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The effects of hydroxyflutamide on action and production of androgens in rats induced to superovulate Yu, Frank Hong 1990

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THE EFFECTS OF HYDROXYFLUTAMIDE C N ACTION A N D PRODUCTION OF ANDROGENS IN RATS INDUCED TO SUPEROVULATE By FRANK HONG YU B.Sc, University of British Columbia, 1988 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE STUDIES (Human Reproductive Biology Programme) W e accept this thesis as conforming to the required standard THE UNIVERSITY OF BRITISH COLUMBIA August 1990 © Frank Hong Yu, 1990 In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission. Department of (OBST&TI^ICS toib ^r^^C-OLD^y The University of British Columbia Vancouver, Canada Date -S^pTgMfc&R- >3 > 1^0 DE-6 (2/88) ii ABSTRACT In two experiments, immature female Sprague-Dawley rats treated with superovulatory doses of pregnant mare serum gonadotropin (PMSG) were used to study the effects of antiandrogen, hydroxyflutamide, on steroid production, particularly the biologically active androgens, testosterone, 5a-dihydrotestosterone, and androstenedione. In the first experiment, the animals were given either 5 mg hydroxyflutamide or vehicle alone at 30 and 36 h following 40 IU PMSG. Compared to the vehicle group, hydroxyflutamide treatment significantly reduced the percentage of degenerate oocytes recovered from oviducts (p<0.05). Serum levels of aromatizable androgens, testosterone and androstenedione, and their aromatized product, estradiol-17fi significantly decreased (p<0.05) in hydroxyflutamide-treated group; however, the serum concentrations of nonaromatizable androgen, 5a-dihydrotestosterone, was not statistically different between the two groups. In the second experiment, ovaries stimulated with 4 or 40 IU PMSG were obtained 48 h later and cultured in the presence and absence of hydroxyflutamide (10~5M) and/or testosterone (10~7 M) to study [4-14C] pregnenolone metabolism to major steroids. In 40 IU stimulated ovaries, hydroxyflutamide significantly decreased the metabolism of pregnenolone to progesterone (p<0.01) and androstenedione (p<0.01) while the production of estradiol-17fi increased significantly (p<0.05); however, pregnenolone conversions to testosterone and 5a-dihydrotestosterone were not statistically different between the untreated and hydroxyflutamide-treated cultures. Testosterone completely reversed the hydroxyflutamide-induced alteration of pregnenolone metabolism. In contrast, there was no difference in the pregnenolone conversion patterns between the untreated and hydroxyflutamide or hydroxyflutamide plus testosterone groups in the culture of iii ovaries stimulated with 4 IU PMSG. Present results confirm previous reports that antiandrogen, hydroxyflutamide, decreases the percentage of abnormal oocytes recovered from superovulating rats, and indicates that this hydroxyflutamide effect may be partly mediated by altered ovarian steroidogenesis, specifically the reduced hypersecretion of aromatizable androgens, testosterone and androstenedione, and/or estradiol-17fi. iv TABLE OF CONTENTS Page Abstract ii Table of Contents iv List of Tables vi List of Figures vii Glossary viii Acknowledgements x INTRODUCTION 1 REVIEW OF LITERATURE 3 I. Ovary A. General 3 B. Follicle growth 4 C. Ovarian steroidogenesis 7 D. Regulation of steroidogenesis by gonadotropins 9 II. Superovulation A. Induction of superovulation 12 B. PMSG and superovulatory response 13 C. Steroids and oocyte quality 15 Ul. Androgens A. Role in ovarian function 17 B. Antiandrogen: hydroxyflutamide 18 V MATERIALS AND METHODS 22 I. Animals 22 II. Experimental Design 22 III. Assessment of Ovulation 23 IV. Organ Culture 23 V. High Performance Liquid Chromatography 24 VI. Measurement of Steroid Hormones 28 VII. Statistical Analysis 29 RESULTS 30 I. Experiment I. A. Ovulation and Oocyte Quality 30 B. Serum Steroid Levels 30 II. Experiment II. A. Pregnenolone Metabolism in Organ Culture 33 DISCUSSION 38 REFERENCES 42 vi LIST OF TABLES Table Page I. Retention times for the HPLC separated steroids 27 II. Effects of hydroxyflutamide on ovulation and oocyte quality 31 vii LIST OF FIGURES Figure Page 1. The principal ovarian steroidogenic pathway for synthesis of progestogens, androgens, and estrogens 6 2. "Two cell-type and two gonadotropin theory" of follicular steroidogenesis 10 3. The structural formula of androgens, testosterone and 5a-dihydrotestosterone, and antiandrogens, flutamide and hydroxyflutamide 20 4. Chromatogram of major steroids separated with HPLC 26 5. Serum contents of estradiol-17fi, progesterone, and androgens after administration of 4 IU PMSG alone and 40 IU PMSG plus vehicle or hydroxyflutamide to immature rats 32 6. Serum contents of androgenic steroids, androstenedione, testosterone, 5a-dihydrotestosterone after administration of 4 IU PMSG alone and 40 IU PMSG plus vehicle or hydroxy-flutamide to immature rats 34 7. Conversion of [14C]pregnenolone to major steroids by culture of 4 IU PMSG stimulated ovaries 35 8. Conversion of [14C]pregnenolone to major steroids by culture of 40 IU PMSG stimulated ovaries 36 GLOSSARY ACAT acyl coenzyme A: cholesterol-acyl transferase °C degree Celsius 1 4 C a radioactive isotope of carbon cAMP cyclic adenosine-3',5'-monphosphate Ci Curie (3.7 X10 1 0 disintegrations per minute) dpm disintegrations per minute FSH follicle stimulating hormone g gram gem twin (geminus) GnRH gonadotropin releasing hormone h hour 3 H tritium, a radioactive isotope of hydrogen hCG human chorionic gonadotropin FfDL high-density lipoprotein HMG CoA 3-hydroxy-3-methylglutaryl coenzyme A HPLC high performance liquid chromatography 3a-HSD 3a-hydroxysteroid dehydrogenase 3fi-HSD 36-hydroxysteroid dehydrogenase 17C-HSD 17fs-hydroxysteroid dehydrogenase i.e. that is (id est) IU international unit LDL low-density lipoprotein LH luteinizing hormone M molar mg milligram min minute ml millilitre m m millimetre mM millimolar mmol millimole NAD+ nicotinamide adenine dinucleotide NADPH nicotinamide adenine dinucleotide phosphate n g nanogram nm nanometre P statistical probability P450 cytochrome P-450 PBS Dulbecco's phosphate buffered saline pH - Log of H + concentration in solution Pg picogram PLSD protected least significant difference PMSG pregnant mare serum gonadotropin PST Pacific standard time RIA radioimmunoassay sec side chain cleavage SEM standard error of means sp. act. specific activity UV ultraviolet radiation v/v/v volume ratio microlitre |im micrometre times gravity X ACKNOWLEDGEMENTS I would like to express my sincere gratitude to my supervisor, Dr. Y.S. Moon, for his invaluable advise, support, and critical encouragement in the formation of this thesis. I am very grateful to Drs. Y.W. Yun and B. Ho Yuen for their support, patience and guidance. I also wish to thank Drs. J. Skala and A.F. Burton for serving on my supervisory committee. My deep appreciation is also extended to Miss S. Leung, Mrs. P. Rajamahendran, and Drs. T.B. Feng, A. Britton , and S. Ma for their concern, encouragement, and emotional support. I am also indebted to Dr. D.T. Armstrong for the generous gift of steroid hormone antisera. This study was supported by the research grants awarded to Drs. Y.S. Moon and B. Ho Yuen from the Medical Research Couincil of Canada and the British Columbia Health Care Research Foundation. I also wish to acknowledge the studentship provided by the Medical Research Council of Canada. 1 INTRODUCTION Superovulatory doses of pregnant mare serum gonadotropin (PMSG) have been shown to induce ovulation over an extended time period ranging from 24 to 72 h following treatment (Walton et al., 1983; Yun et al., 1987). These ovulations are characterized by a PMSG-dose-dependent increase in the percentage of degenerate oocytes as well as a coincidental marked alteration of ovarian steroid pattern (Yun et al., 1987). In addition, a significant positive correlation between androgen levels and the percentage of degenerate oocytes following superovulatory treatment has been noted (Yun et al., 1987). However, the role of androgens in intrafollicular oocyte and follicular maturation is not well understood. Results of in vitro studies have implicated a role of an aromatizable androgen, testosterone, in the atretic process of preantral follicles in several species (McNatty, 1978; Louvet et al., 1975; Carson et al., 1981). In contrast, effects on prevulatory follicles by androgens have been shown to be stimulatory by maintaining its "ovulabilty" through stimulation of granulosa cell mitosis, inhibition of lysosomal enzyme, acid phosphatase, and enhancement of LH binding to LH receptors of preovulatory follicles (Peluso et al., 1980), as well as its direct effects on the ovulatory process in rats treated with low doses of PMSG (Mori et al., 1977). It is also known that androgens are essential for follicular development and ovarian function by serving as a substrate for aromatization to estrogens. Furthermore, it has been suggested that the resumption and progression of meiosis in hamster oocytes may be associated with reduced levels of aromatizable androgens (Mandelbaum et al., 1982). In a recent study from this laboratory, flutamide, an antiandrogen, given to rats previously stimulated with superovulatory doses of PMSG reduced the total androgen levels in blood and 2 increased the percentage of oocytes appearing morphologically "normal" (Yun et al v 1988). To extend the previous study, the present experiment was undertaken employing hydroxyflutamide, a more potent hydroxylated derivative of flutamide (Simard et al., 1986), to determine the effects of this antiandrogen on the metabolism of the biologically active androgens, namely testosterone, androstenedione, 5a-dihydrotestosterone, with special reference to ovulation and oocyte quality in rats induced to superovulate. 