Oocyte maturation.

INTERNATIONAL REVIEW OF CYTOLOGY. VOL . 57

Oocyte Maturation YOSHIOMASUI AND

HUGHJ . CLARKE Department of Zoology. University of Toronto. Toronto. Ontario. Canada

I . Introduction

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A . Concept of Maturation . . . . . . . . . . . . . . . B . Maturation and Ovulation . . . . . . . . . . . . . . C . Maturation and Fertilization . . . . . . . . . . . . . Hormonal Control of Maturation . . . . . . . . . . . . A . Gonadotropins . . . . . . . . . . . . . . . . . . B . Amphibian Oocyte Maturation . . . . . . . . . . . . C . Starfish Oocyte Maturation . . . . . . . . . . . . . D. Fish Oocyte Maturation . . . . . . . . . . . . . . E . Mammalian Oocyte Maturation . . . . . . . . . . . F . Oocyte Maturation in Other Animals . . . . . . . . . G . The Role of Follicles in Oocyte Maturation . . . . . . . Progression of Maturation . . . . . . . . . . . . . . . A . Chronology . . . . . . . . . . . . . . . . . . . B . Morphological Changes . . . . . . . . . . . . . . C . Biochemical Changes . . . . . . . . . . . . . . . Initiation of Oocyte Maturation . . . . . . . . . . . . . A . Maturation-Inducing Substance . . . . . . . . . . . . B . The Role of Ca Ions . . . . . . . . . . . . . . . C . Changes in Electrophysiological Properties . . . . . . . Cytoplasmic Control of Oocyte Maturation . . . . . . . . . A . Maturation-Promoting Factor . . . . . . . . . . . . B . Phosphorylation of Proteins . . . . . . . . . . . . . C . Arrest of Meiotic Division . . . . . . . . . . . . . Nucleocytoplasmic Interaction during Oocyte Maturation . . . A . Chromosome Condensation . . . . . . . . . . . . . B . Development of the Pronucleus . . . . . . . . . . . C . Development of Motile Systems . . . . . . . . . . . Control of Meiosis and Mitosis2oncluding Remarks . . . . A . RoleofCa . . . . . . . . . . . . . . . . . . . B . RoleofcAMP . . . . . . . . . . . . . . . . . . C . Phosphorylation of Cellular Proteins . . . . . . . . . . D. SH Cycle . . . . . . . . . . . . . . . . . . . E . Cytoplasmic Control Factors . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . .

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185 Copyright 0 1979 by Academia press. Inc. All rights of reproduction in any form reserved. ISBN 0-12-364357-0

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I. Introduction A. CONCEPT OF MATURATION Possibly the outstanding phenomenon associated with sexual reproduction in animals is that every cell of an individual derives from a single germ cell. Accordingly, it is thought that germ cells, in contrast to somatic cells, retain developmental totipotentiality throughout the life of the individual. Nonetheless, it is also true that germ cells become as highly differentiated as somatic cells during ontogenesis. In female germ cells, this specialization begins during the very early stages of life. The cells enter meiosis at the fetal or larval stage. Before they begin to grow, meiosis proceeds to the terminal stage of the first meiotic prophase, the diplotene stage. Growing primary oocytes have an enormously enlarged nucleus called the germinal vesicle (GV). Characteristically, it contains lampbrush chromosomes which have been actively engaged in RNA synthesis. Toward the end of the growth period the loops of the lampbrush chromosomes regress after which the oocytes enter a stationary state which persists until ovulation. The duration of the stationary state is consequently dependent on the time of sexual maturity of the animal and on the period of its reproductive cycle. Oocytes in this stationary state have lost their ability to proliferate and will eventually perish if allowed to remain in the ovary. If the oocytes are to continue to live, they must emerge from the stationary state and undergo various changes. These changes occur shortly before or shortly after ovulation. In the normal process of sexual reproduction they occur as a result of maturation and fertilization. In this article, the term “maturation” is used to describe the completion of meiosis as defined by Wilson (1925, pp. 397-398), who stated that maturation is accomplished in the animal oocyte by means of two successive meiotic divisions in the course of which the oocyte buds forth two polar bodies. It represents “the ripening or final stages of the formation of the germ cells. Though often applied to the nuclear changes (meiosis) it properly includes also the cytoplasmic” (p. 1136). Maturation is interrupted by suspension of meiotic division in many species, and its resumption is triggered by insemination. Oocytes which complete maturation independently of sperm penetration must be inseminated in order to begin mitosis. Thus the change oocytes undergo following insemination is a prerequisite for them either to complete maturation or to initiate mitosis. This change, which is caused by insemination, has been designated “activation. Therefore the conversion of quiescent oocytes into active zygotes involves two major pro”

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cesses, maturation and activation. There is, however, one exception to this rule. In parthenogenesis, quiescent oocytes are converted into mitotically active cells without the aid of insemination and often even without two successive meiotic divisions (Tyler, 1941; Graham, 1974). However, we can assume that these oocytes have undergone the same change in physiology that takes place during the maturation and activation of normal oocytes. Although we follow Wilson’s definition of “maturation” throughout this article, we should comment on the usage of this word to avoid confusion in the following discussion. The term “maturation” has been used in various ways by different researchers. Usually, invertebrate zoologists have considered that “maturation” encompasses the entire process of oogenesis (Highman and Hill, 1977). In order to limit the implications of the word, Schuetz (1969) proposed the term “meiotic maturation” to refer to the meiotic process following the release of the oocyte from prophase arrest. In many species, however, meiosis is again arrested at metaphase of the first or second meiotic division (metaphase I or I1 arrest), at which time the oocytes are fully fertilizable. Consequently, many investigators have referred to these oocytes as “mature oocytes, ” implying that they have completed maturation. This usage of the word may be misleading when the progression of oocyte maturation is compared in different species, since oocytes become fertilizable at different stages of maturation in different species. In fact, Wilson (1925) recognized this difficulty, stating that the maturity of the oocyte required for its fertile union with the sperm should not be confused with (the) consequences of maturation (p. 404). Recently the term “prematuration” has been introduced to refer to the process of maturation by which oocytes reach a certain intermediate stage of maturation and become fertilizable (see Schroeder and Hermans, 1975, p. 108). In past years, the problems of oocyte maturation have been reviewed by several investigators from numerous points of view. Early studies of oocyte maturation were reviewed by Schuetz (1969) and Smith and Ecker (1970a). Biochemical aspects of the maturation of the amphibian oocyte have been discussed by Smith (1975) and by Wasserman and Smith (1978a). Reviews by Redshaw (1972), Schuetz (1974), and Baulieu et al. (1978) of amphibian oocyte maturation and those of Channing and Tsafriri (1977) and Tsafriri (1978) concerning mammalian oocyte maturation are particularly relevant to the problems of endocrinological control. Oocyte maturation in fish was reviewed by Jalabert (1976) and Wallace and Selman (1978), and described in starfish by Kanatani (1973, 1975, 1978). Descriptionsof oocyte maturation in marine invertebratesare also found in the books edited by Giese and Pearse (1975). Recently an extensive review of various aspects of oocyte maturation in different species was published in the USSR (Dettlaff, 1977). This article emphasizes cellular and comparative aspects of oocyte maturation.

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B. MATURATION AND OVULATION Various temporal relationships between ovulation and maturation have been observed in different species. In some molluscans, such as clams (Spisula), echiuroids (Urechis), and annelids (Nereis), ovulation occurs before the initiation of maturation. Conversely, in sea urchins, oocytes are not ovulated until after they have completed maturation. However, in many other species the initiation of maturation and ovulation occurs almost simultaneously during normal reproductive periods. In such cases, one might well speculate that ovulation and maturation are closely linked by common systemic factors which may correlate the physiology of the ovary and the oocytes, or that maturation and ovulation are causally linked in some manner. However, it has been shown that maturation and ovulation can occur independently. In mammals, fully grown oocytes often show signs of initiating maturation in the ovary; this is followed, however, not by ovulation, but by atretic degeneration (Baker, 1972). Furthermore, it has been demonstrated (in the LT strain of mice) that oocytes can complete maturation within their follicles and spontaneously develop into embryos (Eppig et ul., 1977). These results indicate that maturation can occur without ovulation. As we discuss later, mammalian oocytes can be induced by gonadotropins to mature in follicles cultured in vitro, but in no case has it been reported that the oocytes have been ovulated. Conversely, Ryan and Grant (1940) reported that follicles of Rana pipiens cultured in Ringer’s solution containing a pituitary suspension ovulated oocytes which showed no signs of maturation. Similar results were reported by Subtelny et al. (1968). And recently, Mom11 and Bloch (1977) found that an antiovulatory drug, ethynylestradiol, blocked maturation but not ovulation when it was applied together with progesterone to isolated follicles of R. pipiens. In the fish Salmo, maturation without ovulation can be induced in cultured follicles by various steroid hormones, but when follicles are exposed to coelomic fluid collected from females which have previously been induced to ovulate by gonadotropins, these follicles ovulate oocytes showing no signs of maturation (Jalabert et al., 1972). Ovulation of immature oocytes can also be induced by exposing follicles to prostaglandin F,, (Jalabert, 1976). Dissociation of ovulation from oocyte maturation has also been reported in starfish. In Asterias, ovarian extracts prepared out of the spawning season were found to contain steroid glycosides (asterosaponin A and B) which suppressed ovulation (Ikegami, 1976). Ikegami also reported that asterosaponin A and B suppressed ovulation but not maturation of ovarian oocytes treated with 1-methyladenine(1-MA) when both chemicals were applied to an ovary obtained during the spawning season. However, ovaries exposed to Ca-free seawater for a short time ovulate immature oocytes upon their return to normal seawater. All these observations provide clear evidence that the processes of ovulation

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and maturation are not causally connected in a wide variety of species, although under normal circumstances these two events are regulated by common systemic factors such as gonadotropins. C. MATURATION AND FERTILIZATION The timing of sperm entry or insemination in relation to the progression of meiotic maturation under normal conditions is characteristic for any given species. Animals may be classified into four groups (Rothschild, 1956) according to the stage of maturation at which insemination normally occurs. In class I, which is represented by Echiuroidea and Platyhelminthes, oocytes are inseminated before maturation begins. For these oocytes, activation by the sperm triggers maturation. Class I1 includes animals in which oocyte maturation proceeds to metaphase I before the sperm enters the oocyte. Insects and ascidians are members of this class. The animals belonging to class I11 are vertebrates whose oocytes are inseminated at metaphase 11. In the last-mentioned two groups (classes I1 and 111), activation causes the resumption of meiotic progression which had been arrested at metaphase. Animals belonging to class IV include sea urchins and coelenterates. Their oocytes complete maturation before sperm entry occurs. Here, activation triggers the initiation of mitosis by the zygotes. Finally, it has been noted that oocytes of some animals, such as starfish, are inseminated at any stage of maturation after GV breakdown (GVBD) (Rothschild, 1956; Stevens, 1970). This classification of animals appears to have no correlation with their phylogenetic order. For instance, Ascaris (nematode), Spisula (mollusc), and Myzostoma (annelid) all belong to class I, and ascidians and insects to class 11. At the same time, two closely related animals, the polychaetes Nereis and Chaetopterus are classified in classes I and 11, respectively. Among the mammals, canines are known to belong to class I, while the others are members of class 111. A most intriguing finding is that two species of the same genus, Arenicolu crisrata (Okada, 1941) and A. marina (Howie, 1963) belong to classes I and 11, respectively. Changes in the fertilizability of oocytes during the course of maturation have been studied using artificial insemination in mice (Iwamatsu and Chang, 1971), pigs (Leman and Dziuk, 1971; Motlik and Fulka, 1974), dogs (Mahi and Yanagimachi, 1976), rabbits (Overstreet and Bedford, 1974), hamsters (Usui and Yanagimachi, 1976), sea urchins (Franklin, 1965; Longo, 1978), frogs (R. pipiens) (Elinson, 1977), and toads (Bufo bufo) (Katagiri, 1974). All these investigations have indicated that sperm can penetrate oocytes at any stage of maturation. However, the incidence of polyspermy is higher in oocytes inseminated at earlier stages of maturation than in those inseminated at the stage at

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which fertilization normally takes place. But no significant correlation has been demonstrated between the number of sperm per oocyte and the stage of maturation of the oocyte. The consonant results obtained by these investigators suggest that the development of the mechanism for blocking polyspermy concomitant with oocyte maturation is a general phenomenon. Niwa and Chang (1975), working on rats, found that polyspermy did not occur at any stage of maturation but pointed out that this might be due to the relatively low concentration of sperm suspension used in their experiment. Mahi and Yanagimachi (1976) also found no evidence of polyspermy, using canine oocytes. Since in dogs the sperm normally enters the oocyte before the onset of maturation, a blockage of polyspermy would not be expected to require maturation. The mechanism underlying developmentof the polyspermy blockage in maturing oocytes is not fully understood at present. However, an increasing capacity of oocytes undergoing maturation to support formation of the fertilization membrane or the zona reaction upon insemination has been generally found to play an important role in polyspermy blockage. In mice, Iwamatsu and Chang (1971) observed only a weak zona reaction in oocytes which had been penetrated by sperm at early stages of maturation, suggesting that the lack of a zona reaction was responsible for polyspermy. Usui and Yanagimachi (1976) in hamsters, and Soupart and Strong (1974) in humans, demonstrated that zona-free oocytes underwent polyspermy regardless of their stage of maturation. The ability of starfish oocytes to form the fertilization membrane upon insemination develops as maturation progresses. Hirai et al. (1971) and Cayer et al. (1975) determined that in Asterias pectinifera this ability appeared at the time of GVBD. Although Schuetz (1975b) in Asteriasforbesii, and Lee et al. (1975) in Pisaster giganteus, found that elevation of the vitelline membrane could be induced in oocytes with an intact GV by insemination, Schuetz (1975b) also demonstrated that these oocytes could not develop normally after the induction of maturation by 1-MA. Possibly oocytes with an intact GV failed to block polyspermy. Rosenberg et al. (1977) have noted that oocytes of Pisaster, exposed to 1-MA, undergo structural changes in the surface of the vitelline membrane as maturation progresses, changes which they postulate to be responsible for development of the blockage of polyspermy. Thus it is likely that development of the polyspermy-blockingmechanism during oocyte maturation is associated with development of the ability of the oocyte to give rise to a genuine fertilization membrane upon insemination. The change in the vitelline membrane following insemination has been regarded as a result of oocyte activation by sperm entry, possibly caused by enzymes released from the cytoplasm (Carroll and Epel, 1975a). Ample evidence has accumulated to indicate that in nonmammalian species (Vacquier et al., 1972; Schuel et al., 1973; Carroll and Epel, 1975b) and in the rabbit (Flechon et al., 1975) cortical granule breakdown (CGBD)plays a major role in changing the properties of the vitelline membrane following insemination. Therefore it seems

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reasonable to ascribe the failure of precociously inseminated oocytes to develop effective polyspermy blockage to the absence of a cortical response to sperm entry. There is evidence that, at least in sea urchins, a close association of the cortical granules (CGs) with the plasma membrane is important for CGBD to occur (Millonig, 1969; Longo and Anderson, 1970). Longo (1978) found that the CGs in sea urchin oocytes did not move to the plasma membrane until maturation had reached a certain stage and suggested that this was at least partly responsible for the absence of CGBD in inseminated immature oocytes. Experiments with frogs by Btlanger and Schuetz (1975) and by Cloud and Schuetz (1977) showed that oocytes which had not matured to metaphase I1 did not activate after insemination or pricking. Yet the divalent cation ionophore A23187 can induce activation reactions such as CGBD and vitelline membrane elevation in these oocytes. It is known that this ionophore is a ubiquitous parthenogenetic agent (Steinhardt et al., 1974), and that it induces Ca release from the egg (Steinhardt et al., 1977), thereby causing CGBD (Vacquier, 1975). Therefore it seems likely that development by the oocyte of the ability to block polyspermy also depends on its increasing capacity to release Ca in response to sperm entry. The variability among species in the timing of sperm entry into the oocyte relative to the meiotic progression of the nucleus may indicate that these cytoplasmic changes, occurring during the course of maturation, do so rather independently of the nuclear changes. Iwamatsu (1966, 1971) showed that this was the case in the fish Oryzias lutipes (medaka). He demonstrated that oocytes could become fertilizable and develop into haploid embryos even when the GV was kept intact in the yolk mass by centrifugal displacement. Amphibian oocytes from which the GV has been removed also develop the ability to undergo parthenogenetic activation when they are treated with progesterone (Smith and Ecker, 1969; Skoblina, 1969). Skoblina (1974) and Katagiri and Moriya (1976) further demonstrated that sperm could enter and activate these enucleated oocytes. Experiments on starfish oocytes by Hirai et ul. (1971) also indicated that enucleated oocytes became activatable upon insemination 20-30 minutes after I-MA treatment. All the results cited above strongly suggest that cytoplasmic maturation can occur independently of the presence of the nucleus (the GV). 11. Hormonal Control of Maturation A. GONALIOTROPINS

In most animals, other than those whose oocytes are induced to mature by insemination, the initiation of oocyte maturation is dependent on ovarian function. Generally, only oocytes which are fully grown mature in response to ovari-

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an stimulus. The systemic factors controlling oocyte maturation are gonadotropins. It is well known that the administration of gonadotropins to female animals having fully grown oocytes in their ovaries causes oocyte maturation, with concomitant ovulation. In vertebrates, gonadotropins are secreted by the pituitary gland (hypophysis). Two different polypeptides, luteinizing hormone (LH) and follicle-stimulating hormone (FSH), can be distinguished in mammals (Papkoff et al., 1973), birds (Farmer et al., 1975), reptiles (Licht and Papkoff, 1974a), and amphibians (Licht and Papkoff, 1974b), but only one gonadotropic hormone (GTH) has been found in fish (Fontaine and Gerard, 1963; Donaldson et al., 1972). The gonadotropin(s) found in invertebrates has not been chemically characterized, except for that of starfish. In starfish, a peptide hormone analogous to vertebrate gonadotropin, known as gamete-shedding substance (GSS), is secreted by the supporting cells of the radial nerves (Kanatani et al., 1971). Gonadotropins act directly on the ovary. Heilbrunn et al. (1939), using R. pipiens, demonstrated for the first time that oocyte maturation and ovulation could be induced in ovarian fragments incubated in Ringer’s solution containing pituitary extracts. This method has been widely used since then to study the effects of gonadotropins on oocyte maturation and ovulation in numerous species. It will become apparent in the following sections that, because the role of gonadotropins in oocyte maturation has been studied in many different species by many different investigators, conflicting results have been obtained, making it difficult to define a generalized mode of gonadotropin action. Nevertheless, information gained from studies using frogs and starfish, which are probably the most thoroughly analyzed animals with regard to this problem, points to a rather simple common principle, namely, that gonadotropins primarily act on the follicle (granulosa) cells to induce maturation of the oocytes in these follicles. B. AMPHIBIAN OOCYTEMATURATION

The experiment by Heilbrunn and his associates (1939) showed that gonadotropins acted on the ovarian follicle to induce maturation in R. pipiens. However, if follicles are incubated with progesterone, ovulation (Wright, 1961) and oocyte maturation (Schuetz, 1967a) are induced. In R. pipiens, it was found that gonadotropin had no effect on the oocyte when the follicle cells were completely removed following treatment with Ca-free medium (Masui, 1967) or pronase (Smith et al., 1968). However, these follicle-free oocytes were shown to undergo maturation in response to progesterone (Masui, 1967; Schuetz, 1967b; Smith et al., 1968) or, when they were incubated with follicle cells, to gonadotropin (Masui, 1967). Hence it has been hypothesized by Masui (1967) and by

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Smith et al. (1968) that gonadotropins induce the follicle cells to secrete a hormone, possibly progesterone-like, which in turn acts on the oocyte to initiate! maturation. Although progesterone is the most potent steroid inducer of oocyte maturation, other hormones such as deoxycorticosterone (DOC) and testosterone, but not estradiol or its derivatives, are also effective in R. pipiens (Schuetz, 1967a; Smith et al., 1968) and in Xenopus laevis (Jacobelli el al., 1974; Schorderet-Slatkine,-1972). The above hypothesis has been substantiated by recent work. Fortune ef al. (1975) and Thibier-Fouchet et al. (1976) demonstrated in Xenopus that both Xenopus gonadotropin and human chorionic gonadotropin (HCG) induced follicles to convert pregnenolone into progesterone and induced oocytes to mature. However, isolated oocytes did not carry out this steroid conversion (ThibierFouchet et al., 1976). Furthermore, drugs which suppress this steriod conversion, such as cyanoketone and eliptin, inhibit gonadotrophicinduction of maturation in follicleenclosed oocytes of R. pipiens (Wright, 1971; Snyder and Schuetz, 1973) and X. Zaevis (Fortune et al., 1975). But these inhibitors do not affect progesterone-induced oocyte maturation. Information concerning hormonal control of oocyte maturation in urodeles is rather scarce. In these animals, complete control of ovulation and oocyte maturation in vitro has not yet been achieved. Methods which induce oocyte maturation in anurans are not successful when applied to urodeles. Two examples should suffice. Progesterone is an effective inducer of maturation in Notophthalmus viridescens only when isolated oocytes are primed with gonadotropin (Pilone and Humphries, 1975). And only oocytes obtained from Pleurodeles waltlii collected during the breeding season are responsive to progesterone treatment (Brachet, 1974; Ozon et aZ., 1975). Perhaps the inefficiency of progesterone in these urodeles can be explained by a consideration of their natural breeding habits, which are different from those of anurans. In many urodele species, only a few eggs, which have completed growth, are spawned each day; those remaining in the ovary quite possibly have not yet grown sufficiently to be able to initiate the maturation process. C . STARFISH OOCYTEMATURATION

Studies by Chaet (1966) and by Kanatani and Ohguri (1966) demonstrated that isolated fragments of the starfish ovary ovulated upon transfer to seawater containing radial nerve factor (RNF or GSS) and that the oocytes underwent maturation concomitant with ovulation. Further research (Schuetz and Biggers, 1967; Kanatani and Shirai, 1967) indicated that oocytes freed from follicle cells did not respond to GSS. At the same time, these investigators also showed that isolated oocytes matured when they were exposed to seawater in whichisolated follicles

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had been incubated with GSS. Thus it has become clear that GSS, or RNF, acts by inducing follicles to secrete a maturation-inducing substance (MIS). Kanatani et al. (1969) purified MIS and identified it as 1-MA. At a concentration of lo-' M, this substance can induce oocyte maturation in all species of starfish so far examined (see Kanatani, 1973). 1-MA is also produced in the ovary of echinoids other than starfish, but an effect on the oocytes of these species has not been confirmed (Kanatani, 1975). Investigation into the mechanism of 1-MA production in the starfish ovary has disclosed that this substance is a derivative of a purine compound which contains a methyl group at the N- 1 position. GSS stimulatesmethylationof this compound,utilizingS-adenosylmethionine (Shirai et al., 1972), to form 1-methyladenosinemonophosphate, which is then hydrolyzed to give rise to 1-MA through the formation of 1-MA riboside (Schuetz, 1970; Shirai and Kanatani, 1972).

D. FISHOOCYTE MATURATION Research on oocyte maturation in fish has produced complicated results with respect to the role of gonadotropins. This may be due primarily to the fact that, excepting studies on the sturgeon and the trout, few experiments have been performed using follicle-free oocytes. Sturgeon oocyte maturation has been investigated by Dettlaff and her colleagues (Dettlaff and Skoblina, 1969). Fully grown sturgeon oocytes are invested with a jelly coat secreted by the follicles and can easily be separated from them without damaging the oocytes. Progesterone treatment induces maturation both in follicle-enclosed and follicle-free oocytes, while gonadotropin is effective only in follicle-enclosed oocytes. Similarly, trout oocyte maturation is induced in isolated follicles by steroids and by gonadotropin (Jacobelli et al., 1974). And, as in the sturgeon, trout gonadotropin is effective only in follicle-enclosed oocytes. Not many steroid hormones affect follicle-free oocytes; 17a,20fl-dihydroxyprogesteroneand 20phydroxyprogesterone are the only potent steroid inducers of maturation of isolated oocytes (Fostier et al., 1973; Jalabert, 1976). Some steroids are effective to a certain extent when applied to follicle-enclosedoocytes of trout, goldfish, or pike. Consequently, it has been suggested that these steroids are metabolized to 17a,20P-di- or 20P-hydroxyprogesterone (Jalabert, 1976). In other fish species, the effects of hormones on oocytes have not been conclusively resolved. In the medaka, cortisol appears to be a more potent inducer of maturation than progesterone when intact follicles are cultured until maturation in a medium containing the steroid (Hirose, 1972). But progesterone is the most potent inducer when the follicles are only briefly exposed to steroids (Iwamatsu, 1974). Recently, Hirose (1976) reported that neither gonadotropin nor corticosteroids were effective in inducing maturation of medaka oocytes

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cultured in vitro after removal of the follicular tissue by ethylenediamine tetraacetic acid (EDTA) or trypsin treatment. Pointing out that cortisol can be synthesized by the ovary in some fish (Colombo et al., 1973;Hirose et al., 1975),he concluded that the maturation process in the medaka, stimulated by the pituitary-ovarian axis, may be mediated by a second substance produced in the ovary during corticosteroid metabolism. Effects of mammalian and fish gonadotropins and various steroid hormones on fish ovarian follicles were investigated by Goetz and Bergman (1978a,b) using yellow perch (Perca flavescens), walleye (Stizostedion vitreum), and brook trout (Salivelinus fontinalis). These investigators found that most of the hormones tested were effective in inducing maturation of follicle-enclosed oocytes, and also suggested that corticosteroids facilitated the efficacy of gonadotropins. An enhancing effect of corticosteroids on the efficacy of gonadotropin and I7~~,20/3-dihydroxyprogesteroneaction in the induction of maturation of follicle-enclosed trout oocytes has been noted by Jalabert (1976).Dettlaff and Davydova (1974)observed that triiodothyronine increased the gonadotropin sensitivity of follicles isolated from cold-stored sturgeon. These results strongly suggest that hormones other than gonadal steroids serve to sensitize follicles to gonadotropins. Recently, Wallace and Selman (1978) succeeded in inducing maturation of follicle-enclosed oocytes of the marine fish Fundulus hereroclitus in vitro using HCG, DOC, and progesterone. These workers consider that the physiological condition of the cultured follicle is an important factor determining its response to hormones. Although the studies discussed above appear to indicate that fish gonadotropin stimulates the ovarian follicle to produce progesterone-like steroids which in turn trigger oocyte maturation, another route of hormonal action has been suggested in catfish (Sundararaj and Anand, 1972). Isolated follicles of this fish respond neither to gonadotropins nor to gonadal steroids, although in vivo administration of LH effectively induces oocyte maturation and ovulation. The effect of the gonadotropin is, however, attenuated by the drug Metopirone which blocks corticosteroid synthesis in the interrenal gland; this inhibition can be overcome by hydrocortisone (HC) or DOC administration. These corticosteroids are also effective in inducing the maturation of follicle-enclosed oocytes in vitro (Goswami and Sundararaj, 1971), while gonadotropin is effective only when the follicles are also incubated with interrenal tissue (Sundararaj and Goswami, 1974). And minced interrenal tissue is reported to increase corticosteroid synthesis, primarily that of DOC, in the presence of gonadotropic hormone (Sundararaj and Goswami, 1969).Finally it was recently reported that the ovary of the catfish was unable to synthesize DOC (Unger et al., 1977). Taken together, these results have been interpreted as indicating that gonadotropic hormone stimulates the interrenal gland to secrete corticosteroids which induce oocyte maturation (Sundararaj and Goswami, 1977).

