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Morphological changes during maturation of starfish oocytes: Surface ultrastructure and cortical actin
DEVELOPMENTAL
BIOLOGY
s&373-334
(1983)
Morphological Changes during Maturation of Starfish Oocytes: Surface Ultrastructure and Cortical Actin’ THOMASE. Friday Harbm Laboratories,
SCHROEDER
AND
STEPHEN
A. STRICKER
University of Washington, Friday Harbor, Washington
Received November
11, 1982;
accepted in revised
form March
98250
1, 1983
The cell surface and extracellular investments of oocytes of the starfish Pisaster ochrawvs are analyzed by Nomarski differential interference contrast microscopy and by scanning electron microscopy. The investing coats include a thin sheet of follicle cells, a jelly coat, and a vitelline layer; their morphologies are described. Methods are outlined for systematically removing them without altering the behavior of the oocyte so that the cell surface can be examined directly. The topography of denuded oocytes changes dramatically when they are treated with the maturation-inducing hormone, 1-methyladenine. The major topographical change is the early and transient formation of prominent surface spikes. These structures arise due to the rapid, reversible polymerization of actin into stout bundles. Polymerization and subsequent depolymerization of cortical actin is monitored by epifluorescence microscopy of oocytes stained with NBD-phallacidin, a stain which is specific for polymerized actin. Based on scanning electron microscopy, spikes apparently utilize preexisting plasma membrane of microvilli, and plasma membrane is apparently lost when spikes collapse. Long after microvilli are eliminated due to spike formation, the number of microvilli is somewhat restored, especially around the animal pole where the polar body forms. A chronology of events observed during oocyte maturation is discussed with reference to the possible mechanisms and implications of polymerization and depolymerization of cortical actin. INTRODUCTION
In starfish (Echinodermata: Asteroidea), as in most animals including the vertebrates, eggs are stored in the ovary as immature oocytes that are arrested at meiotic prophase. Maturation of starfish oocytes is reinitiated under hormonal stimulation in synchrony with ovulation, and the oocytes become fertilizable at spawning, or soon afterward. Accordingly, starfish oocytes taken from the ovary represent an important model system for investigating the processes of maturation. Cytological studies of maturation of starfish oocytes are facilitated by a rich background of information. The hormonal triggers for maturation are known with certainty (Kanatani, 1973). The receptors for the maturation-inducing hormone, 1-methyladenine (l-MA), have been localized at the plasma membrane surface (Kanatani and Hiramoto, 1970; Guerrier and Doree, 1975); suggestions have been made about their morphological manifestation (Schroeder, 1981); and isolation of the putative receptors has been achieved (Morisawa and Kanatani, 1978; Ikadai and Kanatani, 1982). In addition, progress is being made toward defining the various physiological steps in the transduction process from hormone to cytoplasmic alteration (Guerrier et aL, 1977; Moreau and Guerrier, 1980; Mazzei et aL, 1981; Meijer ‘The authors wish to dedicate this paper to the memory fessor Robert L. Fernald (1914-1933).
of Pro-
and Guerrier, 1981; Doree et al, 1981a; Johnson and Epel, 1982) and determining the cytoplasmic factors essential for maturation (Kishimoto et aL, 1981). To this background of knowledge, we add observations of dynamic changes in the surface topography and the organization of cortical actin during maturation. Our objectives are (1) to describe methods for handling starfish oocytes for successful scanning electron microscopy (SEM), (2) to document morphological changes in maturing oocytes as a possible basis for understanding some of the physiological changes mentioned above, and (3) to supplement previous observations concerning “spikes,” the prominent structures that appear transiently on the oocyte surface in response to l-MA (Schroeder, 1981). To test a hypothesis that cortical actin plays a role in spike formation (Schroeder, 1981), we have stained cortical actin with NBD-phallacidin. This fluorescent stain is specific for polymerized actin (Barak et aL, 1980) and has been used to study actin organization in various cell types (Nothnagel et aL, 1981; Schroeder and Christen, 1982; Schroeder, 1982). MATERIALS
AND
METHODS
Adult specimens of Pisaster ochraceus were collected intertidally around San Juan Archipelago, Washington, and were maintained until needed in running seawater at ll-15’C. All operations and observations were con-
373 0012-1606/83 $3.00 Copyright All rights
0 1983 by Academic Press, Inc. of reproduction in any form reserved.
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DEVELOPMENTALBIOLOGY
ducted at seawater temperatures or at 20-22°C (room temperature), as specified. To obtain ovaries, an arm was severed from the base, both ovaries in the arm were removed at the short oviducts, and they were then stored in seawater. Gravid ovaries typically measured 5-8 cm in length and were obtained from April to early August. Immature oocytes with their investing follicle cells were obtained by teasing several ovarian lobules (about 1 cm long). Fragments of ovarian tissue were removed and the oocyte-follicle cell complexes were washed in seawater. This method resulted in the “spontaneous” breakdown of germinal vesicles in -5% of the oocytes, probably as a consequence of mechanical trauma. To remove follicle cells, oocyte-follicle cell complexes were washed in five changes of calcium-free seawater (CaFSW) consisting of 0.38 M NaCl, 49 mM MgClz, 26 mM Na2S04, 8.5 mM KCI, 2.25 mlM NaHC03, 2.5 m&Z EGTA, and 10 mM Tris-HCl buffer with the final pH adjusted to 8.0-8.2. Oocytes free of follicle cells were also obtained by directly rinsing ovarian lobules and fragments in several changes of CaFSW for 1 hr. By adding natural seawater (containing calcium), the fragments contracted, and oocytes without follicle cells were extruded (Kanatani and Shirai, 1969; Cloud and Schuetz, 1979). Very little ‘*spontaneous” breakdown of germinal vesicles was observed using this method. Ovarian fragments were removed by passage through cheesecloth; the oocytes were then rinsed a few times with CaFSW and stored. As monitored by Nomarski (Fig. 2b) and SEM (Fig. 3b), CaFSW alone does not remove any investing coat other than the follicle cells. In our preparations the thick jelly coat and vitelline layer were routinely dissolved together by protease digestion after pretreatment with 10 PM ionophore A23187 (Eli Lilly & Co., Indianapolis, Ind.) which causes premature elevation of the vitelline layer. A 25-~1 volume of ionophore A23187 (1 mg/ml in ethanol) was first added to -50,000 oocytes without follicle cells in 5 ml of CaFSW in a Syracuse watch glass. At 11-15°C vitelline layers were fully elevated by 15 min, and the medium was then mostly removed. Protease solution in CaFSW (5 ml) was added. The oocytes were quickly stirred and then allowed to remain undisturbed as a monolayer for 10 min. The enzyme solution in our early studies was 1% protease (Type V, Sigma Chemical Co., St. Louis, MO.; No. P-5005, 1100 PU/g), but when this product became unavailable we used 0.1% Pronase (Calbiochem-Behring Co., La Jolla, Calif.; No. 53702,56,000 PU/g) with equal success. We never observed that digestion caused parthenogenetic activation, as reported by Bryan and Sato (1970), or that it significantly altered oocyte sensitivity to lMA (Jeffery, 1977). After 10 min of digestion the enzyme
VOLUME98.1983