3 LITERATURE REVIEW I. Ovary A. General Production of mature eggs is the primary function of the ovary. The maturation of oocytes from the start of oogenesis until ovulation occurs in a series of specialized structures, each with its own precisely regulated hormonal milieu. The endocrine functions of the ovary provide for the regulation of the reproductive system in addition to having effects on the pituitary gonadotropin release, secondary sex characters, and metabolism as well as the mating behaviour in some species. Hence, the integration of the gametogenic and endocrine functions, which are cyclic processes, ensures the regular production of healthy oocytes at a time when they are most likely to be fertilized. The periodicity of sequential changes in ovarian structure and function is called the menstrual cycle in primates, and estrous cycle in nonprimate species. These cycles result from complex interactions of various hormones secreted from the hypothalamus, pituitary, and ovaries. The gonadotropin releasing hormone (GnRH) of the hypothalamic origin causes the gonadotrophs of the anterior pituitary to synthesize and release follicle stimulating hormone (FSH) and luteinizing hormone (LH) into the blood stream. Via the circulatory system, FSH and LH reach the ovaries where the former stimulates the growth of ovarian follicles, and the latter is involved in final oocyte maturation, ovulation, and formation and regulation of the corpus luteum function. These gonadal hormones, including various peptide hormones such as inhibins released in response to FSH and LH, in turn influence the function of hypothalamus and/or pituitary by negative or positive feedback mechanisms to regulate the release of GnRH, FSH, or LH to the circulation. 4 B. Follicle growth Follicle growth is an exceedingly complex process beginning with a primordial follicle, a complex of an oocyte and associated spindle-shaped cells enclosed by the basal lamina, and progressing through the various antral stages into the mature Graafian pre-ovulatory follicle. Follicular maturation is initiated when the single layer of spindle-shaped granulosa cell precursors in some primordial follicles differentiates into cuboidal cells, which then begin to divide (van Wagenen and Simpson, 1965). In such follicles, termed primary follicles, granulosa cells mitotically proliferate to result in multiple layers of cells within the basal lamina. These cells synthesize and secrete mucopolysaccharides, which give rise to zona pellucida that surrounds the oocyte. Outside the basal lamina, the vascular thecal layer develops shortly after the granulosa cells begin to proliferate. Here, the spindle-shaped cortical stromal cells layer concentrically around the primary follicle, and a plexus of lymph and blood vessels becomes established without piercing the basal lamina. As the follicle matures, progressively more of the spindle-shaped theca cells migrate to the basal lamina, become epitheloid in shape and acquire organelles characteristic of steroid hormone-secreting cells (Harrison, 1962); these cells are called theca interna. More peripheral theca cells, known as theca externa, retain their spindle-shaped configuration and merge with stromal cells. Increase in oocyte size, granulosa cell proliferation, and differentiation and hypertrophy of theca cells - all contribute to the enlargement of the maturing follicle. Among the granulosa cells of these growing primary follicles, local pockets of fluid accumulate and coalesce to form a central fluid-filled cavity, the antrum; this event transforms the primary follicle into a Graafian follicle. The oocytes of these follicles occupy an eccentric position, enveloped by cumulus 5 oophorus. These few layers of granulosa cells are contiguous with the mural granulosa cells that line the antrum, and function to support, protect, and nourish the oocyte. The granulosa cells and the oocyte are bathed in follicular fluid; constituents of the fluid, protein bound and free sex steroid hormones, plasma proteins, mucopolysaccharides, and electrolytes (Edwards, 1974), nourish and regulate the Graafian follicle as its maturation progresses. Either atresia or ovulation is the end point of follicular growth. The former is a degenerative process whereby the developing oocyte is lost from the ovary prior to ovulation; it has been estimated that over 99 per cent of the 500,000 oocytes present in the human ovary at birth are removed through atresia (Ingram, 1962). Whereas oocyte death and follicle degeneration are an inevitable consequence of atresia, ovulation normally results in the production of mature, healthy oocyte(s). In response to the LH surge, biochemical and structural changes in the pre-ovulatory Graafian follicle result from the effects of hydrolytic enzymes, such as the plasminogen activator and collagenase, acting on the protein substrates in the antral fluid and basal lamina (Beers, 1975; Beers et al., 1975; Espey, 1974)); when the weakened follicle wall ruptures, the oocyte and adhering cumulus oophorus are extruded to culminate the process of ovulation. Immediately following ovulation, capillaries and fibroblasts from the theca interna proliferate and invade the collapsed follicular cavity, thus forming a new temporary endocrine gland, the corpus luteum. Morphological changes of the follicular granulosa and theca cells give rise respectively to large and small luteal cells in the corpus luteum; these transformed cells secrete progesterone and other sex steroid hormones. OH Testosterone HO Cholesterol (1) CH 3 HO " ' HO Dehydroepiandrosterone Pregnenolone (2) CH 3 i c=o CH, H-C-OH (7) Androstenedione Progesterone 20a - Hydroxyprogesterone (4) HO HO 17/3 - Estradiol Estrone 5a - Dihydrotestosterone Legend 1: cholesterol side-chain cleavage P-450 2: A5-3B-hydroxysteroid dehydrogenase: A 5 -4 isomerase 3: aromatase 4: 5a-reductase 5: 20ot-hydroxysteroid dehydrogenase 6: 17a-hydroxyla8e: C-17,20 lyase 7: 17B-hydroxysteroid dehydrogenase Figure 1. The principal ovarian steroidogenic pathway for synthesis of progestogens, androgens, and estrogens 7 C. Ovarian steroidogenesis The ovary elaborates all three major classes of steroids: progestogens, androgens, and estrogens (Figure 1). The common biosynthetic precursor of all steroids is a C27 compound, cholesterol, which can be synthesized de novo or taken up from the blood where it exists bound to lipoproteins. Although cholesterol from both low-density lipoproteins (LDL) and high-density lipoproteins (HDL) has been implicated as steroidogenic precursor (Gwynne and Strauss, 1982), HDL appear to be the more important source of cholesterol in rodents while in other species, it is LDL. Within the ovarian cell, cholesterol can be synthesized de novo from acetates; the rate-limiting enzyme in cholesterol biosynthesis is 3-hydroxy-3-methylglutaryl coenzyme A-reductase (HMG CoA-reductase). Be it from de novo synthesis or lipoprotein uptake, excess cholesterol is stored in intracellular lipid droplets as esters of long-chain fatty acids. Hydrolysis of these esters restores the free form of cholesterol. The level of intracellular cholesterol is maintained by two enzymes involved in the synthesis and hydrolysis of cholesterol esters, respectively acyl coenzyme A: cholesterol-acyl transferase (ACAT) and sterol ester hydrolase, in addition to HMG CoA-reductase; not surprisingly, the activities of these enzymes are under hormonal control, as well as being regulated by the intracellular levels of cholesterol (Strauss et al., 1981). Conversion of cholesterol to pregnenolone is the first, rate-limiting, and hormonally regulated step in steroidogenesis (Caron et al., 1975; Simpson, 1979). A multienzyme complex termed cholesterol side-chain cleavage P-450 (P-450scc) responsible for this conversion is located in the inner mitochondrial membrane (Farkash et al., 1986) and catalyzes three distinct chemical reactions: 20a-hydroxylation, 22-hydroxylation, and cleavage of C-20,22 bond of cholesterol side-8 chain. The C 2 1 compound, pregnenolone, formed from such reactions is the key-intermediate of all classes of steroid hormones. The most abundant product of pregnenolone metabolism is progesterone since it is produced as a biosynthetic intermediate in follicles at all stages of development, and released as an end product during peri- and postovulatory periods. A microsomal enzyme complex, A5-3fi-hydroxysteroid dehydrogenase (3fi-HSD):A 5 - 4 isomerase catalyzes the conversion of pregnenolone to progesterone (Hall, 1984). This physiologically irreversible reaction requires, an electron acceptor, nicotinamide adenine dinucleotide (NAD+). Both pregnenolone and progesterone can serve as precursors in androgen biosynthesis of respective products, dehydroepiandrosterone and androstene-dione; these two alternative pathways are known as 5-ene-3fi-hydroxy (or A5) pathway and 4-ene-3-oxo (or A4) pathway, respectively. In either case, the microsomal enzyme complex, 17cc-hydroxylase:C-17,20 lyase, catalyses the rate-limiting step of androgen production and requires the cofactor, nicotinamide adenine dinucleotide phosphate (NADPH). Dehydroepiandrosterone can be converted to androstenedione by the same enzyme complex, 3fi-HSD:A5"4 isomerase, which produce progesterone from pregnenolone. Metabolism of androstenedione to testosterone is possible via a reversible reaction mediated by 17fi-hydroxysteroid dehydrogenase (17fi-HSD). Reduction of 4-ene of testosterone by 5cc-reductase produces 5oc-dihydrotestosterone; NADPH is the source of reducing protons in this reaction. Aromatizable androgens, androstenedione and testosterone, are the substrates for the production of estrogens, respectively estrone and estradiol-1713. This conversion imparts the characteristic aromatic structure to the estrogens, and is catalyzed by an appropriately named enzyme complex, aromatase, which is 9 located in the membranes of the agranular endoplasmic reticulum. Aromatase requires 3 moles each of NADPH and molecular oxygen: 2 for hydroxylations at C-19 to form a gem-diol and the third for hydroxylation at C-2, resulting in the loss of the angular methyl group as a formate, and subsequent aromatization of the A-ring of the steroid (Goto and Fishman, 1981). D. Regulation of steroidogenesis by gonadotropins The primary regulators of ovarian steroidogenesis are the pituitary gonadotropins, FSH and LH; many of their actions are influenced by the intracellular levels of steroids. The mechanism of action of these peptide hormones is via the classic cyclic AMP-second messenger system; interaction of gonadotropins with specific receptors on target cell surface generate an intracellular rise in cAMP thereby initiating a cascade of cAMP-dependent phosphorylation/dephosphorylation of proteins to modulate the activities of steroidogenic enzymes (Kuo and Greengard, 1969). The response of the ovary to gonadotropins depends on the presence of specific receptors on the membranes of the different cell types. Granulosa cells are the only ovarian cell type that possess FSH receptors and consequently, FSH action is limited