Ovarian steroidogenesis in vitro during the first month posthatching in the domestic chick: Gonadotropin responsiveness and [3H]progesterone metabolism

GENERAL

AND COMPARATIVE

ENDOCRINOLOGY

62, 62-69 (1986)

Ovarian Steroidogenesis in Vitro during the First Month Posthatching in the Domestic Chick: Gonadotropin Responsiveness and [3H]Progesterone Metabolism BABETTA L. MARRONE~ Department of Obstetrics and Gynecology, St. Loais University School of Medicine, St. Louis, Missouri 63104 Accepted October 15, 1985 Ovarian steroidogenesis was examined in domestic chicks during the first month posthatching. In vitro production of androstenedione during a 4-hr incubation was enhanced in a dose-dependent manner by O.l- 100 rig/ml of chicken LH (cLH). The greatest response was observed in ovaries from Day 1 chicks (eightfold), but cLH was also effective in Day 7, 14, 21, and 28 ovaries (three- to sixfold increase). Estrogen production in response to cLH was increased significantly only in Day 1 and 7 ovaries. A similar trend in androstenedione and estrogen production was seen in response to S-bromo CAMP, oLH, and oFSH. In an additional experiment, in vitro metabolism of [jH]progesterone by ovaries from Day 1, 7, 14, and 21 chicks was examined during a 4-hr incubation. The major metabolite, comprising 18-19% of the radioactivity, coeluted with 5B-pregnan-3,20-dione. This is the first report of gonadotropin-stimulated steroidogenesis and progesterone metabolism in the chick during the first month post-hatching. The results show that (1) the responsiveness to gonadotropins decreases during this period and (2) the profile of 13Hlprogesterone metabolism does not resem.ble that seen in adult granulosa and theta cells. o 1986 Academic PXSS. 1~.

Steroidogenesis by the granulosa and theta cells during follicular development in the ovulating hen has been well characterized (Huang et al., 1979; Marrone and Hertelendy, 1983a, b). Furthermore, there is more known about ovarian steroidogenesis in the domestic hen than in any other avian species. In light of this, it is curious that there have been so few investigations on the development of ovarian regulation in this species. In chick embryos, LH stimulates estrogen production (Woods et al., 1981) and in vitro both the right and left ovaries of chick embryos respond to LH stimulation with increased estrogen and testosterone production, although the right ovary regresses just before hatching (Teng and Teng, 1977; Teng et al., 1982). The regulation of ovarian isteroidogenesis in chicks

from the time of hatching until maturity has received little attention. In the present study, aspects of ovarian steroidogenesis by chicks during the first month post-hatching are investigated. Gonadotropin regulation of steroidogenesis using a homologous gonadotropin (chicken LH), which has been shown previously to be effective in stimulating progesterone production in granulosa cells of adult quail (Asem et aZ., 1985) and hen (unpublished observations), is investigated. In addition, PHIprogesterone metabolism by chick ovaries is examined. The information gained from these investigations should be valuable to our understanding of the comparative endocrinology of reproductive development and mechanisms of reproductive development in general.

I Presents address: Los Alamos National Laboratory, Life Sciences Division, MS-M881, Los Alamos, N. Mex. 87545.

Animals. White Leghorn pullets were obtained on the day of hatching from Key-Roy Hatcheries (Berger, MO.). They were housed in brooders under a 14:10,

METHODS

62 00166480186 $1.50 Copyright All rights

0 1986 by Academic Press, Inc. of reproduction in any form reserved.