solution was cautiously removed, CaFSW was gently added, and the CaFSW was changed twice. The oocytes were somewhat adherent soon after digestion; if disturbed at this stage, they tended to lyse, until they had been thoroughly washed. Dose dependencies for germinal vesicle breakdown (GVBD) in oocytes treated with l-MA were determined at selected temperatures by adding 25 ~1 of oocyte suspension containing 100-200 oocytes to 0.2-ml volumes of serially diluted l-MA solutions in seawater or CaFSW. Oocytes must be exposed to l-MA for a definite period of time in order to undergo GVBD, as first suggested by the work of Schuetz (1969) and then substantiated by Guerrier and Doree (1975). The hormone-dependent period (HDP) is defined as the minimum time of exposure to a threshold concentration for 50% GVBD to occur. The HDP for Pisaster oocytes was determined by (1) treating about 2000 oocytes in 5 ml of medium with 10m6Ml-MA at selected temperatures, (2) removing 50~1 aliquots (containing at least 200 oocytes) at 1 to 2min intervals, and (3) rapidly diluting each aliquot into 4 ml of seawater or CaFSW. The oocytes were immediately sedimented by hand centrifugation and resuspended in another 4 ml of seawater. The extent of GVBD was determined by counting at either 90 min (11-15’C) or 60 min (20-22°C) after the onset of l-MA treatment. For scanning electron microscopy, oocytes were fixed first in 1% glutaraldehyde-90% seawater (or CaFSW) at pH 7.4 for 1 hr at room temperature. Oocytes were then rinsed briefly in seawater (or CaFSW) and postfixed in 1% osmium tetroxide-75% seawater (or CaFSW) at pH 8.0-8.2 for 0.5-l hr. Specimens were partially dehydrated in an ethanol series and then completely dehydrated in dimethoxypropane (Maser and Trimble, 1977), critical point dried from carbon dioxide, cemented to specimen stubs with silver paint, and sputter-coated with gold-palladium (60-40%). A JEOL JSM35 scanning electron microscope was used for observations. For staining polymerized actin in the cortex, oocytes were prefixed in 1% formaldehyde-90% CaFSW at pH 8.0-8.2 for 10 min. A single drop containing about 100 oocytes was then added to 2 drops of stain made up as follows: 25 ~1 of 1 mg/ml palmitoyl lysolecithin, 25 ~1 of 3 pg/ml nitrobenzoxydiazole-conjugated phallacidin (NBD-phallacidin, Molecular Probes, Inc., Plano, Tex.) in methanol, and 0.45 ml of CaFSW. After staining for 10 min, the oocytes were mounted under supported coverslips to avoid compression. The oocytes were viewed by epifluorescence microscopy, primarily by focusing on the upper surfaces. Fluorescence was detected by incident illumination at 490 nm supplied by a 100-W tungsten-halogen lamp in a Zeiss Universal microscope equipped with a 500-nm dichroic reflector, a 63X (N.A.
SCHROEDER
AND
STRICKER
CeU Surface
1.25) objective lens, and a 52%nm barrier filter. Fluorescent images were recorded on Kodak Tri-X Pan film (ASA 400) with 15-set exposures and were printed under identical conditions. RESULTS
A ripe female Pisaster ochraceus yields a large number of oocytes which are uniform in appearance and quite transparent. When still surrounded by follicle cells, the oocytes are irregular in shape (Fig. la), but they become spherical soon after being washed in CaFSW, a process that also removes the follicle cells. The fully grown immature oocyte is about 160 pm in diameter
and Cortical Actin of Starfish Oocytes
375
and contains a large clear germinal vesicle that is about ‘75pm in diameter. The germinal vesicle is eccentrically situated so that it lies 3-5 pm from the oocyte plasma membrane at one point (the animal pole), and contains a prominent nucleolus (Fig. 2). By focusing on the uppermost surface of an oocyte with the Nomarski microscope (not shown), the oocyte cortex is seen to contain a layer of clear vacuoles -2 pm in diameter with smaller cortical granules interspersed among the vacuoles. Many, but frequently not all, of the cortical granules are released by activation with ionophore A23187, but the vacuoles do not change. Other organelles and inclusions appear to be uniformly distributed throughout the remaining cytoplasm. Investing Coats
FIG. 1. Low-power
photomicrographs of oocytes from the starfish oocytes and their investing follicle cells as they appear shortly after dissection from the ovary. At this stage oocytes are irregular in shape and the follicle cell investments (arrowheads) are often incomplete. (b) Mature oocytes at the time of first polar body formation. In this preparation the investing coats had been removed and the oocytes were then treated with l-methyladenine to reinitiate meiosis. Maturation events proceed with high synchrony, although a few cells (*) in a batch often fail to mature. Scale bar = 100 pm. Magnifications = 59X.
Pisaster ochramus. (a) Immature
Follicle ce2Ls.A single layer of follicle cells (Figs. la and 2a) surrounds immature oocytes that are dissected from ovaries into seawater. This layer is often incomplete; the larger gaps may be produced during the dissection procedure. Each follicle cell is less than 0.5 pm thick except at its thicker perikaryon from which a slowly beating flagellum emerges. There appear to be about 25 to 75 follicle cells associated with each oocyte. They are roughly polygonal in shape and measure about 25 X 45 pm, according to superficial views by Nomarski microscope (Schroeder, 1981) and by SEM (Fig. 3a, inset). Follicle cells often do not exhibit broad areas of contact along their lateral margins; consequently, the underlying jelly coat is sometimes visible between the cells (Fig. 3a). As shown in Fig. 2a, the follicle cells are separated lo-15 pm from the oocyte surface by the intervening jelly layer. Each follicle cell is connected to the underlying oocyte by over 100 thin follicle cell processes which traverse the jelly layer (Runnstrom, 1944; Schroeder et ah, 1979; Schroeder, 1981). These can sometimes be seen by SEM (Fig. 3a). Insufficient washing in CaFSW causes some of these processes to remain embedded in the jelly coat, where they can be confused for spikes when seen in the light microscope; remnant processes are also sometimes seen by SEM (Fig. 3~). Although intercellular contacts between follicle cells and oocytes occur while the oocytes are still in the ovary (unpublished observations), the attenuated shape of the follicle cell processes probably results from the swelling of the jelly coat. During natural spawning, oocytes are released into seawater without investing follicle cells, which are retained in the ovary after follicle cell-oocyte contacts are separated. Jelly coat. The jelly coat swells to a thickness of lo15 pm upon contact with seawater. Most of the jelly coat appears fibrous in organization by Nomarski microscopy (Figs. 2a and b), although an extremely trans-
FIG. 2. Nomarski photomicrographs of starfish oocytes at 12°C after stepwise removal of investing coats (a-d) and progressively after lMA treatment (e-h). In each case only the animal hemisphere of the spherical oocyte is shown. The large germinal vesicle (gv) with its prominent nucleolus is visible (a-f), until later stages of maturation when these structures have disintegrated (g, h). (a) An oocyte dissected from the ovary into seawater with its fibrous jelly coat (j) and investing layer of follicle cells (fc). (b) In calcium-free seawater the follicle cells dissociate and the jelly coat swells but remains present. (c) Ionophore A23187 causes the vitelline layer (VI) to elevate. (d) Protease digestion in calcium-free seawater removes the vitelline layer and jelly coat, thus exposing the plasma membrane directly to the medium. (e) At 10 min after application of lo-’ Ml-MA, radiating projections called spikes (sp) are visible on the surface. (f) By 22 min after l-MA treatment, spikes at the surface have vanished. (g) By 65 min after l-MA treatment the germinal vesicle has broken down leaving in its place a somewhat yolk-free region (*). (h) The first polar body (pbl) forms at about 165 min after l-MA treatment; in this figure at 215 min the second meiotic apparatus is visible as a clear zone beneath the first polar body. The scale bar = 10 pm. Magnification = 335X.