to granulosa cells. In marked contrast, many cell types express receptors for LH, e.g. the stroma, theca interna, granulosa cells, and luteal cells. Although the granulosa cells at all stages of follicular development appear to possess FSH receptors, they acquire LH receptors only after they have matured to pre-ovulatory follicle. Theca cells acquire LH responsiveness at considerably earlier stages of follicular development than do granulosa cells. One of the first effects of FSH in growing follicle is to increase the activity of aromatase. As the granulosa cells lack 17a-hydroxylase:C-17,20 lyase, the LH 10 cAMP Theca Cells 1 Cholesterol © *\ . * * " * Progesterone I Androstenedione (Circulation) 7 / Basement Membrane Progesterone Granulosa Cells k Androstenedione Cholesterol ' . * ATP cAMP Aromatase Estrogen FSH (Follicular Ruid) Figure 2. "Two cell-type, two gonadotropin theory" of follicular steroidogenesis" 11 synthesis and secretion of estrogens following FSH stimulation at early stages of follicular development are dependent upon the availability of aromatizable androgens. FSH also influences the production of progestogens in these granulosa cells by modulating the enzymes which convert cholesterol to progesterone, namely P-450scc a n d 3fi-HSD:A5*4 isomerase. As the follicle matures, FSH induces the expression of LH receptors in granulosa cells. Thereafter, LH response becomes an additional stimulus to cAMP-regulated processes, especially on the enzymes of progesterone biosynthesis. In addition, the steroidogenic actions of LH on theca cells apparently increase the activities of 17a-hydroxylase:C-17,20 lyase in ovaries (Bogovich and Richards, 1982). Previous studies have demonstrated a "two cell-type, two gonadotropin theory" (Figure 2); there are two principal cell types involved in follicular steroidogenesis: (1) LH-responsive secretory cells comprising the theca interna cells of the follicular envelope and the interstitial cells of the ovarian stroma, and (2) FSH-responsive cells which are the granulosa cells. According to this model, theca interna cells are stimulated by LH to produce androgen from cholesterol, which diffuse across the basal lamina to be used in FSH-stimulated estrogen synthesis by granulosa cells (Moon et al., 1975; Makris and Ryan, 1975; Moon et al., 1978; Erickson, 1978; Tsang et al., 1979). Since granulosa cells secrete progesterone in response to gonadotropins, it is also possible that progesterone of granulosa cell origin may diffuse into the theca layer to serve as a substrate for androgen synthesis. Theca interna cells convert progesterone to androstenedione by 17tx-hydroxylase:C-17,20 lyase. In contrast, granulosa cells do not have significant activities of P-450 enzymes and consequently produce little or no androgens from either progesterone or pregnenolone (Short, 1962; Fowler et al., 1980). However, granulosa cells do possess considerable 176-HSD activity 12 (Makris and Ryan , 1980; M o o n and Duleba , 1982), w h i c h results i n the conversion of androstenedione and testosterone to estrone and estradiol-17fi, respectively. Al though androstenedione is the major ovarian androgen in most species, equil ibrium of 176-HSD activity favours the production of estradiol-17fi as the major estrogen. These interactions between L H and F S H together wi th the cyclical changes in plasma concentrations of L H and F S H provide a mechanism to account for the regulation of ovarian steroidogenesis and follicular growth. II. Superovula t ion A . Induction of superovulation Attempts to increase the number of mammal ian oocytes capable of fertilization and normal development began more than 60 years ago wi th the discovery of pituitary gonadotropins. Studies of Smith and Engle (1927), and Engle (1927) are some of the first accounts which describe the phenomenon of superovulation. Using daily implantation of anterior pituitary tissues in mice, they were able to produce excessive numbers of oocytes (20-48) and implantation sites (19-29). Cole (1936) simplified the procedure of superovulation induction by g iv ing a single injection of unfractionated pregnant mare's serum which produced a maximum of 54 oocytes. Subsequently, investigators found a greater sensitivity to exogenous gonadotropins in immature mice (>80 oocytes ovulated, Gates, 1971) than in mature mice (30-40 oocytes ovulated, Fowler and Edwards, 1951; Biggers et al., 1971). Two possible explanations have been suggested: (1) the mature mice have fewer large follicles capable of responding to gonadotropins than immature mice (Jones and Krohn , 1961), and (2) difficulty i n t iming the administration cycle (McLaren and Mich ie , 1959). Thus, superovulation i n 13 immature females induced with exogenous gonadotropins have become a convenient, and economical model for the study of reproduction. A principal action of exogenous gonadotropins in the induction of superovulation is now known to be the recruitment of a non-growing pool of primordial follicles into a more mature-size class (Chatterjee et al., 1977; Hirshfield, 1989) and/or the rescue of follicles at an early stage of atresia (Peters, 1979; Braw and Tsafriri, 1980). Although a variety of mechanisms may account for the overall increase in the number of healthy follicles and a simultaneous decrease in the number of atretic follicles, these two concepts are not mutually exclusive. Indeed, several investigators have reported evidence that gonadotropins stimulate a cohort of new follicles, but prevent or reverse atresia in mouse follicles in vivo (Peters et al., 1975; Byskov, 1979) and in cultured sheep follicles (Hay et al., 1979). Considering the synergistic action of FSH and LH on folliculogenesis (Falck, 1959; Short, 1962), superovulatory potency appear to depend on the mixture of FSH and LH activities of gonadotropin preparations. Thus, the use of PMSG, primarily a long-acting FSH-like hormone preparation with some LH-like activity, has been developed to induce superovulation. B. PMSG and superovulatory response PMSG is one of the most widely used gonadotropin preparations to induce superovulation in laboratory animals and large domestic species (Cole, 1975). This glycoprotein hormone is structurally similar to human chorionic gonadotropin (hCG, Moor et al., 1980); it contains a hormonally non-specific domain (PMSG-a) and a hormone specific domain (PMSG-fi) which expresses both FSH- and LH-like activities (Gospodarowicz and Papkoff, 1967; Papkoff et al., 1978). The inherent FSH- and LH-like activities, coupled with long half-life, 14 attributable to its high sialic acid content (Schams et al., 1978) contribute to its efficacy in stimulating superovulation. Results of numerous studies using immature rats indicate that low doses (4-8 IU) of PMSG induce a synchronized ovulation (8-12 oocytes) within 72 h by eliciting an endogenous LH surge between 54 and 57 h after the gonadotropin injection (Kostyk et al., 1978; Walton et al., 1983; Yun et al., 1987) and produce normal pregnancy without significantly increasing embryonic or fetal wastage (Nuti et al., 1975). With this regimen, the patterns of circulating steroid hormones (estradiol-17fi and progesterone) and LH during the 24 h period preceding the ovulation at 72 h (Wilson et al., 1974), and the temporal relationship between LH levels and oocyte maturation (Hillensjo et al., 1974) are comparable to those observed at proestrus and estrus of cycling adult rats (Linkie and Niswender, 1972). In contrast, superovulatory doses (20 or 40 IU) of PMSG induce prolonged multiple ovulations that average over 50 oocytes released per rat between 60 and 72 h (Yun et al., 1987). Similar observations of multiple waves of premature or asynchronous ovulations have been made in several species including other rodents (Stern and Schuetz, 1970; Hoffmann et al., 1985), cattle (Bellows and Short, 1972; Callesen et al., 1987), and humans (Navot et al., 1984). In superovulating rats, the first ovulation is induced as early as 24 h with two distinct increases in the number of released oocytes: one prior to 36 h and the other after 48 h (Yun et al., 1987). Two different mechanisms have been suggested to explain the two discrete sets of ovulations in rats induced to superovulate with PMSG: early ovulation within the first 48 h is due to the exogenous LH-activity of PMSG, whereas the second ovulation results from the pituitary-dependent endogenous LH surge (De la Strata et al., 1972; Yun et al., 15 1989). Thus, a prolonged exposure to LH stimulus derived from PMSG (Schams et al., 1978) may explain the precocious ovulation which has been frequently associated with interference of reproductive performance. C. Steroids and oocyte quality Superimposed on the problems of excessive variability of ovulatory response are those associated with inferior oocyte quality in superovulation. Oocytes require a specific intra-follicular steroid milieu for the completion of the full maturational process. A shift in follicular steroid pattern from estrogen- to progesterone-dominance following the LH surge has been demonstrated in regular cycles of many species including cattle (Fortune and Hansel, 1985; Hyttel et al., 1986), sheep (Moor et al., 1973), and humans (McNatty et al., 1979; Moon et al., 1985). Furthermore, the dependence of steroidogenic pattern during the peri-ovulatory period on the size of ovarian follicles, degree of atresia, and number of LH-receptors on granulosa and theca cells is well established (Fortune and Hansel, 1985; Hyttel et al., 1986; McNatty et al., 1984). Therefore, the nature of exogenous gonadotropin-induced follicle growth may affect the final oocyte quality by changing the steroidal microenvironment of the follicle. Abnormal steroid patterns in many species given superovulatory treatment is well known. In sheep, the steroid profile following superovulation with PMSG is characterized by the hypersecretion of follicular estradiol-17fi (Moor et al., 1984). Similarly, excessive rises in both ovarian and serum estradiol-17fi have been noted prior to and subsequent to the normally expected time of ovulation in superovulated rats (Miller and Armstrong, 1981; Walton and Armstrong, 1981). Using cattle given superovulatory treatment, Callesen et al. (1986) have shown an abnormally low and high progesterone/estrogen ratio 16 in a majority of follicles during the pre- and peri-ovulatory periods. In a study of overall ovarian steroid responses (estrogen, progesterone and androgens) following