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However, this interpretation should not be considered definitive. The results obtained with the catfish may be interpreted in a different way. The failure of gonadotropin to induce oocyte maturation in isolated follicles may be attributable to a low sensitivity of the tissue per se toward the hormone; corticosteroids may act to increase the sensitivity of the follicular tissue. Second, it should be noted that the action of steroids in the induction of oocyte maturation generally is not as specific as that of most hormonal action. For example, although DOC is not produced by the amphibian ovarian follicle, it has been found to be as potent an inducer of oocyte maturation as progesterone (Smith et al., 1968; SchorderetSlatkine, 1972). All in all, we feel it would be premature to consider the female of this species an exception to the pituitary-gonadalaxis principle of reproductive control which has been so often demonstrated-in fact, even in the male catfish (Sundararaj and Nayar, 1967).

OOCYTEMATURATION E. MAMMALIAN Maturation of mammalian oocytes enclosed in follicles is induced under the influence of LH, both in vivo and in vitro. As shown by Ayalon et al. (1972) in rats, follicle-enclosed oocytes remain at the dictyate (diplotene) stage if the follicles are explanted from the female before the preovulatory LH surge and cultured in a hormone-free medium, whereas oocytes undergo maturation if the follicles are isolated after the LH surge. Later, Hillensjo et al. (1974) confirmed these results using prepubertal rats injected with pregnant mare serum (PMS). In vitro experiments by Baker and Neal (1972) with mice, Tsafriri et al. (1972) with rats, Hay and Moor (1973) with sheep, Thibault and Gerard (1973) with rabbits, Thibault et al. (1975a) with monkeys and calves, and Gwatkin and Andersen (1976) with hamsters have all indicated that addition of LH to the medium in which follicles are cultured causes the oocytes within these follicles to initiate maturation. However, the ability of LH to induce follicle-enclosedoocytes to mature does not appear to be due to its specific steroidogenic action on the follicles. Experiments using rats (Tsafriri et al., 1972) and rabbits (Thibault and Gerard, 1973) have demonstrated that FSH is also capable of inducing maturation of follicleenclosed oocytes cultured in vitro. The possibility that the effect of FSH might be caused by contaminating LH in the hormone preparation can be ruled out, since antibody made against the p chain of LH does not abolish the effect of FSH, but only that of LH (Tsafriri et al., 1972; Hillensjo et al., 1976). It is also known that the effects of FSH and LH on steroidogenesis in the granulosa cells are different (see Channing and Tsafriri, 1977). Therefore the effect of gonadotropin on the induction of maturation in follicleenclosed oocytes does not appear to be mediated by hormone-stimulated steroidogenesis.

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Studies using rats (hfriri e l al., 1972) and cows and pigs (Foote and Thibault, 1969) have shown that the addition of steroids to follicle culture medium has no effect on oocyte maturation. Furthermore, inhibition of LHinduced steroidogenesis in rat follicles by cyanoketone or aminoglutethiimide does not prevent LH induction of oocyte maturation in the follicles (Lieberman er al., 1976). These results suggest that the maturation of mammalian oocytes is not necessarily induced by steroid production following follicular stimulation by gonadotropins. The initiation of oocyte maturation appears rather to be dependent on the general physiological condition of the follicles. In the monkey, while LH normally induces only oocytes enclosed in fully grown follicles to mature, it also induces maturation of oocytes in small follicles if the follicles have undergone atresia (Thibault et al., 1975b). It is known that oocytes in atretic follicles even spontaneously initiate maturation (Foote and Thibault, 1969). Recently, Moor and Trounson (1977), using sheep ovaries, showed the LH induced oocyte maturation when it was applied to nonatretic large follicles or to atretic small follicles, but that it had no effect on atretic large follicles or on nonatretic small follicles. They also found that oocytes in atretic follicles, both large and small, did not undergo maturation spontaneously when the follicles were incubated under hyperbaric conditions. It was further shown that oocytes within atretic follicles which had been cultured in the presence of FSH and 17P-estradiol possessed the same capability as those in nonatretic follicles, in spite of follicular deterioration, to mature and develop normally following insemination in the oviduct of recipient animals. These results indicate that LH-induced oocyte maturation in atretic small follicles is not due to deterioration of the oocytes, but primarily to deterioration of the follicles. Thus it can be inferred that induction of oocyte maturation within the follicle is brought about by perturbation of the follicular physiology. One of the first indications of atresia is degeneration of the cumulus cells (Hay et al., 1976). Perhaps significantly, the effect of a gonadotropin on follicles first appears in the cumulus cells. It induces both the dispersion of (Thibault et al., 1975a) and hyaluronic acid secretion by (Hillensjo et al., 1976) the cumulus cells. Thus it may be that gonadotropins alter the physiology of the cumulus cells, producing a change in their relationship with the oocytes, which in turn stimulates the latter to initiate maturation. In mammals, it is now well known that oocytes removed from the follicular environment always tend to undergo maturation. This phenomenon was first demonstrated in 1935 by Pincus and Enzmann, using rabbit oocytes, and later confirmed by Chang (1955). Edwards (1965) pointed out that the maturation of oocytes following isolation from their follicular environment was almost universal among mammals-found in the mouse, sheep, cow, pig, monkey, and human. The following species have been added to this list since then: rat (Magnus-

198


son et al., 1977), guinea pig (Jagiello, 1969; Yanagimachi, 1974), hamster (Haidri et al., 1971), and dog (Mahi and Yanagimachi, 1976). In most species, maturation of follicle-free oocytes can take place in a simple medium such as Krebs-Ringer solution containing pyruvate, kept in a 5 % COP95% air mixture (Biggers et al., 1967; Donahue, 1968). Oocytes of some species such as hamsters (Gwatkin and Haidri, 1973, 1974) and sheep (Hay and Moor, 1973) require a lower 0, pressure or addition of amino acids to the culture medium. However, neither hormones nor tissue factors are necessary. These findings might lead one to speculate that the intrafollicular milieu plays an important role in maintenance of the physiological stability of oocytes at the dictyate stage. In this respect, any gonadotropin-induced alteration in the physiological activity of the cumulus cells might cause an interruption of the stabilizing function of the follicles, resulting in maturation of the oocytes. Such physiological perturbation of the follicles might be brought about by gonadotropic activity stimulating follicular activities other than steroidogenesis. For instance, LH stimulates prostaglandin (PG) production in the follicle (Chasalov and Pharriss, 1972; Bauminger et al., 1975), and application of PGE, to cultured follicles induces oocyte maturation (Tsafriri et al., 1972). However, indomethacin, which suppresses PGE, production and also inhibits ovulation in rats (Armstrong and Zamecnik, 1975), fails to suppress LH-induced oocyte maturation (Tsafriri et al., 1972). Thus it is clear that the action of LH on the induction of maturation in follicle-enclosed oocytes is not mediated by PG production, but rather that both act simultaneously and independently to cause physiological changes in the follicles. Both LH and PG are known to stimulate adenyl cyclase to increase the level of cyclic adenosine monophosphate (CAMP) in the follicle (Lamprecht et al., 1973). Reasoning that this might provide a clue as to their function, several groups have tested the effects of cAMP on rat oocytes. Tsafriri et al. (1972) reported that, while both cAMP and its dibutyryl derivative (dbcAMP) exerted no inductive effect on oocyte maturation when applied extrafollicularly, dbcAMP had a positive effect when injected into the follicle. The possibility that the injection procedure itself may have induced the oocytes to mature was ruled out by control experiments showing that injected 5'-AMP had no effect on the oocytes. Moreover, cholera toxin, known to stimulate cAMP production in various types of cells, also effectively induced maturation of follicle-enclosed rat oocytes (Tsafriri et al., 1972). However, derivatives of cAMP have been found to have an inhibitory effect on oocyte maturation at concentrations greater than M. Hillensjo et al. (1978) in rats, and Nekola and Smith (1975) in mice, showed that addition of dbcAMP to the medium in which follicles were cultured inhibited the inducing effect of LH on oocyte maturation. Nonetheless, the changes in the cumulus cells which occurred in the presence of dbcAMP, with or without gonadotropin, were identi-

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cal to those observed during LH-induced maturation (Hillensjo, 1977). An inhibitory effect of dbcAMP on spontaneous maturation of follicle-free oocytes has also been observed in mice (Stem and Wassarman, 1974; Cho et al., 1974) and rats (Magnusson and Hillensjo, 1977). Therefore it is highly probable that the inhibition of maturation of follicle-enclosedoocytes caused by dbcAMP is due to direct action on the oocytes, an action counteracting signals from the cumulus cells to release the oocytes from meiotic arrest. If so, it is conceivable that CAMP or its derivatives could induce maturation of follicle-enclosed oocytes if their action is localized in the cumulus cells. A key to understanding the mechanism which initiates maturation of follicular oocytes is a determination of the nature of the follicular milieu which prevents oocytes from maturing. Analyses of the intrafollicular factors responsible for the stabilization of dictyate oocytes have been carried out by several groups. The possibility that follicular factors which prevent oocytes from initiating maturation affect the physiological activity of the oocytes in general has been speculated upon by Zeilmaker et al. (1972). He points out that spontaneous maturation of rat oocytes can be prevented by hypoxia, suggesting that the reduced oxygen supply to follicular oocytes may be the factor responsible for the suspension of meiosis. However, this simple hypothesis cannot explain the complicating results obtained in other species. For example, oxygen supplied at its atmospheric partial pressure impedes the maturation of follicle-free hamster oocytes (Gwatkin and Haidri, 1974) and causes spontaneous maturation and atretic changes in follicleenclosed sheep oocytes (Hay and Moor, 1973). The existence of a specific follicular factor which prevents oocytes from spontaneously maturing was suggested by Chang (1955) based on his experiments in which isolated rabbit oocytes were cultured in a medium containing follicular fluid. Foote and Thibault (1969) observed that, while follicle-free porcine oocytes spontaneously underwent maturation, oocytes cultured with follicle wall hemispheres failed to do so. Experiments by Tsafriri and Channing (1975a,b) demonstrated that a graded addition of cumulus cells to a culture of follicle-free oocytes inhibited their maturation in a dose-dependent manner but that the inhibition could not be removed by LH. However, Tsafriri et al. (1977), carrying out a similar experiment, found that LH was indeed capable of reversing the inhibitory effect of cumulus cells. Using porcine oocytes, Tsafriri and Channing (1975a,b) determined that isolated oocytes, cultured in a medium containing 50% porcine follicular fluid, underwent maturation with a frequency about half that of oocytes cultured without follicular fluid. The inhibitory effect of follicular fluid on the spontaneous maturation of follicle-free oocytes has been reported to be non-species-specific. Oocytes of the mouse (Channing and Tsafriri, 1977), rat Vsafriri et a/., 1977), and sheep and cow (Jagiello et al., 1977) cultured in media containing porcine follicular fluid were found to undergo maturation with a low frequency compared to those

200


cultured without follicular fluid. In addition, the frequency of maturation of hamster follicle-free oocytes cultured in medium containing bovine follicular fluid was found to be decreased to the same extent as those cultured in medium containing hamster follicular fluid (Gwatkin and Andersen, 1976). The effects of bovine follicular fluid on hamster oocytes (Gwatkin and Andersen, 1976) and of porcine follicular fluid on porcine flsafriri et al., 1976b) and rat (Tsafriri et al., 1977) oocytes have recently been examined under a variety of physical and chemical conditions. These experiments revealed several facts concerning the nature of the maturation inhibitor. Its ability to block oocyte maturation is dose-dependent and is abolished by the addition of LH but not by the addition of dbcAMP. It is associated with a substance whose molecular weight lies between lo00 and 2000 daltons. Finally, it is trypsin-sensitive but resistant to heating at 60°C for 20 minutes. Yet, up to now, no tests have been carried out on the physiological activity of oocytes exposed to the follicular fluid inhibitor. With regard to the reversibility of the inhibitory effect of follicular fluid, some preliminary work by Stone et al. (1978) revealed that, among rat oocytes exposed to porcine follicular fluid for 24 hours, a certain proportion was capable of resuming maturation, but of those exposed to the inhibitory influence for 28 hours or more none resumed maturation. All in all, the evidence cited above indicates that oocyte maturation is caused by an abrupt cessation of the follicular function acting to stabilize the oocyte physiology. In view of the facts that mammalian oocytes spontaneously undergo maturation when they are freed from surrounding follicle cells, and that the tight junctions between oocytes and cumulus cells are lost when oocytes begin to mature following gonadotropin action (Szollosi, 1978; Gilula et al., 1978), it appears likely that maturation of follicle-enclosed oocytes also results from the loss of communication between oocytes and cumulus cells, which may follow an alteration in physiological activities of follicle cells by the gonadotropin action. From this point of view the recent finding by Tsafriri and Bar-Ami (1978) that maturation of follicleenclosed rat oocytes can be induced by Ca-deficient media without hormones may be interpreted in such a way that Ca deficiency brings about oocyte maturation by destabilizing the association between oocytes and cumulus cells. If so, the implication of this finding may be that no inhibitor exists in the follicular fluid to prevent oocytes from maturation or, if the inhibitor is present, its effect must be Ca-dependent. The latter possibility may be tested. However, in view of the following observations, it would be premature to deny totally the possibility of an action of gonadotropin and steroids on oocyte activity in relation to maturation. Recently, Jagiello and Ducayen (1977) demonstrated, in human, monkey, and sheep ovaries, a dense accumulation of HCG1251and LH on oocyte chromatin and in the cytoplasm, using autoradiographic and immunocytochemical techniques, and that these hormones also accumulated in follicle-free oocytes in vitro. Furthermore, it has been demonstrated in rats

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201

that there is a significant difference in the timing of GVBD in follicle-free oocytes cultured in the presence of LH and in those cultured in its absence (hpata et al., 1977). In the former, GVBD occurs less than 1 hour after isolation but not until after at least 2 hours in the latter case. The role of steroid hormones secreted by the follicle cells in promoting oocyte maturation has been studied by several workers. Baker and Neal (1972) noted the synergistic action of estrogen and gonadotropin. Hunter et al. (1976) found that pig oocytes ovulated following HCG administrationoften failed to undergo maturation if ovulation was induced when 17pestradiol levels in the follicle were low (day 17 of the estrous cycle), while those ovulated when hormone levels were high successfully completed maturation. Corroborative evidence comes from Moor and Trounson (1977) who found that the addition of 17P-estradiol to a culture of sheep follicles, in the presence of low levels of gonadotropin, increased the percentage of oocytes which could support normal embryonic development, although it did not affect the percentage which underwent maturation. McGaughey (1977), using pig oocytes, also reported that spontaneous maturation of follicle-free oocytes progressed beyond metaphase I more frequently in the presence of 170-estradiol and progesterone than in their absence. These observations seem to favor the view that follicular steroids also play a significant role in the maturation of mammalian oocytes, although the ability of gonadotropins to alter the physiological condition of the follicle is sufficient to allow the oocyte, meiotically arrested at the diplotene stage, to resume meiosis by initiating maturation.

F. OOCYTEMATURATION IN OTHER ANIMALS In many invertebrates, oocyte maturation is triggered by sperm penetration, but in some species the oocytes begin to mature following completion of their growth period, usually coincident with ovulation. The mechanism which induces oocyte maturation has not been elucidated except in the starfish (see Section KC). In many marine invertebrates, oocytes complete their growth in the coelom before beginning maturation. Oocyte maturation in polychaetes and sipunculoids begins and progresses to metaphase I while the oocytes are in the coelom. In Arenicola marina (a polychaete), it has been shown that removal of the brain (prostomiurn) 3 weeks before spawning prevents the onset of maturation, while the injection of brain homogenate into decerebrated animals causes the resumption of oocyte maturation as well as spawning (Howie, 1963, 1966). Recently Meijer and Durchon (1977) have reported that immature oocytes isolated from the coelom are induced to mature in vitro by exposure to seawater containing brain (prostomiurn) extracts in a dose-dependent manner.

202


In Pectinaria gouldii (Cistemides), Tweedell (1962, 1966) found that oocyte maturation normally occurred only during the natural spawning season; it was characterized by oocytes undergoing GVBD after they were collected in the nephromixium. However, in oocytes collected during nonspawning periods, maturation can be induced by the injection of extracts obtained from various tissues into the coelom (K. S. Tweedell, personal communication). Since tissue extract injection induces the oocytes first to move into the nephromixium, where GVBD later occurs, it may be possible that the tissue factor changes the properties of the oocytes so that they become capable of being drawn into the nephromixium. Although these observations suggest that factors residing in the nephromixium play an important role in triggering oocyte maturation, the factors may not be specific ones since, when the animals are shaken mechanically in seawater, immature oocytes are ovulated and these oocytes spontaneously undergo GVBD in seawater 12-18 minutes after ovulation (Tweedell, 1962; Austin, 1963). Similar observations by Rice (1966, 1975) have been recorded in Sipuncula oocytes. In this species, oocytes begin to mature inside the coelom during the breeding season. Oocytes collected from the coelom of animals during nonbreeding periods will undergo maturation if treated with extracts from various tissues. Although it is premature to speculate on the mechanism initiating oocyte maturation in these species, it is possible that the coelomic fluid of these animals exerts an inhibitory effect on oocytes which prevents them from maturing during nonbreeding seasons. The inhibitory activity is removed as a result of tissue secretion during the breeding season, and consequently maturation is triggered. The lack of tissue specificity shown by the maturation-inducing factor suggests that it is a substance distributed to all tissues. Recent experiments by Peaucellier (1977) indicate that this is indeed the case in the polychaete Subellaria alveoluta. When oocytes of this species are carefully isolated, so that they have no contact with the cloaca, they fail to mature in seawater. But the oocytes thus isolated mature upon exposure to cloaca1 secretion. The factor responsible has been identified as a protease; apparently proteases from any source are effective.

G . THEROLEOF FOLLICLES IN

OOCYTE

MATURATION

Maturation does not normally take place during the growth period of oocytes, with the exception of a few groups of animals. In these species, maturation begins before the oocytes are fully grown. Oocytes of the spmge Hippospongia communis undergo maturation before the period of major growth begins (Tuzet and Pavans de Caccaty, 1958), and those of the insect Drosophila reach metaphase I before their growth ceases (Mahowald, 1977).

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203

Oocytes in the growth phase rarely respond to maturation signals from the outside. Reynhout et al. (1975) reported that follicle-enclosed oocytes of Xenopus must be at least 1.2 mm in diameter to be responsive to gonadotropic or HCG stimulation. However, smaller oocytes, having diameters as small as 0.9 mm, can be induced to mature in vitro by progesterone, provided they have been removed from their follicles. Small follicleenclosed oocytes become responsive to gonadotropins after repeated progesterone administration. Similar observations were recorded by Sakum (1961, 1972, 1975) in the trout. She found that repeated gonadotropin administration caused growing small oocytes to begin maturation, although the maturation process was generally abnormal and resulted in atypical condensation and arrangement of the oocyte chromosomes. In mammals, the ability to initiate maturation appears to be restricted to oocytes enclosed in follicles of some minimum size. No oocytes removed from preantral follicles of prepubertal mice (younger than 14 days) undergo maturation (Szybek, 1972; Erickson and Sorenson, 1974). A graded increase in the tendency of oocytes to undergo maturation was found to occur with the progression of follicular growth in pigs (Tsafriri and Channing, 1975b). As pointed out by Iwamatsu and Yanagimachi (1975) in the hamster and by Sorensen and Wassarman (1976) in the mouse, the tendency of a follicle-free oocyte to begin maturation is directly related to oocyte growth. According to their observations, the frequency with which spontaneous maturation occurs increases linearly in proportion to the diameter of the oocytes, up to a certain size, 80 p m in the hamster and 68 p m in the mouse. Furthermore, maturation of the smaller oocytes is often abortive, being arrested at metaphase I. Development of the tendency toward spontaneous maturation of mouse oocytes occurs in the follicles in the absence of gonadotropic influences. According to Eppig (1977), follicles of 8-day-old mice, cultured for 1 week in hormone-free media, contain growing oocytes capable of initiating maturation upon isolation from the follicles. He also pointed out the importance of a close relationship between the cumulus cells and the oocytes for the latter to grow and mature. The spontaneous maturation of follicle-free oocytes in mammalian species may be indicative of a certain physiological instability of oocytes which have reached a certain stage of growth. Perhaps the physiological state of these oocytes is so unstable that, upon removal from the stabilizing influence of the follicles, they tend to undergo changes which result in maturation. In nonmammalian species, fully grown oocytes appear to be relatively stable, so that external stimuli are needed to initiate maturation. However, their responsiveness to such stimuli may be conditioned by some degree of intrinsic physiological instability. Thus it is possible that the increasing tendency of growing oocytes to undergo maturation is a reflection of increasing physiological instability. From studies of the gonadotropin control of oocyte maturation in various

204


forms of animals we have learned that changes in the physiological activities of the follicle cells are crucial for the initiation of oocyte maturation. However, the maturation process of amphibian, starfish, and sturgeon oocytes can take place without follicles following treatment with follicular hormones. In these animals, follicular hormones such as progesterone and 1-MA act on the oocyte in the same manner regardless of the presence or absence of the follicle; that is, both folliclefree and follicle-enclosed oocytes, once induced to mature, exhibit the same developmental capacity as naturally ovulated oocytes. In R. pipiens, both oocytes which have been ovulated in vitro from isolated follicles after gonadotropin treatment (Ryan and Grant, 1940) and those induced to mature by progesterone following defolliculation (Smith et al., 1968) develop normally if they are inseminated after passing through the oviduct of recipient females. Similar results were obtained with sturgeon oocytes (Dettlaff and Skoblina, 1969). Finally, in starfish, oocytes matured either with or without follicles cleave normally (Kishimoto and Kanatani, 1973) and develop at least to the bipinnaria stage (Guenier et al., 1978). However, oocyte maturation in mammalian species is highly dependent on the follicle. Follicle-free oocytes which have undergone spontaneous maturation are found to lack complete developmental potentiality as compared with follicleenclosed oocytes induced to mature by gonadotropin. Thibault (1972) found that follicle-free oocytes of the rabbit underwent maturation spontaneously in culture but that they did not exhibit the ability to transform sperm nuclei into pronuclei after insemination. Oocytes matured within follicles cultured in the presence of gonadotropin acquired the ability to form male pronuclei. It has also been reported, in the pig, that meiosis which takes place in folliclefree oocytes during spontaneous maturation usually results in considerable chromosomal aberration (McGaughey and Polge, 1971). Studies using rabbits (Chang, 1955) and mice (Cross and Brinster, 1970; Mukherjee, 1972) have shown that oocytes which have undergone maturation without follicles rarely develop beyond the early cleavage stages after insemination, and Van Blerkom and McGaughey (1978b) observed that only a small proportion (13%) of rabbit oocytes matured in vitro could reach the blastocyst stage. Recently Eppig (1978) using oocytes of LT/Sv hybrid mice has noted that follicle-free oocytes, which spontaneously matured in vitro and activated, can initiate cleavage parthenogenetically, but fail to develop further. To attain the potential to develop to blastocysts the oocytes must mature within follicles for 8-9 hours after gonadotropin administration. In sheep, Moor and Trounson (1977) demonstrated that oocytes cultured without follicles failed to develop beyond the blastocyst stage, whereas those matured within follicles, which were cultured in the presence of LH, FSH, and 17P-estradio1, developed normally. When the oocytes of this latter group were fertilized following transfer into recipient females, 73% developed to the blastocyst stage and 63% into newborn lambs. These observations

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205

clearly indicate that an indispensable role is played by the follicles in the process of normal oocyte maturation in mammals. 111. Progression of Maturation A. CHRONOLOGY

Traditionally the process of oocyte maturation has been described referring to changes in chromosome morphology during meiotic progression. Before maturation starts, the oocyte contains a GV with a few large or many small nucleoli (Fig. 1). Chromosomes in the GV are extended and widely dispersed (Fig. 2). With the initiation of maturation, the GV breaks down and the chromosomes begin to condense. As maturation progresses, the chromosomes further contract and become arranged in the middle of a spindle in pairs-metaphase I (Fig. 3). Separation of the paired homologous chromosomes (Fig. 4) is followed by formation of the first polar body (Fig. 5 ) . Then the chromosomes remaning in the oocyte become aligned in the metaphase plate-metaphase I1 (Fig. 6). When the second meiotic division begins, the daughter chromosomes separate (Fig. 7) and become partitioned between the oocyte and the second polar body. The chromosomes remaining in the oocyte after separation decondense to form a nucleusthe pronucleus stage (Fig. 8). Any comparison of the time course of oocyte maturation in different species or under different conditions imposes several difficulties. First, determination of the time at which maturing oocytes move from one stage to another is entirely dependent on one's definition of the two stages, which necessarily involves a certain degree of arbitrariness. From a morphological standpoint, metaphase is the only stage which can be clearly defined. Even if each stage could be clearly defined, arbitrarily or otherwise, abnormalities in the maturation process, which often occur under experimental conditions, make it difficult to judge whether or not the oocytes have reached a certain stage. In addition, since morphological changes in an oocyte are continuous, it is not clear exactly when transitions between stages occur. Second, the progression of maturation in members of an oocyte population is not synchronous. Therefore a statement regarding its time course can be made only on a statistical basis. For example, the time at which oocytes reach a certain transient stage of maturation can only be defined as the time when the proportion of the population at that stage reaches a maximum. Or, the time at which oocytes reach a stage where meiosis is suspended indefinitely, such as metaphase I1 of the vertebrate oocyte, may be defined as the time when half the population has reached that stage. Similar but more complicated considerations may be neces-

206


FIGS.1-8. Chromosome changes during progression of oocyte maturation. Follicle-free mouse oocytes were fixed at different times following culture in an alcohol-acetic acid mixture and stained with Giemsa. FIG. 1. An oocyte with an intact GV and a nucleolus. FIG. 2. Chromosomes in the GV. FIG. 3. Condensed chromosomes at metaphase I. FIG.4. Expulsion of polar body I and oocyte chromosomes and a spindle (arrow). FIG. 5 . Segregation of oocyte chromosomes (aggregated) and polar body I chromosomes (scattered).

