days pp. The medullary region (md) is completely devoid of follicles, and occupied by remnants of sex cords. Primordial and growing follicles are seen in the cortex region (cr). Magnification, same as Fig. 12.

Fig. 15.

The medullary region of the XY ovary shown in Fig. 14. Sex cords are composed of epithelial cells (ep) enclosed in basement membranes (bm). No germ cells are seen inside the sex cord. A follicle near the cortex region is undergoing atresia (indicated with *). Magnification, same as Fig. 13.

Fig. 15.

The medullary region of the XY ovary shown in Fig. 14. Sex cords are composed of epithelial cells (ep) enclosed in basement membranes (bm). No germ cells are seen inside the sex cord. A follicle near the cortex region is undergoing atresia (indicated with *). Magnification, same as Fig. 13.

XY ovary since the medullary region became occupied by large follicles of the cortex origin (data not shown). The number of follicles in the XY ovary was smaller than that in the XX ovary. Around 2 months pp., only a few follicles were seen in the XY ovary whereas many follicles at various stages were present in the control XX ovary.

3β -HSDH

In the control XX ovary, the 3β -HSDH activity appeared first in stromal cells around growing follicles in the medullary region around 10 days pp. (Fig. 16). In contrast, the XY ovary acquired the 3β -HSDH activity in epithelial cells of the oocyte-free medullary cords, but not in the interstitial cells in the medullary region (Fig. 17). Weak activity was also seen in stromal cells around growing follicles in the cortex region.

Fig. 16.

An XX ovary at 10 days pp. stained for 3β -HSDH. The enzymatic activity is localized in stromal cells around growing follicles in the medullary region (md) in contrast to the cortex region (cr). See Fig. 12 for details of structures. The bar indicates 0·1 mm.

Fig. 16.

An XX ovary at 10 days pp. stained for 3β -HSDH. The enzymatic activity is localized in stromal cells around growing follicles in the medullary region (md) in contrast to the cortex region (cr). See Fig. 12 for details of structures. The bar indicates 0·1 mm.

Fig. 17.

An XY ovary at 10 days pp. stained for 3β -HSDH. The enzymatic activity is localized in epithelial cells forming the sex cords (see arrowheads) in the medullary region (md). Weak staining is also seen around growing follicles in the cortex region (cr). See Fig. 14 for details of structures. Magnification, same as Fig. 16.

Fig. 17.

An XY ovary at 10 days pp. stained for 3β -HSDH. The enzymatic activity is localized in epithelial cells forming the sex cords (see arrowheads) in the medullary region (md). Weak staining is also seen around growing follicles in the cortex region (cr). See Fig. 14 for details of structures. Magnification, same as Fig. 16.

Development of fetal gonads and testosterone production in vitro

When gonads were dissected from fetal offspring of 16 d.g. and cultured for 3 days, they showed morphological changes comparable to those of the newborn offspring, i.e. loss of oocytes from the medullary cords of the XY ovary (data not shown). The amount of testosterone secreted from gonadal expiants of 14 d.g. was positively correlated to the ratio of testicular structures (Fig. 18). Testosterone production from the control B6 testis was proportional to that from the B6.YDOM ovotestis. None of the XY ovaries (testicular area/gonad = 0) produced testosterone throughout the culture period (ten ovaries were examined). Similar results were obtained with expiants of 16 d.g.

Fig. 18.

The relation between the amount of testosterone secreted by XY ovotestes, XY ovaries, and B6 XY testes of 14d.g. and the ratio of testicular structures in whole gonad. *, ten XY ovaries without testicular components (testicular area/gonad = 0·0) jverc examined.

Fig. 18.

The relation between the amount of testosterone secreted by XY ovotestes, XY ovaries, and B6 XY testes of 14d.g. and the ratio of testicular structures in whole gonad. *, ten XY ovaries without testicular components (testicular area/gonad = 0·0) jverc examined.

In the present study, the chromosomal sex was determined by dot hybridization with Y-specific DNA probes. We assume that the hybridization with DNA probe AC1L or 145SC5 indicates the presence of the Y chromosome. We have previously found that the DNA probe AC11- and 145SC5-related sequences are repeated several hundred times over a wide range in the mouse Y chromosome (Nishioka & Lamothe, 1986; Nishioka, 1988; Nishioka, unpublished observations). If translocation had occurred, it would have involved a large segment of the Y chromosome. In a previous report by Eicher and others of cytogenetic studies, the presence of the Y chromosome was confirmed while XO cells or chromosomal abnormalities were not found in the B6.YDOM female mouse (Eicher et al. 1982).

Thus determining the chromosomal sex, we have shown that when the B6.YDOM male mouse was crossed with B6 females, half of the XY progeny developed bilateral ovaries and female internal and external genitalia whereas the other half developed testicular components (bilateral testes or hermaphrodites). This result is consistent with a previous report (Eicher & Washburn, 1983). The mature XY female copulated normally, but none of them became pregnant. Although Eicher et al. (1982) reported one exceptional fertile XY female, we could not confirm their results.