superovulatory treatment with PMSG in immature rats, marked elevations of progesterone and androgens, and consistent disruptions of sequential changes in overall ratios of androgens/estradiol-17fi, progesterone/estradiol-17fi, and progesterone/androgens as compared to the control regimen have been reported (Yun et al., 1989). The effects of estrogen and progesterone on mammalian oocyte maturation in vitro have been previously documented. Progesterone has been reported to enhance the maturation of denuded bovine and rabbit oocytes (Robertson and Baker, 1969), cumulus-enclosed rabbit oocytes (Bae and Foote, 1975), and denuded oocytes from gonadotropin-treated pre-pubertal rhesus monkeys (Gould and Graham, 1976). In contrast, estradiol-17fi has been reported to exhibit inhibitory effects on the maturation of cumulus-enclosed bovine and rabbit oocytes (Robertson and Baker, 1969), denuded porcine oocytes (McGaughey, 1977) , and denuded mouse oocytes (Nekola and Smith, 1974; Eppig and Koide, 1978) . Although there is some evidence indicating that the early sequence of maturational changes in the nucleus leading to germinal vesicle breakdown and formation of the first metaphase plate are independent of steroid support (Lieberman et al., 1976), Moor et al. (1980) have shown in vivo that alterations in the steroid profile during maturation, especially in pre-ovulatory follicles, induce morphological changes in the oocyte which are expressed as gross abnormalities at fertilization including polyspermy, arrested sperm condensation, and fragmented male pronuclei. Discordance of follicular development and oocyte maturation following superovulatory treatment with PMSG has been shown in sheep (Moor et al., 17 1984), cattle (Greve et al., 1984), and rats (Yun et al., 1989). The asynchronous nuclear maturation following superovulation as evidenced by the recovery of oocytes at various stages of nuclear maturation in the superovulated rats (Yun et al., 1989) has been attributed to meiotic arrest before or at the diakinesis stage or after the germinal vesicle breakdown and premature meiotic activation of oocytes from certain follicles (Callesen et al., 1986). The study of Yun et al. (1987) has noted the association of these atypical ovulations of premature oocytes with a deviating course of intra-follicular steroid profiles following superovulation with PMSG. Deleterious effects on the normal development of superovulated oocytes in relation to the process of nuclear or cytoplasmic maturation requires clarification. However, the oocytes with faulty maturation following superovulation may be susceptible to a hostile oviductal environment influenced secondarily by altered follicular steroid response (Walton and Armstrong, 1981). III. Androgens A. Role in ovarian function With the establishment of the obligatory role of androgens as substrates for aromatase enzymes to produce estrogens, other possible roles of androgens in ovarian function received little attention since androgens were not considered as one of the "female steroid hormones." However, the discovery of specific androgen binding sites in granulosa cell compartment of estrogen-primed, hypophysectomized immature rats (Schreiber and Ross, 1976) has provided a convincing evidence for other regulatory functions of androgens in the female. In granulosa cells, pre-treatment of intact rats with 5a-dihydrotestosterone 18 inhibits the induction of LH-receptors by FSH and this effect is antagonized by estrogen treatment (Farookhi, 1980). In synergism with FSH, androgens induce granulosa cell aromatase (Daniel and Armstrong, 1980) and stimulate progesterone production in cultured rat granulosa cells (Armstrong and Dorrington, 1976; Nimrod and Lindner, 1976) probably by stimulation of P450scc and 312-HSD (Nimrod, 1977; Welsh et al., 1982). There has been disparate evidence for the role of androgens in intra-follicular oocyte and follicular maturation. Aromatizable androgen, testosterone, has been shown to inhibit estrogen-induced preantral follicle growth (Louvet et al., 1975; Hillier and Ross, 1979), and induce atresia of antral follicles (Zeleznik et al., 1979). In contrast, androgens exert stimulatory effects on pre-ovulatory follicles as testosterone antiserum (Mori et al., 1977) or a non-steroidal antiandrogen (Peluso et al., 1980) was found to decrease the number of follicles ovulating in response to an ovulatory injection of hCG in PMSG-primed immature rats. Although both inhibitory and stimulatory effects of 5a-dihydrotestosterone on follicular development have been reported (Bagnell et al., 1982; Kohut et al., 1985), Smith and Tenney (1978) have demonstrated that this non-aromatizable androgen does not inhibit the maturation or increase the incidence of mouse oocyte degeneration in vitro. B. Antiandrogen: hydroxyflutamide Antiandrogens are of great importance in clinical conditions caused by absolute or relative excess of androgens. The ideal antiandrogen should be non-toxic, effective at low concentrations and devoid of intrinsic hormonal activity (Mainwaring et al., 1974). To date, the presently available antiandrogens failed to wholly satisfy these exacting criteria; however, flutamide, antiandrogen first 19 reported by Neri et al. (1972), possesses two desirable features of ideal antiandrogen: low toxicity and lack of intrinsic hormonal activity (Mainwaring et al., 1974). Flutamide (a,a,a-trifluoro-2-methyl-4'-nitro-m-propionotoluidide) is a non-steroidal antiandrogen which upon inspection of its chemical structure apparently lacks structural homology with 5a-dihydrotestosterone and testosterone (Neri, 1977). It has been demonstrated that this antiandrogen is devoid of hormonal activity and exerts its effects by competition for androgen binding to the receptor, in particular by inhibiting the nuclear binding of the androgen-receptor complex (Neri et al., 1972; Peets et al., 1974; Mainwaring et al., 1974). When administered to human males as a single dose, flutamide was found to be rapidly metabolized to a hydroxylated derivative, a,a,oc-trifluoro-2-methyl-4'-nitro-m-lactotoluidide, which was suggested to be the active antiandrogen (Katchen and Buxbaum, 1975). In vivo studies have shown 1.5-fold antiandrogenic potency of this compound compared to flutamide, and a lack of hormonal activity (Neri, 1977). In terms of inhibiting cytoplasmic and nuclear binding of androgens, Simard et al. (1986) have demonstrated that hydroxyflutamide has approximately 1% of the potency of testosterone, compared to an approximate potency of 0.01% for flutamide, in competing for the binding to the adenohypophysial androgen receptor. Past investigations of antiandrogenic effect of this antiandrogen have focussed on its ability to displace testosterone from the specific androgen receptor in various target tissue. Recent studies have indicated that in rat liver, flutamide may affect the enzymes involved in androgen metabolism, namely 5oc-reductase and 3oc-HSD (Lax and Schriefers, 1981; Graef et al., 1987). Furthermore, alterations OH Flutamide Hydroxyflutamide Figure 3. The structural formula of androgens, testosterone and dihydrotestosterone, and antiandrogens, flutamide and hydroxyflutamide. 21 in the levels of steroid hormones following flutamide treatment to rats stimulated with superovulatory doses of PMSG support the possibility of flutamide effect on androgen metabolism; however, the specifics of the modulation of steroid metabolic pathway by flutamide is not known. 22 MATERIALS AND METHODS I. Animals Immature female Sprague-Dawley rats were purchased from Charles-River Canada Inc. (St. Constant, Quebec) at 22 days of age. The animals were housed in temperature-controlled (20°-25°C), and air-conditioned room under a photoperiod of 12 h light: 12 h dark (lights on 07:00-19:00, PST). Throughout the treatment period, all animals had free access to standard rat chow and water. At the age of 28 days, the animals were injected subcutaneously with either 4 IU (control dose) or 40 IU (superovulatory dose) PMSG between 09:00 and 10:00 h. PMSG (Equinex, Ayerst Labs, Montreal, Quebec) was given in 0.4 ml of 0.9% NaCl solution. II. Experimental Design In experiment I, at 30 and 36 h after treatment with 40 IU PMSG, rats received subcutaneous injections of 5 mg hydroxyflutamide suspended in 0.4 ml of ethanol-saline solution (1:1, v/v) or vehicle alone. The rats were sacrificed by cervical dislocation at 48, and 60 h, and trunk blood was collected for steroid radioimmunoassays (RIA). Ovaries were dissected free of bursae, connective tissue, and fat under a stereo dissecting microscope. Oviducts were separated from the uterine horns at the uterotubal junction and flushed with a small volume of Dulbecco's phosphate buffered saline (PBS), pH 7.4 to recover the ovulated oocytes and assess their gross morphology. In experiment II, PMSG treated rats were killed at 48 h, and the ovaries were collected for organ culture. 