parent component of the jelly gradually expands beyond the fibrous zone after oocytes remain in seawater or CaFSW for a few hours. In the SEM the jelly coat resembles a tangled mat of fine fibers (Figs. 3a and b). We do not know, however, how much of this organization is a product of specimen preparation. In our experience neither CaFSW or acidification to pH 4.15 dissolves the jelly coat, contrary to reports elsewhere (Jeffery, 19’77; Shoji et aZ., 19’78; Nakamura and Hiramoto, 19’78; Nemoto et aZ., 1980). Lowered pH does cause the jelly coat to dissipate temporarily, but it reappears when the pH is again raised. We find that it is necessary to remove the jelly coat along with the vitelline layer by digestion in protease, as described under Materials and Methods. Vitelline layer. The vitelline layer of Pisaster oocytes is about 0.25 pm thick, according to transmission electron micrographs (Schroeder, 1981), and thus is much thicker than its counterpart on sea urchin eggs (Schroeder, 1979). This layer was seldom observed by SEM, because we did not specifically endeavor to expose it by removing the jelly coat alone; nevertheless, in some
preparations the protease treatment digested only the jelly coat, while leaving the vitelline layer relatively intact (Fig. 3~). In such preparations, the vitelline layer appears quite smooth on its outer surface, although it has been shown that microvilli are embedded in the vitelline layer from below, before elevation occurs (Hirai et al., 1971; Cayer et ak, 1975; Schroeder, 1981). Sometimes we see small pores in the vitelline layer (Fig. 3~) through which follicle cell processes apparently project as they course toward the oocyte surface. These holes, however, are usually not visible. Thus, it is likely that the vitelline layer material can rearrange itself and obscure the holes left when the follicle cell-oocyte contacts are broken and the follicle cell processes withdraw. We have had little success using 1 M urea to remove the vitelline layer, in contrast with studies of oocytes from other species (Dan-Sohkawa, 1976). In our hands, urea only caused an occasional rupture of the vitelline layer, thereby revealing a portion of the oocyte surface. The microvilli in these cases appeared somewhat longer and more erect than after protease digestion.
SCHROEDER AND STRICKER
Cell Surface
and Cortical
Actin
of Starfish
Oocytes
FIG. 3. Scanning electron micrographs of various surfaces exposed during the removal of investing coats surrounding Pim.steroocytes. The insets in the center illustrate the corresponding surfaces at low magnification. (a) Each follicle cell (fc) surrounding the oocyte possesses a single flagellum (fg); from the undersides of these cells, thin follicle cell processes (arrowheads) project toward the oocyte through the fibers of the jelly coat (jc) which is visible in the gaps between follicle cells. (b) In calcium-free seawater the jelly coat is entirely exposed and appears to be composed of fibers that form a thick mat. (c) In a few specimens, digestion in protease removed the jelly coat without removing the vitelline layer; wrinkles in the vitelline layer may be due to its elevation and expansion caused by ionophore A23187 treatment and to its subsequent collapse during preparation. Several long processes pass through small pores in the vitelline layer (arrowheads) are presumed to be fragmentary follicle cell processes still in contact with the oocyte; these are not usually seen and probably result from inadequate rinsing in CaFSW. Microvilli on the oocyte surface (mv) can be seen through a small tear in the vitelline layer. (d) After the complete digestion of the vitelline layer, the oocyte plasma membrane is entirely exposed; numerous short microvilli cover its surface. Scale bar = 1 pm (insets, 10 Mm). Magnification = 5400X (insets, 660X).
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DEVELOPMENTAL BIOLOGY
Hormone Requirements and the Course of Maturation The threshold concentration of l-MA for Pisaster oocytes in vitro, i.e., the concentration at which 90+% undergo germinal vesicle breakdown (GVBD), is typitally 1 X lop6 M. It is temperature independent from 11 to 22°C and is the same in natural seawater or ar-
VOLUME98,1983
tificial CaFSW. The schedule for meiotic maturation, however, is distinctly temperature dependent. At -12°C GVBD begins at 55 min; the long-lasting first meiotic spindle is set up somewhat later; and, the first polar body is emitted at -165 min (Fig. 2). At -21°C GVBD begins 26 min after treatment with l-MA. The hormone-dependent period for Pisaster oocytes
FIG. 4. Changes in surface topography after application of l-MA at -12°C. (a) Before l-MA treatment the oocyte surface is uniformly covered with short microvilli. (b) Within 1 min after l-MA treatment, triangular projections -2 pm long emerge from the surface as microvilli begin to disappear. (c) By 10 min after l-MA the projections are transformed into definitive spikes measuring 3-5 pm in length (they are up to 10 pm long in living oocytes) and microvilli have been reduced to tiny bumps. (d) By 30 min after l-MA the formerly erect spikes have been replaced by thinner, longer, and flaccid processes which are interpreted as being the membranous remnants of the spikes without their rigid contents. Microvilli are virtually absent at this stage. Several visible depressions, which can also be seen before l-MA treatment, probably represent collapsed vacuoles that occupy the cortex of these oocytes. Scale bar = 1 pm. Magnification = 6790X.
SCHROEDER AND STRICKER
Cell Surface
the HDP is 11-12 at -12°C is 27-31 min. At -2l”C, min. These values suggest that the HDP is a constant fraction of the time to GVBD. At a given temperature, the HDP is the same for oocytes in seawater or CaFSW. The HDP is unchanged by the presence or absence of oocyte investing coats and is not affected by higher concentrations of l-MA.
Early Surface Responses to l-MA The surface topography of Pisaster oocytes is rapidly and dramatically altered by treatment with l-MA. Qualitatively, these changes include the appearance of long, straight spikes, the simultaneous loss of microvilli, resorption of the spikes, and eventually a partial restoration of microvilli. Some of these events are discernible by light microscopy (Fig. 2), while others can only be appreciated by SEM (Figs. 4 and 5). Accurate quantification of these changes is complicated by the shrinkage of the specimens that typically
and Cortical Actin of Starjsh
Oocytes
379
occurs during preparation for SEM, as discussed fully elsewhere (Schroeder, 1979). Shrinkage reduces the dimensions of structures but presumably does not effect their numbers. In our SEM micrographs the overall oocyte diameter is -35% less than it is in life (100 pm vs 160 pm). We do not know, however, if this shrinkage is isometric at the level of the microvilli. Thus, we will simply cite measurements obtained from representative SEM micrographs, without correcting for shrinkage. Before treatment with l-MA the cell surface is uniformly covered with about 2.5 X lo5 short microvilli. In the SEM these microvilli measure -0.1 pm in diameter and -0.35 pm in length (Figs. 3d and 4a). Very soon after l-MA application, the oocyte surface begins to undergo changes that culminate in the transient appearance of spikes (Schroeder, 1981). The following description refers to the schedule of events at -12’C. Several minutes elapse before spikes can be seen by Nomarski microscopy (Fig. 2e), but at even earlier
FIG. 5. Fluorescence micrographs of l-MA-treated oocytes that have been fixed and stained with NBD-phallacidin, a stain that is specific for polymerized actin. The times after l-MA correspond to those illustrated in Fig. 3 by SEM. (a) An extremely low level of fluorescence is detectable before l-MA. (b) Within 1 min after l-MA application an extensive meshwork of thin polymerized actin bundles -2-pm-long begins to appear. (c) By 10 min after l-MA, when spikes are longest and most prominent, numerous bold actin bundles up to 10 pm long appear all over the oocyte surface. (d) By 30 min after l-MA, there is no evidence of polymerized actin by fluorescent staining with NBDphallacidin. Scale bar = 10 pm. Magnification = 1455~.