OVARIAN

STEROIDOGENESIS

1ight’:dark cycle, at 27”, with Purina Chick Starter and water provided ad libitrtm. Hormones and chemicals. Chicken LH (cLH) was generously provided by Dr. Peter J. Sharp. Ovine LH (oLN; NIAMDD-LH-22; 2.3 NIH-LH-Sl/mg) and ovine FSH (oFSH; NIAMDD-oFSH-14; 9 NIH-FSHSlimg) were gifts from the National Pituitary Agency of the National Institutes of Health. The steroids, 8-bromo cyclic AMP, and 3-isobutylI-methyl-xanthine (IBMX) were obtained from Sigma Chemical Company (St. Louis, MO.). Medium 199 with Hank’s salts was purchased from Grand Island Biological (New York). [1.2,6,7-3HIAndrostenedione (sp act, 80-115 Ciimmol), [2,4,6,7-3H]estradiol-17p (sp act, 90-115 Ciimmol), and [1,2,6,7-iH]progesterone (sp act, 90- 115 Ciimmol) were purchased from New England Nuclear Corporation (Boston, Mass.). High-performance liquid chromatography (HPLC)grade solvents for the HPLC analysis and solvents for thin-layer chromatography (TLC) were purchased from Fisher Scientific (Pittsburgh, Pa.) and Baker Chemical Company (Phillipsburg, N.J.). Tissue collection and incubation. On Day 1, 7, 14, 21, or 28 post-hatching, chicks were killed by cervical dislacation. The ovaries were removed, cleaned of connective tissues, and placed into cold Medium 199. Each’ ovary was weighed, cut into 4- 10 pieces, and placed into individual polystyrene culture tubes containing 0.1 mM (IBMX) in Medium 199. There were five ovaries per treatment group. Following a 30-min preincubation, the treatment was added. The final volume in each tube was 2 ml. Ovaries were incubated open to the air, at 38” in a shaking water bath for 4 hr, unless specified otherwise. The incubations were terminated by immersion of the tubes into an ice-water bath. The medium was transferred to extraction tubes and frozen at -4” until extraction for radioimmunoassay or HPLC analysis. In some cases, after removal of the medium the ovarian pieces were washed twice with saline and stored frozen until protein assay (Lowry et al., 1951). Radioimmunoassays. The medium was extracted twice with 2 vol of diethyl ether. The ether extract was evaporated to dryness under a stream of nitrogen in a 45” water bath. The sample was redissolved in 1 ml ethanol and lOO+p,laliquots were taken for radioimmunoassays of progesterone, androstenedione, and estradioliestrone. The efficiency of the extraction procedure for each steroid was >95%. All samples from one experiment were analyzed in the same radioimmunoassay. ‘The androstenedione antiserum was obtained from Endocrine Sciences (Tarzana, Calif.). The cross-reactivity and inter- and intraassay CVs, and sensitivity have been reported previously (Marrone and Hertelendy, 1985). Parallelism was determined by measuring the androstenedione concentration in different

IN THE

DOMESTIC

CHICK

63

volumes of the extracted sample. In 50, 100, 150. and 200 p+l the concentrations of androstenedione were 1.44 t 0.037, 1.41 * 0.084, 1.35 I 0.108, and 1.3i I 0.076 pgipl, respectively. The estrogen antiserum was the gift of Dr. B. V. Caldwell. It is specific for estradiol-17P and es&one. The cross-reactivity, inter- and intraassay CVs, and sensitivity, have been reported previously (Marrone and Hertelendy, 1985). Parallelism was detemined by measuring the estrogen concentration in different volumes of the extracted sample. In 50. 100. 150, and 200 ~1, the concentrations of estrogen were 0.89 2 0.16, 0.84 IC,0.05. 0.76 i .06, and 0.67 2 .04 pgipl, respectively. The cross-reactivity of the progesterone antiserum (Marrone and Hertelendy, 1983a), the inter- and intraassay CVs. and sensitivity (Hammond et nl., 1980) have been reported. Most samples in the present study were at the lower limits of detectability of the progesterone radioimmunoassay. HPLC. For determination of [3H]progesterone metabolism, ovaries from chicks on Day 1, 7, 14, and 21 were prepared as above and incubated with 1 kg/ml [3H]progesterone in 2 ml Medium 199. The incubations were carried out open to the air at 38” in a shaking water bath for 5 hr. The media and .homogenized ovarian pieces were extracted twice with 2 vol of diethyl ether and the extract was evaporated to dryness under a nitrogen stream in a 45” water bath. The details of the HPLC analysis have been reported previously (Marrone and Hertelendy. 1983b). The sample was dissolved in 25 ~1 of HPLC-grade methanol and 20 ~1 was injected onto a 5 km reversedphase Ci8 Altex Ultrasphere HPLC column (1.5 cm )( 4.6 mm). The mobile phase was tetrahydrofuran:methanol:water (16:28:56) at a flow rate of 1 mlimin. about 3200 psi. A Beckman pump (Model IlOA) and microprocessor (Model 420) were used. Fractions (0.5 ml) were collected for scintillation counting and the profile was visualized by uv spectroscopy using a Hitachi spectrometer (Model 110-10) at 240 or 210 nm. A Shimadzu C-RIA integraterirecorder recorded the spectrometry data. Fractions were collected for 90 min after injection and the total radioactivity in 190 fractions was calculated. The production of each steroid metabohte was based on elution of radioactivity with a known steroid standard. In some cases TLC was done on fractions after HPLC to further characterize a metabolite. The radioactivity in each peak from the HPLC analysis was expressed as a percentage of the total radioactivity. Only fractions with radioactivity twice that of background were used in the calcuiations. Data analysis. Cotiparisons between two groups were analyzed by St,udent’s t test. Comparisons among more than two groups were analyzed by oneway analysis of variance followed by Neuman-Keuls