23 III. Assessment of Ovulation Oviducts were severed from the uterine horns at the utero-tubal junction and placed in a few drops of PBS in 35X10 mm tissue culture dish (Falcon Labware, Becton Dickinson Canada Inc., Mississauga, Ontario). Under a stereo dissecting microscope (20X magnification, Nikon SMX-10), the cumulus-enclosed oocyte was readily visible in the enlarged and translucent ampulla of the oviduct. The oocyte mass was expelled from the oviduct by inserting a blunt-ended 30 gauge needle through the infundibulum and flushing with 0.3 - 0.5 ml of PBS. Subsequently, in order to facilitate the oocyte counting, the extracoronal cumulus cells surrounding the oocytes were dispersed using 0.1% of hyaluronidase (ovine type II, Sigma Chemical Co., St. Louis, MO) in PBS for 5-10 min. The recovered oocytes were counted under the stereo dissecting microscope (40x magnification) and examined without staining using a phase-contrast microscopy (100X magnification, Nikon Optiphot). As described elsewhere (Elsden et al., 1978; Miller and Armstrong, 1981; Yun et al., 1987), the oocytes were scored as abnormal when the occurrence of fragmentation and other degenerative changes such as irregular cell mass with debris, an amorphous opaque mass of vitelline material, and empty zona pellucida were evident. Fragmented oocytes constituted a majority of oocytes that were scored as abnormal. IV. Organ Culture In the incubation medium, each ovary was divided into approximate thirds by cutting twice with respect to hilus . Ovary pieces were randomly distributed among 24-well polystyrene plates (Falcon Labware, Becton Dickinson 24 Canada Inc.) with each well containing the equivalent of one ovary (2 ends, and 1 centre-piece). Medium used was Eagle's minimum essential medium supplemented with nonessential amino acids (0.1 mM), L-glutamine (2 mM), penicillin (100 U/ml), streptomycin (100 mg/ml), and fungizone (250 ng/ml) - all obtained from Grand Island Biological Company (Grand Island, NY). Incubations were carried out in Water-Jacketed Incubator (Forma Scientific Division, Mallink rodt Inc., Marietta, OH) at 37°C under an atmosphere of 5% C O 2 in air. The tissues were preincubated for 6 h with and without hydroxyflutamide (10"5 M) and/or testosterone (10"7 M), washed in fresh medium, and re-incubated with the same concentrations of antiandrogen and testosterone for additional 4 h in the presence of [14C] pregnenolone (0.5 mM; sp. act. 55 Ci/mol, New England Nuclear Research Products, Du Pont Canada Inc., Mississauga, Ontario). The incubations were carried out in quadruplicates for each treatment group and the experiment was repeated three times. At the end of incubations, the harvested media were stored at -20°C until subsequent analysis, and the protein content of homogenized tissues was determined by the method of Lowry et al. (1951) V. High Performance Liquid Chromatography The 1.0 ml aliquots of sera were extracted twice with five volumes of diethyl ether (anhydrous, BDH Chemicals Ltd., Toronto, Ontario) by vortexing for 2 min. The extracts were evaporated to dryness under nitrogen flow at 35°C and reconstituted with 100 pi of methanol. An integrated HPLC system from Bio-Rad (Richmond, CA) consisting of Model 700 Gradient Module, Model 1350 Soft-Start Pumps, dynamic gradient mixer (stainless steel), Model 7125 syringe loading injector fitted with 50 pi 25 injection loop, and Model 700 Data Station was used to separate the steroids. Elutions were performed on a ODS-5S reversed-phase column (240x4.5 mm, Bio-Rad) using a gradient of acetonitrile and methanol (Omnisolv chromatography grade, BDH Chemicals) in highly purified water (Milli-Q Reagent Water System, Millipore Corporation, Bedford, MA). All solvents were passed through 0.22 um filter (Millex-GV, Millipore Corporation) and de-gassed everyday prior to use. To ensure that injection loop was loaded to full capacity, an overfill technique was employed. After injection of the sample, the column was eluted with 20% methanol in water for 3 min. Then an increasing gradient of acetonitrile from 60% to 85%, and a decreasing gradient of methanol from 20% to 5% in water were applied for 26 min. The flow rate was 1 ml/min, and the temperature was ambient. The effluent hormone fractions were collected in ether-rinsed glass culture tubes (13X150 mm) at retention times previously determined from an elution profile utilizing a standard mixture of steroids containing about 1 mg each of estradiol-17fi, testosterone, androstenedione, and progesterone (Figure 3). The elution of steroid standards was monitored using an UV absorption detector (UVIS 204, Linear Instruments, Reno, NV) at 230 nm, and this profile was checked each morning prior to HPLC separation of serum samples. In order to determine the retention time of 5a-dihydrotestosterone (non-UV-absorbing steroid), cold and tritium labelled 5cc-dihydrotestosterone and the four steroid standards listed above were separated by HPLC. Using UV230 and continuous flow radioactivity detections (Flo-one\fieta, Radiomatic Instruments and Chemical Co., Tampa, FL) in series, the retention time of cold 5a-dihydrotestosterone was estimated (Table I). The collected fractions were -GRAPH KEY. 360 MV 100 X 36.00 MIN. XB H #A Y/ 0 XC 0B B/ 0 A D J 41_ .121- .161. _20J_ -24J_ -281. -32J_ .361 Figure 4. Chromatogram of major steroids separated with HPLC. Separation with a gradient of increasing acetonitrile concentration from a solvent (CH3CN: CH3OH: H20) ratio of 60:10: 30, v /v /v at 5 min to 85: 5: 10 at 26 min. E2=estradiol, T=testosterone, A=androstenedione, D=5a-dihydrotestosterone, P4=progesterone. Table I. Retention times for the HPLC separated steroids Trivial Name Retention Time (min) Estradiol-17fi 12.53 Testosterone 13.86 Androstenedione 14.86 5a-dihydrotestosterone ~15.9* 20a-hydroxyprogesterone 17.74 Progesterone 20.14 Pregnenolone ~21.7* * Estimated from the chromatography of radiolabeled steroids employing acontinuous flow radioactive detector connected in series to a UV detector 28 evaporated under nitrogen flow at 35°C and reconstituted with 1.0 ml of absolute ethanol for RIA. To determine the procedural loss from HPLC separation of major steroids, steroid free serum was used. Pooled, normal rat serum was incubated with activated Norit-A charcoal (Fisher Scientific Co., Fair Lawn, NJ) employing a ratio of 2 g to 10 ml of serum for 24 h at 4°C. The resulting slurry was centrifuged 4 times at 4700 Xg for 30 min. It has been reported that this procedure removes over 99% of steroids from serum without affecting the total protein concentration or pH (Hollander and Shenkman, 1974). Subsequently, known quantities of the five steroid standards were added to the serum and separated by HPLC as described above. The procedural loss from HPLC separation was less than 20% for all steroids studied. VI. Measurement of Steroid Hormones To the evaporated fractions from organ culture experiment, Scinti-Verse E (Fisher Scientific Co.) was added, and radiolabeled steroid content was determined by counting on LKB 1217 Rackbeta liquid scintillation counter. Specific antibodies, kindly donated by Dr. D.T. Armstrong, University of Western Ontario, London, Ontario, were used for each radioimmunoassays (RIA) of estradiol-17fi, progesterone, androstenedione, and testosterone. Cold steroid standards, estradiol-17fi, progesterone, testosterone, androstenedione, and 5oc-dihydrotestosterone, were obtained from Sigma Chemical Co., and the radiolabeled steroid tracers, [2,4,6,7,16,17-3H] estradiol-17fi (sp. act. 140 Ci/mmol), [1,2,6,7,16,17-3H] progesterone (sp. act. 112 Ci/mmol), [2,6,7-3H] testosterone (sp. act. 80 Ci/mmol), [1,2,6,7-3H] androstenedione (sp. act. 110 Ci/mmol), were purchased from Amersham Co. (Arlington Heights, IL). 29 In the assay procedure, a range of 50-200 pi aliquots of the HPLC-separated steroid fractions, if necessary, after 10-fold dilution with absolute ethanol, were dried down and assayed in duplicate. Approximately 10,000 dpm of tracer was added to each tube. The unbound hormone was removed by the dextran-coated charcoal adsorption method, and the bound steroid was counted on the liquid scintillation counter. The assays were linear at 25-300 pg/tube. The binding efficiency of the steroid antibodies was 40-60%, and the non-specific binding was less than 5%. The inter- and intra-assay coefficients of variation for all steroid RIA's were less than 12% and 10% respectively. Because of the relatively high cross-reactivity of testosterone antibody to other androgens, including 5oc-dihydrotestosterone (75%), RIA measurements of 5a-dihydrotestosterone contents were carried out using this antibody. Furthermore, this testosterone antibody was employed to determine the total androgens from ether extracted serum samples which had not been separated by HPLC. VII. Statistical Analysis All results in this study are presented as means ± SEM. Statistical analysis of data was achieved employing the Student's t-test, or when appropriate, the analysis of variance followed by Fisher's PLSD test. Differences were considered significant at p<0.05. 30 R E S U L T S I. Experiment I: A. Ovulation and oocyte quality The ovulatory response in animals given 40 IU PMSG was not affected by hydroxyflutamide treatment (Table II). In both the hydroxyflutamide and vehicle groups, the mean number of oocytes recovered at 48 and 60 h were 2.1- to 4.2-fold (both p<0.05) greater than the number ovulated by 60 h from rats injected with 4 IU PMSG, indicating a good superovulatory stimulation. The slightly lower mean number of oocytes in hydroxyflutamide group compared to vehicle regimen was not statistically significant at 48 and 60 h. However, the reduction in the percentage of degenerate oocytes recovered from hydroxyflutamide-treated animals compared to vehicle group was significant at both time periods studied. B. Serum steroid levels Serum steroid contents of androgens, estradiol-176, and progesterone after treatment with hydroxyflutamide or vehicle are shown in Figure 5. At 48 h, the levels of estradiol-17fi in hydroxyflutamide and vehicle groups were comparable to rats given only 4 IU PMSG. But by 60 h, the elevated estradiol-17fi concentration due to superovulatory dose of PMSG in vehicle group (0.682±0.145 ng/ml) significantly declined (p<0.05) following hydroxyflutamide treatment to a value (0.269±0.053 ng/ml) similar to rats treated with control regimen (0.272±0.113 ng/ml). Serum progesterone levels were not affected by hydroxyflutamide in superovulating rats. In contrast, the total androgens determined by radioimmunoassay without prior HPLC separation showed a very large increase above the control levels in both hydroxyflutamide and vehicle groups. By 60 h, hydroxyflutamide-treated s uperovulating rats exhibited a 31 Table II. Effects of hydroxyflutamide on ovulation and oocyte quality. Time (h) after PMSG a 48 60 Control No. of oocytes - 7.0 ±1 .3 % degenerate oocytes - 0.5 ± 0.3 Vehicle0 No. of oocytes 18.4 ± 2.1 29.3 ± 3.7 % degenerate oocytes 26.7 ± 3.7 34.7 ± 6.0 Hydroxyflutamide^ No. of oocytes 16.8 ± 2.3 27.4 ± 5.6 % degenerate oocytes 3.0 ± 1.76 12.1 ± 3.4* a Results are expressed as means ± SEM for the number of ovulated rats (n = 6 or 7). b Rats were given 4 I U PMSG. C/d Rats were treated with vehicle or 5 mg hydroxyflutamide at 30 and 36 h afterinjection of 40 I U PMSG. e/f p< 0.05, compared to corresponding vehicle treatment group E o> CO I -J o B CO ESTRADIOL-178 1.0 0.8 0.6 0.4 0.2 0.0 • 4 IU PMSG Q 40 IU PMSG+Veh S 40 IU PMSG+Flu 48 PROGESTERONE 70 80 -SO -40 30 H 20 10 48 ANDROGENS s.o 4.0 -3.0 2.0 1.0 H 0.0 60 60 48 60 Time (h) after PMSG Figure 5. Serum contents of estradiol-17fi, progesterone, and androgens after administration of 4 IU PMSG alone and 40 IU PMSG plus vehicle or hydroxyflutamide to immature rats. Values are means ± SEM (n=6 or 7). The means with no letters in common are significantly different (p<0.05). 