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DEVELOPMENTAL BIOLOGY
times there are topographical changes which are obvious by SEM. Within 1 min after treatment, the surface displays broad leaf-like projections (Fig. 4b); these are more or less triangular in shape and -2 pm on a side, bearing on their edges short projections that are reminiscent of microvilli. Microvilli are still visible between these projections (Fig. 4b), but they are much reduced in length. At about 10 min after l-MA application, the leaf-like projections are gone, and the oocyte surface is covered by about 6500 spikes. These structures are very straight, abruptly pointed, -0.3 pm in diameter, and 3-5 pm long when viewed by SEM (Fig. 4~). In living oocytes they are frequently 10 pm long. In Fig. 4c the density of fully formed spikes is about -0.3/pm2. By this stage, microvilli are further reduced to small, sparse bumps. Spikes can be seen to withdraw soon after they are fully formed. By 20 min after l-MA application very few spikes are visible by Nomarski (Fig. 2f), and by 30 min none are visible even by SEM (Fig. 4d). The stout, straight spikes are replaced by thin, curved, and seemingly flaccid projections that lie along the oocyte surface (Fig. 4d). These are not as numerous as spikes were at 10 min, and they soon disappear entirely. They undoubtedly represent membranous remnants of spikes that have lost their rigidity. Between these flaccid projections there are a few small bumps that probably represent shortened microvilli.
Cortical Actin aJter Treatment with l-MA The early-surface responses to l-MA, described above, are correlated with dramatic changes in actin of the oocyte cortex, as indicated by patterns of staining with NBD-phallacidin. Before l-MA is applied, the cortex displays only a very low level of fluorescence (Fig. 5d). Within 1 min after application of l-MA, an elaborate meshwork of very thin fluorescent lines is present throughout the cortex (Fig. 5b). These lines are -2 pm long and apparently represent bundles of polymerized actin in an early stage of formation. They do not occur in the deeper cytoplasm. By 10 min after treatment with l-MA, the putative actin bundles are fewer in number, but they are longer, thicker, and more boldly fluorescent (Fig. 5~) than at 1 min. These bright needles of fluorescence are 4-12 pm in length and occur only in the cortex as projections from the oocyte surface. Their location, density, and time of appearance correlate with those of the definitive spikes seen by Nomarski or SEM. By 30 min after lMA application, when spikes are completely withdrawn, no fluorescence can be detected in whole starfish oocytes stained with NBD-phallacidin (Fig. 5d). Thus, there is evidence of even less polymerized cortical actin at this stage than before l-MA is applied.
VOLUME 98, 1983
Polar Body Formation Very little change in the topography of starfish oocytes occurs between elimination of the flaccid projections and formation of the first polar body (Fig. 6). Microvilli gradually reappear, although never to the density seen before application of l-MA (Figs. 3d and 4a). The microvilli become slightly polarized in distribution on the oocyte shortly before polar body formation. Microvilli are longer and slightly more dense within a circular zone about 75 pm in diameter around the animal pole where the polar body forms (Figs. 6a and b). Within this zone (comprising less than 10% of the oocyte surface), microvilli are 0.5 pm long and occur at a density of -3.5/pm2 (Fig. 6b). The polar body itself usually exhibits microvilli that are about the same length but are sparser (Fig. 6b). Outside this polar zone, microvilli are only -0.15 pm long and occur at a density of -3.0/pm2 (Fig. 6~). Staining with NBD-phallacidin at polar body stages yields a mottled pattern of weak fluorescence surrounding the polar body (Fig. 6d). A minor topographical feature that is also affected by l-MA is an array of surface depressions which are -2 brn in diameter and visible at all stages of maturation (Figs. 4a-d, and 6~). They presumably represent the clear vacuoles situated in the cortex and have collapsed during peparation for SEM. The surface overlying these depressions sometimes becomes perforated during specimen preparation, particularly at the time of germinal vesicle breakdown and at polar body stages (Fig. 6~). DISCUSSION
Surface Ultrastructure and the Control of Actin Polymerixation Using SEM to observe surface changes of experimentally denuded oocytes, we find that the immature oocyte is thoroughly covered with short microvilli. The hormone l-MA induces a rapid reorganization of the surface which culminates after 10 min in long spikes, previously described elsewhere (Rosenberg, 1978; Schroeder, 1981). Microvilli are quantitatively eliminated as spikes form. The loss or retraction of microvilli during oocyte maturation has been reported for other species of starfish (Hirai et aL, 1971; Cayer et aL, 1975; Rosenberg, 1978; Hirai and Shida, 1979). Based on the numbers and dimensions of microvilli and spikes (see Results), the total surface area of an oocyte is constant during spike formation, suggesting that spikes “borrow” plasma membrane from the microvilli. Our observations with NBD-phallacidin confirm an earlier hypothesis that spikes contain dense bundles of polymerized actin (Schroeder, 1981). Indeed, actin polymerization begins within a minute, and the forces which
SCHROEDER AND STRICKER
Cell ,%&ace and Cortical Actin of Star$sh
Ooc@es
FIG. 6. First polar body stage at 165 min after l-MA treatment. (a) Survey SEM view of the small first polar body on the surface of the oocyte. (b) A first polar body (pbl) as it is pinching off from the rest of the oocyte; this area corresponds to the trapezoidal box in a. Relatively long microvilli are present in this region of the oocyte surface. (c) At a location 90” away from the first polar body, in a region corresponding to the rectangular box in a, microvilli are much shorter and slightly less numerous; depressions similar to those in Figs. 4a and d sometimes appear to be perforated at this stage (arrowheads). (d) Staining with NBD-phallacidin reveals a weak fluorescence that outlines the polar body (pbl). Scale bars a and d = 10 pm; b and c = 1 Frn. Magnifications: a = 64 OX; b and c = 6790X; d = 1455X.