64

BABETTA

L.

test. Probability values of <0.05 were considered to be statistically significant.

RESULTS

cLH dose vespome. Chick ovaries, Day I, 7, 14, and 21, were incubated with 0.01, 0.1, 1, 10, or 100 ngiml of cLH for 4 hr. Androstenedione production was stimulated in a dose-dependent manner by cLH in chick ovaries from each age group examined (Fig. 1). The effect of cLH on androstenedione production was maximal at a dose of IO-100 &ml. The EDso was 1-2 rig/ml and was the same for chick ovaries from each age group. Estrogen production was stimulated significantly by cLH in ovaries from Day 1 and Day 7 chicks only (Fig. 1). Ovarian progesterone production was not consistently increased by cLH at any age (data not shown). In a separate experiment, the effects of a maximally stimulating dose of cLH (100 ngiml) on ovaries from chicks at Day 1, 7,

%+

.Ol

.l

MARRONE

14, 21, and 28 were compared (Fig. 2). Androstenedione production was significantly elevated at all age groups, but the maximal response was greater in Day 1 ovaries (8 fold increase) as compared to the others (3to 6-fold increases). As in the previous experiment, estrogen production was significantly elevated only in chick ovaries from Day 1 (Zfold increase) and Day 7 (1.4-fold increase). The chick ovary increases greatly in size as the chick matures. Based on a sample of 70 to 80 chicks per each age group, the average ovarian weight (mg/ovary i. SEM) was 8.93 ? 0.55, 15.6 t 0.91, 28.6 t 1.69, and 41.7 ? 1.81 in Day 1, 7, 14, 21 ovaries, respectively. Therefore, it should be pointed out that the same pattern of results was seen when the steroid production data were expressed in picograms per ovary, rather than in picograms per milligram ovary, as in Figs. I-5. Maximal steroid responses by Day 1 ovaries were greater than

1

10

100

cLH (rig/ml) FIG. 1. Effect of cLH on production of androstenedione (top panel) and estrogen (bottom panel) by ovaries from chicks on Day 1, 7, 14, and 21 post-hatching during a 4-hr incubation. Each point represents the mean F SEM of five ovaries. (0) Day 1, (A) Day 7, (II) Day 14, (0) Day 21.

OVARIAN STEROIDOGENESIS

IN THE DOMESTIC CHICK

65

sponse to 100 rig/ml cLH. In drostenedione production by ovaries was increased from pg/pg protein to 1.03 t 0.10 in response to 100 ngiml cLH.