33 significant 45% (p<0.05) decrease in total androgens to 2.07510.237 ng/ml, a value which was still considerably elevated above the control level (0.355±0.102 ng/ml). Decline in total androgens observed by 60 h in hydroxyflutamide treated rats is due to reduction in aromatizable androgens (Figure 6). Compared to vehicle, androstenedione and testosterone levels decreased 67% (p<0.01) and 62% (p<0.01) in hydroxyflutamide-treated animals. The concentration of the third biologically active androgen, 5a-dihydrotestosterone, was not significantly changed by the antiandrogen treatment. I. Experiment II: A. Pregnenolone metabolism in organ culture The ability of hydroxyflutamide to alter the metabolism of [14C]pregnenolone in culture of ovaries stimulated with 4 or 40 IU PMSG are shown in Figures 7 and 8 respectively. Incubations of 4 IU PMSG stimulated ovaries were unresponsive to hydroxyflutamide (10~5 M) alone or with testosterone (10~7 M). However, significant decline in the conversion of pregnenolone to progesterone (p<0.01) and androstenedione (p<0.01) was observed in the incubations with hydroxyflutamide in 40 IU PMSG stimulated ovaries (Figure 8). The presence of the antiandrogen also significantly increased the accumulation of radiolabeled estradiol-17fi (p<0.05) without influencing the levels testosterone and 5oc-dihydrotestosterone. The percentage of substrate recovered in these incubations were 13% compared to 50% and 45% in control and hydroxyflutamide plus testosterone groups respectively (data not shown), indicating that a portion of radiolabeled pregnenolone was probably metabolized to other steroid fraction(s) not collected in this study. Interestingly, the addition 34 ANDROSTENEDIONE OB c 0) 2 2 CD (0 2.0 1.5 1.0 0.5 -0.0 • 4 IU PMSG Q 40 IU PMSG+Veh m 40IUPMSG+FIU 46 60 TESTOSTERONE 3.0 i b 2.5 i 2.0 1.5 1.0 0.5-0.0 48 60 5o-DIHYDROTESTOSTERONE 0.8-48 60 Time (h) after PMSQ Figure 6. Serum contents of androgenic steroids, androstenedione, testosterone, 5a-dihydrotestosterone after administration of 4 IU PMSG alone and 40 IU PMSG plus vehicle or hydroxyflutamide to immature rats. Values are means ± SEM (n=6 or 7). The means with no letters in common are significantly different (p<0.05). 35 50 -i Androstenedione Testosterone Dlhydrotestosterone Estradiol Progesterone Figure 7. Conversion of P'KTlpregnenolone to major steroids by culture of 4 IU PMSG stimulated ovaries. 36 Figure 8. Conversion of p4C]pregnenolone to major steroids by culture of 40 IU PMSG stimulated ovaries. * p<0.05, ** p<0.01, compared to corresponding steroid levels in CONTROL and OH-FLU+T groups of testosterone completely reversed the antiandrogen-induced alterations pregnenolone metabolism (Figure 8). 38 DISCUSSION The results of the present study show that hydroxyflutamide treatment reduced the androgens level, specifically the levels of aromatizable androgens (androstenedione and testosterone), following the administration of superovulatory doses of PMSG to immature female rats. Additionally, this study confirms the earlier reports by our laboratory which indicated that antiandrogen flutamide reduces oocyte degeneration in the superovulation rat model (Yun et al., 1988) The synthetic nonsteroidal antiandrogen, hydroxyflutamide, is a pharmaceutically potent compound of great importance in clinical conditions caused by absolute or relative excess of androgens. It has been demonstrated that the primary mode of action of this antiandrogen is via the inhibition of androgen uptake and/or nuclear binding of androgens without other hormonal activities (Neri et al., 1972; Peets et al., 1974). Indeed, twice daily injections of 5 mg flutamide for 4 cycles in intact adult female rats have been demonstrated to have no effect on the estrous cycle, with no significant change in the plasma concentration of estradiol-17fi (Luthy et al., 1987). But recentiy, flutamide effect of decreasing in vivo production of androgens and estradiol-17fi has been reported in rats treated with superovulatory doses of PMSG (Yun et al., 1988). In addition, in vitro results of Hillier et al. (1977) show that the addition of flutamide to rat granulosa cell cultures alters their steroidogenic capacity to produce progesterone. In the present study, hydroxyflutamide significantly reduced the serum levels of estradiol-17fi and its aromatizable androgen precursors, testosterone and androstenedione, by 60 h after PMSG in immature rats. These results suggest that hydroxyflutamide, in addition to its action at the receptor level, may also inhibit the synthesis of androgens and estrogens in rats 39 treated with a superovulatory dose of PMSG (40 IU) by a specific reduction of aromatizable androgens. Since the serum levels of progesterone or 5a-dihydrotestosterone were not affected by the antiandrogen treatment, it thus appears that the site of inhibition in the steroidogenic pathway by hydroxyflutamide is at 17a-hydroxylase and/or C-17,20-lyase. Indeed, enzyme assays using microsomal preparations of rat testis have shown that flutamide and hydroxyflutamide competitively inhibit 17a-hydroxylase and C-17,20-lyase (Ayub and Levell, 1987). The reduction of aromatizable androgens and estradiol-171S by hydroxyflutamide in superovulating rats is in contrast to a previous report which showed no effect on ovarian steroid levels by flutamide treatment to immature rats primed with 5 IU PMSG (Peluso et al., 1980). This discrepancy may be accounted for by the different doses and batches commercial preparations of PMSG employed (Murphy et al., 1984); superovulatory regimen may provide a different response to antiandrogen at the ovarian level. In fact, our previous study has shown that a superovulatory dose of PMSG significantly increases ovarian androgen levels, while control dose significantly decreases the levels after 36 h (Yun et al., 1987). Results of experiment II seem to support this concept. Hydroxyflutamide was without effect on the metabolism of pregnenolone in the culture of 4 IU PMSG stimulated ovaries; however, in the cultures of 40 IU stimulated ovaries, it markedly altered the pattern of pregnenolone metabolism which was completely restored by the addition of testosterone. Although the disparate effects of hydroxyflutamide on in vivo and in vitro production of progesterone and estradiol-176 (no effect in vivo, decreased progesterone production in vitro; decreased estradiol-17fi levels in vivo, increased production in vitro) is very puzzling and requires further study, 40 the in vitro results clearly demonstrate that hydroxyflutamide alters steroidogenesis in ovaries given superovulatory dose of PMSG. Furthermore, it suggest that there may exist functional differences between ovaries treated with control and superovulatory regimens of PMSG in their response to hydroxyflutamide. Marked elevation of ovarian and serum steroid hormone (particularly androgens) following superovulation with PMSG in rats has been associated with high incidence of degenerate oocytes prior to fertilization (Yun et al., 1987). Flutamide treatment of these superovulating rats decreased the production of ovarian androgens and profoundly reduced the oocyte and embryo degeneration (Yun et al., 1988). Results of the present study showing reduced aromatizable androgen production associated with reduced oocyte degeneration in hydroxyflutamide-treated superovulating rats confirm and extend our previous studies. Favourable follicular microenvironment of antiandrogen-treated animals due in part to the reduced hypersecretion of aromatizable androgens may perhaps partly account for the recovery of larger percentage of oocytes with improved gross morphology. Indeed, in superovulating hamster, resumption of meiosis and oocyte maturation were concomitant with changes in follicular fluid steroid concentrations, including a decline in testosterone, androstenedione, and estradiol-176 (Mandelbaum et al., 1982). In contrast, disruption of follicular steroid patterns, asynchronous follicular development and intrafollicular oocyte maturation in superovulating sheep (Moor et al., 1984, 1985) and cattle (Greve et al., 1984; Callesen et al., 1986) have been reported. Furthermore, a recent-finding of various stages of nuclear maturation in superovulated oocytes from PMSG treated rats indicate that meiotically aberrant oocytes may be ovulated from physiologically unfit follicles; these atypical ovulations of superovulated oocytes 41 with premature or asynchronous nuclear maturations have been closely related to abnormal follicular steroid profiles (Yun et al., 1989). Another alternative mechanism by which hydroxyflutamide may reduce oocyte degeneration is by enhancing the viability of ovulated oocytes by improving the steroid-affected oviductal environment (Yun et al., 1988). Deleterious effects of excessively elevated estradiol-17fi on oocyte aging and early embryo development have been widely documented (Cline et al., 1977; Stone and Hamner, 1977; Miller and Armstrong, 1981). Decreased estradiol-17fi production by 60 h in the present study due to the limited availability of aromatizable androgens in animals treated with the antiandrogen may contribute to an establishment of less hostile oviductal environment for the ovulated oocytes. However, in view of the present data which indicate that reduced oocyte degeneration occurred by 48 h without changes in serum estradiol-17fi or androgens levels, alterations in steroid hormones (including estradiol-17fi) may not be the only factor in the consideration of hydroxyflutamide effects in superovulating immature rats. In summary, hydroxyflutamide reduced oocyte degeneration in superovulating rats, confirming the previous-findings of this laboratory. In view of its effects on in vivo production of testosterone and androstenedione, this hydroxyflutamide effect may be mediated in part by reduced hypersecretion of ovarian steroids, particularly aromatizable androgens and/or estradiol-17fi. However, the observed improvement in the perturbation of oocyte normality prior to changes in steroid hormones indicates that other mechanisms may be involved in this antiandrogen effect. Further study is required to elucidate the precise mode of hydroxyflutamide effect at the ovarian level. 