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207

s q if a proportion of the population fails to complete maturation because of physiological deterioration. Third, inaccuracy in determining the time course of oocyte maturation could be introduced by ignorance of its exact time of initiation. In nonmammalian species this initiation time may be defined as the time when a maturationinducing agent, such as a hormone or sperm, is applied to the oocytes in vitro. Similarly, in mammals the time at which follicle-free oocytes initiate maturation in vitro may be defined as the time at which oocytes are isolated from their follicles or at which a maturation inhibitor, such as dbcAMP, is withdrawn. Determination of the time at which follicle-enclosed oocytes initiate maturation is a more complex problem. It is generally agreed that, in the normal estrous cycle of mammals, maturation begins at the time of the LH surge, since oocytes enclosed in follicles fail to initiate maturation if they are removed before the LH surge occurs (Ayalon et al., 1972; Hillensjo, 1976; see also Section 11,E). The timing of the LH surge and of the initiation of maturation may be determined at the same time if LH secretion is suppressed at various times using a drug such as Nembutal. The time at which Nembutal administration fails to inhibit oocyte maturation can be considered the point at which oocyte maturation begins. During natural ovulation, rat oocytes reach metaphase I1 10 hours after the LH surge (hfriri and Kraicer, 1972). When ovulation is induced by HCG treatment, oocytes also reach metaphase I1 about 10 hours after HCG injection ( a i l maker et al., 1974). These observations suggest that any delay in the initiation of oocyte maturation following HCG administration is negligible. In fact, it was found that, when HCG was injected into animals, the hormone could be detected within 5 minutes of its administration (Jagiello and Ducayen, 1977). Therefore it may be assumed that the administration of a gonadotropin, as well as the natural surge in vivo, immediately stimulates follicles to initiate oocyte maturation. On the basis of the considerations above, we discuss differences in the time course of oocyte maturation observed under different experimental conditions and among various species. In rodents, the interval between the initiation of maturation and metaphase I1 is similar in both hormone-induced maturation of follicle-enclosed oocytes and spontaneous maturation of follicle-free oocytes (Table I). Thus it may be assumed that the initial events of maturation occur virtually simultaneously in hormonally stimulated follicles and in isolated oocytes. However, a considerable delay (McGaughey and Polge, 1971) or acceleration (Motlik and Fulka, 1976) in the progression of maturation was reported for spontaneously maturing pig ooFIG.6. Metaphase II chromosomes. FIG. 7. Segregation of oocyte chromosomes and polar body I1 chromosomes following activation with ionophore A23187 (anaphase II). FIG. 8. Female pronucleus and polar body II chromosomes. (Brazill, 1977.)

208


TIME

TABLE I COURSE OF OoCYTE MATURATION^

In vivo maturation of follicleenclosed oocytes (hours)

Animal: Stimuli: Stage GV PMI MI

Rabbit Coitus

Rat HCG

Rat

LH surge

Mouse HCG

Hamster HCG

Ram pipiens Pituitary

0 0 0 0 0 0 4.0-7.0 3.0-3.5 4.0-5.0 6.5 6.0 6.5 7.0-8.0 22 MII 9.0-9.5 10.5 10.00 13.00 11.0-12.0 38 MVMII 0.44-0.51 0.62 0.60 0.49 0.62-0.66 0.58 Reference Thibault Zeilmaker Tsafriri and Edwards Usui and Smith et al. (1972) et al. Kraicer and Gates Yanaghachi (1966) (1 976) (1974) (1972) (1959) In v i m maturation of oocytes (hours) Animal: Rabbit Rabbit Mouse Hamster Sturgeon Starfish Stimuli: Isolation Isolation Isolation Isolation Gonadotropin 1-MA Stage 0 0 0 0 0 GV 0 6.0-7.8 6.0-9.0 12.2-12.6 PMI MI 3.3-3.6 5.0-6.0 9.0 9.0 13.4-14.8 1 .o MII 8.1-8.5 9.0-11.0 13.0 c12.0 18.4 11.5 MI/MII 0.41-0.42 0.55 0.69 (0.78) 0.72 0.67 Reference Thibault Chang Donahue Iwamatsu and Vassetzky Ikeda et al. (1972) (1955) (1968) Yanagimachi (1970) ( 1976) ( 1975) Relative duration (7) of different phases in first meiosis and first mitosis**.‘ First mitosis

First meiosis Animal Stage

PMI

MI

Al TI Total

~~~~

Rat

Hamster 1.5 (20) 2.0 (27) 2.0 (27) 2.0 (27) 7.5 (100)

1.0 (21) 2.3 (48) 0.25 (5) 1.28 (27) 4.8 (100) ~~~~

~~~

Mouse 1.5 (18)

4.4-4.5 (55)

1.5-2.0 (21) 0.5 (6) 7.5-9.5 (100) ~~~

Acipenser

2.2 (25) 3.7 (43) 1.7 (20) 1.0 (12) 8.6 (100) ~

~

Oyster

Acipenser

13-14 (26-27) 12-19 (25) 13-14 (29) 7-9 (19)

0.1 (20) 0.2 (40) 0.14 (28) 0.05 (12) 0.5 (100)

-

~

“GV, Immature stage; PMI, prometaphaseI; MI, metaphase I; Al,anaphase I; TI, telophase I; MII, metaphase II, 7 , duration of the first cleavage cycle. bVassetzky ( 1977). CPercentageis given in parentheses.

cytes as compared with hormone-induced in vivo maturation. The time required for follicle-free oocytes to reach metaphase I1 in in vitro culture following their isolation ranges from 42.5 to 55 hours (McGaughey and Polge, 1971), whereas that for follicle-enclosedoocytes maturing in vivo following HCG administration lies between 36 and 37 hours (Hunter and Polge, 1966).

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209

In different species, oocyte maturation proceeds at different rates, even in members of the same phylogenetic class (Table I). However, the relative times required for oocytes to reach different stages of maturation appear to be fairly constant among species. For example, the ratio between the times required for oocytes to reach metaphases I and 11, recorded from the onset of maturation, is similar in different species. This trend seems to suggest that each step of oocyte maturation is coordinated under a principle common to oocytes of different species. This notion may be strengthened by the observation that the relative durations of prometaphase, metaphase, anaphase, and telophase are approximately the same among species and even in meiosis and mitosis (Vassetzky, 1970, 1973, 1977). In addition, evidence has been presented that coordination also exists between different maturational events occumng in the cytoplasm of the oocyte. In X. Zuevis, the time of GVBD, again taken from the onset of maturation, that is, progesterone application to the oocytes, and the time required for progesteronetreated oocytes to become resistant to protein synthesis inhibitors have been found to be variable among oocytes from different females, but the ratio between these two values is always constant, that of the former to the latter being 0.65:l.OO (Wasserman and Masui, 1975a). In R. pipiens oocytes, the times required for GVBD to occur and for an oocyte to become responsive to activation stimuli vary among oocytes obtained from different females. There is also a seasonal dependence. In the fall or early winter, GVBD takes place 20-24 hours following progesterone application at 18"C, while in the spring or early summer it occurs after only 10-14 hours. Likewise, the time required for oocytes to become responsive to activation stimuli varies between 30 and 50 hours. Nonetheless, the ratio of the time elapsed before the appearance of the ability to activate to that elapsed before GVBD was found to remain fairly constant at 2.7 ? 0.2 (Lohka, 1978). These observations may indicate that sequential events taking place during oocyte maturation, either cytoplasmic or nuclear, are closely coordinated within each oocyte. B. MORPHOLOGICAL CHANGES

The progression of oocyte maturation is accompanied by fundamental changes in cytoplasmic as well as in nuclear structures. The recent technical development of oocyte culture has enabled investigators to obtain oocytes at various stages during maturation, allowing detailed study of these changes with light and electron microscopy in mammalian, amphibian, and fish oocytes. Information from these studies has been carefully compared with that from oocytes at the diplotene stage in the ovary. The chromosome configuration of oocytes at the diplotene stage varies from species to species. In mammals, which have been the most extensively studied,

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the chromosomes, condensed during the early phase of meiosis, become more or less diffuse when the oocytes enter the stationary phase. In mouse oocytes at this stage, the nucleus (the GV) contains thin chromatin strands which are occasionally heterochromatinized, clusters of granules ranging between 400 and 900 8,in diameter, and a large spherical nucleolus (Szollosi et al., 1972; Calarco et al., 1972). However, the chromosomes of primate oocytes remain in a partially condensed state and are scattered in the GV (Zamboni et al., 1972). Baker and Franchi (1966, 1972) have observed many short loop projections on these chromosomes, similar to those of lampbrush chromosomes of amphibian oocytes. The chromosomes of fully grown oocytes of lower vertebrates remain in a more condensed state than those of mammalian oocytes and are grouped in a small area of the GV. Light microscopy of chick oocytes has revealed that these discrete chromosomes are confined to an area 20 p m in diameter within the discoidal GV which itself measures approximately 100 p m in depth and 300-400 p m in diameter (Olsen and Fraps, 1950). In amphibian oocytes the GV contains lampbrush chromosomes with many projecting loops. Chromosomes in anuran (R. pipiens) fully grown oocytes are more contracted and project shorter loops (Duryee, 1950) than those of urodele (Norophthalmus viridescens) oocytes (Pilone and Humphries, 1975). This difference in chromosomal morphology between the two amphibian species may reflect a difference in the state of growth of preovulatory oocytes, which might be due to their different breeding habits (see Section 11,B). Studies of fish oocyte chromosomes during oocyte maturation are rather limited. In sturgeons, Dettlaff and Skoblina (1969) showed that the chromosomes in fully grown oocytes were partially condensed and aggregated near the center of the GV. Observations of invertebrate oocyte chromosomes during oocyte maturation are also few, though many investigators have studied the chromosomes of growing oocytes. Das (1976) noted that, in Urechis caupo oocytes, meiosis progressed from the diplotene stage to diakinesis before fertilization triggered maturation. Transition from the diplotene stage to the first meiotic division is initiated by condensation of the chromosomes in the GV. In mammals, chromosome condensation begins near the inner membrane of the nuclear envelope where the chromatin first adheres (Calarco et al., 1972; Motlik et al., 1978). In lower vertebrates, the chromosomes gather in the middle of the GV, where they begin to condense. The process of condensation, which in urodeles involves rapid regression of the loops of the lampbrush chromosomes (Pilone and Humphries, 1975) and progressive contraction of the entire chromosome, has been observed in anurans by Dettlaff and Skoblina (1969) and Brachet et al. (1970) and in fish by Aisenstadt and Dettlaff (1976). These condensing chromosomes also migrate toward the animal pole concurrently with GVBD. Light microscope observations by Vassetzky (1973) of oyster oocyte maturation showed similar chromosome behavior.

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The behavior of nucleoli during the early phase of GVBD has been described both in mice (Zamboni, 1972) and in amphibians (Brachet et al., 1970). In mammals the nucleolus increases in size and accumulateselectron-dense material before disintegrating, while in amphibians the nucleoli condense into a giant nucleolus before disintegration occurs. It was pointed out by Brachet (1965) and Brachet et al. (1970) that DNA cores in the nucleoli remained after nucleolar disintegration, being discharged into the cytoplasm as Feulgen-positive bodies. Undulation of the nuclear envelope is observed shortly before GVBD in oocytes of Chaetopterus (Merriam, 1961), mice (Szollosi et al., 1972), amphibians (Brachet et al., 1970), and sturgeons (Dettlaff and Skoblima, 1969). This undulation of the nuclear envelope heralds GVBD. Cinematographic observations of rat oocyte maturation by Lopata et al. (1977) clearly indicate that only oocytes showing this nuclear envelope undulation undergo GVBD. According to Szollosi et al. (1972), the undulating nuclear envelope in mouse oocytes is eventually folded to form nuclear envelope doublets which are then fragmented. Most of the fragments thus formed later separate into single cistemae at or before metaphase I and eventually become indistinguishablefrom smooth endoplasmic reticulum. In amphibian oocytes, this undulating movement of the nuclear envelope, as well as its disintegration, begins in the vegetal half of the GV (Dettlaff and Skoblina, 1969; Brachet et al., 1970). A classic study of GVBD in chick oocytes by Olsen and Fraps (1950) reported further that the nuclear envelope, once disintegrated at the time of GVBD, was reconstituted during diakinesis as a smaller envelope surrounding an aggregation of the chromosomes discharged from the GV. In the process of GVBD, varied interactions occur between the GV and the cytoplasm. In mice (Zamboni, 1972), amphibians (Brachet et al., 1970), and loach (Iwamatsu and Ohta, 1977), it has consistently been observed that mitochondria increase in number, particularly in the area close to the actively undulating nuclear envelope-more specifically, in the indentations of the envelope. In amphibian oocytes, cytoplasmic granules rich in /3-glycogen accumulate near the disintegrating nuclear envelope and, as GVBD progresses, the glycogen granules, as well as the mitochondria move into the nucleus. Here they surround the meiotic spindle which has a high &glycogen content (Brachet et al., 1970). At the same time, in both mouse (Calarco et al., 1972) and amphibian oocytes, the development of prominent microtubules running perpendicularly to the envelope can be observed. These later penetrate the nuclear envelope. In addition, numerous cytasters form in oocytes of these species. In amphibians (Balinsky and Dans, 1963), sturgeon (Aisenstadt and Dettlaff, 1972), and mouse and human (Zamboni, 1972) oocytes the stacked lamellae disintegrate concomitantly with dissolution of the nuclear envelope. The vesicles formed from them migrate peripherally via Golgi apparatus to give rise to secretory granules. The cortical granules formed inside the cytoplasm before oocyte maturation then migrate toward the periphery of the oocyte as maturation

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proceeds. A similar observation has been reported in sea urchins (Long0 and Anderson, 1970). Changes in the peripheral structures of oocytes undergoing maturation have been studied in amphibians (Van Gansen and Schram, 1968; Kemp and Istock, 1967) and in mammals (Zamboni, 1972). In amphibian oocytes, the number and size of the surface microvilli are progressively reduced during GVBD. This change in surface fine structure may cause macroscopic changes as GVBD progresses, which include increased surface luster and weakened attachment to the vitelline membrane (Schuetz, 1974). However, in mammalian oocytes, the surface microvilli increase and are maintained during the period of maturation (Zamboni, 1972). C. BIOCHEMICAL CHANGES

1 . Energy Metabolism The respiration of oocytes was one of the classic subjects which attracted the attention of “chemical embryologists” a few decades ago (see Needham, 1942; Brachet, 1950). However, until recently, when extensive mammalian research was undertaken, the field remained undeveloped. Much of the original work on oocyte respiration was done using marine invertebrates. Ironically, however, this remains the most confusing area when one attempts to determine the relationship between maturation and respiration changes. Boell et al. (1940) found no change in the O2 consumption rate of starfish ( A . forbesii) oocytes during maturation. But Lindahl and Holter (1941) and Borei (1948) reported a decrease in the case of sea urchin (Psammechinus, Paracentrotus) oocytes. A similar decrease in O2consumption during maturation was reported for the medaka (Nakano, 1953) and recently for the loach (Ozernyuk et al., 1973). Conversely, an increase in the level of respiration during maturation was observed in starfish by Borei (1948), Houk (1974), and Schulz and Lambert (1973), and in sea urchins (Arbacia) by Boell et al. (1940). Among the vertebrates, Brachet et al. (1975a) found that the O2consumption of progesterone-stimulated oocytes of the toad X. laevis gradually increased following GVBD. Spontaneous maturation of follicle-free oocytes of the rat is also accompanied by a gradual increase in the O2 consumption of the oocytes following GVBD; this rise does not occur in oocytes which fail to undergo GVBD (Magnusson et al., 1977; Magnusson and Hillensjo, 1977). These conflicting results, especially those concerning marine invertebrates, may in part be explained by the fact that the oocytes used in some experiments were completely devoid of follicles, while in others they may have been invested with follicle cells. In the latter instance, the values obtained for 0, consumption clearly reflect changes occurring both in the oocyte and in the surrounding cells.

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In fact, as shown by Hillensjo et al. (1975) and Dekel et a f . (1976), the rate of O2 consumption of rat follicle-oocyte complexes decreases during oocyte maturation following LH stimulation. Since by far the greater proportion of the respiratory activity of the complex is due to respiration of the follicle cells surrounding the oocyte (Hillensjo et al., 1975; Magnusson et al., 1977), it is important to realize that, while the 0, consumption of the oocyte may increase during maturation, this would be hidden by a greater decrease in the O2consumption of the surrounding follicle cells. Experiments with mammalian follicles have demonstrated an increase in the rate of lactic acid release as well as of glucose uptake by the follicles after LH stimulation (Nilsson, 1974; Hillensjo, 1976; Tsafriri et al., 1976a). However, ailmaker and Verhamme (1977) recently measured the lactate present within the follicle itself and found that there was no significant change in its level as maturation proceeded to metaphase I. Thus it appears that the level of lactate present in the follicle before LH stimulation is sufficient to support the initiation of oocyte maturation. In other words, increased production of lactic acid by follicles stimulated by LH is not obligatory for the initiation of oocyte maturation. In fact, the lactic acid production of follicles following LH stimulation can be suppressed by treatment with iodoacetate without inhibiting the initiation of oocyte maturation (Tsafriri et al., 1976a). Thus it seems that the stimulatory effect of LH on oocyte maturation is independent of any enhancement of glycolytic activity of the follicle. A wide variety of glucose metabolites, such as pyruvate, lactate, oxaloacetate, succinate, and fumarate, as well as glucose itself, can be used by the follicle to support oocyte maturation, though with varying degrees of effectiveness. However, as first pointed out by Biggers et al. (1967) in mice, follicle-free oocytes can effectively utilize only pyruvate and oxaloacetate as substrates. Lactate and other glucose derivatives can be utilized effectively provided the oocytes are incubated with follicle cells, which are capable of metabolizing lactate. This is apparently a universal phenomenon found not only in the oocytes of other mammalian species such as the monkey (Brinster, 1971), cow (Rushmer and Brinster, 1973), and rat (ailmaker and Verhamme, 1974) but also in amphibians such as X. laevis (Eppig and Steckman, 1974, 1976). In fact, Zeilmaker and Verhamme (1974) suggested that, because of the limited availability of 0, in the follicle, lactate rather than pyruvate can be expected to be the major energy source available to follicle-enclosed oocytes. It has been suggested (Sorensen, 1972; Zeilmaker et al., 1972) that the inability of follicle-free oocytes to consume lactate may result from a lack of endogenous nicotinamide adenine dinucleotide phosphate (NAD) necessary to convert lactate to pyruvate via the activation of lactic acid dehydrogenase (LDH). This suggestion is based on observations that addition of NAD causes the maturation of follicle-free oocytes in a medium containing only lactate as an energy source. However, Eppig and Steckman (1976)

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raise the possibility that the ineffectiveness of substrates other than pymvate and oxaloacetate as energy sources in Xenopus oocytes in culture may be due to differences in the permeability of the oocyte to different substrates, in view of the fact that pyruvate is taken up 30 times faster than any of the other glucose metabolites. In their experiments, these investigators noticed that several days are required before the differential effects of various energy sources became manifest, indicating that the oocytes were using an endogenous energy source. Legname and Buhler (unpublished) reported that Bufo ovarian oocytes contained more citrate than fumarate during the winter months, but that the ratio was reversed in the spring or when winter oocytes were induced to mature by progesterone treatment in the absence of exogenous nutrients. Although there is not yet complete agreement with respect to the mechanisms underlying the energy metabolism of maturing oocytes, there is no doubt that oocyte maturation is an energy-consumingprocess. Most investigators agree that GVBD cannot take place under conditions inhibiting aerobic metabolism, such as lack of oxygen (Zeilmaker and Verhamme, 1974; Gwatkin and Haidri, 1974) or the presence of a respiratory inhibitor such as KCN (Brachet et al., 1975a). Schulz and Lambert (1973) noted that the maturation of starfish oocytes involved a decrease in AMP and ATP levels, but an increase in the level of ADP, and suggested that the increased 0, consumption observed could be interpreted as an indicator of enhanced oxidative phosphorylation acting to restore ATP to its prematuration level. This increase in 0, consumption of the oocyte has been found to occur after GVBD in starfish (Schulz and Lambert, 1973; Houk, 1974), Xenopus (Brachet et al., 1975a), and rats (Magnusson et al., 1977). In this connection, it is interesting that neither anaerobic conditions nor the presence of KCN interfere with progression of the second meiotic division in rat oocytes (Zeilmaker and Verhamme, 1974). Perhaps the increased level of O2 consumption following GVBD is a result of mechanisms operating to replenish ATP consumed during GVBD, thus enabling oocytes to proceed with meiotic division. One of the energy-requiring processes occumng during oocyte maturation is the synthesis of protein. This dependence seems logical intuitively; two observations are mentioned here. First, there is a substantial reduction in the level of protein synthesis during maturation of R. pipiens oocytes under anaerobic conditions (Smith and Ecker, 1970b). Second, the rise in O2consumption observed in maturing X . laevis oocytes coincides with the time of increased amino acid incorporation into oocyte protein (Brachet et al., 1975a). 2. Protein Synthesis An increase in the protein synthesis activity of oocytes during maturation is apparently a general phenomenon among animals. However, the fact that protein synthesis can be measured in a variety of ways under a variety of conditions makes it difficult to compare accurately the results obtained by different inves-

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tigators. Smith et al. (1966) injected R. pipiens oocytes with radioactive leucine and then extracted the oocytes with a saline solution following incubation for a given period of time. The extracts were then precipitated with hot trichloroacetic acid ("CA), and the leucine incorporated into this fraction measured. Obviously, this approach gives no information concerning proteins insoluble in the saline solution. Some investigators, however, have exposed oocytes to a labeled amino acid for a certain length of time, collected all the material precipitated by TCA directly from the oocytes, and measured the radioactivity in the precipitate. This fraction may represent the total protein content of the oocytes. However, our experience has led us to believe that with amphibian oocytes this method often introduces some error into the determination, since free leucine in TCA precipitates is difficult to remove, while the method using perchloric acid (PCA) instead of TCA gives more accurate results. Moreover, it should be noted that an accurate determination of the size of the amino acid pool in the oocyte, which may change during maturation, is difficult and yet crucial for comparison of the rates of protein synthesis at different stages of maturation. Nevertheless, some reports provide no data concerning the amino acid pool size in maturing oocytes at different stages. In spite of these shortcomings, we state as a general rule that changes in protein synthesis activity occur during the maturation of oocytes of all species studied to date. The rate of synthesis usually increases as a result of treatment with a maturation-inducing hormone. In R. pipiens, protein synthesis increases by a factor of about 10, about 18-24 hours after hormone treatment, following GVBD, and reaches its maximum level after metaphase I (Smith et ul., 1966; Smith and Ecker, 1970a). However, recent reinvestigations of protein synthesis in X. laevis (O'Connor and Smith, 1976) and R. pipiens (Shih et al., 1978) oocytes have indicated that the increase in the rate of total protein synthesis during maturation is at most two-fold when amino acid pool sizes and diffusion rates of labeled amino acids are taken into consideration. Protein synthesis activity in oocytes of A. forbesii (Wassarman, 1971) and U.caupo (Blankstein and Kiefer, 1977) increases two to four times, again after GVBD. However, it was reported that although treatment of X . laevis oocytes resulted in a stimulation of protein synthesis 1-4 hours after treatment, which was before GVBD, this stimulation was only temporary and protein synthesis was sharply reduced, eventually to 50% of the maximum level attained (Baltus et al., 1973; Brachet et al., 1974). Invertebrate studies by Houk and Epel (1974), using Patiria niniata, demonstrated that protein synthesis in starfish oocytes began to increase 12 minutes after 1-MA stimulation, before GVBD occurred. By prophase 11, it had reached a level five times that of untreated oocytes. This change in protein synthesis activity following 1-MA stimulation is unaffected by fertilization. Changes in the pattern of oocyte protein synthesis also occur as maturation progresses. Smith and Ecker (1971), using R. pipiens oocytes at different stages of maturation, analyzed the incorporation of radioactive leucine into various

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protein fractions following separation by one-dimensional gel electrophoresis and discovered characteristic changes occurring during the progression of maturation. Pennequin et al. (1975) and Schorderet-Slatkine and Beaulieu (1977), using the technique of double-labeling electrophoresis, observed differences in the synthetic activity of various protein fractions extracted from hormone-treated and untreated Xenopus oocytes labeled with 3H and '*C, respectively. Protein synthesis by mammalian oocytes has been studied by one- or two-dimensional gel electrophoresis followed by autoradiography in mice (Golbus and Stein, 1976; Schultz and Wassarman, 1977a,b; Schultz et al., 1978), pigs (McGaughey and van Blerkom, 1977), sheep (Warnes et al., 1977), and rabbits (van Blerkom and McGaughey, 1978a). While the studies done on amphibian (Smith and Ecke, 1971; Pennequin et al., 1975) and mouse (Golbus and Stein, 1976; Schultz and Wassarman, 1977a,b) occytes showed significantchanges in protein synthesis patterns only after GVBD, the study of sheep oocytes revealed changes occuring before GVBD (Warnes et al., 1977). These conflicting results may lead us to question whether the oocyte protein synthesized during maturation is required for the progression of maturation. Early studies showed that protein synthesis inhibitors, such as cycloheximide and puromycin, acted as strong inhibitors of oocyte maturation in amphibians (X. laevis, Brachet, 1967; R . pipiens, Smith and Ecker, 1969; R . temporaria and B . bufo, Dettlaff, 1966) and in fish (Acipenser, Dettlaff and Skoblina, 1969). These reports consistently indicated that GVBD, chromosome condensation, and nucleolar dispersion were all inhibited at concentrations of the inhibitor which suppressed virtually all protein synthesis. However, Smith and Ecker (1970b) and Ecker and Smith (1971a) showed that a substantial reduction in the level of protein synthesis under anaerobic conditions did not prevent oocytes from maturing, provided they had undergone GVBD. Baltus et al. (1973) also obtained evidence that maturation could proceed when protein synthesis was 50% inhibited by a 5-hour cycloheximide treatment. In mammals, treatment with protein synthesis inhibitors such as puromycin or cycloheximide arrests oocytes at metaphase I or at the circular bivalent chromosome stage, but GVBD and chromosome condensation are unaffected (Stem et al., 1972; Golbus and Stein, 1976; Wassarman and Letourneau, 1976b; Schultz and Wassarman, 1977b). However, it should be noted that protein synthesis was not completely inhibited in these experiments. It is possible that the small amount of synthesis still occurring in the presence of the inhibitors may provide the proteins necessary for the early events of maturation. The results obtained by researchers working with invertebrates are not quite so uniform. Houk and Epel (1974) found pactamycin to be a very effective inhibitor of protein synthesis in P. rniniata oocytes-a dose of 100 pg/ml eliminated all incorporation into TCA-precipitable protein within 15 minutes of its application. Despite the complete lack of protein synthesis, GVBD occurred and the oocytes