cause spikes to protrude may derive from this polymerization and assembly into bundles. Prior to l-MA treatment, fluorescence due to stained polymerized actin is extremely low, though present; perhaps it is attributable to actin in microvilli, although our transmission EM observations have not yet revealed polymerized actin at this stage. Spikes are withdrawn by 30 min and no detectable sign of polymerized actin remains. When spikes collapse there appears to be a loss of cell surface (i.e., plasma membrane). The cell surface remains nearly smooth until microvilli partially reappear by the time of first polar
body formation. Even then, microvilli are still fairly sparse and are distributed in a polarized fashion, perhaps reflecting cortical movements (Hamaguchi and Hiramoto, 1978). Our rough calculations indicate that the total cell surface area at this stage is still 40% less than in the immature oocyte. The mechanisms by which l-MA causes actin polymerization or depolymerization are not known. There is evidence that an elevation of intracellular pH is at least part of the trigger for actin polymerization in cells (Tilney et ak, 19’78; Carron and Longo, 1982; Begg et al, 1982; Schroeder and Christen, 1982). In starfish oocytes,
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DEVELOPMENTALBIOLOGY
however, treatment with l-MA does not cause intracellular pH to rise (Johnson and Epel, 1982), yet it triggers the actin polymerization involved in spike formation. Furthermore, application of permeant weak bases which can elevate intracellular pH fails to induce the formation of spikes (Schroeder, unpublished observations); indeed, rather than mimicking l-MA, such weak bases actually inhibit l-MA-induced germinal vesicle breakdown (Cloud and Schuetz, 1979; Guerrier et al, 1978; Doree et ah, 1982). Thus, the triggering of actin polymerization in starfish oocytes may involve complex factors.
Identijcation
and Efects of Removing Investing Coats
In order to study the surface ultrastructure of oocytes, we devised methods for systematically removing investing layers (follicle cells, jelly coat, and vitelline layer) without altering the responses of oocytes to lMA. We have cleared up previous misinterpretations of these investments (Rosenberg et aL, 1977; Lee et al, 1977) which apparently arose through the unwarranted belief that CaFSW removes the jelly coat. On the other hand, we do confirm the findings of Cayer et al. (1975) and Cloud and Schuetz (1979) that CaFSW can completely remove the follicle cells and their processes, provided the oocytes are rinsed sufficiently. In the case of Pisaster oocytes, complete removal of the jelly coats and vitelline layers requires protease digestion after ionophore treatment. Responses of P&aster oocytes to l-MA proved to be identical before or after removing the investments and regardless of being cultured in natural seawater or CaFSW. These responses include (1) surface spike formation and withdrawal; (2) the time course, rate, and extent of maturation; (3) the threshold concentration of l-MA; and (4) the duration of the hormone-dependent period. We fail to confirm observations by Rosenberg and Lee (1981) that both the threshold concentration and the hormone-dependent period are specifically and significantly altered by the calcium content of the medium.
Relationship of Cortical Actin to Other Ph@ological Events Relating spike formation to other l-MA responses or to oocyte maturation is problematical. Spike formation is not necessary for germinal vesicle breakdown (Schroeder, 1981), yet both events share l-MA as their specific inducer; in fact, a single class of l-MA receptors may be involved in both responses, since the hormone specificities (using analogs of l-MA) and hormone sensitivities (using experimentally altered oocytes) of the two responses are indistinguishable (T. E. Schroeder and H. Ikadai, unpublished observations).
VOLUME 98. 1983 Ca2+ burst Actin polymerization
Actin depolymerization Phosphorylation of cortical proteins
x 4 $ kJ I
50-
End of hormone dependent period
60-
Membrane depolarization
k4
70-
w I I=
80-
OOJ
Cortical relaxation
Germinal vesicle breakdown
FIG. ‘7. A timetable of molecular and biophysical events that occur during maturation of starfish oocytes. The data are drawn from published reports of various species studied under a variety of conditions. The time scale is normalized (Time 0 corresponds to the application of l-MA and Time 100 represents the beginning of germinal vesicle breakdown) so that the timing of various events can be compared.
A normalized timetable of starfish oocytes is presented in Fig. 7. It summarizes the temporal relationship between actin polymerization and various other changes that occur during maturation. We have assumed (perhaps simplistically) that chronological events between l-MA application and germinal vesicle breakdown can be proportionalized and that comparisons between species are valid. The earliest known response to l-MA is a burst of intracellular free calcium ions that occurs within seconds (Moreau et ak, 1978). Putative actin polymerization and the associated topographical changes begin soon after the calcium burst (this study; Schroeder, 1981). When surface spikes reach their maximum extension, several events begin almost simultaneously: actin depolymerization starts (this report); protein kinase activity rises and phosphorylation of cortical proteins reaches a plateau (well before a steady state is achieved in the endoplasm) (Guerrier et aZ., 1977); the membrane potential declines from -70 to less than -20 mV by the time that germinal vesicle breakdown occurs (Miyazaki et al, 1975); and mechanical rigidity of the cortex rapidly declines (Nakamura and Hiramoto, 1978, Nemoto et aL, 1980). These changes appear to begin before the end of the hormone-dependent period, the time at which
SCHROEDER AND STRICKER
CeU Surface
the cell becomes irreversibly committed to undergo germinal vesicle breakdown. Very little is known about events between the end of the hormone-dependent period and the germinal vesicle breakdown; high activity of maturation-promoting factor appears only after germinal vesicle breakdown occurs (Kishimoto et al, 1981). The timetable presented in Fig. 7 provides a basis for exploring causal mechanisms in the cellular responses to l-MA, although we recognize that sequential events need not be causally linked. With regard to possible mechanisms of actin polymerization, the fact that a calcium burst is the only cytoplasmic event known to precede actin polymerization suggests that these two events may be linked; actin polymerization could be controlled by a calcium-sensitive regulator. On the other hand, spikes do not form merely by treating oocytes with ionophore A23187, so the potential role of calcium is not simple. Actin depolymerization and spike withdrawal coincide temporally with several known biochemical changes (Fig. 7). Phosphorylation of diverse proteins in the cortex seems to be a particularly important event during the hormone-dependent period (Guerrier et cd, 1977). It is possible that actin depolymerization results from phosphorylation of a key protein, for example, by activating a regulatory protein. Such a mechanism for regulating actin is not incompatible with the idea that protein phosphorylation is also involved in reinitiating meiosis. It is noteworthy that :Mabuchi (1981) has recently isolated from starfish oocytes a 17,000-dalton protein called “depactin” which causes rapid actin depolymerization in vitro. It is not yet known if depactin’s activity is altered by phosphorylation. Analyses of the phosphoproteins which appear soon after l-MA application (Preddie and Abastado, 1979; Doree et al., 1981b; Mazzei and Guerrier, 1982) sometimes reveal a protein about the size of depactin, but they are otherwise somewhat conflicting. Such studies set the stage for further work that could clarify the mechanisms and interrelations of spike formation, actin assembly-disassembly, and the processes leading to meiotic reinitiation in starfish oocytes. This study was supported by the U. S. Public Health Service in a suballocation of a Biomedical Research Support Grant RR 07096 to the University of Washington and by NIH Research Grant GM 19464. REFERENCES BARAK, L. S., YOCVM, R. R., NOTHNAGEL, E. A., and WEBB, W. W. (1980). Fluorescence staining of the actin cytoskeleton in living cells with 7-nitrobenz-2-oxa-3,3-diazole phallacidin. Proc Nat Acad. Sci USA 77.980-984. BEGG, D., REBHUN, L. I., and HYA’IT, H. (1982). Structural organization of actin in the sea urchin egg cortex: Microvillar elongation in the absence of actin filament bundle formation. J. CeU. Biol 93, 24-32.