When the XY female had been ovariectomized and grafted with XX ovaries, regular estrous cyclicity was initiated. In contrast, all XX females that had been ovariectomized and grafted with XY ovaries stopped estrous cyclicity. These results suggest that the pituitary of the XY female can respond to normal ovarian signals to induce estrous cyclicity. Therefore, the XY female is infertile due to a defect inside the XY ovary. This result does not exclude the possibility that an abnormality outside the gonad results in the defect of the XY ovary during development. Oocytes of the XY female could be made to ovulate after injection with PMSG followed by hCG (Eicher & Washburn, 1986; present results). It is, therefore, likely that one of the possible causes of the infertility is abnormal endocrine function of the XY ovary.

We examined the morphological development of the XY ovary. In previous studies, we investigated only XY ovaries from hermaphrodites, which possessed contralateral ovotestes or testes (Nagamine et al. 1987a). In the present study, we ruled out the possible influence of testicular tissue by examining the XY female with bilateral ovaries. The sex ratio of offspring (XX females: XY females: XY males or hermaphrodites) remained constant throughout the development after 16 d.g. At 14 d.g., although a larger number of the progeny contained testicular tissue, 21% of the XY progeny possessed no trace of testicular components. Since testosterone production was positively correlated with the area of testicular structures, it is unlikely that some cells had differentiated into testicular cell types without characteristic organization. Accordingly, it is reasonable to assume that some XY females had never possessed testicular components during development. Hence, the infertility of the XY ovary could be attributed to an abnormality common to all XY ovaries.

The XY ovary was morphologically indistinguishable from the XX ovary until 16 d.g., when all germ cells had entered meiotic prophase and were distributed in sex cords over the whole gonad. Between 16 and 19 d.g., most oocytes in medullary cords of the XY ovary degenerated. Since synaptonemal complexes were seen in many degenerating oocytes, the degeneration appeared to occur in mid to late pachytene stages. In the normal XX ovary, the first set of oocytes reached the diplotene stage and induced follicle formation in the medullary region. These follicles underwent atresia, and left steriodogenic cells in the interstitium (Merchant-Larios, 1984; present observations). The role of the steroidogenic cells derived from the first generation of atretic follicles is not fully understood. Since the XY ovary lost oocytes in the medullary region prenatally, it never developed follicles in this region. Instead, epithelial cells forming sterile medullary cords acquired the 3/3-HSDH activity while stromal cells remained negative. We have previously reported that in W/Wv mutant mice or busulphan-treated rats, the oocytes-free sex cords remain negative for 3β -HSDH (Merchant-Larios & Centeno, 1981; Merchant-Larios, 1976). It is worth noting that the XX and XY epithelial cells of the medullary cords appear to respond differently to the loss of oocytes. On the other hand, we observed that in the testicular area of the B6.YDOM ovotestis, only the interstitial cells became positive for the 3β -HSDH activity (data not shown). Hence, the XY epithelial cells of the oocyte-free medullary cords are also different from the XY epithelial cells forming the seminiferous tubules.

Degeneration of oocytes is common during normal ovarian development (Baker, 1963; Speed, 1988). However, the XY female lost far more oocytes than the control XX female. It has been reported that the XO female mouse also loses many oocytes after late pachytene stage (Burgoyne & Baker, 1985). Most degenerating oocytes appear to be located in the cortex region of the XO ovary shown in this paper while occasional oocytes are seen in the medullary region. Since the XO female mouse is usually fertile (Lyon & Hawker, 1973), the infertility of the B6.YDom’ female cannot be explained by a single dose of the X chromosome. It is also difficult to attribute the oocyte loss to the genetic element of XY oocytes because the fate of the XY oocyte is very different in two regions. It is more likely that the presence of the Y chromosome in the epithelial cells disturbs the normal ovarian development of the medullary region. Although epithelial cells in the medullary and cortex regions are similar at the ultrastructural level, the interaction of epithelial cells with oocytes is clearly different between these two regions.

From the present study, we postulate that the precocious death of oocytes in the medullary cords of XY ovaries prevents follicular formation from these cords, leading to abnormal differentiation of steroidogenic cells. This altered developmental pathway may be responsible for abnormal endocrine functions of the XY ovary. Therefore, in the XX ovary, the growth and atresia of follicles from the medullary cords may be critical for the development of normal ovarian functions.

We are grateful to Dr J. F. Nelson for valuable discussions, and Jose Guadalupe Baltazar and Jamilah P. Saeed for excellent technical assistance. This study was supported in part by grants from the Medical Research Council of Canada (MA-9740 to T.T. and MT-6809 to Y.N.).

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