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developed a spindle capable of organizing chromosomes. Maturation was not arrested until metaphase I or anaphase I. Brachet and Steinert (1967), conducting experiments with the starfish Asterias rubens, found that puromycin (50 pg/ml) and cycloheximide (20 pg/ml) did not inhibit GVBD but did prevent chromosome migration and spindle formation. The work of Zampetti-Bosseler et al. (1973), using Asterias glacialis, demonstrated that fusidic acid or puromycin prevented formation of the mitotic apparatus required at metaphase I, but not GVBD. In this case, protein synthesis appears to have been reduced to about 20% of control levels. The same group also studied Chaetopterus oocytes, finding that fusidic acid greatly suppressed protein synthesis and stopped maturation before GVBD. Puromycin, however, while inhibiting protein synthesis by 50%, did not affect maturation, implying that only some of the protein synthesized during maturation was actually required for maturation (cf. amphibian results above). According to Blankstein and Kiefer (1977) incubation of U.caupo with cycloheximide (50 pg/ml) inhibits protein synthesis but does not prevent fertilized eggs from undergoing fertilization membrane elevation, GVBD, two meiotic divisions, and pronuclear formation. These eggs are unable to cleave, though. Apparently, in this species, all the proteins required for meiotic maturation are present before the onset of maturation, and the proteins synthesized during maturation are needed for later developmental events. Thus two questions arise. First, what portion of the protein synthesized by the maturing oocyte is responsible for the progression of maturation and, second, at what time are the proteins necessary for each step of maturation synthesized? Mom11 et al. (1975), using R. pipiens, reported that the proteins responsible for GVBD, which occurs 8-9 hours following gonadotropic stimulation of follicleenclosed oocytes, were synthesized within 5 or 6 hours after hormone treatment. Inhibition of protein synthesis by cycloheximide after this time failed to inhibit GVBD. Wasserman and Masui (1975a) showed that the inhibitory action of cycloheximide on GVBD in X. laevis oocytes was effective when the inhibitor was applied during the first two-thirds of the time period between progesterone stimulation and GVBD. Thus, in amphibians, as suggested by Smith and Ecker (1970b) and Ecker and Smith (1971a), it seems likely that the low level of protein synthesis occurring during the early period before GVBD reflects the production of proteins necessary for the events of maturation per se to occur, while the much higher level observed later represents proteins required for processes occurring later in development. However, it must be pointed out that the effect of protein syndesis inhibitors preventing oocytes from initiating maturation does not necessarily signify a requirement for new protein synthesis by the oocytes to initiate maturation, since it is possible that the inhibitors also interfere with ongoing synthesis of preexisting, but short-lived, proteins in the oocytes to deprive them of the ability to respond to maturation-inducing agents. Schultz and Wassarman (1977a), using mice, have suggested that inhibition of

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protein synthesis during the first 5 hours of maturation-up to the circular bivalent chromosome stage-does not affect maturation, but that inhibition during the second 5-hour phase-from the circular bivalent chromosome stage to metaphase I-prevents any further progression of maturation. These results suggest a situation different from that in amphibians. Nevertheless, the two can be reconciled to the extent that certain proteins are synthesized during specific phases of maturation and that they are essential to the maturation process at some subsequent point. However, in P. miniuta, Houk and Epel (1974) noted that the maturation of pactamycin-treated oocytes, compared to that of controls, was retarded at all stages before finally stopping. Based on this observation, they postulate that the blocking of maturation by protein synthesis inhibition may be a meiosis-specific, but not stage-specific, response. In other words, meiosis is arrested not because a specific protein required at a specific stage is unavailable, but rather because a general lack of protein synthesis causes maturation to slow down as the reserves of available protein are exhausted. Eventually, maturation stops when there is not enough protein available to support any further progression. Changes in oocyte protein synthesis activity have been studied under the influence of other compounds which inhibit maturation. Drugs such as theophylline (O’Connor and Smith, 1976) and papaverine (Bravo et al., 1978) have been shown to inhibit oocyte maturation in X. laevis, as does dbcAMP (Stem and Wassarman, 1974) in mice. According to Bravo et al. (1978), aclose correlation exists between the inhibitory effects of the compounds on Xenopus oocyte protein synthesis and their capacity to block maturation. However, mouse oocytes which fail to undergo GVBD, because of treatment with dbcAMP, also fail to exhibit only the changes in protein synthetic pattern which occur during normal maturation (Stem and Wassarman, 1974; Schultz and Wassarman, 1977b). Stimulation of protein synthesis following the initiation of maturation does not appear to require the presence of the GV in amphibians. Smith and Ecker (1969) and Ecker and Smith (1971a) showed, in R. pipiens, that normal oocytes and those from which the GV had been removed (enucleated)exhibited quantitatively and qualitatively almost identical patterns of leucine incorporation into proteins following progesterone treatment. In mice Schultz et ul. (1978), using labeled methionine and two-dimensional electrophoresis, found that there was no difference in the protein synthesis pattern between nucleated and enucleated oocytes when they were cultured in the presence of dbcAMP which prevents nucleated oocytes from GVBD. However, when oocytes were cultured without dbcAMP, the difference became manifest approximately at the time at which nucleated oocytes underwent GVBD. From these observations they proposed that certain protein synthesis-stimulating factors which have appeared in the oocyte cytoplasm in the initial period of maturation, are stored in the GV until they are released into the cytoplasm at GVBD. Here they direct the observed changes in

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protein synthesis. If, as in enucleated oocytes, these factors remain and accumulate in the cytoplasm, the pattern of protein synthesis in the cytoplasm may be changed when the level of the factors reaches a critical point. This point clearly may be reached earlier than it is reached in intact oocytes. Whatever the case, it would be expected that GV removal just prior to GVBD ought to delay these changes in the protein synthesis pattern until concentrations of the proteinstimulating factors reach the required cytoplasmic levels. At any rate, the fact that the changes in the protein synthesis pattern during cytoplasmic maturation of oocytes can occur independently of the GV indicates that these changes are not dependent on transcriptional activity of the nucleus. This explains why destruction of the chromosomes in the oocyte by X ray (Masui, 1973a,b), or their inactivation by RNA synthesis inhibitors (Smith and Ecker, 1970a; Ziegler and Masui, 1976a), does not affect the course of cytoplasmic maturation in amphibian oocytes. 3. RNA Synthesis Early work with amphibians consistently showed that gonadotropin-induced maturation of follicle-enclosed oocytes was inhibited by the RNA synthesis inhibitor, actinomycin D (Dettlaff, 1966; Brachet, 1967; Brachet and Steinert, 1967; Schuetz, 1967b). Another inhibitor, a-amanitin, is also effective, reducing RNA synthesis in follicles by 80% or more (Wasserman and Masui, 1974). Curiously, though, these RNA synthesis inhibitors occasionally fail to suppress maturation of follicle-enclosedoocytes (Merriam, 1972; Wasserman and Masui, 1974). The reason for this is unclear. It may be that these follicles were exposed to gonadotropins at subthreshold levels in vivo (Wasserman and Masui, 1974). In rats, LH-induced maturation of follicle-enclosed oocytes cannot be inhibited by actinomycin D vsafriri et al., 1972). It is interesting to note that ethidium bromide which is known to inhibit mitochondria1 RNA synthesis in eukaryotic cells is always the most potent inhibitor of gonadotropin-induced maturation of follicle-enclosed oocytes in amphibians (Wasserman and Masui, 1974; Schmerling and Skoblina, 1978). These observations suggest that the effect of gonadotropin on follicles in initiating oocyte maturation may involve the induction of RNA synthesis under certain physiological conditions, since all RNA synthesis inhibitors fail to inhibit maturation when follicle-freeoocytes are induced to mature, as discussed later. Progesterone-induced maturation of follicle-free amphibian oocytes is inhibited neither by actinomycin D (Schuetz, 1967b; Smith and Ecker, 1969; Baltus et al., 1973) nor by a-amanitin (Baltus ef al., 1973; Wasserman and Masui, 1974; Brachet et al., 1974). Although actinomycin D does not prevent oocytes from initiating maturation and has even been observed to exert a favorable effect (Baltus et al., 1973), the progression of maturation is not normal. For instance, the behavior and morphology of the chromosomes in actinomycin D-treated

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oocytes are quite irregular, either becoming pycnotic or failing to condense following GVBD (Baltus et al., 1973; Ziegler and Masui, 1976a). Maturation of follicle-free mouse oocytes proceeds to metaphase I or to the circular bivalent chromosome stage in the presence of low concentrations of actinomycin D, but at concentrations higher than 1 pg/d it is inhibited before GVBD (Bloom and Mukhejee, 1972; Golbus and Stein, 1976) or is accompanied by severe chromosomal aberrations (Alexandre and Gerin, 1977). Isolated oocytes of Asterias and Chaetopterus have been found by different investigators to initiate maturation in the presence of actinomycin D in all species examined. Maturation of A. forbesii oocytes occurs normally at actinomycin D concentrations as high as 10 pg/ml, as scored by the occurrence of normal fertilization (LaMarca et al., 1971). Asterias glacialis and Chaetopterus oocytes undergo normal maturation at actinomycin-D concentrations of 20 pg/ml (Zampetti-Bosseler et al., 1973). However, Brachet and Steinert (1967) found that a concentration of 20 pg/ml prevented polar body formation in A . rubens. It should be noted, though, that actinomycin D reduced RNA synthesis in the oocytes only by about 50% at the concentrations used in the experiments of LaMarca et al. (1971) and Zampetti-Bosseler et al. (1973). It is not known how completely RNA synthesis was inhibited in the experiment of Brachet and Steinert (1967). Although the results cited above appear to indicate that oocyte maturation is not highly dependent on RNA synthesis, this does not necessarily imply that there is no RNA synthesis during oocyte maturation. The pioneering work of Brown and Littna (1964) with X. laevis indicated that fully grown oocytes, stimulated to ovulate by HCG, synthesized not only rRNA but also a significant amount of heterogeneous RNA. The latter may include mRNA and mitochondrial RNA (mtRNA). Oocyte RNA synthesis is stimulated during a brief period between the onset of maturation following progesterone treatment and GVBD in Xenopus (LaMarca et al., 1975; Webb et al., 1975) and R . pipiens (Morrill et al., 1975). However, this progesterone-induced rise in RNA synthesis has not been observed in R. pipiens (Smith and Ecker, 1970a). At the time of GVBD, a sharp decrease in rRNA synthesis occurs, and only the synthesis of heterogeneous RNA, which corresponds to 1-2% of prematurational levels of RNA synthesis, continues (Webb et al., 1975). In mice, Rodman and Bachvarova (1975, 1976) showed that RNA synthesis continued at least up to the last 2 hours before GVBD. Experiments by Wassarman and Letourneau (1976a) also indicate that RNA synthesis occurs in fully grown oocytes containing an intact GV. These results contradict earlier work by Oakberg (1968) and Moore et al. (1974). Wassarman and Letourneau (1976a), who injected radioactive precursor directly into the follicle, suggest that the failure of the earlier workers to detect labeled RNA in fully grown oocytes may

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have been due to low intrafollicular levels of the precursor which was injected intraperitoneally. Alexandre and GBrin (1977) hypothesize that the RNA synthesized early in maturation, before GVBD, may include mRNA. During and after GVBD, there is a dramatic reduction in the rate of RNA synthesis directed by the nucleus (Rodman and Bachvarova, 1976). A similar reduction in nuclear RNA synthesis also occurs in starfish and Chaetopterus oocytes following the onset of maturation (Boylan et al., 1973; Zampetti-Bosseler et al., 1973). With respect to the nature of the RNA synthesized after GVBD, there is evidence suggesting that the RNA is not of ribosomal origin. In Asterias (LaMarca et al., 1971; Boylan et al., 1973), as well as in Xenopus (Webb et al., 1975), the major portion of the RNA synthesized comprises two RNA populations of different S values, 4-5 and 15-19S, while the minor portion is heterodispersed RNA.

4. DNA Synthesis Experiments with X . laevis oocytes by Hanocq et al. (1974) and by Brachet et al. (1974) have shown that there is no detectable DNA synthesis in the nucleus or chromosomes during progesterone-induced maturation, and that various agents which inhibit DNA synthesis, such as hydroxyurea, deoxyadenosine, cytosine arabinoside, ethidium bromide, X rays, and dimethyl B rifampicin, exert no inhibitory effect on maturation. Although treatment with these agents leads to pycnotic chromosomes in the oocytes, it is difficult to assess whether this is due to DNA synthesis inhibition or to other effects of the agents. Huez et al. (1972) and Zampetti-Bosseler et al. (1973), working with starfish oocytes, obtained results similar to those for Xenopus oocytes. In their experiments, maturation, scored by polar body formation, was unaffected by DNA synthesis inhibitors, though synthesis was not completely inhibited. Experiments with U . caupo oocytes by Blankstein and Kiefer (1977) demonstrated that normal chromosome condensation and pronuclear formation were not inhibited by DNA synthesis inhibitors. While it is thus apparent that DNA synthesis is not required during maturation, it is equally true that some amount of DNA synthesis occurs during the maturation process. In amphibian oocytes, Hanocq et al. (1974) noted that DNA synthesis occurred in the cytoplasm, since DNA synthesis could be detected in oocytes from which the GV had been removed. This DNA synthesis is possibly of mitochondria1 origin and a continuation of the low level of mitochondrial DNA synthesis taking place throughout oogenesis. During maturation, DNA synthesis of an undetermined nature has also been reported to take place in starfish (Wassannan, 1971; Huez el al., 1972; Zampetti-Bosseler et al., 1973) and U . caupo (Blankstein and Kiefer, 1977) oocytes. Wassarman (1971) found

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that the DNA synthesized during maturation of A. forbesii was of nuclear origin, whereas mitochondria1DNA synthesis did not occur until the 128-cell stage. We were unable to find any reports which examined DNA synthesis during the maturation of mammalian oocytes, except for two dealing with repair synthesis (Masui and Pedersen, 1975; Brazil1 and Masui, 1978). In conclusion, then, we state that any DNA synthesis which occurs during oocyte maturation is not obligatory for the progression of maturation.

IV. Initiation of Oocyte Maturation A. MATURATION-INDUCING SUBSTANCE 1. Specificity

As discussed in Section I1,A-F, there is a growing body of evidence indicating that hormones play an essential role in the induction of oocyte maturation in a wide variety of animals. As yet, though, there are only a few groups of animals in which a chemically defined substance is known to act directly on oocytes as a MIS. These are amphibians and starfish. In amphibians, progesterone appears to be the natural MIS causing oocyte maturation, although other steroids are known to be as effective as progesterone; these include DOC in R. pipiens (Schuetz, 1967a; Smith and Ecker, 1971) and aldosterone and testosterone in X. laevis (Jacobelli et al., 1974; SchorderetSlatkine, 1972), but not estradiol or its derivatives in either species. A comparison of the potency of various steroids as inducers of maturation in R. pipiens oocytes led Morrill and Bloch (1977) to suggest that a special arrangement of substituents on the upper (p) surface of the steroid molecule was of critical importance with respect to its potency as a MIS, 3,20-dione, 21-01 forms being the most active. However, introduction of a polar group oriented toward the lower (a)surface abolishes the activity of a MIS. A similar comparative study of the maturation-inducing potency of various steroid using medaka oocytes has led Iwamatsu (1978) to the conclusion that the steroids effective in inducing maturation have in common a C=O (or a-OH)group at 3C, and a P-OH group at 17C in the C19-steroids,and a C=20 (or P-OH) group at 3C and a C=O (or a-OH) group at 20C in the C2,-steroids, in addition to a A4 or A5unsaturated or 5-saturated configuration. Progesterone is known to be metabolized by amphibian oocytes. To determine its metabolic products, oocytes were incubated with radioactive progesterone, and the derivatives produced by the oocytes were chromatographicallyanalyzed. As shown in Table II, oocytes of R. pipiens (Reynhout and Smith, 1973), X. laevis (Fouchet et al., 1975), P . waltlii (Fouchet er al., 1975; Ozon et al.,

TABLE II METABOLISM OF PROGESTERONE IN AMPHIBIAN OOCYTES Species R a m pipiens,

R . temporaria Xenopus laevis

Triturus alpestris

Pleurodeles waltlii

Metabolite Sa-Pregnan-20a-ol-3-one Sa-Pregnane-3,20-dione Sa-Pregnane-3P2Oa-diol 17a,2Oa-Dihydroxypregn-4-en-3-one 17a-Hydroxy-4-pregnene-3,20-dione CAndrostene-3,17-dione Sa-Pregnene-3,20-dione SP-Regnene-3.20-dione 17a-Hydroxy-4-pregnene-3,2O-dione Sa-Pregnene-3,2O-dione SP-Pregnene-3,20-dione 2OP-Hydroxypregn-4-en-3-one Sa-Pregnane-3,20-dione 3a-Hydroxy-Sa-pregnan-20-one 3P-Hydroxy-Sa-pregnan-20-one

Enzyme Sa-Reductase 20a-Hydroxylase 17a-Hydroxylase 19,2I-desmolase Sa-Reductase 5P-Reductase 1701-Hydroxylase Sa-Reductase 5P-Reductase 20P-Hydroxysteroidoxidoreductase (soluble) Sa-Reductase (microsomal) 3a-Hydroxysteroidoxidoreductase(microsomal) 3p-Hydrox ysteroid oxidoreductase (microsomal)

Reference Reynhout and Smith, 1973; Antilla, 1977 Fouchet e t a l . , 1975 Antilla, 1977

Antilla, 1977 Ozon et a l . , 1975

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1975), and R. temporaria and Triturus alpestris (Antila, 1977) all possess 5a-or SP-reductase, which converts progesterone into 5a- or SP-reduced derivatives such as 5a-pregnandione. In addition, 17a-hydroxylase was found in Xenopus and Triturus oocytes. Pleurodeles oocytes contain 3a-, 3p-, and 2Qphydroxysteroid oxidoreductases. Thus different steroid derivatives of progesterone are the major products of its metabolism in the oocytes of different species; for instance, Sa-pregnandione in R. pipiens and androstenedione in X. luevis. However, all these derivatives are less potent than progesterone as MISS when applied to the oocytes of these species. Thus it seems quite unlikely that the metabolic conversion of progesterone plays a significant role in the initiation of maturation. In starfish, it has been shown beyond doubt that 1-MA is the natural MIS. Although this chemical possesses the highest maturation-inducing activity among compounds used with Asterius oocytes (Kanatani and Shirai, 1971), recently an artificially synthesized adenine derivative, 1-benzyladenine (1-BA), has been shown to be a more potent MIS in Marthasterias glacialis oocytes (Doree et al., 1976a,b). After comparing the activity of various adenine derivatives in this species of starfish, Doree et al. (1976a) concluded that the nature of the group in the N-1 position, and the absence of additional groups in the N-7 and N-9 positions, were of critical importance for MIS activity. An analysis of the metabolism of 1-MA in the oocytes of A . forbesii was carried out by Toole and Schuetz (1974). They found that radioactive 1-MA applied to oocytes was metabolized into a biologically inactive compound, the nature of which was unknown. Thus it may be concluded that 1-MA acts as a MIS prior to chemical modification.

2. Action Site The necessity for stereochemical specificity of the molecules which act as MIS suggests that these molecules must interact with a certain site on a specific reacting molecule or a molecular unit in the oocyte. In order to determine the location of such reacting molecules in the oocyte, the capacity to bind MIS has been compared in different subcellular components of the oocyte. In amphibian oocytes, Ozon and Belle (1973) found that the component containing the melanosome fraction showed the highest progesterone-binding capacity. Further experiments indicated that progesterone was the steroid hormone with the strongest affinity for melanosomes (Ozon et al., 1975; Belle et al., 1977a). When melanosomes incubated with progesterone are injected illto Xenopus oocytes, some of the changes occurring during the early phase of maturation, such as GV migration toward the animal pole and its partial disintegration, can be seen in the recipient oocytes, although they eventually undergo degeneration (Jacobelli et al., 1974). On the basis of these observations, the suggestion was made that progesterone-bound melanosomes may play a role in

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the initiation of oocyte maturation in amphibians. However, the discovery of a mutant albino strain of X . laevis completely devoid of melanosomes and their precursors, which produces fertile oocytes (Bluemink and Hoperskaya, 1975), has made it difficult to assign any significant role to melanosomes with respect to the initiation of oocyte maturation. The nature of the subcellular components in starfish oocytes which interact with 1-MA has been investigated in A. forbesii by Jeffery (1977). He compared the ability of various enzymes to deprive the oocytes of the capacity to respond to 1-MA, finding RNases and proteases to be the most effective among those tested. He also noted that enzyme-treated oocytes recovered from the loss of reacting capacity within 90minutes following transfer to enzyme-freemedium and that this recovery was not suppressed by an RNA synthesis inhibitor (actinomycin D) or a protein synthesis inhibitor (emetin). Although it would be premature at the present time to ascribe the observed effects of enzyme treatments to the specific action of the enzymes used, it may be suggested that interaction of the molecular unit with 1-MA involves RNA. However, in the starfish M . glacialis it was found that mild treatment of the oocytes with Triton X-100 (0.01-0.02%) abolishes their responsivenessto 1-MA @ode et al., 1976b). This was confirmed by Kanatani (1978), using A . pectinifera. Kanatani (1978) and Morisawa and Kanatani (1978) further reported that a Triton X-100 wash of follicle-free oocytes contained a heat-stable, nonprotein substance, and that oocytes incapacitated following Triton treatment had their responsiveness to 1-MA restored when incubated with this substance. These observations strongly suggest the existence of a substance in starfish oocytes that interacts with 1-MA, thus initiating maturation. Smith and Ecker (1969) and Masui and Markert (1971) using R. pipiens, and Kanatani and Hiramato (1970) using A. pectinaris, showed that a MIS, progesterone or 1-MA, always failed to induce oocyte maturation when it was microinjected into oocytes. These MISS are effective only when applied to the outer surface of oocytes. Accordingly, it has been hypothesized that a MIS primarily interacts with molecules located on or near the outer surface of the oocyte. There is one report indicating that some steroids, for example, hydrocortisone, effectively induce maturation if injected into X. laevis oocytes (Schorderet-Slatkine, 1972). However, this observation does not appear to contradict the above hypothesis, since the possibility that the injected steroid may have leaked from the oocyte and exerted a surface action was not ruled out. The surface action hypothesis has recently gained strong support from the experiments of Baulieu and his associates (Baulieu et al., 1978; Godeau et al., 1978). They succeeded in inducing Xenopus oocytes to mature by exposing them to a polymer-conjugated steroid 3-0x0-4-androstein-17P-amido polyethylene oxide (ACA-poly EO), which has a molecular weight of over 20,000 daltons. This compound is as effective as progesterone in inducing the maturation of

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Xenopus oocytes when it is applied to the oocyte surface, whereas it is totally ineffective when injected into oocytes. In additon, uptake of this chemical by treated oocytes is negligible compared to that of free steroids, and no degradation of the compound occurs during the incubation period with the oocytes. Conversely, neither the carrier polymer (poly EO) alone, nor its conjugate with an inactive steroid, such as estradiol, has a maturation-inducing effect. Therefore it is almost certain that the active polymer-conjugated steroid acts on the surface of the oocytes to induce maturation, although, strictly speaking, proof of its actual surface localization by electron microscope autoradiography is still required. Recent experiments by Shida and Shida (1976) suggest that the molecules reacting with 1-MA may also be localized on the oocyte surface. Their results demonstrate that a-(1+6)-heterogalactan, a polysaccharide, reversibly inhibits the maturation-inducing action of 1-MA. Because of its large size (1.55S), it probably interacts mainly with molecules on the surface of the oocytes to interfere with 1-MA action. The distribution of reaction sites for a MIS on the surface of the oocyte has been investigated in both amphibians and starfish. Recently, Cloud and Schuetz (1977) found that the sensitivity of R. pipiens oocytes to MISS was higher in the animal half than in the vegetal half. This demonstration involved the application of progesterone to a restricted area (ranging from 13 to 15% of the total surface area) of a single oocyte by tightly fitting it into a conical tube with the open end exposed to a medium containing progesterone. During the course of exposure, progesterone was found to accumulate on the exposed hemisphere. When the oocytes were exposed to progesterone at the animal pole, 55% underwent GVBD within 24 hours, whereas none of those exposed to progesterone at the vegetal pole underwent GVBD. This result supports the idea that the molecule or molecular unit responsive to progesterone is more concentrated in the animal hemisphere than in the vegetal hemisphere. Local application of 1-MA to starfish oocytes has been carried out by Shirai (1978). In her experiment, an oocyte was tightly fitted into a capillary tube, and one hemisphere was stained with Nile blue and the other with neutral red. When both hemispheres of the oocytes thus stained were exposed to 1-MA, the polar body was given off by each of the two hemispheres equally frequently. However, when only one of the hemispheres was exposed to 1-MA, the polar body was usually given off by the hemisphere to which 1-MA had been applied. From these observations, it may be concluded that 1-MA acts equally on all regions of the starfish oocyte, but that its differential application determines the site of polar body formation.