1

7

14

21

28

1

AGE

(days)

7

14

21

28

FIG. 2. Androstenedione (left panel) and estrogen (right panel) production in the absence (0) or presence (IB) of a maximally stimulating dose of cLB (100 ngiml) during a 4-hr incubation. Each bar represents the mean + SEM of five ovaries. *P < 0.05 vs control.

those by the more mature ovaries. For example, in data from Fig. 1, 100 ngiml cLH increased pg/ovary androstenedione production by 5.3-fold, whereas a 2.7-fold increase was seen in Day 21 ovaries. Similarly, in Fig. 2, 7-fold and 4.3-fold increases in picograms per ovary androstenedione production were observed in response to 100 rig/ml cLH in Day 1 and Day 21 ovaries, respectively. Ovarian protein content was also measured in the experiments depicted in Figs. 1 and 2 and was found to increase as the age of the chick and the ovarian weight increased. Based on 70 to 80 chicks per each age group, the average protein content of each ovary (mg protein/ovary -+ SEM) was: 0.412 1. 0.065, 0.711 & 0.043, 1.59 2 0.17, and 2.37 t 0.14 for Day 1, 7, 14, and 21 chicks, respectively. Consequently, when steroid production in response to cLH was expressed as pg/mg protein the same pattern of results, was seen as when the data were expressed in terms of picograms per milligrams ovary weight, as in Figs. 1 and 2. For example, in Fig. 2, androstenedione production by Day 1 chick ovaries increased from 1.20 i 0.17 pg/pg protein to 9.15 t 1.82 pg/kg protein in re-

contrast, anDay 21 chick 0.31 & 0.05 pgt*g protein

Time Course of cEH Effects Ovaries from 2-day-old chicks were incubated with 10 rig/ml cLH for 0, 0.5, 1, 2, 4, 6, or 8 hr (Fig. 3). The production ofandrostenedione was doubled by 0.5 hr and was increased lo-fold by 4 hr. Estrogen production was doubled by 2-4 hr and increased fourfold by 6 hr. Progesterone production was unchanged at 4 hr, but increased slightly by 6 and 8 hr. 8-Bromo

CAMP Stimulation

In general, the responsiveness of chick ovaries to stimulation with 8-bromo CAMP paralleled that of LH (Fig. 4). In response to 8-bromo CAMP, androstenedione production was stimulated significantly in ovaries from each age group. Androstene250 200

0

0.8 1 2

4

--8

8

TIME (hrs)

FIG. 3. Time course of cLH (10 ng/mI) on androstenedione (top panel), estrogen (middle panel), and progesterone (bottom panel) production. Each point represents the mean ? SEM of five ovaries from 2-day-old chicks.

66

BABETTA

0

.I

.5

8-bromo

CAMP

1

L.

2

(mM)

FIG. 4. Effect of 8-bromo-CAMP on production of androstenedione (top panel) and estrogen (bottom panel) by ovaries from chicks on Day 1 (O), 7 (A), 14 (U), and 21 (0) post-hatching during a 4-hr incubation. Each point represents the mean -C SEM of five ovaries.

dione production was stimulated about 8fold by 2 mM 8-bromo CAMP in Day 1 and 7 ovaries, 5.6-fold in Day 14 ovaries, and 3.8-fold in Day 2 I ovaries.

MARRONE

[3HjProgesterone Metabolism Ovaries from Day 1, 7, 14, and 21 chicks, incubated with [3H]progesterone, produced small amounts of radioactivity that coeluted with androstenedione and 17-hydroxyprogesterone. However, the major peak of radioactivity at each age examined coeluted with S/3- and Sa-dihydroprogesterone (DHP) with a retention time of about 80 min (Table 1). Further chromatography was used to distinguish whether the radioactivity coeluted with 5l3-DHP or ICYDHP in these samples. Samples of Day 14 chick ovaries were subjected to HPLC. A large fraction was collected from 65 to 85 min following injection. This fraction was then subjected to three TLC systems, in which 5P-DHP was separated from 5~DHP: System 1, chloroform:cyclohexane:N-butylacetate (1: 1: 1); System 2, benzene:ethyl acetate (8:2); System 3, heptane:ethyl acetate (3:l). In each case, the radioactivity in the sample fraction comigrated with authentic 5@DHP. In addition, aliquots of the 65 to 85 min fraction from