and Cortical
Actin
of Star&sh
Oocytes
383
BRYAN, J., and SATO, H. (1970). The isolation of the meiosis I spindle from the mature oocyte of Pisaster ochraceus. Exp. Cell Res. 59, 371-378. CARRON, C. P., and LONGO, F. J. (1982). Relation of cytoplasmic alkalinization to microvillar elongation and microfilament formation in the sea urchin embryo. Dev. Biol 89, 129-137. CAYER, M. L., KISHIMOTO, T., and KANATANI, H. (1975). Formation of the fertilization membrane by insemination of immature starfish oocytes pretreated with calcium-free seawater. Deuelop. Growth D$ feren 17,119-B%. CLOUD, J. G., and SCHUETZ, A. W. (1979). 1-Methyladenine induction of oocyte (starfish) maturation: Inhibition by procaine and its pH dependency. J. Exp. Zool. 210,11-16. DAN-S• HKAWA, M. (1976). A “normal” development of denuded eggs of the starfish, Asterina pectin$ero. Den Grwth D&?ren. 18,439445. DOREE, M., KISHIMOTO, T., LE PEUCH, C. J., DEMAILLE, J. G., and KANATANI, H. (1981a). Calcium-mediated transduction of the hormonal message in meiosis reinitiation of starfish oocytes. Exp. Cell Res. 135, 237-249. DOREE, M., CAVADORE, J.-C., LE PEUCH, C. J., and DEMAILLE, J. G. (1981b). Hormone-induced phosphorylation of a 16,000 dalton polypeptide following meiosis reinitiation in starfish oocytes. Exp. Cell Res. 135, 251-257. DOREE, M., SANO, K., and KANATANI, H. (1982). Ammonia and other weak bases applied at any time of the hormone-dependent period inhibit 1-methyladenine-induced meiosis reinitiation of starfish oocytes. Dev. Biol SO, 13-17. GUERRIER, P., and DOREE, M. (1975). Hormonal control of reinitiation of meiosis in starfish: The requirement of 1-methyladenine during nuclear maturation. Dev. Biol 47. 341-348. GUERRIER, P., MOREAU, M., and DOREE, M. (1977). Hormonal control of meiosis in starfish: Stimulation of protein phosphorylation induced by 1-methyladenine. Mol. Cell Endocrinol. 7, 137-150. GUERRIER, P., MOREAU, M., and DOREE, M. (1978). Control of meiosis reinitiation in starfish: Calcium ion as the primary effective trigger. Ann. Biol. Anim. Bid Biophys. 15,441-452. HAMAGUCHI, M. S., and HIRAMOTO, Y. (1978). Protoplasmic movement during polar-body formation in starfish oocytes. Exp. Cell Res. 112, 55-62. HIRAI, S., KUBOTA, J., and KANATANI, H. (1971). Induction of cytoplasmic maturation by 1-methyladenine in starfish oocytes after removal of the germinal vesicle. Exp. Cell Res. 68, 137-143. HIRAI, S., and SHIDA, H. (1979). Shortening of microvilli during the maturation of starfish oocyte from which vitelline coat was removed. Bull Mar. Biol Sta Asamushi, Tohoku Univ. 16, 161-167. IKADAI, H., and KANATANI, H. (1982). Regeneration of oocyte-surface factor responsible for 1-methyladenine-induced oocyte maturation in starfish. Biomed Res. 3, 378-383. JEFFERY, W. R. (1977). Hormonal action of 1-methyladenine: The effect of enzymatic digestion of intact starfish oocytes on the induction of meiosis. Gen Comp. Endocrinol 31, 259-269. JOHNSON, C. H., and EPEL, D. (1982). Starfish oocyte maturation and fertilization: Intracellular pH is not involved in activation. De-v. Biol 92,461-469. KANATANI, H. (1973). Maturation-inducing substance in starfishes. Int. Rev. Cytd 35, 253-298. KANATANI, H., and HIRAMOTO, Y. (1970). Site of action of l-methyladenine in inducing oocyte maturation in starfishes. Exp. Cell Res. 61, 280-284. KANATANI, H., and SHIRAI, H. (1969). Mechanisms of starfish spawning. II. Some aspects of action of a neural substance obtained from radial nerve. Biol. BuU 137, 297-311. KISHIMOTO, T., HIRAI, S., and KANATANI, H. (1981). Role of germinal
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vesicle material in producing maturation-promoting factor in starfish oocyte. Dev. BioL 81, 177-181. LEE, H. H., ROSENBERG, M. P., and HOESCH, R. (1977). The starfish (Patiria and Pisaster) oocyte: A model system for the study of hormone-surface interaction in oogenesis. Lab. Anim Sci 27,860-865. MABUCHI, I. (1981). Purification from starfish eggs of a protein that depolymerizes actin. J. B&hem 89, 1341-1344. MASER, M. D., and TRIMBLE, J. J. (1977). Rapid chemical dehydration of biologic samples for scanning electron microscopy using 2,2-dimethoxypropane. J. Histochem. Cytochewz. 25, 247-251. MAZZEI, G., MEIJER, L., MOREAU, M., and GUERRIER, P. (1981). Role of calcium and cyclic nucleotides during meiosis reinitiation in starfish oocytes. Cell mere% 10, 139-145. MAZZEI, G., and GUERRIER, P. (1982). Changes in the pattern of protein phosphorylation during meiosis reinitiation in starfish oocytes. Dev. BioL 91, 246-256. MEIJER, L., and GUERRIER, P. (1981). Calmodulin in starfish oocytes. I. Calmodulin antagonists inhibit meiosis reinitiation. Dev. BioL 88, 318-324. MIYAZAKI, S.-I., OHMORI, H., and SASAKI, S. (1975). Potassium rectifications of the starfish oocyte membrane and their changes during oocyte maturation. J PhysioL 246, 55-78. MOREAU, M., and GUERRIER, P. (1980). In vitro interactions between membrane, hormone, and cyclic nucleotides as revealed with aequorin. Dev. BioL 79, 488-492. MOREAU, M., GUERRIER, P., DOREE, M., and ASHLEY, C. C. (1978). Hormone-induced release of intracellular Ca’+ triggers meiosis in starfish oocytes. Nature (London) 272, 251-253. MORISAWA, M., and KANATANI, H. (1978). Oocyte-surface factor responsible for 1-methyladenine-induced oocyte maturation in starfish. Gam. Res. 1, 157-164. NAKAMURA, S., and HIRAMOTO, Y. (1978). Mechanical properties of the cell surface in starfish eggs. Dev. Growth Di&ren. 20, 317-327. NEMOTO, S-I., YONEDA, M., and UEMURA, I. (1980). Marked decrease in the rigidity of starfish oocytes induced by 1-methyladenine. Dev. Growth Di$krm 22, 315-325. NOTHNAGEL, E. A., BARAK, L. S., SANGER, J. W., and WEBB, W. W. (1981). Fluorescence studies on modes of cytochalasin B and phallotoxin effects on cytoplasmic streaming in Chara. J. Cell BioL 88, 364-372.