3. Mode of Action Investigations into the nature of the interaction between a MIS and reacting oocytes molecules appear to have been hampered by limited evidence concerning

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the existence of competitive chemical analogs of MISS. Although it has been reported that the steroid analogs ethynylestradiol and MK 665 inhibit the maturation of progesterone-treated R. pipiens oocytes to some extent (Morrill and Bloch, 1977), the presence of follicle cells in this experiment makes it difficult to assess the direct action of the steroids on the oocytes. In starfish, it was shown that I-MA uptake by oocytes was competitively inhibited to a high degree by its analog, 1,9-dimethyladenine, which itself has little MIS activity @ode et al., 1976b). Nevertheless, no competitive inhibition of 1-MA-induced maturation is caused by this chemical analog. Both in starfish and in amphibians, xanthine derivatives have an inhibitory effect on oocyte maturation. But, although xanthine derivatives are chemically analogous to 1-MA, their inhibitory effect in starfish is observed only when they are applied at concentrations lo4 times higher than 1-MA (Doree et al., 1976a). Similarly, in amphibian oocytes, xanthine derivatives, such as caffeine and theophylline, inhibit maturation at high concentrations (0'Connor and Smith, 1976). Obviously, the inhibition cannot be due to the derivatives acting as competitive analogs of the MIS (progesterone). Rather, it is likely that the inhibitory effect of xanthine derivatives on oocyte maturation, both in amphibians and in starfish, is brought about through their well-known effects on phosphodiesterase and Ca movements in the cell. The reaction of an oocyte to a MIS appears to involve two distinct steps. Marot et al. (1977) found that, when Xenopus oocytes were exposed to progesterone for 20 hours at subthreshold concentrations, the oocytes began GVBD at much earlier times following a second treatment with maturation-inducing levels of progesterone. However, the facilitating effect of subthreshold progesterone treatment does not last more than 24 hours after the oocytes are returned to hormone-free medium. The induction of maturation in the starfish ovary also appears to be a two-step process. Using M. glacialis, Guerrier and Dorke (1975) showed that oocytes exposed to lo-' M 1-MA for at least 4.5 minutes, or for two 2.5-minute periods separated by no more than 7.5 minutes, matured. The short treatments became ineffective when the interval between the two exceeded 7.5 minutes, indicating that the oocyte reaction to subthreshold doses of 1-MA was reversible. These observations seem to indicate that, in amphibians and starfish, the first response of oocytes to a MIS is a reversible reaction which is followed by an irreversible reaction that actually triggers maturation. The reversibility of the initial step of maturational change induced by a MIS in oocytes suggests that this change may not depend on the formation of a stable complex between the MIS and its receptor on the oocyte. If the formation of a receptor-inducer complex is a prerequisite for the subsequent process of maturation, it should be expected that some minimum amount of MIS must be taken up by the oocyte before it initiates maturation. However, at least in the induction of maturation in Xenopus oocytes, this is not the case. Bell6 et al. (1976) found

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that, as the concentration of progesterone to which the oocytes were exposed was decreased, the duration of exposure required to induce maturation increased, but the amount of steroid taken up by each oocyte before initiating maturation was greatly reduced. Apparently, the critical factor for the induction of maturation in this case was not the number of complexes formed between the inducer and its oocyte receptor, but the number of interactions between them. Perhaps the MISreceptor molecule interaction involves the induction of a permanent change in the receptor molecule located on the oocyte surface, without the formation of a stable MIS-receptor complex. Maturation would be induced by the cumulative effect of many molecules undergoing this change. The nature of the change in the oocyte surface molecules necessary to initiate maturation may be inferred from the effects of various chemicals which mimic the effect of a natural MIS. The molecules involved in this change may be proteins with sulfhydryl (SH)or disulfide (S-S) residues. Kishimoto and Kanatani (1973) observed that S-S reducing agents, such as dithiothreitol (D'IT) and 2,3-dimercapto-l-propanol(BAL), acted as MISs in starfish oocytes and that SH-blocking agents such as p-chloromercuribenzoate (PCMB) reversibly inhibited 1-MA-induced maturation. Furthermore, the SH content of the proteins isolated from the oocyte cortex rapidly increased following 1-MA treatment (Kishimoto et al., 1976). Opposing this notion, however, is the work on X. laevis by Brachet et al. (1975b), who found that SH-reducing agents were ineffective in inducing maturation but that some organomercurials, such as p-hydroxymercuriphenylsulfonate (PHMPS) and p-hydroxymercuribenzoate (PHMB), acted as potent MISs. Recently, Pays et ul. (1977) found that the SH-oxidizing agent cysteamine was also an effective inducer of maturation and that the effects of both it and the mercurial compounds were reversed by D'IT, while other SH-oxidizing agents, such as diamide and dithionitrobenzene, were ineffective. It should be noted that SH reagents active as MISs do not induce maturation when they are injected into oocytes, suggesting that their sites of action are on the oocyte surface (Brachet et al., 1975b;Pays et al., 1977). The findings discussed above are consistent with the idea that the reaction of the oocyte to a MIS involves a change in the conformation of the surface protein, a change including either the formation or dissociation of S-S bonds. The notion that the initial oocyte reaction to a MIS consists primarily of conformational changes in the surface protein may be of general significance. It has been shown that Chaetopterus oocytes, which have been prevented from maturing following isolation in Ca-free seawater, begin to mature upon treatment with trypsin (Goldstein, 1953; Ikegami et al., 1976) or with PHMPS (Brachet and Denis-Donini, 1977). Similarly, oocytes of U. caupo can be induced to mature by treatment with trypsin (Paul, 1975) or with mersalyl acid (sarylgan) (Johnston and Paul, 1977). The induction of oocyte maturation by proteases has also been successful in Sabellaria (Peaucellier, 1977a) and Spisulu (Ashton,

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1959). All these observations, taken together, suggest that an alteration in oocyte protein character, induced either by proteolytic enzymes or by SH reagents, may be the initial step of maturation. Thus it is possible that the primary action of a MIS is to cause alterations in the conformation of oocyte surface protein. B. THE ROLEOF Ca IONS The induction of oocyte maturation is dependent on the presence of divalent cations in the external medium to various degrees in different animals. Chaetopterus (Goldstein, 1953; Ikegami et al., 1976; Brachet and Denis-Donini, 1977), Spisula (Schuetz, 1975a), and U . caupo (Johnston and Paul, 1977) oocytes fail to initiate maturation in Ca-free seawater. Oocytes of amphibians, such as X . luevis, fail to undergo maturation when cultured in Ca- and Mg-free Ringer’s solution after treatment with progesterone (Merriam, 1971a,b), while the addition of these or other divalent cations, such as Ba and Sr, has been found to support the maturation of oocytes (Wasserman, 1976; Marot et al., 1976). In R. pipiens, oocyte maturation can be induced by progesterone in Ca- and Mg-free Ringer’s solution if the oocytes are exposed to the hormone shortly after isolation from the follicles (Eicker and Smith, 1971b). However, they become unable to respond to the hormone if they are stored for 2 hours or more in Ca- and Mg-free medium at 4°C (Kostellow and Morrill, 1979). In all these cases, it has been observed that oocytes kept in medium lacking divalent cations become capable of initiating maturation upon their return to normal conditions. That is, oocytes are reversibly incapacitated by being deprived of external divalent cations. However, the oocytes of some marine invertebrates, such as starfish (Shirai and Kanatani, 1974; Guerrier et al., 1978) and Sabellaria (Peaucellier, 1977), can be induced to mature in seawater lacking divalent cations. The role of internal Ca ions in maturation has been tested by injecting ethylene glycol bis(Zaminoethy1ether)-N,N’-tetraacetic acid (EGTA) into oocytes. This type of experiment showed that Xenopus oocytes injected with EGTA were unable to initiate maturation following progesterone treatment even when divalent cations were present in the external medium (Wasserman, 1976; Masui et al., 1977). Similarly, starfish oocytes injected with EGTA failed to respond to 1-MA (Guemer et al., 1978; Moreau et al., 1978). Thus it is clear that removal of internal Ca ions from oocytes inhibits their maturation, whether or not maturation is dependent on the presence of divalent cations in the external medium. This observation implies that intracellularCa ions play an indispensable role in the initiation of oocyte maturation. Apparently, oocytes which do not require the presence of Ca in the external medium for maturation possess sufficient endogenous reserves of Ca ions to provide that necessary when the oocytes are stimulated by a MIS. It may be that these oocytes release Ca from an internal

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reservoir upon stimulation by a MIS. However, the initiation of maturation in oocytes which require the presence of Ca in the external medium appears to depend on an influx of ions from the medium following MIS stimulation. The release of Ca ions from the internal reservoir of the oocyte following stimulation by a MIS has recently been examined. O’Connor et al. (1977), using Xenopus oocytes stimulated by progesterone, and Johnston and Paul (1977) using inseminated or trypsin-treated oocytes of U.caupo, demonstrated that oocytes preloaded with 45Carapidly released a considerable amount of Ca into the external medium when they were stimulated to initiate maturation. However, the rate of Ca release soon decreased, returning to a level comparable to that occurring in unstimulated oocytes. Eventually, shortly before GVBD, the rate of efflux of Ca from the preloaded stimulated oocytes became less than that from the unstimulated oocytes indicating a greater degree of sequestration of Ca in the stimulated oocytes. The rate of Ca influx into oocytes was also studied in stimulated and unstimulated oocytes of both species. The data indicate that the amount of 45Cataken up from the external medium increases at a much faster rate in stimulated oocytes than in unstimulated controls, reaching the maximum rate shortly before GVBD. These observations indicate that oocytes induced to mature first release stored Ca ions but, as maturation progresses, sequester Ca ions more actively than those which have not been induced to mature. This may explain an early observation that R. pipiens oocytes which had been ovulated contained more Ca than those in the ovary (Morrill et al., 1971). Marot et al. (1976) showed that Xenopus oocytes undergoing maturation following stimulation by mercurial compounds also accumulated Ca from the external medium, although no significant change in the influx or efflux of Ca was observed in progesterone-stimulated oocytes. It appears likely that oocytes, when stimulated by a MIS, activate a mechanism which mobilizes Ca ions. Direct evidence of this Ca mobilization was recently provided by Moreau et al. (1978), who injected a Ca-sensitive luminescent protein, aequorin, into oocytes. They found that aequorin injected into starfish (M.glacialis) oocytes kept in Ca-free seawater luminesced less than 1 second after the oocytes were exposed to 1-MA. This suggests a rapid release of endogenous Ca, sequestered in the oocytes before their exposure to 1-MA, into the free space in the cell where the aequorin was introduced. When the oocytes were immersed in Ca-free seawater containing aequorin, no light flash was detected following the addition of 1-MA to the seawater, indicating that there was no release of endogenous Ca into the external medium upon stimulation by 1-MA. When EGTA was injected into oocytes before or during the appearance of the Ca peak, the emission of light by previously injected aequorin and GVBD were both suppressed, whereas when EGTA was given after the Ca peak had appeared, GVBD took place.

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A similar approach was used by Belle et al. (1977b) to study Ca mobilization in Xenopus oocytes following MIS stimulation. In this case, however, no significant light emission was observed. This negative result may have been due to inadequate optical properties of the surface of Xenopus oocytes with respect to the detection of light emitted from the inside. The observation of Ca mobilization using the aequorin method in starfish oocytes strongly suggests that an increase in the internal concentration of Ca is a prerequisite for the initiation of maturation. In this respect, the Ca efflux observed in Xenopus and Urechis oocytes following MIS stimulation can be regarded as representing a discharge of excess ions released during the transient Ca surge. An active role for Ca ions in the initiation of oocyte maturation was suggested by classic experiments on parthenogenesis in marine invertebrates such as Hydroida (Pasteels, 1935) and Cumingia (Hollingsworth, 1941). However, the exceedingly high Ca concentrations, usually over 100 mM, required to stimulate the oocytes in these experiments makes doubtful the interpretation that the Ca ions applied to the oocytes acted in a specific way to control cell physiology. In fact, Guerrier et al. (1978) found that starfish (M. glacialis) oocytes isolated from the ovary in Ca-free seawater could be induced to mature without 1-MA stimulation if they were exposed to high Ca concentrations (75-300 mM). Thus it appears that Ca ions applied at high concentrations act as a nonspecific triggering agent which secondarily causes the release of internal Ca. Evidence for a specific role of Ca ions in the initiation of oocyte maturation has, however, been obtained from studies of the effects of certain chemicals known to interfere with the physiological action of Ca in a variety of cells, including neural, muscle, and secretory cells. In Xenopus oocytes, amphiphilic cations, including phenothiazine neuroleptics, tricyclic antidepressants, anorexiants, local anesthetics, P-adrenergic blocking agents such as propanolol and the tertiary amines D200 and D600, have been found to be capable of inducing maturation (Schorderet-Slatkine and Schorderet, 1976; SchorderetSlatkine er al., 1977a). It is known that these agents concentrate in cellular membranes, disturbing their phospholipid turnover and resulting in a concomitant release of Ca from its binding site in phosphatidic acid (Feinstein, 1964; Seeman, 1972). In addition, La ions, known to displace external membrane Ca, thus blocking Ca flux and releasing sequestered internal Ca, also effectively induce oocyte maturation in X . laevis (Schorderet-Slatkineet al., 1976). La ions have also been found to act as a MIS in R. pipiens, but D600 has neither an inhibitory nor an inducing action on oocyte maturation, although both effectively block Ca uptake by oocytes (Kostellow and Momll, 1979). Recently, gammexane, which specifically suppresses Ca mobilization in excitatory cells, has been found to inhibit effectively maturation of progesterone-treated Xenopus

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oocytes (Schorderet-Slatkineet al., 1977b). D600, isoptin, procaine, Mn ions, and methylxanthine all inhibit 1-MA-induced maturation in starfish oocytes, and these agents have also been found to suppress the light emission usually seen following 1-MA stimulation of aequorin-injectedoocytes (Moreau et al., 1978). Finally, oocyte maturation in Chaetopterus can be induced by tetracaine (Ikegami el al., 1976). All these findings, taken together, indicate beyond doubt that the mobilization of Ca is deeply involved in the initiation of oocyte maturation in many species. The mode of Ca action, however, is not yet fully understood. It may differ among species, depending on the specific molecular organization of the oocyte membranes. For instance, the antibiotic, ionophore A23187, which facilitates the transportation of divalent cations across membranes (Reed and Lardy, 1972), acts as a MIS for the oocytes of several species, but under differing ionic conditions. Among marine invertebrates, the ionophore is known to induce oocytes of Nereis (Chambers, 1974), SpisuZa (Schuetz, 1975a), and Chaetopterus (Brachet and Denis-Donini, 1977) to mature, provided that Ca ions are present in the ambient medium. The ionophore can also induce oocyte maturation in X . laevis, if the Mg or Ca ion concentration in the external medium is greater than 10 mM, but it cannot induce R. pipiens oocytes to mature (Wasserman and Masui, ,1975b). In this connection, it is interesting to note the recent finding by Baltus et al. (1977) that fully grown Xenopus oocytes, when cultured in a medium supplemented with an excess of Ca (20 mM) or Mg (40 mM) for 9-10 hours, could initiate maturation without the addition of ionophore or hormones to the medium. Furthermore, they reported that medium-sized oocytes, normally unresponsive to progesterone, initiated maturation in response to the hormone when they were cultured in Ca- or Mg-fortified medium. Although the observations cited above appear to indicate that inward mobilization of Ca or Mg ions from the external medium into oocytes, which can be facilitated by ionophore A23 187, effectively induces maturation, it should be noted that the ionophore cannot induce Xenopus oocytes to mature at excessively high external Ca levels, though its activity is unaffected by high Mg levels (Masui et al., 1977). Peaucellier (1977b) found that Sabellaria oocyte maturation could be induced by the ionophore in the absence as well as in the presence of divalent cations, and that simple treatment with EDTA also induced maturation. Furthermore, Ca-mobilizing agents which have been found to be effective in inducing oocyte maturation in X . laevis had no effect on Sabellaria oocytes at nontoxic doses except for La ions (Peaucellier, 1978). Finally, it should be pointed out that the ionophore is completely ineffective as an inducer of oocyte maturation in starfish (Schuetz, 1975b), even though photometric determination of the intracellular Ca ion level using aequorin has revealed that the ionophore actually induces a sizable Ca surge, to a level 20 times higher than that induced by 1-MA (Moreau et al., 1978). These facts may imply that the transient increase in the intracellular

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level of Ca ions preceding the initiation of maturation must be regulated within an appropriate range in order for the actual process of maturation to take place. The site of action of Ca ions in triggering oocyte maturation may be near the surface of the oocyte. Moreau et al. (1976b) reported that a very small amount of Ca introduced by iontophoresis into areas less than 0.2 mm below the surface induced maturation of Xenopus oocytes, while introduction into areas deeper than that was ineffective. They also found that this effect of iontophoresis was ion-specific; that is, no ions other than Ca were able to induce maturation, and EGTA iontophoresis counteracted the effect of Ca. In this experiment, the effectiveness of Ca iontophoresis depended on the presence of relatively high Mg concentrations (10-20 mM), suggesting a synergism between Ca and Mg in the initiation of oocyte maturation. Furthermore, with respect to the surface action hypothesis, it is interesting to note that La, which may mobilize Ca in oocytes, can induce maturation only when applied to the external surface of Xenopus oocytes, failing to do so if injected into the oocytes (Schorderet-Slatkine et al., 1976). Consideration of these results, together with those from the experiments using Ca iontophoresis, certainly suggests that the site of the reaction of Ca ions with oocytes is near the surface membrane. IN ELECTROPHYSIOLOGICAL PROPERTIES C. CHANGES

1. The Membrane Potential of Immature Oocytes In all animals studied to date, measurements of the electropotential difference between the outside and inside of an oocyte, called the membrane potential, have shown that the inside of a fully grown ovarian oocyte is negatively charged with respect to the outside; that is, the membrane is inwardly negative. Recent studies of the membrane potential in amphibian oocytes have revealed that the resting potential changes when oocytes are isolated from their follicles. In R. pipiens, Ziegler and Mom11 (1977) showed that the membrane potentials of follicle-enclosed and of follicle-free oocytes are -36 5 2 and -77 -+ 2 mV, respectively. Similar results were obtained in X. Zaevis by Wallace and Steinhardt (1977), who reported that the membrane potentials of follicleenclosed and follicle-free oocytes were -27 -t 2 and -65 f 2 mV,respectively. The hyperpolarization caused by removal of the follicles has been attributed to the activation of Na,K-dependent ATPases in the oocyte membrane, since ouabain significantly inhibits the hyperpolarization (Wallace and Steinhardt, 1977). However, in Xenopus it was found that there were oocytes in the ovary which did not undergo hyperpolarization following defolliculation. Thus two types of fully grown oocytes must exist in the ovary. Generally, oocytes which are hyperpolarized by defolliculation are of average size, less than 1.25 mm in diameter, while those whose membrane potential is the same before and after defollicula-

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tion tend to be larger, greater than 1.25 mm in diameter (Wallace and Steinhardt, 1977). Such variations in the physiological properties of oocytes have not been observed in R . pipiens, the oocytes of which grow synchronously in the ovary. Both in Rana (Tupper and Maloff, 1973; Ziegler and Morrill, 1977) and in Xenopus (Wallace and Steinhardt, 1977) it was observed that, when the concentration of K ions in the external medium was increased, the membrane was gradually depolarized, and no difference in the electropotential between the inside and outside was observed when the K concentration was increased to 200 mM. Thus the internal concentration of K ions in the amphibian oocyte may be estimated to be 200 mM. This concentration is significantly higher than that calculated from measurements of K and water content of the oocyte using atomic absorption methods (130-140 mM), as well as that determined by measurements of intracellular K activity with a K-selective electode (106 mM) (Ziegler and Morrill, 1977). Ziegler and Morrill (1977) hence suggested that one factor responsible for this discrepancy might be a nonuniform distribution of K ions in the oocyte, as suggested by Horowitz and his associates (Century and Horowitz, 1974; Horowitz and Paine, 1976). However, substitution of tris(hydroxymethy1 amino)methane (tris) for Na in the external medium causes hyperpolarization (Tupper and Maloff, 1973), indicating that a Na influx contributes to depolarization of the oocyte membrane. Studies of Na ion exchange using radioactive isotopes have shown that there is indeed a constant exchange of Na taking place in the oocytes of R . pipiens (Mom11 et al., 1977b) and X. laevis (O’Connor et al., 1977). When the oocytes are maintained in Ca-free solution, the Na influx is increased, but the K influx is decreased (Ecker and Smith, 1971b; Tupper and Maloff, 1973; Morrill et al., 1977b). Consistent with this observation, marked depolarization of the membrane has been observed in oocytes following the removal of Ca from the external medium in the case of both Rana (Tupper and Maloff, 1973) and Xenopus (Bell6 et al., 1977b). These results strongly suggest that the selective permeability of amphibian oocytes to K and Na ions is highly dependent on external Ca. Furthermore, Bell6 et a1. (1977b) showed that the depolarization induced by deprivation of external Ca exceeded that induced by ouabain, indicating that a major portion of Na-K transport in amphibian oocytes was regulated by Ca. In the starfish Nordora punctiformis (Hagiwara and Takahashi, 1974) and A. pectinifera (Miyazaki et al., 1975a) it was found that, as in X. Laevis, two types of oocytes, differing in membrane potential characteristics, existed in an oocyte population. The depolarized type had a low membrane potential, -14.6 -+ 5.5 mV, and the hyperpolarized type had a high membrane potential, -72.7 +- 3.6 mV. The membrane potential of oocytes of the depolarized type, however, can be induced to become more negative by replacing Na in the seawater by tris,

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suggesting that the reduced (i.e., less negative) membrane potential of oocytes of this type is due to a change in permeability to Na ions. Miyazaki et a2. (1975a) found that, in the hyperpolarized type, the resting membrane potential was virtually unaffected by changing the levels of Na, Ca, and Mg ions in the external medium but was strongly affected by changing the K levels, and in fact complete depolarization occurred when the K concentration was raised to 196 mM. This suggests that the internal K concentration of oocytes is about 200 mM. Thus it appears that the membrane potential of starfish oocytes is due almost exclusively to K diffusion across the membrane and, as opposed to the situation in amphibian oocytes, is not influenced by external divalent cations. Measurements of steady currents passing through the oocyte membrane at various voltage levels have indicated that there is an ingoing rectification (i.e., current flowing into the oocyte) at voltages near the resting potential (-70 mV), an outgoing rectification (i.e., current flowing out of the oocyte) above -20 mV, and a high negative resistance at voltages between these two rectification regions (Fig. 9). Shen and Steinhardt (1976), studying oocytes of P. miniata, noted that two types of oocytes distinctly different in resting membrane potential existed, as in Asterias and Xenopus. The depolarized type showed membrane potentials ranging from - 10 to -25 mV, and the hyperpolarized type showed potentials ranging from -65 to -90 mV. The membrane potentials of both types of oocytes were also influenced by the concentration of K ions in the external medium. However, these investigators found the oocytes of this species to exhibit peculiar electrophysiological characteristics as compared with those of Asterias. First, since the membrane could not be completely depolarized until the external K concentration approached 400 mM, the internal K concentration of the oocytes was estimated to be about 400 mM. This value is extraordinarily high compared to that of the oocytes of other species. Second, the membrane potential did not change when the Na and Mg concentrations in the external medium were changed but became more negative-that is, the membrane was hyperpolarized when the Ca concentration was increased or when the C1 concentration was decreased. In addition, ouabain had no effect on the value of the membrane potential. Measurement of the steady current passing through the oocyte membrane at different voltages indicated that there was a relatively high resistance near the resting potential level and above 0 mV, and a negative resistance between these two levels (Fig. 9). Data for mammalian oocytes are rather limited. Although some work has been carried out on the membrane potential of metaphase I1 oocytes of the mouse, which has indicated properties similar to those observed in amphibian oocytes (Powers and Tupper, 1974, 1975), no detailed studies on ovarian oocytes have been published.

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n

FIG.9. Current ( I ) and voltage ( V )relationship in immature oocytes and maturing oocytes. A,, Immature oocyte ( A . pectinifera. 0 minute after hormone treatment); A2, maturing oocyte (A. pectinifera. 30 minutes after treatment); P,,immature oocyte (P.mininta, 0 minute after treatment); P,, maturing oocyte ( P . miniata, 30-40 minutes after treatment); M,. maturing oocyte (Mus musculus. metaphase 11). A, and AZ (Miyazaki et al., 1975b); P, and P, (Shen and Steinhardt, 1976); M2 (Powers and Tupper, 1974). Note an increase in the slope of curves in the inward current region, indicating the disappearance of the inward rectification and an increase in the resistance to an inward directed current, occurring during the course of maturation.

From the brief survey of the literature summarized above, it is clear that, before initiating maturation, an oocyte is polarized negatively inward, and this polarization is mainly due to a selective permeability to K ions. However, it should be emphasized that the membrane potential appears to be controlled in a different manner in different species. In amphibians, Ca appears to control Na and K transport, both of which are involved in establishing the resting potential, whereas in starfish, except Patiria, the regulation of the membrane potential is due primarily to K ions.

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2. Changes in Electrical Properties during Maturation Measurements of the membrane potentials in amphibian oocytes have revealed that fully grown oocytes in ovarian follicles are hyperpolarized compared to ovulated oocytes in the uterus (Maeno, 1959; Mom11 and Watson, 1966). This result suggests that depolarization of the membrane occurs during maturation. The membrane potential of progesterone-treatedoocytes of Xenopus (Belle et al., 1976, 1977b; Moreau et al., 1976c; Wallace and Steinhardt, 1977) and Rana (Ziegler and Momll, 1977) becomes less negative, usually to less than -20 mV and sometimes reaching 0 mV, before GVBD. A similar depolarization of the membrane has also been observed when maturation is induced by ionophore A23187 or PHMB (Moreau et al., 1976~).However, very large oocytes of the depolarized type do not undergo further depolarization following progesterone treatment (Wallace and Steinhardt, 1977). When 1-MA is applied to oocytes of the starfish A . pectinifera, the membrane becomes depolarized within 15 minutes (prior to GVBD) and reaches a steady potential ranging from -20 to 0 mV (Miyazaki et al., 1975b). However, in this species, the membrane later hyperpolarizes once again. In P. miniata, the membrane potential becomes more negative with the initiation of maturation, causing hyperpolarization from -80 to -95 mV (Shen and Steinhardt, 1976). Recently, the membrane potential change in mouse oocytes during maturation was examined by Powers (Biggers et al., 1977; Powers and Biggers, 1976). It was found that the resting potential in dictyate stage oocytes was about -35 mV and was reduced to -25 mV at the time of GVBD. Further depolarization occurred as the oocytes proceeded to metaphase 11, at which time it reached - 14 mV (Powers and Tuppers, 1974, 1975). During maturation, membrane resistance is also changed. In Xenopus oocytes the resistance to ingoing current increases well before GVBD, shortly before depolarization takes place (Belle et al., 1977b). Membrane resistance, however, then decreases just before GVBD occurs (Moreau el al., 1976~).As seen in Fig. 9, during starfish ( A . pectinifera) oocyte maturation, the current-voltage slope (resistance) becomes steeper in the rectification regions, especially in the inward rectifying region near the resting potential level, and the negative resistance previously observed at voltages above it disappears before GVBD begins (Miyazaki et al., 1975b). A similar change in membrane resistance has been observed in Patiria oocytes. In this species, although no depolarization of the membrane occurs during maturation, membrane resistance at voltages near the resting potential and in the inward rectifying region increases before GVBD begins (Shen and Steinhardt, 1976). Mouse oocytes at metaphase I1 also exhibit a steep voltage-current slope over a wide range of voltages (Powers and Tupper, 1974).

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3 . Changes in K and Nu Ion Permeability The changes in the electrophysiologicalproperties of oocytes during the initial period of maturation described above may be interpreted to be a result of changes in the membrane permeability to K ions, and possibly also to Na ions, since the resting potential of the oocyte membrane depends mainly on its selective permeability to Na and K, as discussed at the beginning of this section. Studies by O’Connor et al. (1977) of ion transport in Xenopus oocytes, using radioactive K and Na, have shown that both the influx and the efflux of K ions are considerably increased shortly (2 hours) after progesterone treatment; however, this is followed by a continuous decrease beginning before GVBD. However, the rate of Na influx steadily increases, starting almost immediately after exposure to progesterone, but subsequently decreases following GVBD. Na efflux also increases, and it continues to do so for several hours after progesterone treatment. In Xenopus oocytes, the permeability ratio between Na and K ions (PNa/PK), calculated from ion flux rates, first decreases from 0.10 to 0.05 in the period following progesterone stimulation and then increases to 0.43 at GVBD (O’Connor et al., 1977). Similarly, in R. pipiens the rate of Na uptake increases after progesterone treatment but declines to the unstimulated level after GVBD, whereas the rate of K uptake steadily declines (Ziegler and Morrill, 1977), producing overall an increase in the total Na content and a decrease in the total K content per oocyte. However, since oocyte water content increases from 50% to 65% of the total weight of the oocyte when it reaches metaphase 11, the Na concentration does not rise proportionally to its uptake, while the decrease in K concentration is augmented, resulting in depolarization. The depolarization of mouse oocytes during maturation may also be ascribed to an increase in the permeability to outgoing K ions, as suggested by Powers and Biggers (1976). In starfish (A. pectinifera) 1-MA-treated oocytes reduce the conductance of ingoing K current (Miyazaki et al., 1975b). In batstar (Patiria) oocytes, however, the hyperpolarization induced by l-MA is augmented in Na-free media, suggesting an increase in inward-directed Na conductance during maturation, while the internal K concentration is rather increased and no marked change occurs in the ratio between Na and K ion permeability (PNJPK) (Shen and Steinhardt, 1976). This is in contrast to the trend found in oocytes of other animals, in which all evidence suggests a decrease in internal K concentration during maturation. Except for the case of Patiria, changes in ion transport during maturation in various animals may be summarized as follows. The depolarization of the membrane during the initial period of maturation is accompanied by a loss of selective K permeability and perhaps also by an increase in inward directed Na diffusion, which result in a decrease in the internal K concentration and an increase in Na concentration in the oocyte.