Effects of oLH and oFSH on Androstenedione Production In a dose-response study using 100,500, or 1000 rig/ml oFSH or 0.01, 0. I, 10, or 100 rig/ml oLH, both hormones stimulated androstenedione production in ovaries from Day 1 and Day 21 chicks. The magnitudes of the responses to all three doses of oFSH were similar. The stimulation by oLH was dose dependent and similar in degree to that observed in response to cLH. The androstenedione responses to 100 rig/ml oLH 1 21 and 1000 rig/ml of oFSH in Day 1 and Day AGE (days) 21 chicks are shown in Fig. 5. As with cLH FIG. 5. Effect of a maximally stimulating dose of and S-bromo CAMP, greater increases were oLH (m) (100 ngiml) and oFSH (I%) (1000 ngiml) on achieved in Day 1 chick ovaries (- 6-fold androstenedione production by ovaries from chicks on for both oLH and oFSH) than in Day 21 Day 1 and 21 post-hatching. Each bar represents the ovaries (oLH, 4.7-fold; oFSH, 4-fold). mean t SEM of five ovaries.

OVARIAN

STEROIDOGENESIS

ilPLC were subjected to oxidation with 2% 3rO,, extracted, and reapplied to HPLC :Marrone et al., 1985). No change was de:ected in the retention time of the fractions :reated by this method. The greater amount of radioactivity associated with the 80 min peak in Day 14 and 21 chick ovaries than in Day 1 and 7 avaries appears to be due to the greater amount of tissue in the incubations of the more mature ovaries rather than to a true increase in enzyme activity. When ovaries from Day 1 chicks were pooled to equal the weight of a single Day 14 ovary (-30 mg), the percentage of total CPM associated with .this peak was similar to that of a single Day 14 chick ovary (Table 1). DISCUSSION The maximal androstenedione response of chick ovaries to trophic stimulation decreased with increasing age during the first month post-hatching. Likewise, estrogen production in response to trophic stimulation was greater in the younger (Days 1 and 7) ovaries. A similar pattern of steroidogenesis was seen in response to S-bromo CAMP stimulation, suggesting that the gonadotropins increase steroidogenesis in chick ovaries by a mechanism involving CAMP. The effects of CAMP on steroidogenesis in the adult hen theta (Marrone and

IN

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CHICK

Hertelendy, 1985) and granulosa cells (Asem and Hertelendy, 1985) and in embryonic chick ovaries (Teng and Teng, 1977) also parallel the effects of gonadotropins. The reason for increased responsiveness of chick ovaries to gonadotropin stimulation of steroidogenesis during the first week post-hatching is not clear at present. However, the increased responsiveness to younger chick ovaries is apparently not related to the use of homologous gonadotropin, since the same pattern was seen in response to oLH and oFSH. An interesting finding in this study was the pattern of metabolism of C3H]progesterone by post-hatch chick ovaries. S@DHP was identified previously as a progesterone metabolite in embryonic chick ovaries (Gonzalez et al., 1983). In the present study, we have tentatively identified SP-dihydroprogesterone as the major metabolite of progesterone in post-hatch chick ovaries. This tentative identification is based on coelution of radioactivity with S&dihydroprogesterone on HPLC and i’n three separate TLC systems. In addition this metabolite was not susceptible to oxidation by CrO,. However, since &&her mass spectroscopy nor recrystallization to constant specific activity was performed on this compound we can claim only to have partially characterized this metabolite.

TABLE 1 METABOLISM OF[~HIPROCESTERONE BYCXICK OVARIES Percentage total recovered cpm (Z I SEM) Coeluting standard steroid Androstenedione 17-Hydroxyprogesterone Progesterone (substrate) “SO min peak”*

Day 1

Day 1 pooled**

Day 7

Day 14

Day 21

2.65 k 0.12 1.88 it 0.37

2.65 k 0.40 1.60 ? 0.18

3.03 i 0.38 2.31 i 0.41

3.10 i- 0.23 4.62 I 0.53

3.26 r?r 0.31 4.12 I 0.35

67.3 I 4.36 5.08 i- 0.64

45.5 k 4.15 17.9 + 1.89

64.5 i_ 3.41 7.76 k 0.55

48.6 2 4.84 19.6 2 3.72

40.9 t 6.20 17.7 i 2.15

Note. Chick ovaries, 5 per group except as noted*“, were incubated for 5 hr with 1 pgiml [3H]progesterone. Steroids in the sample extracts were subjected to HPLC analysis as described under Methods. * Coelutes with SB-dihydroprogesterone. ** Five incubations, each containing ovaries pooled from 4-5 chicks. Total tissue weight -30 mg, matched to the average weight of a single Day 14 ovary.