42 REFERENCES Armstrong, D.T. and Dorrington, J.H. (1976). Androgens augment FSH-induced progesterone secretion by cultured rat granulosa cells. Endocrinology, 99: 1411-1414. Ayub, M. and Levell, M.J. (1987). Inhibition of rat testicular 17oc-hydroxylase and 17,20 lyase activities by antiandrogens (flutamide, hydroxyflutamide, RU 23908, cyproterone acetate) in vitro. J. Steroid Biochem., 28: 43-47. Bae, LH. and Foote, R.H. (1975). Effects of hormone on the maturation of rabbit oocytes recovered from follicles of various sizes. J. Reprod. Fert, 42: 357-360. Bagnell, C.A., Mills, T.M., Costoff, A., and Mahesh, V.B. (1982). A model for the study of androgen effects on follicular atresia and ovulation. Biol. Reprod., 27: 903-914. Beers, W.H. (1975). Follicular plasminogen and plasminogen activator and the effect of plasmin on ovarian follicular wall. Cell, 6: 379-386. Beers, W.H., Strickland, S., and Reich, E. (1975). Ovarian plasminogen activator: Relationship to ovulation and hormonal regulation. Cell, 6: 387-394. Bellows, R.A. and Short, R.E. (1972). Superovulation and multiple births in beef cattle. 10tn Biennial Symposium on Animal Reproduction. 34 (suppl. 1): 67-79. Biggers, J.D., Whitten, W.K., and Whittingham, D.G. (1971). The culture of mouse embryos in vitro. In Methods in Mammalian Embryology. J.C. Daniel, Jr. ed., Freeman, San Francisco, ch. 6, pp. 86-116. Bogovich, K., and Richards, J.S. (1982). Androgen biosynthesis in developing ovarian follicles: evidence that luteinizing hormone regulates thecal 17a-hydroxylase and C-17,20 lyase activities. Endocrinology, 111: 1201-1208. Braw, R.H. and Tsafriri, A. (1980). Effects of PMSG on follicular atresia in the immature rat ovary. J. Reprod. Fert, 59: 267-272. 43 Byskov, A.G. (1979). Atresia. In Ovarian Follicular Development and Function, A.R. Midgley Jr., and W.A., Sadler Jr. eds., Raven Press, New York, pp. 41-57. Callesen, H., Greve, T., and Hyttel, P. (1986). Preovulatory endocrinology and oocyte maturation in superovulated cattle. Theriogenology, 25: 71-86. Callesen, H., Greve, T., and Hyttel, P. (1987). Premature ovulations in superovulated cattle. Theriogenology, 28: 155-165. Caron, M.G., Goldstein, S., Savard, K., and Narsh, J. (1975). Protein kinase stimulation of a reconstituted cholesterol side chain cleavage system in the bovine corpus luteum. J. Biol. Chem., 250: 5137-5143. Carson, R.S., Findlay, J.K., Clarke, I.J., and Burger, H.G. (1981). Estradiol, testosterone and androstenedione in ovine follicular fluid during growth and atresia of ovarian follicles. Biol. Reprod., 24: 105-113. Chatterjee, A., Pal, A.K., and Gupta, T. (1977). Pregnant mare's serum gonadotropin. III. Hemispaying and the reversal antifertility faculty on pregnant mare's serum gonadotropin. Fertil. Steril., 28: 1101-1103. Cline, E.M., Randall, R.A., and Oliphant, G. (1977). Hormone-mediated oviductal influence on mouse embryo development. Fertil. Steril., 28: 766-771. Cole, H.H. (1936). On the biological properties of mare gonadotropic hormone. Am. J. Anat., 59: 299-331. Cole, H.H. (1975). Studies on reproduction with emphasis on gonadotropins, antigonadotropins and progonadotropins. Biol. Reprod., 12: 194-211. Daniel, S.A.J, and Armstrong, D.T. (1980). Enhancement of follicle-stimulating hormone-induced aromatase activity by androgens in cultured rat granulosa cells. Endocrinology, 107: 1027-1033. De la Strata, M , Forcelledo, M.L., and Serrano, C. (1972). Influence of the hypophysis on pregnant mare's serum gonadotropin-induced ovulation in immature rats. J. Reprod. Fert., 31: 23-28. 44 Edwards, R.G. (1974). Follicular fluid. J. Reprod. Fert, 37:189-219. Elsden, R.P., Nelson, L.D., and Seidel, G.E., Jr. (1978). Superovulating cows with follicle stimulating hormone and pregnant mare serum gonadotropin. Theriogenology, 9: 17-26. Engle, E.T. (1927). Pregnancy following superovulation in the mouse. Proc. Soc. Exp. Biol. Med., NY, 25: 84-85. Eppig, J.J. and Koide, S.L. (1978). Effects of progesterone and estradiol-17fi on the spontaneous meiotic maturation of mouse oocytes. J. Reprod. Fert., 53: 99-101. Erickson, G.F. (1978). Normal ovarian function. Clin. Obstet. Gynecol. 21: 31-42. Espey, L.L. (1974). Ovarian proteolytic enzymes and ovulation. Biol. Reprod. 10: 216-235. Falck, B. (1959). Site of production of estrogen in rat ovary as studied in micro-transplants. Acta Physiol. Scand. (Suppl), 47: 163. Farkash, Y., Timberg, R., and Orly, J. (1986). Preparation of antiserum to rat cytochrome P-450 cholesterol side chain cleavage, and its use for ultrastructural localization of the immunoreactive enzyme by protein A-gold technique. Endocrinology, 118:1353-1365. Farookhi, R. (1980). Effects of androgen on induction of gonadotropin receptors and gonadotropin-stimulated adenosine 3',5'-monphosphate production in rat ovarian granulosa cells. Endocrinology, 106: 1216-1223. Fortune, J.E. and Hansel, W. (1985). Considerations of steroids and gonadotropins in follicular fluid from normal heifers and heifers primed for superovulation. Biol. Reprod., 32: 1069-1079. Fowler, R.E. and Edwards, R.G. (1957). Induction of superovulation and pregnancy in mature mice by gonadotrophins. J. Endocrinol., 15: 374-384. 45 Fowler, R.E., Fox, N.L., Edwards, R.G., Walters, D.E., and Steptoe, P.C. (1978). Steroidogenesis by cultured granulosa cells apirated from human follicles using pregnenolone and androgen as precursors. J. Endocrinol. 77: 171-183. Gates, A.H. (1971). Maximizing yield and developmental uniformity of eggs. In Methods in Mammalian Embryology. J.C. Daniel, Jr. ed., Freeman, San Francisco, ch. 4, pp. 64-75. Gospodarowicz, D. and Papkoff, H. (1967). A simple method for the isolation of pregnant mare serum gonadotrophin. Endocrinology, 80: 699-702. Goto, J. and Fishman, J. (1977). Participation of nonenzymatic transformation in the biosynthesis of estrogens from androgens. Science, 195: 80-81. Gould, K.G. and Graham, C.E. (1976). Maturation in vitro of oocytes recovered from pre-pubertal rhesus monkeys. J. Reprod. Fert, 46: 269-270. Graef, V., Golf, S.W. and Tiischen, M. (1981). Sex-specific action of antiandrogens on androgens induced changes in hepatic microsomal 3fi-hydroxysteroid dehydrogenase and 5a-reductase activity in the rat. J. Steroid Biochem. 14: 883-887. Greve, T., Bousquet, D., King, W.A., and Betteridge, K.J. (1984). In vitro fertilization and cleavage of in vivo matured bovine oocytes. Theriogenology, 22: 151-165. Gwynne, J.T. and Strauss III, J.F. (1982). The role of lipoproteins in steroidogenesis and cholesterol metabolism in steroidogenic glands. Endocr. Rev., 3: 299-329. Hall, P.F. (1984). Cellular organization for steroidogenesis. Int. Rev. Cytol., 86: 53-95. Harrison, R.J. (1962). The structure of the ovary, In The Ovary. S. Zuckerman, A.M. Mandl, and P. Eckstein eds., Academic Press, London, pp. 143-182. Hay, M.F., Moor, R.M., Cran, D.G., and Dott, H.M. (1979). Regeneration of atretic ovarian follicles in vitro. J. Reprod. Fert., 55: 195-207. 46 Hillensjo, T., Barnea, A., Nillson., L., Herlitz, H., and Ahren, K. (1974). Temporal relationship between serum LH levels and oocyte maturation in prepubertal rats injected with pregnant mare's serum gonadotropin. Endocrinology, 95:1762-1766. Hillier, S.G., Knazek, R.A., and Ross, G.T. (1977). Androgenic stimulation of progesterone production by granulosa cells from preantral ovarian follicles: further in vitro studies using replicate cell cultures. Endocrinology, 100:1539-1549. Hillier, S.G. and Ross, G.T. (1979). Effects of exogenous testosterone on ovarian weight, follicular morphology, and intraovarian progesterone concentration in estrogen-primed, hypophysectomized immature female rats. Biol. Reprod., 20: 261-268. Hirshfield, A.N. (1989). Rescue of atretic follicles in vitro and in vivo. Biol. Reprod., 40: 181-190. Hoffmann, J.C., Yanagimachi, R., Peter, J., and De Feo, V.J. (1985). Multiple causes of pregnancy failure in hamsters precociously ovulated by human chorionic gonadotropin. Biol. Reprod., 33: 1147-1157. Hollander, C.S. and Shenkman, L. (1974). Thyroxine and triiodothyroxine. In Methods of Hormone Radioimmunoassay. B.M. Jaffe and H.R. Behrman eds., Academic Press, New York, p. 218. Hyttel, P., Callesen, H., and Greve, T. (1986). Ultrastructural features of preovulatory oocyte maturation in superovulated cattle. J. Reprod. Fert., 76: 645-656. Ingram, D.L. (1962). Atresia. In The Ovary. S. Zuckerman, A.M Mandl and P. Eckstein eds., Academic Press, London, pp. 247-273. Jones, E.C., and Krohn, P.L. (1961). The relationship between age, number of oocytes and fertility in virgin and multiparous mice. J. Endocrinol., 21: 469-495. 47 Katchen, B. and Buxbaum, S. (1975). Disposition of a new, nonsteroid, antiandrogen, a,a,a-trifluoro-2-methyl-4'-nitro-m-propionotoluidide (flutamide), in men following a single oral 200 mg dose. J. Clin. Endocrinol. Metab., 41: 373-379. Kohut, J.K., Jarrel. J.F., and YoungLai, E.V. (1985). Does dihydrotestosterone induce atresia in the hypophysectomized immature female rats treated with pregnant mare's serum gonadotropin? Am. J. Ob. Gy., 151: 250-205. Kostyk, S.K., Dropcho, E.T., Moltz, H., and Swartwout, J.R. (1978). Ovulation in immature rats in relation to the time and dose of injected human chorionic gonadotropin or pregnant mare serum gonadotropin. Biol. Reprod., 19: 1102-1107. Kuo, J.F. and Greengard, P. (1969). Cyclic nucleotide-dependent protein kinases. IV. Widespread occurrence of adenosine 3',5'-monophosphate-dependent protein kinases in various tissues and phyla of the animal kingdom. Proc. Natl. Acad. Sci. USA, 64: 1349-1355. Lax, E.R. and Schreifers, H. (1981). NADPH: 4-ene-3-oxosteroid-5a-reductase and NADH: 4-ene-3-oxosteroid-5a-reductase in liver microsomes of different species of animals. Acta Endocrinol. 98: 261-266. Lieberman, M.E., Tsafriri, A., Bauminger, S., Collins, W.P., Ahren, K., and Lindner, H.R. (1976). Oocytic meiosis in cultured rat follicles during inhibition of steroidogenesis. Acta Endocrinol. (Kbh.), 83: 151-157. Linkie, D.M. and Niswender, G.D. (1972). Serum levels of prolactin, luteinizing hormone, and follicle stimulating hormone during pregnancy in the rat. Endocrinology, 90: 632-637. Louvet, J.P., Harman, S.M., Schreiber, J.R., and Ross, G.T. (1975). Evidence for a role of androgens in follicular maturation. Endocrinology, 97: 366-372. Lowry, O.H., Rosebrough, N.S., Farr, A.L., and Randall, R.S. (1951). Protein measurement with the Folin phenol reagent. J. Biol. Chem., 193: 265-275. 