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PREDDIE, E., and ABASTADO, R. (1979). A phosphorylated protein in KCl-Triton extracts of 1-methyladenine treated cortices from oocytes of the starfish Marthasterias glacialis. Dev. BioL 73, 345-352. ROSENBERG,M. P. (1978). Relationship of changes in the cortical layer to the resumption of meiosis in starfish oocytes: Hormonal and cation requirements. Ph.D. dissertation. Univ. Toledo, Ohio. ROSENBERG,M. P., HOESCH, R., and LEE, H. H. (1977). The relationship between 1-methyladenine induced surface changes and fertilization in starfish oocytes. Exp. Cell Res. 107, 239-245. ROSENBERG, M. P., and LEE, H. H. (1981). The roles of Ca and Mg in starfish oocyte maturation induced by 1-methyladenine. J. Exp. ZooL 217,389-397. RUNNSTROM, J. (1944). Notes on the formation of the fertilization membrane and some features of the early development of the Aster&s egg. Acta ZooL 25, 159-167. SCHROEDER, P. C., LARSEN, J. H., and WALDO, A. E. (1979). Oocytefollicle cell relationships in a starfish. Cell. Tissue Res. 203, 249256. SCHROEDER, T. E. (1979). Surface area change at fertilization: Resorption of the mosaic membrane. Dev. BioL 70, 306-326. SCHROEDER, T. E. (1981). Microfilament-mediated surface change in starfish oocytes in response to I-methyladenine: Implications for identifying the pathways and receptors for maturation-inducing hormones. .I Cell BioL SO, 362-371. SCHROEDER,T. E. (1982). Novel surface specializations on a sea anemone egg: “Spires” of actin-filled microvilli. J. Morphol. 174, 207-216. SCHROEDER, T. E., and CHRISTEN, R. (1982). Polymerization of actin without acrosomal exocytosis in starfish sperm. Exp. Cell Res. 140, 363-371. SCHUETZ, A. W. (1969). Induction of oocyte shedding and meiotic maturation in Pismter ochraceus: Kinetic aspects of radial nerve factor and ovarian factor induced changes. BioL Bull. 137, 524-534. SHOJI, Y., HAMAGUCHI, M. S., and HIRAMOTO, Y. (1978). Mechanical properties of the endoplasm in starfish oocytes. Exp. Cell Res. 117, 79-87. TILNEY, L. G., KIEHART, D. P., SARDET, C., and TILNEY, M. (1978). Polymerization of actin. IV. Role of Ca++ and H+ in the assembly of actin and in membrane fusion in the acrosomal reaction of echinoderms. .J. Cell BioL 77, 536-550.
BIOLOGY
s&373-334
(1983)
Morphological Changes during Maturation of Starfish Oocytes: Surface Ultrastructure and Cortical Actin’ THOMASE. Friday Harbm Laboratories,
SCHROEDER
AND
STEPHEN
A. STRICKER
University of Washington, Friday Harbor, Washington
Received November
11, 1982;
accepted in revised
form March
98250
1, 1983
The cell surface and extracellular investments of oocytes of the starfish Pisaster ochrawvs are analyzed by Nomarski differential interference contrast microscopy and by scanning electron microscopy. The investing coats include a thin sheet of follicle cells, a jelly coat, and a vitelline layer; their morphologies are described. Methods are outlined for systematically removing them without altering the behavior of the oocyte so that the cell surface can be examined directly. The topography of denuded oocytes changes dramatically when they are treated with the maturation-inducing hormone, 1-methyladenine. The major topographical change is the early and transient formation of prominent surface spikes. These structures arise due to the rapid, reversible polymerization of actin into stout bundles. Polymerization and subsequent depolymerization of cortical actin is monitored by epifluorescence microscopy of oocytes stained with NBD-phallacidin, a stain which is specific for polymerized actin. Based on scanning electron microscopy, spikes apparently utilize preexisting plasma membrane of microvilli, and plasma membrane is apparently lost when spikes collapse. Long after microvilli are eliminated due to spike formation, the number of microvilli is somewhat restored, especially around the animal pole where the polar body forms. A chronology of events observed during oocyte maturation is discussed with reference to the possible mechanisms and implications of polymerization and depolymerization of cortical actin. INTRODUCTION
In starfish (Echinodermata: Asteroidea), as in most animals including the vertebrates, eggs are stored in the ovary as immature oocytes that are arrested at meiotic prophase. Maturation of starfish oocytes is reinitiated under hormonal stimulation in synchrony with ovulation, and the oocytes become fertilizable at spawning, or soon afterward. Accordingly, starfish oocytes taken from the ovary represent an important model system for investigating the processes of maturation. Cytological studies of maturation of starfish oocytes are facilitated by a rich background of information. The hormonal triggers for maturation are known with certainty (Kanatani, 1973). The receptors for the maturation-inducing hormone, 1-methyladenine (l-MA), have been localized at the plasma membrane surface (Kanatani and Hiramoto, 1970; Guerrier and Doree, 1975); suggestions have been made about their morphological manifestation (Schroeder, 1981); and isolation of the putative receptors has been achieved (Morisawa and Kanatani, 1978; Ikadai and Kanatani, 1982). In addition, progress is being made toward defining the various physiological steps in the transduction process from hormone to cytoplasmic alteration (Guerrier et aL, 1977; Moreau and Guerrier, 1980; Mazzei et aL, 1981; Meijer ‘The authors wish to dedicate this paper to the memory fessor Robert L. Fernald (1914-1933).
of Pro-
and Guerrier, 1981; Doree et al, 1981a; Johnson and Epel, 1982) and determining the cytoplasmic factors essential for maturation (Kishimoto et aL, 1981). To this background of knowledge, we add observations of dynamic changes in the surface topography and the organization of cortical actin during maturation. Our objectives are (1) to describe methods for handling starfish oocytes for successful scanning electron microscopy (SEM), (2) to document morphological changes in maturing oocytes as a possible basis for understanding some of the physiological changes mentioned above, and (3) to supplement previous observations concerning “spikes,” the prominent structures that appear transiently on the oocyte surface in response to l-MA (Schroeder, 1981). To test a hypothesis that cortical actin plays a role in spike formation (Schroeder, 1981), we have stained cortical actin with NBD-phallacidin. This fluorescent stain is specific for polymerized actin (Barak et aL, 1980) and has been used to study actin organization in various cell types (Nothnagel et aL, 1981; Schroeder and Christen, 1982; Schroeder, 1982). MATERIALS
AND
METHODS
Adult specimens of Pisaster ochraceus were collected intertidally around San Juan Archipelago, Washington, and were maintained until needed in running seawater at ll-15’C. All operations and observations were con-
373 0012-1606/83 $3.00 Copyright All rights
0 1983 by Academic Press, Inc. of reproduction in any form reserved.