68

BABETTA

Nevertheless, the difference between the metabolism of [sH]progesterone by the chick ovary and the adult hen ovary is particularly noteworthy. In adult hen theta cells, [3H]progesterone is converted mainly to 20@dihydroprogesterone (20@DHP) (Llewelyn, 1981; Marrone et al., 1985); granulosa cells do not metabolize appreciable amounts of progesterone (Marrone and Hertelendy, 1983b). SP-DHP is not a metabolite of progesterone in either theta or granulosa cells of the yolk-filled follicles (Marrone and Hertelendy, 1983b, Marrone et aZ., 1985). Conversely, in the present study, 20&DHP was not a metabolite of [3H]progesterone in chick ovaries. Therefore, at some time between the first month post-hatch and the onset of reproductive activity there is a shift in the nature of ovarian metabolism of [3H]progesterone toward production of %OP-DHP. Although the granulosa and theta cells of the yolkfilled follicles do not produce SP-DHP, in preliminary experiments using ovarian stroma from adult hens there appeared to be some metabolism of [3H]progesterone to a metabolite that coelutes on HPLC with SP-DHP. Therefore, in the adult hen, the immature fohicles that are imbedded in the stroma may possess SP-reductase activity. In the immature rat ovary, progesterone is converted mainly to .5a-reduced metabolites. In particular, 5a-androstane-3or,l7@ diol (3a-diol) is a major metabolite of progesterone (Eckstein et al., 1976; Karakawa et al., 1976). 3ol-Diol formation decreases at around the time of puberty and is not a significant ovarian product in adult rats (Karakawa et al., 1976; Eckstein and Ravid, 1979). However, in parallel to what may be the case in the chicken, in the rat there is a shift from biosynthesis of Scx-reduced products during the prepubertal/pubertal period to biosynthesis of 20a-reduced products of progesterone in the adult ovary (Eckstein and Ravid, 1979). Although the physiological significance of this shift in steroidogenesis is not resolved,

L. MARRONE

it has been suggested that 3or-diol may play a role in maintaining tonic inhibition of gonadotropins in immature rats. 3o-diol administration delays sexual maturation (Kraulis et al., 1981; Kramer and MeijsRoelofs, 1982) and inhibits the ovulatory release of gonadotropins during the pubertal period (Eckstein et al., 1976; Kraulis et al., 1981). However, whether 3ar-diol plays a key role in the onset of puberty is arguable, because only high doses of 301diol are able to delay puberty, and in addition, inhibition of 5c-w-reductase activity does not advance puberty onset (Ojeda et IKE., 1984). Nevertheless, the fact that both the chicken and the rat exhibit shifts in steroid biosynthesis during maturation suggests that the role of these shifts may be similar in each species. Whether this role is important to vertebrate reproductive development is an intriguing question to pursue. ACKNOWLEDGMENTS Supported by NIH Grant HD-16623. The generous gifts of chicken LH, by Dr. Peter Sharp of the Agricultural Research Council, Midlothian, U.K., and ovine LH and FSH, by Dr. S. Raiti of the NIAMDD, are gratefully acknowledged. Ms. Wafaa Eleissawy provided technical assistance and Ms. Charlene Sandier and Ms. Martha Maes provided secretariaf assistance.