48 Luthy, I., Caron, S., Belanger, A., and Labrie, F. (1987). Effects of flutamide, a pure antiandrogen, on endocrine parameters in the adult female rat. Gynecol. Endocrinol., 1: 151-168. McGaughey, R.W. (1977). The culture of pig oocytes in minimal medium, and the influence of progesterone and estradiol-17fi on meiotic maturation. Endocrinology, 100: 39-45. McLaren, A., and Michie, D. (1959). Superpregnancy in the mouse. I. Implantation and fetal mortality after induced superovulation in females of different ages. J. Exp. Biol., 36: 281-300. McNatty, K.P. (1978). Follicular fluid. In Vertebrate Ovary. R.E. Jones ed., Plenum Press, New York, pp. 215-259. McNatty, K.P., Heath, D.A., Henderson, K.M., Lun, S., Hurst, P.R., Ellis, L.M., Montgomery, G.W., Morrison, L., and Thurly, D.C. (1984). Some aspects of thecal and granulosa cell function during follicular development in the bovine ovary. J. Reprod. Fert, 72: 39-53. McNatty, K.P., Smith, D.M., Makris, A., Osathanondh, R., and Ryan, K.J. (1979). The microenvironment of the human antral follicle: Interrelationships among the steroid levels in antral fluid, the population of granulosa cells, and the status of the oocyte in vivo and in vitro. J. Clin. Endocrinol. Metabol., 49:851-860. Mainwaring, W.I.P., Mangan, F.R., Feherty, P.A., and Freifield, M. (1974). An investigation into the anti-androgenic properties of the non-steroidal compound, SCH 13521 (4'-nitro-3'-trifluoromethylisobutyrlanilide). Mol. Cell. Endocrinol., 1: 113-128. Makris, A. and Ryan, K.J. (1975). Progesterone, androstenedione, testosterone, estrone, and estradiol synthesis in hamster ovarian follicle cells. Endocrinology 96: 694-701. Makris, A. and Ryan, K.J. (1980). The source of follicular androgen in the hamster follicle. Steroids 35: 53-64. 49 Mandelbaum, J., Plachot, M., and Mowszowicz, I. (1982). Follicular sex steroids and prostaglandins during oocyte maturation in hamsters. In Follicular  Maturation and Ovulation. R. Rolland, E.V. van Hall, S.G. Hillier, K.P. McNatty, and J. Schoemaker eds., Excerpta Medica, Amsterdam, pp. 276-281. Miller, B.G. and Armstrong, D.T. (1981). Effects of a superovulatory dose of pregnant mare serum gonadotropin on ovarian function, serum estradiol, and progesterone levels and early embryo development in immature rats. Biol. Reprod., 25: 261-271. Moon, Y.S., Dorrington, J.H., and Armstrong, D.T. (1975). Stimulatory action of follicle stimulating hormone on estradiol-17fi secretion by hypophysectomized rat ovaries in organ culture. Endocrinology, 97: 244-247. Moon, Y.S. and Duleba, A.J. (1982). Comparative studies of androgen metabolism in theca and granulosa cells of human follicles in vitro. Steroids 39:419-430. Moon, Y.S., Ho Yuen, B., Pride, S.M., Rowe, T.C., Poland, B.J., McComb, P.F., and Gomel, V. (1985). A preliminary report on the establishment of pregnancies in an in vitro fertilization (IVF) programme at the University of British Columbia (UBC). Gamete Res., 11: 289-296. Moon, Y.S., Tsang, B.K., Simpson, C , Amstrong, D.T. (1978). 17fi-Estradiol biosynthesis in cultured granulosa and theca cells of human ovarian follicles: stimulation by follicle stimulating hormone. J. Clin. Endocrinol. Metab. 47:263-267. Moor, R.M., Hay, M.F., Mcintosh, J.E.A., and Caldwell, B.V. (1973). Effect of gonadotrophins on the production of steroids by sheep ovarian follicles cultured in vitro. J. Endocrinol., 58: 599-611. Moor, R.M., Kruip, Th.A.M., and Green, D. (1984). Intraovarian control of folliculogenesis: limits to superovulation? Theriogenology, 21: 103-116. 50 Moor, R.M V Osborn, J.C., and Crosby, I.M. (1985). Gonadotrophin-induced abnormalities in sheep oocytes after superovulation. J. Reprod. Fertil., 74: 167-172. Moor, R.M., Polge, C , and Willadsen, S.M. (1980). Effect of follicular steroids on the maturation and fertilization of mammalian oocytes. J. Embryol. Exp. Morph., 56:319-335. Mori, T., Suzuki, A., Nishimura, T., and Kambegawa, A. (1977). Evidence for androgen participation in induced ovulation in immature rats. Endocrinology, 101: 623-626, Murphy, B.D., Mapletoft, R.J., Manns, J., and Humphrey, W.D. (1984). Variability in gonadotrophin preparations as a factor in the superovulatory response. Theriogenology, 21: 117-125. Navot, D., Margalioth, E.J., Laufer, N., Mor-Yosef, S., and Schenker, J.G. (1984). Asynchronous ovulation in human menopausal gonadotropin induction" of ovulation for in vitro fertilization. Fertil. Steril., 42: 806-807. Nekola, M.V. and Smith, D.M. (1974). Steroid inhibition of oocyte maturation in vitro. Endocrinology, 94A: 165, Abstr. Neri, R.O. (1977). Studies on the biology and mechanism of action of nonsteroidal antiandrogens. In Androgens and Antiandrogens. L. Martin and M. Motta eds., Raven Press, New York, pp. 179-189. Neri, R., Florance, K., Koziol, P., and van Cleave, S. (1972). A biological profile of a nonsteroidal antiandrogen, SCH 13521 (4'-nitro-3'-trifluoromethyl-isobutyranilide). Endocrinology, 91: 427-437. Nimrod, A. (1977). Studies on the synergistic effect of androgen on the stimulation of progesterone secretion by FSH in cultured rat granulosa cells: A search for the mechanism of action. Mol. Cell. Endocrinol., 8: 201-211. Nimrod, A. and Lindner, H.R. (1976). A synergistic effect of androgen on the stimulation of progesterone secretion by FSH in cultured rat granulosa cells. Mol. Cell. Endocrinol., 5: 315-320. 51 Nuti, K.M., Sridharan, B.N., and Meyer, R.K. (1975). Reproductive biology of PMSG-primed immature female rats. Biol. Reprod., 13: 38-44. Papkoff, H., Bewley, T.A., and Ramachandran, J. (1978). Physicochemical and biological characterization of pregnant mare serum gonadotrophin and its subunits. Biochem. Biophys. Acta, 532: 373-377. Peets, E.A., Henson, M.F., and Neri, R.O. (1974). On the mechanism of the antiandrogenic action of flutamide (a,a,a-trifluoro-2-methyl-4'-nitro-m-propionotoluidide) in the rat. Endocrinology, 94: 532-540. Peluso, J.J., Stude, D., and Steger, R.W. (1980). Role of androgens in hCG-induced ovulation in PMSG-primed immature rats. Acta Endocrinol., 93: 505-512. Peters, H. (1979). Some aspects of early follicular development. In Ovarian  Follicular Development and Function. A.R. Midgley Jr. and W.A., Sadler Jr. eds., Raven Press, New York, pp. 1-13. Peters, H., Byskov, A.G., Himelstein-Braw, R., and Faber, M. (1975). Follicular growth: The basic event in the mouse and human ovary. J. Reprod. Fert., 45: 559-566. Robertson, J.E. and Baker, R.D. (1969). Role of female sex steroids as possible regulators of oocyte maturation. Proc. 2 n d Annu. Meet. Soc. Study for Reprod., Davis, Abstr. 57, p.29. Schams, D., Menzer, C., Schallenberger, E., Hoffman, B., and Hahn, R. (1978). Some studies on pregnant mare serum gonadotropin (PMSG) and on endocrine responses after application for superovulation in cattle. In Control of Reproduction in the Cow. J.M. Sreenan ed., The Hague, Martinus, Nijhoff, pp. 122-143. Schreiber, J.R. and Ross, G.T. (1976). Further characterization of rat ovarian testosterone receptor with evidence for nuclear translocation. Endocrinology, 99: 590-596. 52 Short, R.V. (1962). Steroids in the follicular fluid and the corpus luteum of the mare: "A two-cell type" theory of ovarian steroid synthesis. J. Endocrinol. 24: 59-63. Simard, J., Luthy, I., Guay, J., Belanger, A., and Labrie, I. (1986). Characteristics of interaction of the antiandrogen flutamide with the androgen receptor in various target tissues. Mol. Cell. Endocrinol., 44: 261-270. Simpson, E.R. (1979). Cholesterol side-chain cleavage, cytochrome P-450 and the control of steroidogenesis. Mol. Cell. Endocrinol., 13: 213-227. Smith, P.E. and Engle, E.T. (1927). Experimental evidence regarding the role of the anterior pituitary in the development and regulation of the genital system. Am. J. Anat., 40: 159-217. Smith, D.M. and Tenney, D.Y. (1978). Mouse oocyte maturation in vitro: effects of steroids. Biol. Reprod., 18: 69, Abstr. Stern, S. and Scheutz, A.W. (1970). Asynchrony of ovulation and mating in mice treated with gonadotropins. J. Reprod. Fert., 23: 257-261. Stewart, F., Allen, W.R., and Moor, R.M. (1976). Pregnant mare serum gonadotropin: Ratio of follicle stimulating hormone and luteinizing hormone activities measured by radioreceptor assay. J. Endocrinol., 71: 471-482. Stone, S.L. and Hamner, C.E. (1977). Hormonal and regional influences of the oviduct on the development of rabbit embryos. Biol. Reprod., 16: 636-646. Strauss III, J.F., Schuler, L.A., Rosenblum, M.F., and Tanaka, T. (1981). Cholesterol metabolism by ovarian tissue. Adv. Lipid Res., 18: 99-157. Tsang, B.K., Moon, Y.S., Simpson, C. and Amstrong, D. T. (1979). Androgen biosynthesis in human ovarian follicles; cellular source gonadotropic control, and adenosine 3',5'-monphosphate mediation. J. Clin. Endocrinol. Metab. 48: 153-158. 53 van Wagenen, G. and Simpson, M.E. (1965). Embryology of the Ovary and Testis: Homo sapiens and Macaca mulatta, Yale University Press, New Haven. Walton, E.A. and Armstrong, D.T. (1981). Ovarian function and early embryo development in immature rats given superovulatory dose of PMSG, later neutralized by antiserum. Biol. Reprod., 25: 271-280. Walton, E.A., Evans, G., and Armstrong, D.T. (1983). Ovulation response and fertilization failure in immature rats induced to superovulate. J. Reprod. Fert, 67: 91-96. Welsh, T.H., Jr., Jones, P.B.C., Ruiz de Galarreta, C.M., Fanjul, L.F., and Hsueh, A.J.W. (1982). Androgen regulation of progestin biosynthetic enzymes in FSH-treated granulosa cells in vitro. Steroids, 40: 691-700. Wilson, C.A., Horth, C.E., Endersby, C.A., and McDonald, P.G. (1974). Changes in plasma levels of estradiol, progesterone, and luteinizing hormone in immature rats treated with pregnant mare serum gonadotrophin. J. Endocrinol., 60: 293-304. Yun, Y.W., Ho Yuen, B., and Moon, Y.S. (1987). Effects of superovulatory doses of pregnant mare serum gonadotropin on oocyte quality and ovulatory and steroid hormone responses in rats. Gamete Res., 16: 109-120. Yun, Y.W., Ho Yuen, B., and Moon, Y.S. (1988). Effects of an antiandrogen, flutamide, on oocyte quality and embryo development in rats superovulated with pregnant mare's serum gonadotropin. Biol. Reprod., 39: 276-286. Yun, Y.W., Weick, R.F., Yu, F.H., Ho Yuen, B., and Moon, Y.S. (1989). Serum luteinizing hormone response in pregnant mare serum gonadotropin-treated rats. 45 th Annual Meeting of the American Fertility Society, San Francisco. Yun, Y.W., Yu, F.H., Ho Yuen, B., and Moon, Y.S. (1989). Effects of superovulatory doses of pregnant mare serum gonadotropin on follicular steroid contents and oocyte maturation in rats. Gamete Res., 23:1-10. 54 Zeleznik, A.J., Hillier, S.G., and Ross, G.T. (1979). Follicle stimulating hormone-induced follicular development: examination of the role of androgens. Biol. Reprod., 21: 673-681. 

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