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4. Significance of Changes in Electrophysiological Properties

The question of whether changes in ion flux into oocytes are a necessary step for the initiation of maturation has been examined by several groups of investigators. Dorke et al. (1976b) showed that the maturation of starfish oocytes following 1-MA treatment was inhibited by treatment with tetraethylammonium PEA), which is known to block K channels in the cell. In the mouse (Powers and Biggers, 1976) it was found that treatment with valinomycin, which acts as an ionophore for monovalent cation transport, induced hyperpolarization of the oocytes and strongly retarded the progression of maturation. This effect was interpreted as being a result of the increase in membrane permeability to K ions induced by the ionophore, which may temporarily increase internal K levels. In accordance with this interpretation is the finding by Baltus et al. (1977) that Xenopus oocytes exposed to valinomycin in a K-free medium could initiate GVBD without stimulation by a MIS. In their experiment, the K ion content of gocytes was reduced by 40% as compared with that of controls kept in normal Ringer’s solution. Vitto and Wallace (1976) found that removal of K ions from the external medium or treatment with ouabain to inhibit the selective permeability to K of Xenopus oocytes facilitated their maturation. Conversely, maturation of R. pipiens treated with progesterone can be inhibited by increasing the K concentration in the external medium in the absence of divalent cations (Ecker and Smith, 1971b). All this evidence is consistent with the view that a reduction in internal K concentration following the loss of selective K permeability is a necessary step in the initiation of maturation by a MIS. The question of whether changes in electrical properties of oocytes are necessary for them to initiate maturation has been examined. Doree et aE. (1976b), who voltage-clamped starfish oocytes following 1-MA treatment, found that oocytes clamped at -40 mV were unable to undergo maturation. They also observed that depolarization of the membrane induced in the starfish oocyte following a short treatment with 1-MA (2.5 minutes) was reversible and that oocytes with the membrane potential reversed did not initiate maturation. These results appear to suggest a correlation between the depolarization induced by a MIS and the initiation of maturation. However, there is evidence that apparently contradicts this view. In starfish (M.glacialis), oocyte maturation can be induced with 1-ethyladenine, as well as with 1-isopropyladenine, without changing the membrane potential of oocytes (Doree et al., 1976b). Moreover, the fact that Patiria oocytes can be induced to mature by 1-MA treatment without undergoing any depolarization, but rather hyperpolarization, may present the most serious challenge to the theory that assumes the existence of a strict causal relationship between changes in the membrane potential and the initiation of maturation. The observation that a reduction in Na concentration of external media brings

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about a retardation of maturation in Xenopus (O’Connor et al., 1977) and mouse (Powers and Biggers, 1976) oocytes suggests that not only K but also Na ions play a role in the regulation of ionic conditions necessary for oocyte maturation. Therefore, it may be assumed that the intracellular ionic conditions of an oocyte required for the initiation of maturation can be fulfilled by changing the membrane conductance to these two ion species without causing observable depolarization of the membrane potential.

5. Roles of ATPase Changes in the rate of K and Na ion flux occurring early in maturation suggest that corresponding changes in membrane ATPase activity occur in oocytes, which regulate the transport of these ions. Mom11 et al. (1974) found that the ATPase activity of whole homogenates of R. pipiens oocytes increased sharply prior to GVBD. They noted that this rise in ATPase activity appeared to coincide in time with the characteristic membrane potential change. However, they also found that cycloheximide treatment of oocytes had no effect on the rise in ATPase activity, while it completely inhibited GVBD. It may be that oocyte protein which must be synthesized for GVBD to occur is not related to the measured ATPase activity. According to Mom11 et al. (1971, 1974), before the initiation of maturation R. pipiens oocytes possess ATPases which can be activated by Mg, Na, and K and strongly inhibited by ouabain and by strophantidin, whereas ATPases in oocytes at metaphase I1 are activated by Ca in the presence of Mg. However, in X . laevis, Pays et al. (1977) found that ATPase activities detectable both on the surface of intact ovarian oocytes and in homogenate preparations were virtually unaffected by ouabain and strongly activated by Ca and Mg but not by Na or K. This observation apparently contradicts those of Morrill et al. (1971). Pays et al. (1977) also found that the ATPase activities of homogenates of progesteronetreated oocytes and of oocytes treated with PHMPS sharply rose before GVBD took place, while the surface ATPase activity of intact oocytes did not show any significant change during the course of maturation. In view of these results, it may be premature to speculate on the correlation between changes in the ATPase activity of oocytes and in their ion transport activities during the process of maturation.

V. Cytoplasmic Control of Oocyte Maturation A. MATURATION-PROMOTING FACTOR 1. Origin and EfSects

In amphibian and starfish oocytes it has been repeatedly shown, as reviewed in Section IV,A, that a MIS is effective only when it is applied externally so as to

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act on the mcyte surface. In addition, discovery of the early change in the membrane potential of oocytes following MIS Stimulation strengthens the View that the initial maturational event takes place in the plasma membrane of the wcyte. Therefore it appears that the action of a MIS is primarily directed toward the surface membrane, as opposed to the cytoplasm or the nucleus (the GV). A logical consequence of this notion is that the signal given by a MIS to the oocyte surface must be transmitted to the nucleus by a cytoplasmic messenger. Whatever the mechanism of transmission, there is evidence that transmission of the signal through the cytoplasm of the oocyte occurs. Iwamatsu (1966), using fish (medaka) oocytes, demonstrated that GVBD could be inhibited when the GV was displaced from the hyaline cytoplasm near the surface of the animal pole into the yolk-filled vegetal hemisphere. Masui (1972) obtained similar results with R. pipiens oocytes. In his experiment oocytes were constricted in the equatorial zone to various degrees, using thread, after the GV had been displaced into the vegetal hemisphere by centrifugation. The oocytes were then exposed to progesterone. The occurrence of GVBD when the GV was in the vegetal hemisphere was considerably delayed compared to when it was in the animal hemisphere, and the frequency of GVBD was markedly decreased as the cytoplasmic connection between the animal and vegetal hemispheres was made narrower by increased constriction. Thus it seems likely that the cytoplasmic activity causing GVBD develops first in the animal hemisphere and then spreads to the vegetal hemisphere, especially in view of the fact that the animal hemisphere is more sensitive to progesterone than the vegetal hemisphere (Cloud and Schuetz, 1977). It was at one time thought that oocyte maturation could be initiated by the direct action of gonadotropin on the GV. Dettlaff and her associates (1964), using amphibians, demonstrated that substances in the GV of follicleenclosed oocytes treated with gonadotropin became capable of breaking down the GV itself when injected into untreated oocytes. They cautioned, however, that the active factor inducing GVBD might be a cytoplasmic product, since the material transferred into the recipient oocytes contained a small amount of cytoplasm surrounding the GV. Gurdon (1967) found that gonadotropin failed to induce GVBD when it was injected directly into oocytes, suggesting that its action was indirect. Evidence of indirect hormonal action on the GV has been produced by Masui and Markert (1971), who demonstrated that the cytoplasm of progesteronestimulated R. pipiens oocytes exhibited the ability to induce GVBD when injected into untreated oocytes before the donor oocytes underwent GVBD. They also found that this cytoplasmic activity appeared in oocytes treated with progesterone after their GV had been removed. It thus became clear that the cytoplasm of progesterone-treated oocytes developed the ability to induce GVBD independently of the presence of the GV. Further observations of the oocytes induced to

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undergo GVBD by the injection of cytoplasm from progesterone-treated oocytes have revealed that the recipient oocytes undergo further maturational changes, including development of surface contractility, condensation of chromosomes to the metaphase state, and polar body formation. Thus it has been postulated that a cytoplasmic factor responsible for promoting maturational events in general appears in the oocyte as a result of its surface stimulation by a MIS; this cytoplasmic factor has been designated maturation-promoting factor (MPF) (Masui and Markert, 1971). The GV-independent origin of MPF strongly suggests that it is a product of cytoplasmic activities which do not require genomic function. MPF has been found in oocytes of other amphibian species, such as X. laevis (Schorderet-Slatkine and Drury, 1973) and Ambystoma mexicanum (Reynhout and Smith, 1974), and also in the sturgeon (Dettlaff et al., 1977). In starfish oocytes, Kishimoto and Kanatani (1976) found that the cytoplasm of 1MA-treated oocytes, injected into untreated oocytes, was capable of inducing maturation of the recipient oocytes, and that the maturational events induced in the recipients followed the normal time course of 1-MA-treated oocytes. Reynhout and Smith (1974) showed that oocytes of a given species of amphibians could be induced to mature by microinjection of an appropriate amount of cytoplasm from maturing oocytes of other species. This finding demonstrates that the MPF of oocyte cytoplasm is effective in other species, suggesting that the active factor is not species-specific. Similarly, Kishimoto and Kanatani (1977) demonstrated that the interspecific transfer of cytoplasm from 1-MA-treated oocytes among different species of starfish resulted in the induction of maturation in the recipients. MPF can be found in the cytoplasm of oocytes induced to mature by MISS other than steroid hormones. It has been detected in Xenopus oocytes treated with mercurials (Brachet et al., 1975b), ionophore A23187 (Wasserman and Masui, 1975b), valinomycin (Baltus et al., 1977), and La ions (SchorderetSlatkine et al., 1976). These results suggest that MPF is not a product of the reaction between receptor molecules on the oocytes and steroid hormones. The effects of MPF on young oocytes at various stages of oogenesis which are unable to respond to MIS have been tested by Hanocq-Quertier et al. (1976) using X. laevis. It was found that MPF obtained from maturing oocytes caused GVBD and chromosome condensation in these young oocytes, though meiotic division could not take place because of failure to form a spindle. Similar effects of maturing oocyte cytoplasm on small, immature oocytes have been demonstrated in mice by Balakier (1978), who fused maturing oocyte cytoplasm with small oocytes, using inactivated Sendai virus, and observed GVBD as well as chromosome condensation to the metaphase state. These results clearly indicate that MPF is an ubiquitous cytoplasmic factor which promotes meiotic changes in the nucleus.

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The effect of MPF has been shown to be dose-dependent by Masui and Markert (1971) using amphibian oocytes, and by Kishimoto and Kanatani (1976) using starfish oocytes. Both these groups found that the frequency with which GVBD occurred in oocytes following the injection of maturing oocyte cytoplasm was almost linearly proportional to the volume of cytoplasm injected. This dose dependence of the effect of MPF has enabled us to assay its activity in a semiquantitative fashion.

2 . Time Course In amphibian oocytes, MPF appears shortly before GVBD and attains its maximum activity at GVBD. The activity remains at a high level until the oocytes are activated, at which time it begins to decrease rapidly (Masui and Markert, 1971). However, it should be emphasized that MPF activity can be detected even in the cytoplasm of cleaving blastomeres, although by this time it is quite weak (Masui and Markert, 1971). In starfish (A. pectinifera), Kishimoto and Kanatani (1976) found that MPF activity appeared 13 minutes after I-MA stimulation of oocytes. It reached its highest level between 20 and 40 minutes and then declined rapidly, disappearing by 80 minutes after hormone treatment. Since GVBD and formation of the second polar body take place 30 and 80 minutes after treatments, respectively, in the oocytes of this species, the change in MPF activity relative to meiotic progression shows a similar time course in Rana and Asterias; that is, MPF rises during karyokinesis and falls during pronuclear formation. The maturation of Xenopus oocytes induced by the injection of maturing oocyte cytoplasm containing MPF cannot be inhibited by protein synthesis inhibitors (Wassennan and Masui, 1975a; Drury and Schorderet-Slatkine, 1975). The time at which the maturation process becomes resistant to the inhibition of protein synthesis coincides with the time at which MPF activity can first be detected in the oocytes (Wasserman and Masui, 1975a). The time required for oocytes to develop MPF activity, measured from the time of progesterone stimulation, is approximately 65% of the time required to initiate GVBD. Since MIS-induced maturation of amphibian oocytes is always inhibited by protein synthesis inhibitors (Section III,C), it seems likely that the process preceding the appearance of MPF is the one which requires protein synthesis. Thus Wasserman and Masui (1975a) hypothesize that the process leading to the initial increase in MPF activity involves synthesis of a new protein, called the “initiator,” but that the action of MPF to induce GVBD does not. Furthermore, MPF action does not appear to be dependent on Ca ions. Masui et al. (1977) showed that the injection of EGTA into progesterone-stimulated oocytes did not inhibit their maturation when the injection took place after MPF had appeared in the oocytes. Guerrier et al. (1976) found that maturation of Xenopus oocytes

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induced by MPF injection was also resistant to proteolytic enzyme inhibitors such as antipain and leupeptin, both of which inhibit the maturation of progesterone-treated Xenopus oocytes and 1-MA-treated starfish oocytes (Clark and Kanatani, 1975). Thus it may be concluded that the action of MPF involves neither the synthesis nor the degradation of proteins.

3. Amplification MPF has been found to develop in oocytes induced to mature by the injection of cytoplasm from maturing oocytes. In order to examine the mechanisms which underly this secondary development of MPF activity in the recipient oocytes, a serial transfer of cytoplasm was carried out in R. pipiens oocytes (Masui and Markert, 1971). In this experiment, 60 nl of cytoplasm, which represents about 3% of the volume of an oocyte, was transferred from progesterone-treateddonor oocytes to untreated recipient oocytes, and then from these first recipients to the second recipients and from the second to the third, at 24-hour intervals. The cytoplasm thus transferred was able to induce maturation in the recipients after every transfer with similar frequencies, ranging from 75 to 90%, despite the fact that the cytoplasm of the original progesterone-treateddonors was clearly extensively diluted by the serial transfers. Therefore the cytoplasm transferred appears to stimulate development of MPF in the recipient oocytes, increasing its activity to the same level as that of the donor within the following 24 hours. Based on this observation, it has been hypothesized that MPF is produced by autocatalytic amplification (Masui and Markert, 1971). Similar experiments have been carried out with oocytes of X . laevis (Reynhout and Smith, 1974; Drury and Schorderet-Slatkine, 1975), the starfish A . pectinifera (Kishimoto and Kanatani, 1976), and the sturgeon Acipenser stellatus (Dettlaff et al., 1977). In all these experiments, cytoplasm of maturing oocytes could be serially transferred to recipient oocytes as many as 5 to 10 times without any resulting loss of MPF activity. In order to examine the requirement of the secondary development of MPF activity for protein synthesis in recipient oocytes injected with MPF, experiments involving the transfer of cytoplasm from progesterone-treated oocytes into a series of cycloheximide-treated recipients have been carried out. Drury and Schorderet-Slatkine(1975), using X . laevis oocytes, transferred the cytoplasm at 2-hour intervals. Their results showed that the frequencies with which recipient oocytes underwent GVBD following cytoplasmic injection declined during the transfer, and that GVBD could not be induced after three successive transfers. In contrast to this result, Wasserman and Masui (1975a), also using Xenopus oocytes, showed that serial transfers caused no appreciable decrease in the frequency of GVBD among cycloheximide-treated recipient oocytes when the transfers were carried out at 7-hour intervals. The latter result was recently confirmed by Dettlaff et al. (1977), using X . laevis oocytes. In their experiment, each cyto-

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plasmic transfer was carried out after the donoroocyte had undergone GVBD, and it was found that high MPF activity could still be detected in the cycloheximidetreated recipient oocyte cytoplasm even after cytoplasm was transferred five times. The experiments cited here were performed under similar conditions except for the difference in the duration of the interval between successive transfers. The amount of cytoplasm injected in each transfer ranged from 4 to 5% of the average volume of the oocyte, and the doses of cycloheximide applied to recipient oocytes were found to be sufficient to inhibit more than 90%of the protein synthesis occurring in untreated oocytes undergoing maturation. Therefore it may be that the apparent discrepancy between the results of the first experiment and those of the second and third can be reconciled if the rate of MPF amplification is taken into account. Since cycloheximide-treated recipient oocytes may take more time than untreated recipients to amplify MPF to the same level as that in the donors, it is reasonable to expect that a continuous decrease in MPF activity might result from successive cytoplasmic transfers taking place at short intervals insufficient for the cycloheximide-treated oocytes to produce a maximum concentration of MPF. In starfish and mammals, it may be supposed that the MPF amplification process, if it occurs, is independent of protein synthesis, since in these animals oocyte maturation is not inhibited by protein synthesis inhibitors up to metaphase 1 (see Section 111,C). However, it has been reported that MPF-induced maturation in sturgeon oocytes is sensitive to protein synthesis inhibitors (Dettlaff et al., 1977). In this fish, although MPF amplification as well as GVBD fails to occur following injection of maturing oocyte cytoplasm into recipients treated with cycloheximide, the oocytes acquire the ability to undergo cortical changes, such as vitelline membrane elevation and CGBD, following activation. 4. Nature In order to determine the cytoplasmic localization of MPF in R. pipiens oocytes, Masui (1972) assayed MPF activities in different fractions of oocytes stratified by a mild centrifugal force. In his experiments, oocytes which had been induced to mature by progesterone were placed in the interface between a 40% Ficoll solution and Ringer’s solution and centrifuged. The cytoplasmic contents of the oocytes separated into five layers-the lipid, fluid hyaline, gel hyaline, pigment, and yolk layers. MPF activity was found mainly in the fluid hyaline and gel hyaline layers. A similar experiment with starfish oocytes treated with 1-MA has recently been carried out by Kishimoto and Kanatani (1977), who found that MPF activity was primarily localized in the hyaline layer, which consists of multivesicular bodies, Golgi apparatus, and homogeneous cell sap. The extraction of MPF from amphibian oocytes was hampered by unexpected difficulties. MPF activity is so unstable that it is easily lost if the oocytes are homogenized. Baltus et al. (1973), however, found that extracts from homoge-

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YOSHIO MASUI AND HUGH J. CLARlCE

nates of Xenopus oocytes exhibited the ability to induce atypical reactions, called pseudomaturation, in oocytes injected with the extracts. Electron microscope studies by Steinert et al. (1974) revealed that the pseudomaturation thus induced involved lobulation of the cortical cytoplasm, hyperdevelopment of internal membranes, and folding and disruption of the nuclear envelope following the migration of the GV toward the animal pole. These pseudomaturational events were not followed by meiotic progression, however. In order to avoid the adverse effect of homogenization on MPF activity a new extraction procedure was developed (Masui, 1974; Wasserman and Masui, 1976). This procedure consists of a rapid crushing of oocytes in a tube by direct application of a centrifugal force to the oocytes, followed by a quick separation of the extract from the particulate fractions. Wasserman and Masui (1976) found that MPF remained active in R. pipiens oocyte extracts thus prepared with a phosphate-buffered sucrose solution containing NaCl and MgSO,. Furthermore, they found that the activity became relatively stable when EGTA was added to the extraction medium, and that the presence of Mg was essential for its maintenance; the presence of Ca at a concentration as low as M rapidly inactivated MPF. These investigators suggest that MPF activity is associated with macromolecules of 4, 13, and 30S, which are heat-labile, protease-sensitive, and RNase-resistant. Further stabilization of MPF was recently achieved by Drury (1978), who extracted MPF from homogenates of Xenopus oocytes using a new extraction medium which consisted of a glycerophosphate-buffered sucrose solution containing NaF or ammonium molybdate. The presence of these chemicals is essential for stabilizing MPF activity. Furthermore, he also found that MPF was irreversibly destabilized by dilution in the absence of ATP. These results may indicate that MPF activity is maintained by phosphorylation of its molecules, and that NaF and ammonium molybdate, which are known to be inhibitors of phosphatases, may act as inhibitors of dephosphorylation of MPF, while ATP may enhance its phosphorylation. Gel filtration experiments by Druly (1978) have indicated that MPF is a macromolecule having a molecular weight between 0.6 and 1.0 x lo6 daltons. Furthermore, he found that the molecules could be sedimented from Xenopus oocyte extracts by a 30% saturated solution of (NH4)$04. Thus tentatively the nature of MPF may be assumed to be that of a phosphoprotein. B . PHOSPHORYLAT~ON OF PROTEINS 1. Phosphokinesis The characteristics of MPF revealed by recent experiments appear to point to a key role for protein phosphorylation in oocyte maturation. According to Monill

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and Murphy (1972), incorporation of 3'P into the protein of R. pipiens oocytes rapidly increases during maturation. Maller et al. (1977) found that protein phosphorylation of Xenopus oocytes rose sharply a few hours after progesterone treatment and reached its highest level at the time of GVBD. The protein phosphorylated during oocyte maturation is not yolk protein (Wallace, 1974). According to Maller et al. (1977), the predominant fraction is a protein molecule of 5.5 x lo4daltons. These workers note that the rise in protein phosphorylation in individual oocytes after progesterone treatment is always followed by GVBD, suggesting a close relationship between protein phosphorylation and GVBD. Further, they found that protein phosphorylation occurred in enucleated oocytes treated with progesterone and that not only progesterone, but also the injection of maturing oocyte cytoplasm, induced a burst of protein phosphorylation in normal oocytes which was followed by GVBD. Maller et al. (1977) also showed that protein phosphorylation induced by progesterone was inhibited by cycloheximide, whereas that induced by cytoplasm injected from maturing oocytes was not. Since protein synthesis of the oocytes had been suppressed by 95%, this result clearly indicates that phosphorylation occurred on preexisting protein, though its initiation by progesterone may require protein synthesis. These features of protein phosphorylation, which are quite similar to those of MPF activity, may be indicative of a close relationship between protein phosphorylation and the activation of MPF molecules. A similar pattern of protein phosphorylation has been discovered by Guerrier et al. (1975) in 1-MA-stimulated oocytes of M. glacialis. In this member of the starfish family, protein phosphorylation in the oocyte increases dramatically 5 minutes after hormone treatment. Their results further indicate that there is a differential distribution of phosphokinase activities in the oocyte, that is, histone and nonhistone phosphokinase activities are detected in the soluble and particulate fractions, respectively. Later experiments by Guerrier et al. (1977) revealed that protein phosphorylation following 1-MA treatment first increased in the cortex where it reached a maximum 8- 10 minutes after hormone treatment, but it increased rather slowly in the endoplasm, reaching a maximum 30 minutes after treatment, after which phosphorylation activity decreased until it reached control levels 1 hour after treatment. Interestingly, the time course of the phosphorylation activity appeared to follow that charted by Kishimoto and Kanatani (1976) for MPF. There is a close relationship between the level of protein phosphorylation in a given oocyte population and the frequency with which GVBD is induced in that population. Furthermore, various maturation inhibitors, such as SH oxidants, inhibit protein phosphorylation in oocytes treated with 1-MA (Guerrier et al., 1977). That oocyte maturation is induced by stimulation of protein phosphokinase is a possible interpretation of the results of Wiblet et al. (1975) in Ambystoma mexicanum and of Moreau et al. (1976a) in X . laevis, both of whom injected a

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phosphokinase preparation obtained from heterologous tissues into oocytes and succeeded in inducing GVBD in the recipients.