REFERENCES Asem, E. K., and Hertelendy, E (1985). Influence of follicular maturation on luteinizing hormone, cyclic 3’-5’-adenosine monophosphate, forskolin and cholesterol stimulated progesterone production in hen granulosa cells. Biol. Reprod. 32, 257-268. Asem, E., Marrone, B. L., and Hertelendy, F. (1985). Steroidogenesis in ovarian cells of the Japanese quail (Coturnix coturnix japonica). Gen. Comp. Endocrinol. 60, 353-360. Eckstein, B., and Ravid, R. (1979). Changes in pathways of steroid production taking place in the rat ovary around the time of first ovulation. J. Steroid Biochem. 11, 593-597. Eckstein, B., Yehud, S., Shari, J., and Goldhaber, G. (1976). Suppression of luteinizing hormone release by 5e+androstane-3a,17B-diol and its 3pepimer in immature ovariectomized rats. J. Endocrinol. 70, 25-30. Gonzalez, C. B., Cozza, E. N., DeBedners,

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M. E. O., Lantos, C. P., and Aragones, A. (1983). Progesterone and its reductive metabolism in steroidogenic tissues of the developing hen embryo. Gen. Comp. Endocrinol. 51, 384-393. Hammond, R. W., Todd, H., and Hertelendy, F. (1980). Effects of mammalian gonadotropins on progesterone release and cyclic nucleotide production by isolated avian granulosa cells. Gen. Comp. Endocrinol. 41, 4X1-476. Huang, E. S.-R., Kao. K. J., and Nalbandov, A. V. (1979). Synthesis of sex steroids by cellular components of chicken follicles. Biol. Reprod. 20, 454-461. Karakawa, T., Kurachi, K., Aoma, T., and Matsumoto, K. (1976). Formation of S&-reduced Clpsteroids from prbgesterone in vitro by pathway through &reduced C,,-steroids in ovaries of late pre-pubertal rats. Endocrinology 98, 571-579. Kramer, P., and Meijs-Roelofs, H. A. M. (1982). Retardation of first ovulation in pubertal rats after treatment with .5ol-androstane-3a,l7@-diol or its 3@-epimer. J. Endocrinol. 92, 31-35. Kraulis, I., Naish,,S. J., Gravenor, D., and Ruf, K. B. (1981). So.-Androstane-3cx,l7@-diol: Inhibitor of sexual maturation in the female rat. Biol. Reprod. 24,445-453,

Llewelyn, C. A. (1981). Conversion of progesterone to 20p-hydroxy-4-pregnene-3-one by the preovulatory foilicle in the hen (Gallus domesticus). ARCS Med. Sci. 9, 859. Lowry, 0. H., Rosebraugh, N. J., Farr, A. L., and Randall, R. J., (1951). Protein measurement with the folin phenol reagent. J. Biol. Ckem. 193, 265-275.

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Marrone, B. L., and Hertelendy, E (1983a). Steroidogenesis by avian ovarian cells: Effects of LH and substrate availability. Amer. J. Pkysioi. 244, E487-E493. Marrone, B. L., and Hertelendy, F. (1983b). Steroid metabolism by avian ovarian cells during follicular maturation. Biol. Reprod. 29, 953-962. Marrone, B. L., and Hertelendy, E (1985). Decreased androstenedione production with increased foilicular maturation in theta cells from the domestic hen (Gallus domesticus). J. Reprod. Fertil. 14, 543-550. Marrone, B. L., Wiebe, J. P., Buckingham. K. D., and Hertelendy, I? ( 1985). Analysis of steroid metabolites produced by theta cells from the adul? domestic hen. J. SreroidBiockem. 23, 395-37’8. Ojeda, S. R., Katz, K. H., Costa, M,-E., and A&is. J. P. (1984). Effect of experimental alterations in serum levels of Sa-androstone-3ol,I7P-diol on the timing of puberty in the female rat. Net!raenductinology 39, 19-24. Teng, C. T. and Teng, C.-S. (1977). Studies on sexorgan development. The hormonal regulation of steroidogenesis and adenosine 3’:5,‘-cyclic ,monophosphate in embryonic-chick ovary. Biockem. J. 162, 123-134.

Teng, C. T., Teng, C.-S., Bousfield, G. R., Liv. W.-K., and Ward, D. N. (1982). Differential responsqof @,owifig and regressing chicken ovaries to gonadotropic hormones. Gen. :Cor7~p’. :&docrinol. 48, 325-332. Woods, J. E., Mennella. J. A., and Thommes, R. C. (1981). The hypothalamic-adenahjpophyseal-gonadaI axes in the developing chick embryo. I. LH sensitivity. Gen. Co&p. Endocrindl. 45. 66-73.