2. Role of CAMP The phosphorylation of certain proteins has been shown to be dependent on the level of cAMP in various types of cells. Thus it appears quite likely that changes in protein phosphorylation activity occurring in an oocyte during the course of maturation are correlated with changing levels of cAMP in the oocyte. Early experiments by Pays-de Schutter et al. (1975) to determine the levels of cyclic nucleoside monophosphates (CAMPand cGMP) in Xenopus oocytes revealed no marked differences between oocytes in the ovary and those at metaphase 11. However, the work of Speaker and Butcher (1977), using R. pipiens, showed that cAMP and cGMP levels, which are 0.6 and 0.8 pM respectively in fully grown ovarian oocytes, decrease to about half that 5 hours after hormone treatment, but then increase in the next 5 hours until shortly before GVBD occurs. The cAMP level decreases again after GVBD, reaching a minimum level when the first polar body is given off, and then returns to the same level as that of ovarian oocytes when they reach metaphase 11. The cGMP level, however, does not change as markedly as the cAMP level during the post-GVBD maturation period. Similar changes in cAMP levels in follicle enclosed oocytes of R. pipiens have been reported by Mom11 et al. (1977a). The decrease in cAMP level immediately following oocyte stimulation may be necessary for the initiation of maturation. As indicated in Section II,E, in mammals, high concentrations of dbcAMP inhibit maturation of both follicle-free oocytes and those enclosed in follicles. In amphibians, O’Connor and Smith (1976) showed that maturation of Xenopus oocytes failed to occur when cAMP degradation was inhibited by a xanthine derivative, theophylline, but this inhibitory effect was not observed when the chemical was injected into the oocytes. Morrill et al. (1977a), using follicle-enclosed oocytes of R. pipiens, reported that dbcAMP and theophylline inhibited maturation, without suppressing the protein synthesis stimulated by progesterone, when these chemicals were applied to oocytes incubated in a Ca-containing medium. These results, though difficult to interpret at present, seem to imply that a decrease in the cAMP level of oocytes is required for the initiation of maturation. Corroborating this notion is the recent finding by Godeau et al. (1977) that cholera toxin, known to activate cellular adenyl cyclase to increase cAMP levels in the cell, antagonizes the action of progesterone in Xenopus oocytes, thus inhibiting their maturation when applied externally. In starfish, it is known that xanthine derivatives have an inhibitory effect on the maturation of 1-MA-treated oocytes (Dorke et al., 1976a). It has been pointed out that cAMP derivatives added to mammalian oocyte culture do not interfere with general biochemical activities, such as protein synthesis (Stem and Wassarman, 1974; Schultz and Wassarman, 1977a,b) or

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respiration (Magnusson and Hillensjo, 1977; Magnusson et al., 1977; Hillensjo et al., 1977), of oocytes occurring prior to the initiation of maturation, and that theophylline applied to amphibian oocytes similarly does not interfere with protein synthetic activity prior to the onset of maturation (Morrill et d.,1977a). Rather, these compounds affect only the changes in biochemical activities which accompany the initiation of maturation. Therefore it appears likely that the maturation-inhibiting effect of these chemicals is brought about by raising cAMP levels in oocytes. If so, a high level of cAMP in ovarian oocytes probably prevents them from altering their biochemical activities in a manner which would initiate maturation. It has been shown beyond doubt that the role of CAMPin cellular activities lies mainly in regulation of the activities of protein phosphokinases. Maller and Krebs (1977) demonstrated that the catalytic (C) subunit of CAMP-dependent protein kinase, when injected into X. laevis oocytes treated with progesterone, inhibited the maturation of these oocytes, whereas the regulatory (R) subunit, as well as the protein inhibitor (I) of the enzyme, when injected into untreated oocytes, triggered the initiation of maturation. Since C subunits increase CAMP-dependent protein phosphorylation in the cell, while R subunits and I protein inhibit it by interacting with the C subunits, Maller and Krebs (1977) hypothesize that a high level of cAMP liberates C subunits from binding with R subunits and thus enhances the protein phosphorylation responsible for preventing oocytes from maturing. Conversely, inhibition of the C subunit, either by a decrease in the cAMP level or by the addition of R subunits or I protein, stops this protein phosphorylation and releases the oocytes from arrest at the diplotene stage. Ozon et al. (1978), using X . laevis and Discoglossus pirtus, demonstrated that the injection of I protein into oocytes initiated maturation. These experiments seem to indicate that an alteration in the pattern of protein phosphorylation in oocytes is the factor initiating their maturation. Thus the decrease in the CAMP level in oocytes in response to a MIS may lead to an alteration in their protein phosphorylation pattern by causing, on the one hand, suppression of cAMP-dependent protein kinase and, on the other, enhancement of phosphorylation of different kinds of protein. The initial decrease in the cAMP level of oocytes in response to a MIS may be caused by an increase in phosphodiesterase (PDE) activity. The observation that the PDE inhibitor, theophylline, has no inhibitory effect on oocyte maturation when it is injected into oocytes (O’Connor and Smith, 1976) suggests that the PDE responsible for the initiation of oocyte maturation is localized near the surface of oocytes. In this regard, Ca ions appear to play an important role in activating surface membrane-bound PDE. Maller and Krebs (1977) have postulated the existence of a Ca-dependent regulatory (CDR) protein that regulates PDE activity in amphibian oocytes. This CDR protein, upon reacting with Ca, changes its conformation and becomes capable of activating PDE. According to

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J. L. Maller (personal communication) the CDR protein isolated from beef heart can induce maturation in Xenopus oocytes when it is injected into oocytes after treatment with Ca. At this point it might be useful to summarize the foregoing discussion into some form of a working hypothesis in order to explain the initiation mechanism of oocyte maturation (Fig. 10). (1) The action of a MIS on the surface of oocytes causes a release of Ca ions which have been sequestered in the oocytes. (2) The released Ca ions act on the CDR protein which in turn causes activation of PDE. (3) The activated PDE degrades CAMP, resulting in a decreased CAMPlevel. (4) Thus CAMP-dependent protein phosphorylation is discontinued, and CAMPindependent protein phosphorylation increases. (6) This phosphorylated protein itself may represent active MPF, and its nonphosphorylated form the inactive precursor of MPF. If so, the protein is a phosphorylase kinase which catalyzes the phosphorylation of its own precursor and activates the precursor, thus autocatalytically amplifying its own activity. A protein of this kind may actually exist (Belle et al., 1976; Singh and Wang, 1977). (7) In amphibian oocytes, the

MIS

1

1

Gy

Protein 6-P

2

MWactiye)

8

;

ProteinA-p (inactive)

.L

GV FIG. 10. Scheme of the initiation of maturation in Xenopus oocytes. 1, MIS induces a conformational change in surface receptors; 2, Ca is released; 3, PDE is activated, thus lowering the level of CAMP; 4, CAMP-dependent protein kinase, which has been phosphorylating protein A, is inactivated; 5, phosphorylation of protein A, which has been constantly synthesized and degraded in the oocyte is terminated, and thus free protein A (the initiator) appears; 6, the initiator catalyzes phosphorylation of protein B (MPF precursor) to form active MPF; 7, active MPF phosphorylates autocatalytically precursor MPF (protein B); 8, MPF acts as a GVBD inducer. The scheme is a synthesis of previous hypotheses (Wasserman and Masui, 1975a; Maller and Krebs, 1977).

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25 1

CAMP-independent phosphorylation of MPF may require synthesis of a protein (initiator), as hypothesized by Wasserman and Masui (1975a). However, there are at present no theories concerning exactly how MPF causes GVBD. C. ARRESTOF MEIOTICDIVISION 1. Metaphase Arrest

Except for species whose oocytes complete maturation before fertilization (coelenterates and echinoids) and those in which maturation is initiated by fertilization (echiuroids and some marine invertebrates), oocyte maturation is generally arrested either at metaphase I or 11. The arrested meiotic process is resumed following fertilization. These facts have been interpreted as suggesting that oocytes build up a self-inhibitory factor during the course of their maturation, and that their activation can be regarded as a mechanism for removing the inhibitory factor (see Monroy, 1965). Early investigators (Bataillon and Tchou-Su, 1930; Dalcq et al., 1936) considered the oocytes of amphibians to be “intoxicated” with COz. Heilbrunn and his associates (Heilbrunn et al., 1954) and Osanai (1967) suggested that an accumulation of acid polysaccharide was responsible for the arrest of meiotic progression prior to fertilization. However, the mechanisms underlying the meiotic block at metaphase I or I1 have not been the subject of intensive experimental analysis until recently. Humphries (1961) observed that coelomic oocytes of Triturus viridescens which had initiated maturation during ovulation sometimes advanced past metaphase I1 unless they were invested with a jelly coat while passing through the oviduct. Thus he suggested the potential significance of a contribution from the oviducal secretion to establishment of the meiotic block. However, since a majority of the oocytes without jelly investment still remained at metaphase 11, he acknowledged the possibility that a metaphase-blocking mechanism intrinsic to the oocytes existed.

2. Cytostatic Factor If an inhibitory factor blocking meiotic divisions at metaphase exists in unfertilized eggs, and its removal allows the eggs to proceed toward the completion of meiosis and the initiation of mitosis, it might be expected that the reintroduction of suck an inhibitory factor into fertilized eggs would arrest the progression of their cell cycles. And, in fact, Masui and Markert (1971), using R. pipiens, injected unfertilized egg cytoplasm (in volumes ranging from 30 to 120 nl) into one of the blastomeres of two-cell embryos and demonstrated that the cell cycle of the recipient emZlryos was indeed amsted a€ metaphase (Figs. 11 and 12). They found that the frequency with which metaphase arrest was induced and the

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FIG.11. An arrested blastomere of a two-cell embryo injected with the extract of an unfertilized egg (R. pipiens). (Meyerhof and Masui, 1977.) FIG.12. Metaphase arrest in a blastomere injected with the extract of an unfertilized egg (R. pipiens). (Meyerhof and Masui, 1977.) FIG. 13. Dark-field photomicrographs showing grain density over chromosomes induced to condense in enucleated oocytes injected with leucine-labeled GV contents. (Ziegler, 1979.) FIG.14. Bright-field photomicrographs showing the absence of grain accumulation over chromosomes induced to condense in enucleated oocytes injected with ovarian oocyte cytoplasm. (Ziegler, 1979.)

time at which the division of the recipient blastomeres was arrested were both correlated with the amount of cytoplasm injected and with the time of injection during the cleavage cycle; that is, the larger the cytoplasmic volume injected, and the earlier the stage of the recipient blastomeres, the earlier the blastomeres were arrested-hence the larger the arrested blastomeres. In a control series of experiments, cytoplasm obtained either from activated eggs which had been pricked with a glass needle, or from two-cell embryos, was injected into recipient blastomeres. Since cytoplasm from the activated donor eggs exhibited little ability to arrest the cell cycle in the recipient blastomeres, they concluded that unactivated eggs possessed a cytoplasmic factor which caused metaphase arrest of the cell cycle during the mitotic division of blasto-

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meres, and that its inhibitory activity could be removed by the activation process. This cytoplasmic factor was designated “cytostatic factor” (CSF) by Masui and Markert (1971). Furthermore, they postulated that it was CSF which was responsible for the metaphase block during meiosis, since its activity disappeared concomitantly with the resumption of meiosis triggered by egg activation. According to these investigators, CSF activity develops in the oocyte during maturation. It is undetectable in the oocyte before GVBD but appears shortly after GVBD and remains at a high level until activation. The development of CSF does not depend on the presence of the GV, since oocytes enucleated prior to the initiation of maturation also develop CSF activity following progesterone treatment. Meyerhof (1978) found that Xenopus oocytes induced to mature in vitro by progesterone developed CSF activity following GVBD; this activity was detectable when the cytoplasm was injected into blastomeres of R. pipiens two-cell embryos. No activity could be detected after the donor oocytes were activated by electric shock. However, very little CSF activity could be demonstrated in Xenopus eggs ovulated in vivo when their cytoplasm was injected into blastomeres of two-cell embryos of the same species. Also, failure to find evidence of CSF activity has been reported by Chulitzkaya (1970) and Chulitzkaya and Feulgengauer (1977), who injected cytoplasm from unfertilized eggs of R. temporaria and A . stellatus into eggs of the same species at various times following fertilization. Recently, however, Meyerhof (1978) found that cytoplasm of unfertilized Xenopus eggs obtained following HCG-induced ovulation consistently showed CSF activity when injected into blastomeres of two-cell embryos of the same species, provided the donor eggs were injected with a small quantity of EGTA (50 nmoledegg) prior to withdrawal of the cytoplasm to be tested. In control experiments cytoplasm from two-cell blastomeres which had been injected with the same dose of EGTA exhibited no CSF activity when injected into recipient blastomeres. Therefore the previous failure to demonstrate CSF activity in naturally ovulated Xenopus eggs can be attributed to a loss of CSF activity by the cytoplasm, probably caused by a release of Ca by the egg cytoplasm during the injection procedure. Similar reasoning may also explain the failure of Chulitzkaya and her associate to detect CSF activity in R. temporaria and Acipenser eggs. Interestingly, evidence supporting the existence of CSF activity in mammalian oocytes may be found in the results of recent experiments by Balakier and Czolowska (1977) who fused fragments of cytoplasm of mouse oocytes, which had been induced to mature in vitro, with blastomeres of two-cell embryos using Sendai virus. They found that the blastomere cells began to undergo mitosis following cytoplasmic fusion and reached metaphase, but the majority of these cells arrested at this point and failed to divide.

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Interspecific transfer of unfertilized egg cytoplasm into blastomeres of twocell embryos has been carried out between R . pipiens and X . laevis by Meyerhof (1978), whose technique included EGTA injection into donor eggs prior to the transfer. He found that cytoplasm of unfertilized Xenopus eggs strongly inhibited the cell cycle of R . pipiens blastomeres at the metaphase stage, whereas the cytoplasm of unfertilized Rana eggs generally failed to arrest Xenopus blastomeres. As expected, the cytoplasm of each species showed a CSF effect on cleaving blastomeres of its own species. It may be conjectured that Xenopus eggs have relatively higher CSF activity than Rana eggs or that Xenopus blastomeres have a stronger resistance to the activity of CSF than Rana blastomeres. In any case, this observation, taken together with previous results, suggests the possibility that CSF is a cytoplasmic factor involved in the arrest of both meiosis and mitosis at the metaphase stage in oocytes and embryonic cells of various species.

3. C a Effects on CSF CSF has been extracted from unfertilized eggs of R . pipiens by crushing the eggs using centrifugal force in a manner identical to that used for MPF extraction. Masui (1974) noted that CSF activity in the extracts was as unstable as MPF, disappearing within 24 hours on cold storage; but a more stable CSF activity, lasting at least a few weeks in cold storage, was obtained when Ca was added to the extraction medium. The effects of Ca ions on CSF activity have been further investigated by Meyerhof and Masui (1977). In contrast to the previous result, they found that the high CSF activity in the extracts made with freshly ovulated R . pipiens eggs could be maintained by increasing the Mg concentration in the extracts as well as by the addition of EGTA. However, the addition of EDTA was found to abolish CSF activity completely. Thus it became apparent that CSF activity of fresh oocyte extracts was dependent on Mg ions and became relatively stable after the removal of Ca ions. At the same time, Ca ions apparently destabilized the activity, since addition of Ca to the extracts at a concentration as low as M rapidly abolished CSF activity (Meyerhof and Masui, 1977). This finding, an apparent contradiction to the previous one, indicates that Ca-sensitive CSF activity is a entity different from that previously found to be stabilized by Ca. Study of the time course of the CSF activity in the extracts prepared from freshly ovulated R . pipiens eggs has revealed that the extract loses CSF activity, either in the presence or absence of Ca, and then regains this activity during cold storage. The second CSF activity is resistant to Ca, and its development is accelerated when Ca ions are present in the extract. Meyerhof and Masui (1977) named the CSF in fresh egg extracts, whose activity is Casensitive, “primary” CSF, distinguishing it from that developed secondarily in aged extracts, “secondary” CSF, whose activity is Ca-resistant. Although secondary CSF may be an artifact created during the aging process

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of the extract, the effects of primary and secondary CSF on progression of the blastomere cell cycle are indistinguishable, arresting mitosis at metaphase in the same fashion. Because of its highly stable nature, secondary CSF has been partially characterized and found to be associated with macromolecules which can be precipitated with @H4)2S04 at 30% saturation, can be passed through BioGel 15M, and are RNase- and heat-sensitive (Masui, 1974; Y. Masui, unpublished). 4. Inactivation of CSF and E g g Activation During fertilization, eggs arrested at metaphase I1 resume meiosis, forming the pronucleus and then proceeding to mitosis. The process of egg activation must involve inactivation of CSF. The ability of activated oocytes to inactivate CSF has been tested by injecting unfertilized egg cytoplasm or its fresh extracts into fertilized eggs at various times following insemination. Meyerhof and Masui (1977) found that cleavage and further development of the recipient eggs were unaffected when the injection took place within the first 45 minutes following insemination, indicating that the eggs were capable of inactivating CSF in the initial 30 minutes of activation, since all eggs had been penetrated by sperm by 15 minutes after insemination. The ability of activated oocytes to inactivate CSF appears to develop in the cytoplasm without participation of the nucleus, since the CSF developed in oocytes which had been induced to mature after removal of the GV disappeared following pricking with a glass needle or electric shock (Masui and Markert, 1971). This demonstration of cytoplasmic autonomy in the process of egg activation is consistent with the observations by Smith and Ecker (1969) and Skoblina (1969) that oocytes induced to mature after GV removal underwent the same surface changes as observed in normal eggs when they were activated. The high sensitivity of primary CSF to Ca may be an indication that its inactivation during egg activation is caused by Ca ions, whose concentration rapidly increases at the time of activation, as demonstrated by Steinhardt er al. (1977) in sea urchin eggs, and by Ridgway et al. (1977) in medaka eggs. The Ca release induced by sperm penetration appears to be a factor causing egg activation. In fact, the resumption of meiosis, arrested at metaphase 11, can be induced by ionophore A23 187 which releases Ca sequestered by the membrane system in the eggs of various species (Steinhardt et al., 1974). Also injection of Ca ions by iontophoresis activates mouse oocytes arrested at metaphase I1 to resume meiosis (Fulton and Whittingham, 1978). However, it has been noted that egg activation in the mouse is difficult to induce by ionophore treatment when Mg ions are present in the external medium (Masui et aE., 1977). In this mammal, activation of eggs by the ionophore is highly dependent on the relative strengths of the antagonistic effects of Ca and Mg present in the external medium, and only when the Ca effect exceeds the Mg effect can eggs be activated (Masui and Miller,

256


unpublished). This antagonism between these two divalent cations may be attributed to the opposite effects exerted by these ions on the activity of CSF, Mg having a stabilizing effect and Ca a destabilizing one. Thus it may be concluded that the decrease in the free Ca level by cytoplasmic sequestration during oocyte maturation in vertebrates favors the development of MPF and CSF, both of which are inactivated by Ca and stabilized by Mg, whereas its increase during activation favors the disappearance of both factors upon fertilization.

5 . Metaphase Arrest in Other Species Meiotic arrest at metaphase I is a widely observed phenomenon in oocyte

maturation in invertebrates. In these animals, whether or not the arrest is brought about by mechanisms similar to those postulated in vertebrate oocytes is open to speculation. It has been suggested by von Borstel (1957) that metaphase arrest during oocyte maturation in the parasite wasp Habrabracon juglandis is a genetically controlled process. His hypothesis is based on the following observations. In this species, maturation is initiated shortly after the oocytes are ovulated from the ovariole, but meiotic division is arrested at metaphase I during the period when the oocytes are in the uterine sac. At the time of oviposition, meiosis is resumed and the eggs develop parthenogenetically into haploid males (telytokous). However, in animals of a certain genetic strain, the oocytes continue meiosis, without metaphase arrest, and develop parthenogenetically in the uterine sac. The mechanism responsible for resumption of the meiotic division arrested at metaphase I has been analyzed recently by Pijnacker and Ferwerda (1976) in another telytokous insect, Carausius morosus (stick insect). In this animal meiosis is normally arrested at metaphase I for 5.5 days until the eggs are released from the uterine sac by oviposition. However, as long as the eggs are held in the uterine sac artificially by obstruction of the cloaca, they never resume meiosis. Only when they are exposed to the air can meiosis be resumed. In fact, meiosis in eggs laid in an atmosphere lacking 0, is arrested until the eggs are exposed to air. Thus it may be concluded that the factor overcoming meiotic arrest is the activation of aerobic respiration in oocytes.

VI. Nucleocytoplasmic Interaction during Oocyte Maturation A. CHROMOSOME CONDENSATION 1. Chromosome Condensation Activity

Cytoplasmic transfer experiments involving immature and maturing oocytes have demonstrated that the behavior of the GV during maturation is under the

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control of oocyte cytoplasmic activities which occur independently of nuclear functions. The effects of oocyte cytoplasm can be observed not only in the GV but also in nuclei introduced from other cells. Classic observations by Brachet (1922) on sea urchin oocytes, and by Bataillon and Tchou-Su (1934) on amphibian oocytes, showing that sperm nuclei introduced into oocytes by precocious insemination conformed to the behavior of the female nucleus in the oocyte, suggested the significant influence of the cytoplasm on the nucleus. The effects of oocyte cytoplasm on somatic cell nuclei were first investigated by Gurdon (1967, 1968) using X. Zaevis. He found that nuclei isolated from blastulas or from adult brain and exposed to oocyte cytoplasm underwent changes in morphology similar to those occurring in the GV. This was confirmed by Ziegler and Masui (1973), using R. pipiens, who found that the changes induced in the foreign nuclei followed the same time course as those in the oocyte nucleus. These observations strongly suggest that cytoplasmic control over the nucleus is not tissue-specific. The ability of oocyte cytoplasm to condense chromosomes to the metaphase state has been studied by Ziegler and Masui (1973, 1976a,b), who introduced nuclei isolated from adult R. pipiens brain into oocytes which had been treated with progesterone but prevented from undergoing activation by the use of a phosphate buffer @H 6.2 or lower) in the injection procedure. They found that the cytoplasmic activity causing condensation of the chromosomes appeared in the oocytes shortly after GVBD and persisted until they were activated. Using sperm nuclei, Moriya and Katagiri (1976) investigated the time course of chromosome condensation activity (CCA) in B . bufo oocytes undergoing maturation. These workers treated sperm with a dilute solution of Triton X-100, making them permeable to cytoplasmic factors, before introduction into the oocytes. They found that the sperm chromosomes condensed to the metaphase state upon exposure to the cytoplasm of oocytes which had undergone GVBD, but that sperm injected into activated eggs failed to do so. This CCA of the oocyte cytoplasm has not been demonstrated in normally ovulated uterine eggs. In this connection, it was noted by Brun (1974) that X. laevis oocytes ovulated in vivo were unable to condense chromosomes of transplanted nuclei, in contrast to oocytes induced to mature in vitro after progesterone treatment. He suggested that the development of the ability of oocytes to respond to activation stimuli is due to an influence of oviducal secretion on the oocytes (Brun, 1975). The fact is that oocytes induced to mature by progesterone in vitro are able to condense the chromosomes of transplanted nuclei when oocyte activation is prevented by external application of EGTA buffer during the transplantation operation, whereas oocytes ovulated in vivo cannot condense chromosomes. However, recently Meyerhof (1978) has discovered that CCA can be demonstrated in oocytes matured in vivo as well as in CSF-arrested blastomeres if nuclei are injected together with EGTA and Mg,

258


whereas CCA is not detectable in oocytes under any conditions after they are activated. Thus it appears that jelly-invested oocytes matured in vivo have a more unstable cytoplasm than jelly-free oocytes induced to mature in vitro by progesterone administration, so that CCA disappears in the former because of cytoplasmic perturbation caused by the injection procedure. Since EGTA stabilizes CCA in normally ovulated oocytes, it is highly probable that this cytoplasmic perturbation includes a discharge of Ca which is responsible for the rapid loss of CCA. The necessity of RNA and protein synthesis for the appearance of CCA has been investigated by Ziegler and Masui (1976a) using R . pipiens oocytes induced to mature in vitro by progesterone. They treated these oocytes with cycloheximide or a-amanitin at concentrations high enough to suppress protein or RNA synthesis, respectively, and at the same time injected the oocytes, which were at various stages of maturation, with isolated adult brain nuclei. The injected nuclei formed metaphase chromosomes in spite of the inhibition of protein and RNA synthesis when the injection took place at metaphase 11 (48 hours after hormone treatment), whereas those injected at metaphase I (24 hours after treatment) failed to undergo chromosome condensation. Since in both cases the recipient oocytes could be shown to have possessed CCA before the nuclei were transplanted, the failure of the nuclei injected into the recipients at metaphase I to condense must be attributed to a deficiency in protein and RNA synthesis which would normally occur following nuclear injection. However, no new protein or FWA synthesis was necessary for the chromosome condensation induced at metaphase 11, indicating that all the proteins and FWA required for chromosome condensation had been provided by the host oocytes by that time.

2 . Roles of the GV The possible importance of substances in the GV in chromosome condensation was suggested by Ziegler and Masui (1973), based on their early observation that no condensed brain nuclei chromosomes could be found 3-4 hours after the nuclei were injected into enucleated oocytes induced to mature by progesterone. It was noted, however, that the number of nuclei remaining in these oocytes was greatly reduced compared to the number in oocytes containing GV material. In later experiments therefore Ziegler and Masui (1976b) examined the number of nuclei remaining in the oocytes as well as the percentage of nuclei with condensed chromosomes at various times following injection. They found that chromosome condensation occurred both in nuclei injected into oocytes lacking GV material and in those injected into normal oocytes containing GV material when the nuclei were first examined, after 2 hours of exposure to oocyte cytoplasm. However, the number of clusters of condensed chromosomes was found to be markedly decreased after a 3-hour exposure, and no condensed chromosomes were found at 4 hours in oocytes lacking GV material, while the number of

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condensed chromosome clusters increased in nOITnal OOCYteS Over this time period. This observation strongly suggests that substances in the GV are not required for the induction of chromosome condensation in transplanted nuclei, but that the maintenance of condensed chromosomes following nuclear brane breakdown is highly dependent on the presence of GV substances. Whether or not material which accumulates in the GV during oogenesis associates with the chromosomes is a question several groups have undertaken to answer. Smith and Ecker (1970a) pointed out that proteins synthesized in ovarian oocytes of hibernating R. pipiens tended to accumulate in the GV, but these proteins were released into the cytoplasm following GVBD and redistributed into the zygote nuclei and into nuclei transplanted into activated eggs. According to Wassarman and Letourneau (1976b), 3H-labeled proteins which accumulate in the GV of mouse oocytes are found to associate with condensing chromosomes when GVBD takes place. They suggest that these proteins are predominantly histones, since protein labeled with try~tophan-~H neither accumulates in the GV nor associates with condensing chromosomes. This observation is in accordance with others indicating that the synthesis of histones is highly stimulated both in Xenopus (Adamson and Woodland, 1977) and mouse (Wassarman and Letourneau, 1976b) oocytes undergoing maturation. In mouse oocytes, Wassarman and Letourneau (1976b), using sodium dodecyl sulfate (SDS) gel electrophoresis, showed that F, histone was synthesized within 5 hours of isolation and was also phosphorylated. Recently Ziegler (1978) and Masui et al., (1979), using R. pipiens oocytes, analyzed the association of GV and cytoplasmic proteins with condensed chromosomes in oocytes. Progesterone-treatedoocytes, from which the GV had been removed, were injected with GV material obtained from immature oocytes previously labeled with Ie~cine-~H. The recipient oocytes were then injected with brain nuclei, while at the same time their protein synthesis was inhibited by cycloheximide. Autoradiographs of the nuclei injected in these oocytes revealed that the labeled material accumulated on the condensed chromosomes of the nuclei (Fig. 13). In the control experiment, labeled cytoplasm, instead of GV material, was injected together with brain nuclei into oocytes at metaphase I1 after protein synthesis had been inhibited. In contrast to the results for the experimental series, there was no accumulation of label on the condensed brain chromosomes (Fig. 14). In a second set of experiments, enucleated oocytes were treated with progesterone and labeled for 48 hours, at which time they reached a stage of maturation equivalent to metaphase 11. Cytoplasm from these oocytes was then injected, together with brain nuclei, into normal metaphase I1 oocytes which had been treated with cycloheximide. Under these conditions, the condensed chromosomes of the brain nuclei accumulated the label. Taken together, these results indicate that chromosomes undergoing condensation in maturing oocytes accumulate not only GV proteins but also cytoplasmic proteins which are

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synthesized by the oocytes during the course of maturation, whereas proteins found in the cytoplasm of immature oocytes with an intact GV do not accumulate on injected chromosomes. The association of GV proteins with condensed chromosomes may reflect a role of GV proteins in maintenance of the chromosomes in the oocyte cytoplasm; more specifically, they may act to protect chromosomal DNA from disintegration by cytoplasmic enzymes. Recently, Wyllie et al. (1977) showed that DNA injected into Xenopus oocytes remained inact when it was inside the GV, whereas DNA injected into the cytoplasm was degraded quickly. In this regard, it is interesting to note a recent report by Clark and Merriam (1977) that the GV contains a large amount of nonpolymerized actin which acts as a strong DNase inhibitor.

B. DEVELOPMENT OF THE PRONUCLEUS 1 . Male Pronucleus Growth Factor There are significant changes in cytoplasmic activities as a result of oocyte activation as discussed in Section V,A, and C. In amphibian oocytes, for instance, CCA disappears during activation, nuclei exposed to activated egg cytoplasm increase their volume, and their chromosomes become diffuse and initiate DNA synthesis (Graham et a l . , 1966; Gurdon, 1967, 1968). Changes in the cytoplasmic activity of mammalian oocytes during the course of maturation and activation have been studied by the insemination of oocytes at various stages of maturation and activation. Usui and Yanagimachi (1976), using hamster oocytes, found that sperm nuclei could be incorporated into zona-free oocytes at any stage of maturation as well as into zygote cells. An examination of the behovior of the incorporated sperm chromatin showed that chromatin incorporated into the oocytes before GVBD remained in a condensed state, whereas that incorporated at any time after GVBD decondensed even though the female chromosomes were undergoing condensation. Similar results have been reported by Iwamatsu and Chang (1972) using mice, and Niwa and Chang (1975) using rats, both of whom observed that decondensation of sperm nuclei incorporated into oocytes generally occurred only after GVBD. Only in the dog (Mahi and Yanagimachi, 1976) is there evidence that sperm nuclei incorporated into oocytes can undergo decondensation before GVBD. Generally, mammalian sperm nuclei incorporated into maturing oocytes at post-GVBD stages undergo decondensation independently of oocyte activation. These results apparently contradict the observations of Skoblina (1974, 1976) and Katagiri and Moriya (1976) that sperm nuclei injected into amphibian 00cytes remained in a condensed state until the oocytes were activated.

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However, in the sea urchin, while Long0 (1978) also observed swellkg of spem nuclei incoprated into meiotically dividing oocytes, he noted that these swelling nuclei lacked a pronuclear envelope and were composed of a central conoid mass of chromatin. This observation suggests that, although sperm nuclei incorporated into meiotically dividing mammalian or sea urchin oocytes may undergo a certain amount of swelling, this does not necessarily imply that a male pronucleus in being formed. On the contrary, recent studies of precocious insemination of R. pipiens OOcytes by Elinson (1975,1977) have revealed that sperm nuclei incorporated before oocytes become activatable develop into metaphase chromosomesembedded in a well-developed spindle following decondensation to a limited extent. He considers that a genuine pronucleus can be formed only after the oocytes have been activated by parthenogenetic stimuli. This has been confirmed by Ziegler (1979) and Lohka (1978), who observed a transient decondensation of sperm nuclei chromosomes injected into unactivated R. pipiens oocytes prior to condensation into the metaphase state. The ability of mammalian oocyte cytoplasm to induce development of the male pronucleus has been found to be dependent on the conditions under which the oocytes are induced to mature. Thibault and his associates (Thibault, 1972; Thibault et al., 1975a,b) showed in a variety of species, including rabbits, cows, and sheep, that foIIicIe-free oocytes which had undergone spontaneous maturation in vitro were unable to induce male pronuclear formation, while follicleenclosed oocytes which had been induced to mature by gonadotropins were able to support development of the pronucleus. They accordingly hypothesized that the failure of spontaneously maturing oocytes to induce pronuclear formation was due to lack of a cytoplasmic factor, named “male pronucleus growth factor” (MPGF). Mandelbaum et al. (1977) found that hamster oocytes developed MPGF only if they were kept in follicles for at least 6 hours following gonadotropin stimulation. Hunter et al. (1976) and Moor and Trounson (1977), working with pigs and sheep, respectively, have suggested that development of MPGF depends on the presence not only of gonadotropin but also of 17P-estradiol in the follicular environment, based on their finding that follicle-enclosed oocytes became capable of inducing male pronuclear formation following fertilization only if the oocytes matured in a medium containing sufficiently high levels of 17pestradiol. 2. Preparation for DNA Synthesis The importance of GV substances in the formation of, and DNA synthesis by, the pronucleus has been demonstrated by Skoblina (1974, 1976) and Katagiri and Moriya (1976) in R. temporaria and B . bufo oocytes, respectively. They found

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that sperm injected into OOCyteS underwent pronuclear transformation and initiated DNA synthesis once the recipients had reached metaphase a and had been activated. However, when the GV of a recipient oocyte was removed prior to injection, the sperm nuclei failed to swell and did not synthesize DNA. Katagiri and Moriya (1976) confirmed the active role of GV substances in promoting sperm nuclear transformation by introducing the karyoplasm of the GV into enucleated oocytes, which restored the ability of the oocytes to induce swelling as well as DNA synthesis in the injected sperm nuclei which would otherwise have remained inactive. They also found that the GV contents exerted the same effect whether they were taken from progesterone-treated or untreated oocytes, indicating that essentially the activity of the nucleoplasm did not change following progesterone stimulation. However, precocious rupture of the GV and subsequent mixing of the nucleoplasm and cytoplasm did not result in acquisition by the oocytes of the ability to promote DNA synthesis in transplanted nuclei (Gurdon, 1967). Clearly then, cytoplasmic maturation is also necessary for development of the oocyte activity promoting DNA synthesis in the nucleus. Gurdon (1967) transplanted adult brain nuclei into maturing oocytes of Xenopus and found that the cytoplasmic activity initiating DNA synthesis in the nuclei appeared after GVBD. This activity was also manifested when purified DNA was introduced into the oocytes (Gurdon and Speight, 1969). Furthermore, it was found that, not only did a marked increase in the overall activity of DNA polymerases in oocytes occur during maturation but, in addition, significant activity of a new DNA polymerase, distinguishable from those existing in immature oocytes, became detectable (Grippo and LoScavo, 1972; Grippo et al., 1975; Benbow et al. 1975). A recent study by Grippo et al. (1977) has indicated that DNA polymerase activity in Xenopus oocytes increases by 50% before GVBD (2-4 hours after hormone treatment), and that the presence of the GV is essential for this increase to occur, since only a 10%increase in activity occurs in oocytes deprived of their GV. Qualitative changes with respect to the factors and conditions controlling the initiation of DNA synthesis have been found to occur in Xenopus oocytes during maturation. Benbow and Ford (1975) found that DNA polymerases in extracts of ovarian oocytes were capable of initiating DNA synthesis only when supplied with denatured (single-stranded) DNA as a template, whereas extracts of uterine eggs could promote DNA synthesis using native (double-stranded) DNA templates. They attributed this capability of uterine oocyte extracts to the presence of a protein factor in the extracts which opened the template DNA and formed the initiation ‘‘eye. ’’ The ability of an oocyte to synthesize DNA for repairing damage to chromosomal DNA changes during maturation. In mouse oocytes, this unscheduled DNA synthesis can be induced at various stages of maturation by ultraviolet light irradiation (Masui and Pedersen, 1975) and by carcinogenic

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agents such as 4-nitroquinoline oxide (4NQO) (Brazil1 and Masui, 1978). Autoradiographic studies have indicated that the highest activity occurs in oocytes with an intact GV, regardless of the agent applied. When oocytes are treated with ultraviolet light or 4NQ0 after GVBD, the levels of DNA synthesis induced in the oocyte chromosomes are reduced by half, and little DNA synthesis is observed in the chromosomes in the polar body. Consequently, it was suggested that the levels of enzymic activities involved in repair DNA synthesis decreased during maturation of mouse oocytes, especially in the polar body. However, some alkylating agents such as methylmethane sulfonate (MMS) induce DNA synthesis at fairly constant levels in the oocyte and the polar body chromosomes throughout the maturation period. These observations may indicate that the activities of various enzymes involved in DNA synthesis in mouse oocytes are differentially altered during the progression of maturation. These changes may represent the initial step in a general decline in DNA repair activity in mouse cells, which continues throughout early life until by adulthood all somatic cells show very little activity in repairing damaged DNA. c . DEVELOPMENT OF MOTILESYSTEMS 1. Surface Contractility In order for a mature oocyte to begin development, it must be able to undergo cleavage following fertilization. Various chemical and physical agents activate unfertilized eggs which subsequently begin to cleave. A growing body of evidence suggests that the primary agent responsible for triggering surface contraction of activated eggs is the C L ion (Gingell, 1970; Schroeder and Strickland, 1974; Brachet, 1977). Classic experiments by Delage (1901), Wilson (1903), Yatsu (1905), Chambers (1921), and Costello (1940) with oocytes of marine invertebrates suggested that the ability of the oocyte cortex to cleave depended on GV substancess distributed throughout the cytoplasm following GVBD. A similar conclusion was reached by Tchou-Su and Yu-Lan (1958), Dettlaff et al. (1964), and Smith and Ecker (1970a), all of whom examined the surface contractility of amphibian oocytes undergoing maturation. Their experiments, essentially the same as the classic ones performed on marine invertebrates, showed consistently that the ability of the oocytes to initiate cleavage upon introduction of a nucleus, either by fertilization or by nuclear transplantation, became manifest only after GVBD had taken place, and that oocytes deprived of GV substances by enucleation failed to develop this ability. However, Smith and Ecker (1970a) noted that an equatorial constriction could occur in enucleated R. pipiens oocytes following activation by pricking with a glass needle, indicating that a certain degree of surface contractility could be

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acquired by the OOCYteS without GV substances. Recently, Hollinger and Schuetz (1976) found that OoCYtes of R . pipiens undergoing maturation could induced to ckave if Ca was locally injected underneath the cortex, and further that the cleavage furrow always occurred in a definite orientation with respect to the location of the Ca injection. This ability of oocytes to react to Ca injection occurred in those which had undergone GVBD but not in those with an intact GV. However, oocytes deprived of GV substances by enucleation also became capable of reacting to Ca injection in the same manner as normal oocytes if they were matured for the same length of time. Also, interestingly, Iwamatsu (1971), using medaka oocytes, found that oocytes with an intact GV, which had been displaced into the yolk mass by centrifugation and consequently failed to break down during maturation, became capable of cleaving and developing into haploid embryos upon fertilization. He obtained similar results using oocytes from which the GV had been surgically removed. These results suggest that, contrary to the classic notion, GV substances are not necessary for the development of surface contractility by oocytes. It has been shown that cytochalasin B (CB) can induce cleavage of oocytes under certain conditions. In Sabellaria alveolata (polychaete), Peucellier et al. (1 974) found that cleavage occurred in unfertilized eggs (arrested at metaphase I) treated with CB, despite the fact that meiosis did not resume. Furthermore, Wassaman et al. (1976) have shown that CB can induce cleavage of fully grown mouse oocytes with an intact GV which have been prevented from undergoing maturation by dbcAMP. Electron microscope studies have revealed that underlying the CB-induced cleavage furrow are microfilaments of a contractile ring similar to those found in normal cleavage furrows in developing embryos (Wassarman et al., 1977). Thus it may be that the surface contraction of oocytes induced by CB requires neither the release of GV substances into the cytoplasm nor activation of the cortical cytoplasm. The observations described here seem to be consistent with the notion that oocytes are equipped with a surface contractile system before they undergo maturation. This contractile system must remain unresponsive to activation stimuli until the oocytes have reached a certain stage of maturation. Maturation of the oocyte cytoplasm somehow induces the surface contractile system to become responsive to the activation stimuli. The fact that oocytes deprived of GV substances are unable to cleave normally upon insemination, as observed in toad (Katagiri and Moriya, 1976), sturgeon (Dettlaff and Skoblina, 1969), and starfish (Hirai et al., 1971; Lee et al., 1975), strongly suggests an indispensability of the contributions from the GV to the development of endoplasmic mechanisms involved in the coordination of karyokinesis and cytokinesis. One key endoplasmic structure involved in this coordination is the aster, as pointed out by Rappaport (1971) and Kubota (1969).

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2 . Aster Formation

In the sea urchin it has been observed that oocytes with an intact GV, either at the previtellogenic or vitellogenic stage, fail to form an aster after insemination. The inability of these growing oocytes to develop an aster has been attributed to a lack of necessary cytoplasmic components (Franklin, 1965). Recent research by Longo (1978) has shown that aster formation can take place only after oocytes begin meiotic division, that is, after GVBD. Skoblina (1974), 1976) in R. temporaria, and Katagiri and Moriya (1976) in B. bufo, consistently observed aster formation in the cytoplasm of oocytes matured after GV removal, following insemination or injection of Triton X100-treated sperm nuclei. Clearly, such aster formation occurred independently of any GV contribution. Heidemann and Kirschner (1975) studied aster formation in Xenopus oocytes microinjected with various cellular components, such as nuclei, basal bodies, or centrioles from a variety of sources. They found that aster formation could be induced in uterine eggs but not in ovarian oocytes possessing an intact GV. According to these investigators aster formation requires the presence of GV substances (Heidemann and Kirschner, 1978). However, Elinson (1977) has suggested that the ability to form asters may be correlated with changes in cytoplasmic conditions induced by egg activation, rather than by maturation itself. He reported that, when maturing oocytes of R. pipiens were inseminated after GVBD, the sperm nuclei incorporated into the oocytes failed to develop asters but instead formed spindles around their chromosomes. However, when these oocytes were activated, large astral rays developed around the spindles. Recently, Hanocq-Quertier et al. (1978) showed that aster formation could also be induced by the presence of DzO in the cytoplasm of Xenopus oocytes undergoing maturation, and that this D,O-induced aster formation could be observed even in medium-sized oocytes, which are unresponsive to progesterone, after GVBD had been induced by MPF injection. Furthermore, these workers found that, while no cytaster or spindle formation could be induced by DzO in small oocytes, even if GVBD and chromosome condensation were induced by MPF injection, the formation of these fibrillar systems could be induced after the injection of protamine. They suggest that the ability of protamine to enhance fibrillation in young oocytes is due to its action in precipitating an excess of soluble RNA present in small oocytes which would otherwise inhibit formation of microtubules (Bryan et al., 1975). However, contrary to the observations of Skoblina (1974, 1976) and Katagiri and Moriya (1976) concerning aster formation by sperm nuclei, these investigators found that DzO-induced cytaster formation did not occur under any circumstances if the GV was removed from the oocytes before treatment, consistent with the observation by Heidemann and Kirschner (1978).

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3 . GV Material Knowledge of the various physiological functions as well as of the molecular characteristics of the GV may be of paramount importance in the study of oocyte maturation in view of the essential role played by the GV during the course of oogenesis and early development of animals. The nuclear envelope of the GV shows a wide variation in permeability to different types of molecules, ranging from ions to macromolecules. Especially important is the fact that proteins of low or medium molecular weight can freely pass through the envelope (Century and Horowitz, 1974; Bonner, 1975a; Feldherr, 1975). Certain classes of proteins have been found to accumulate predominantly in the GV and, when these proteins are labeled and injected into the cytoplasm of recipient oocytes, they again migrate into and remain in the GV (Bonner, 1975b). Recently Feldherr and Pomerantz (1978) have shown that the accumulation of specific nuclear proteins in the GV is not controlled by the nuclear envelope but rather by selective binding within the nucleoplasm, since a loss of the nuclear permeability barrier does not affect the distribution of nuclear proteins. Hill et al. (1974) using Triturus oocytes, and Merriam and Hill (1976) using Xenopus oocytes, resolved the proteins present in the GV into as many as 68 polypeptides by SDS gel electrophoresis. Most (85%) of these proteins can readily be dissolved in saline solution, but the remainder form a gel which can be dissolved only in alkaline salt solutions. One of the two major components of the gel appears to be actin (Clark and Merriam, 1977). This component shows the same electrophoretic mobility and peptide-mapping pattern as muscle actin, reacts with antiactin antibodies, and binds with DNase I to inhibit its activity. According to Clark and Merriam (1977) these actin molecules are present in a nonpolymerized form in intact oocytes but are readily polymerized upon exposure to Mg and K ions. The other major component of GV proteins is a myosinlike protein weighing about 250,000 daltons (Clark and Merriam, 1977). The gel component of the GV contracts slowly in the presence of ATP and a low concentration of Ca, forming microfilaments 70-80 A in diameter which can be decorated by heavy meromyosin. Actin has also been suggested to be present in the GV of mammalian oocytes as a result of studies by Amsterdam et al. (1977) using immunofluorescent-cytochemical techniques. These observations make it tempting to speculate that it is actin in the GV which is responsible for protecting DNA from nucleolytic degradation (Wyllie et al., 1977) and that the GV contributes actin which aids spindle formation and chromosome condensation and protection in oocytes following GVBD, thus making karyokinetic movement of the chromosomes possible, as suggested by Forer (1974). Finally, Burzio and Koide (1976), using Xenopus oocytes, found that the GV contains poly-(ADP-ribose) synthetase having an extremely high activity, and that its activity further rises sharply following progesterone treatment of the oocytes, but before GVBD (Burzio and Koide, 1977). These workers

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postulate that, since this nuclear enzyme adds ADP ribose from NAD to chromosome proteins (Honjo and Hayashi, 1973; Sugimura, 1973), probably causing alterations in chromosomal conformation, it is highly probable that the progesterone-stimulated increase in the activity of the enzyme in the GV is related to the appearance of CCA during oocyte maturation. L. 0. Burzio @ersonal communication) recently found that inhibition of poly-(ADP ribose) synthetase by its substrate (NAD) also inhibits maturation of progesterone-treated oocytes in Xenopus.

VII. Control of Meiosis and Mitosis-Concluding Remarks Oocytes arrested at the diplotene stage are in the G2 phase of the cell cycle. They proceed into the GI phase as zygotes, following maturation and activation. Hence oocyte maturation and activation as a whole appear to be analogous to the transition process of a cell, which includes mitosis, and take it from the G2to the G, phase. This concluding section considers several ways in which the control mechanisms of oocyte maturation and those of mitosis in somatic cells are analogous. A. ROLE OF Ca Although the agents which induce the transition from G2 to GI in oocytes, that is, MISS, are generally ineffective in inducing mitosis in other cells, some of them, for example, proteolytic enzymes (Burger et al., 1972) and ionophore A23187 (Maino et al., 1974) are known to be active mitogenic agents in certain types of cells such as lymphocytes. The common effect induced by all these agents in the cell is a rise in the intracellular Ca level. That Ca plays an important role in regulation of the cell cycle is suggested by the periodic changes in free Ca level in the cell occurring during the cell cycle. Clothier and Timourian (1972), for instance, showed that a cyclic influx and efflux of Ca occurred during the cell cycle of cleaving zygotes of the sea urchin. In this cell system, three peaks of Ca mobilization are observed: the first during interphase, the second just before the beginning of mitosis, and the third at the initiation of cleavage. This observation agrees well with recent observations by Holmes and Stewart (1977) on the cell cycle of Physarumpolycephalum. They showed that a Ca efflux, which involved more than half of the total cellular Ca, occurred over a short period of time, ranging from 3 to 4 minutes, at the beginning of metaphase. This efflux was followed by an influx which continued until the commencement of anaphase. The Ca incorporated into the cell during metaphasic progression was then released.

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In oocytes, such as those of Xenopus and starfish, a release of cytoplasmic bound Ca occurs following MIS treatment (Section IV,B). This release of Ca is soon followed by a continuous uptake of Ca by the cell, which may be interpreted as active sequestration, thus lowering the free Ca level in the oocyte. This condition may be a prerequisite for the appearance of MPF and CSF in the cell, since both these cytoplasmic factors are sensitive to Ca ions. The relationship between the movement of Ca ions during the cell cycle and during mitosis is an interesting topic for speculation. First, Ca ion levels probably regulate the assembly of microtubules in the cell, since it is known that Ca ions inhibit their assembly in vitro (Weisenberg, 1972; Borisy and Olmsted, 1972). Second, Ca ions activate or inactivate numerous cellular enzymes. In oocytes, however, since Ca or Ca-mobilizing agents effectively induce maturation only when they are applied to the surface of oocytes (Section IV,B), it is difficult to ascribe the primary effect of Ca mobilization on oocytes to its action on the microtubule systems in the cell. Rather, it appears more likely that the action of the mobilized Ca is exerted on enzymes associated with the surface membrane of oocytes, causing a change in their activities. These enzymes probably include ATPases, PDE, and adenyl cyclase. At present, we know little about the action of these enzymes in animal oocytes, and the data currently available, for instance concerning ATPase activity of the oocytes, have given us no significant information with respect to the initiation of oocyte maturation. However, the importance of changes in the enzyme activities which regulate cAMP levels in the oocytes is suggested by observations that PDE inhibitors and cAMP derivatives have a marked inhibitory effect on oocyte maturation in numerous kinds of animals. Again, these inhibitors are effective only when applied to oocytes externally. These results strongly suggest that Ca plays an important role in controlling the activities of enzymes located on the surface membrane of oocytes, which are involved in the regulation of oocyte cAMP levels. It may be that, as put forth in Maller and Krebs’ (1977) hypothesis, the role of Ca present on the surface of the oocyte is to directly or indirectly activate PDE. B. ROLEOF cAMP Studies on the level of cAMP during the cell cycle of various cell types have revealed that it always decreases sharply when cells enter mitosis from the G2 phase (Burger et al., 1972; Sheppard and Prescott, 1972; Yasumasu et al., 1973; Zeilig et al., 1974), while the changes in the cAMP levels of cells undergoing transitions between other phases of the cell cycle vary among different cell types

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(see Prescott, 1976). In contrast to the behavior of CAMP,cGMP levels appear to remain rather constant throughout the cell cycle, although some fluctuation has been observed (Seifert and Rudland, 1974). It has been also observed that dbcAMP (Freedman et al., 1975), as well as PDE inhibitor (Burger et al, 1972; Millis et al., 1974), prevents cells at interphase from entering mitosis. Analogies concerning cAMP function can be drawn between mitotically dividing somatic cells in culture and meiotically dividing oocytes, since in the latter it has also been observed that cAMP levels are sharply decreased upon initiation of maturation and that dbcAMP and PDE inhibitors prevent oocytes from undergoing maturation (see Section V,B). C. PHOSPHORYLATION OF CELLULAR PROTEINS The finding that the inhibition of CAMP-dependentprotein kinase by its R subunit leads to the initiation of oocyte maturation, and that addition of the C subunit of the enzyme to the oocyte causes its suppression, strongly suggests that the processes by which primary oocytes eventually become zygotes are initiated by an interruption of the protein phosphorylation catalyzed by CAMP-dependent protein kinases. This interruption may cause an alteration in the pattern of protein phosphorylation in the cell. It has been established that, during mitosis, nuclear or chromosomal proteins are phosphorylated. For example, the phosphorylation of F, histone has been well-documented in various types of cells (Bradbury et al., 1974; Gurley et al., 1974). This phosphorylation has been suggested to be correlated with chromosome condensation. Alterations in the pattern of protein phosphorylation of cells undergoing the transition from a mitotically quiescent to an active state have been investigated in different types of cells. Generally, it has been found that type I and type I1 CAMP-dependentprotein kinases display different activities at different phases of the cell cycle (Costa et al., 1976). For example, in lymphocytes, a mitogen stimulates the cells to activate type I CAMP-dependent protein kinase, whereas dbcAMP activates both type I and type 11, thereby causing an inhibition of cell proliferation (Byus et al., 1977). It appears that type I1 protein kinase exerts a negative influence on the proliferation process of the cell. Corroborating this notion are the recent findings of Kletzien et al. (1977) using the baby hamster kidney cell line. They demonstrated the existence of a protein which was phosphorylated only in mitotically active cells and showed that cAMP suppressed phosphorylation of this protein, while at the same time enhancing phosphorylation of other proteins, the net result being an inhibition of cell proliferation.

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These observations suggest that meiotic divisions in oocytes and mitotic divisions in somatic cells are both controlled by common cellular mechanisms involving phosphorylation of a certain category of proteins.

D. SHCYCLE The finding that oocyte maturation can be induced by SH-blocking or SHreducing agents only when these agents are applied externally suggests that the initiation of maturation involves changes in the SH content of surface proteins (Section IV,A). This has been verified by measurements of the SH content of the cortex of starfish oocytes before and after the induction of maturation by 1-MA or SH-reducing agents (Kishimoto and Kanatani, 1973). Furthermore, Ikeda et al. (1976) showed that the amount of cortical SH reached a peak a few minutes before the expulsion of each polar body and then dropped sharply. This cyclic change in the SH content of the cortex is comparable to that observed during the cleavage cycle in sea urchin zygotes, in which the maximum level occurred shortly before each cleavage division (Sakai, 1968). This parallel change in cortical SH values during the course of oocyte maturation and of zygote cleavage seems to further strengthen the view that the cellular processes of meiosis and mitosis are governed by a common control system involving oxidoreduction of SH groups in the cortex proteins.

E. CYTOPLASMIC CONTROL FACTORS It has been established beyond question that maturing oocytes develop cytoplasmic factors which induce nuclear membrane breakdown, chromosome condensation, and metaphase arrest. These cytoplasmic factors have proven to be effective in inducing the same changes in somatic cell nuclei introduced into oocytes as in the native germ cell nucleus (Section VI, A-C). Furthermore, the existence of similar cytoplasmic factors in somatic cells has been well documented by cell fusion experiments in which nondividing (G or G 2 )cells are fused with cells in mitosis, resulting in premature chromosome condensation (Johnson and Rao, 1971; Matsui et al., 1971). Thus it appears that the nuclear events in oocytes undergoing meiotic divisions, and those in somatic cells undergoing mitotic divisions, are under the control of the same cytoplasmic factors. In this connection, we recall the observation of Masui and Markert (1971) that MPF, which is abundant in maturing oocytes of R. pipiens, can also be detected in blastomeres actively engaged in mitosis. Wasserman and Smith (1978b) also showed that MPF appeared in the cleaving blastomeres of amphibian oocytes

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during mitosis. Furthermore, W. J. Wasserman (personal communication) found that MPF capable of inducing maturation in Xenopus oocytes existed in the cytoplasm of cultured mammalian cells in mitosis. Recent experiments by Y. Masui (unpublished), involving the transfer of cytoplasm from sea urchin (Strongylocentrotuspurpuratus) oocytes into R . pipiens oocytes, have indicated that sea urchin zygotes produce a cytoplasmic factor during mitosis which causes oocyte maturation in frogs.

ACKNOWLEDGMENTS

We thank Mr. William A. Welch for his valuable contribution to the preparation of the manuscript for this article. A portion of the work cited here was supported by grants from the National Cancer Institute of Canada and the National Research Council of Canada, awarded to Y.M. for the period 1972-1977.

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DNA damage response during mouse oocyte maturation.

Embryological aspects of oocyte in vitro maturation.

Hamster oocyte penetrability during preovulatory maturation.

Potassium rectifications of the starfish oocyte membrane and their changes during oocyte maturation.

Porcine oocyte maturation in vitro: role of cAMP and oocyte-secreted factors - A practical approach.

In vitro follicle growth supports human oocyte meiotic maturation.

Multiple requirements of PLK1 during mouse oocyte maturation.

KIF20A regulates porcine oocyte maturation and early embryo development.

Pharmaceutical options for triggering of final oocyte maturation in ART.

Transcriptomic features of Pecten maximus oocyte quality and maturation.

Thyroid stimulating hormone enhancement of bovine oocyte maturation in vitro.

Synthesis and modification of D7 protein during Xenopus oocyte maturation.

Atrial natriuretic peptide inhibits spontaneous rat oocyte maturation.

Development of calcium release mechanisms during starfish oocyte maturation.

Oocyte-secreted factors in oocyte maturation media enhance subsequent development of bovine cloned embryos.

Oocyte maturation in vitro: contribution of the oviduct to total maturation in Xenopus laevis.

Role of arachidonic acid cascade in Rhinella arenarum oocyte maturation.

Artificial control maturation of porcine oocyte by dibutyryl cyclicAMP.

Relationship between growth and meiotic maturation of the mouse oocyte.

Nondisjunction by failures in the molecular control of oocyte maturation.

Fluoride impairs oocyte maturation and subsequent embryonic development in mice.

Reevaluation and evolution of the simulated physiological oocyte maturation system.

[In vitro oocyte maturation for female fertility preservation].

JAK2 inhibitor CEP-33779 prevents mouse oocyte maturation in vitro.