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DEVELOPMENTALBIOLOGY
ducted at seawater temperatures or at 20-22°C (room temperature), as specified. To obtain ovaries, an arm was severed from the base, both ovaries in the arm were removed at the short oviducts, and they were then stored in seawater. Gravid ovaries typically measured 5-8 cm in length and were obtained from April to early August. Immature oocytes with their investing follicle cells were obtained by teasing several ovarian lobules (about 1 cm long). Fragments of ovarian tissue were removed and the oocyte-follicle cell complexes were washed in seawater. This method resulted in the “spontaneous” breakdown of germinal vesicles in -5% of the oocytes, probably as a consequence of mechanical trauma. To remove follicle cells, oocyte-follicle cell complexes were washed in five changes of calcium-free seawater (CaFSW) consisting of 0.38 M NaCl, 49 mM MgClz, 26 mM Na2S04, 8.5 mM KCI, 2.25 mlM NaHC03, 2.5 m&Z EGTA, and 10 mM Tris-HCl buffer with the final pH adjusted to 8.0-8.2. Oocytes free of follicle cells were also obtained by directly rinsing ovarian lobules and fragments in several changes of CaFSW for 1 hr. By adding natural seawater (containing calcium), the fragments contracted, and oocytes without follicle cells were extruded (Kanatani and Shirai, 1969; Cloud and Schuetz, 1979). Very little ‘*spontaneous” breakdown of germinal vesicles was observed using this method. Ovarian fragments were removed by passage through cheesecloth; the oocytes were then rinsed a few times with CaFSW and stored. As monitored by Nomarski (Fig. 2b) and SEM (Fig. 3b), CaFSW alone does not remove any investing coat other than the follicle cells. In our preparations the thick jelly coat and vitelline layer were routinely dissolved together by protease digestion after pretreatment with 10 PM ionophore A23187 (Eli Lilly & Co., Indianapolis, Ind.) which causes premature elevation of the vitelline layer. A 25-~1 volume of ionophore A23187 (1 mg/ml in ethanol) was first added to -50,000 oocytes without follicle cells in 5 ml of CaFSW in a Syracuse watch glass. At 11-15°C vitelline layers were fully elevated by 15 min, and the medium was then mostly removed. Protease solution in CaFSW (5 ml) was added. The oocytes were quickly stirred and then allowed to remain undisturbed as a monolayer for 10 min. The enzyme solution in our early studies was 1% protease (Type V, Sigma Chemical Co., St. Louis, MO.; No. P-5005, 1100 PU/g), but when this product became unavailable we used 0.1% Pronase (Calbiochem-Behring Co., La Jolla, Calif.; No. 53702,56,000 PU/g) with equal success. We never observed that digestion caused parthenogenetic activation, as reported by Bryan and Sato (1970), or that it significantly altered oocyte sensitivity to lMA (Jeffery, 1977). After 10 min of digestion the enzyme
VOLUME98.1983
solution was cautiously removed, CaFSW was gently added, and the CaFSW was changed twice. The oocytes were somewhat adherent soon after digestion; if disturbed at this stage, they tended to lyse, until they had been thoroughly washed. Dose dependencies for germinal vesicle breakdown (GVBD) in oocytes treated with l-MA were determined at selected temperatures by adding 25 ~1 of oocyte suspension containing 100-200 oocytes to 0.2-ml volumes of serially diluted l-MA solutions in seawater or CaFSW. Oocytes must be exposed to l-MA for a definite period of time in order to undergo GVBD, as first suggested by the work of Schuetz (1969) and then substantiated by Guerrier and Doree (1975). The hormone-dependent period (HDP) is defined as the minimum time of exposure to a threshold concentration for 50% GVBD to occur. The HDP for Pisaster oocytes was determined by (1) treating about 2000 oocytes in 5 ml of medium with 10m6Ml-MA at selected temperatures, (2) removing 50~1 aliquots (containing at least 200 oocytes) at 1 to 2min intervals, and (3) rapidly diluting each aliquot into 4 ml of seawater or CaFSW. The oocytes were immediately sedimented by hand centrifugation and resuspended in another 4 ml of seawater. The extent of GVBD was determined by counting at either 90 min (11-15’C) or 60 min (20-22°C) after the onset of l-MA treatment. For scanning electron microscopy, oocytes were fixed first in 1% glutaraldehyde-90% seawater (or CaFSW) at pH 7.4 for 1 hr at room temperature. Oocytes were then rinsed briefly in seawater (or CaFSW) and postfixed in 1% osmium tetroxide-75% seawater (or CaFSW) at pH 8.0-8.2 for 0.5-l hr. Specimens were partially dehydrated in an ethanol series and then completely dehydrated in dimethoxypropane (Maser and Trimble, 1977), critical point dried from carbon dioxide, cemented to specimen stubs with silver paint, and sputter-coated with gold-palladium (60-40%). A JEOL JSM35 scanning electron microscope was used for observations. For staining polymerized actin in the cortex, oocytes were prefixed in 1% formaldehyde-90% CaFSW at pH 8.0-8.2 for 10 min. A single drop containing about 100 oocytes was then added to 2 drops of stain made up as follows: 25 ~1 of 1 mg/ml palmitoyl lysolecithin, 25 ~1 of 3 pg/ml nitrobenzoxydiazole-conjugated phallacidin (NBD-phallacidin, Molecular Probes, Inc., Plano, Tex.) in methanol, and 0.45 ml of CaFSW. After staining for 10 min, the oocytes were mounted under supported coverslips to avoid compression. The oocytes were viewed by epifluorescence microscopy, primarily by focusing on the upper surfaces. Fluorescence was detected by incident illumination at 490 nm supplied by a 100-W tungsten-halogen lamp in a Zeiss Universal microscope equipped with a 500-nm dichroic reflector, a 63X (N.A.
SCHROEDER
AND
STRICKER
CeU Surface
1.25) objective lens, and a 52%nm barrier filter. Fluorescent images were recorded on Kodak Tri-X Pan film (ASA 400) with 15-set exposures and were printed under identical conditions. RESULTS
A ripe female Pisaster ochraceus yields a large number of oocytes which are uniform in appearance and quite transparent. When still surrounded by follicle cells, the oocytes are irregular in shape (Fig. la), but they become spherical soon after being washed in CaFSW, a process that also removes the follicle cells. The fully grown immature oocyte is about 160 pm in diameter
and Cortical Actin of Starfish Oocytes
375
and contains a large clear germinal vesicle that is about ‘75pm in diameter. The germinal vesicle is eccentrically situated so that it lies 3-5 pm from the oocyte plasma membrane at one point (the animal pole), and contains a prominent nucleolus (Fig. 2). By focusing on the uppermost surface of an oocyte with the Nomarski microscope (not shown), the oocyte cortex is seen to contain a layer of clear vacuoles -2 pm in diameter with smaller cortical granules interspersed among the vacuoles. Many, but frequently not all, of the cortical granules are released by activation with ionophore A23187, but the vacuoles do not change. Other organelles and inclusions appear to be uniformly distributed throughout the remaining cytoplasm. Investing Coats
FIG. 1. Low-power
photomicrographs of oocytes from the starfish oocytes and their investing follicle cells as they appear shortly after dissection from the ovary. At this stage oocytes are irregular in shape and the follicle cell investments (arrowheads) are often incomplete. (b) Mature oocytes at the time of first polar body formation. In this preparation the investing coats had been removed and the oocytes were then treated with l-methyladenine to reinitiate meiosis. Maturation events proceed with high synchrony, although a few cells (*) in a batch often fail to mature. Scale bar = 100 pm. Magnifications = 59X.
Pisaster ochramus. (a) Immature
Follicle ce2Ls.A single layer of follicle cells (Figs. la and 2a) surrounds immature oocytes that are dissected from ovaries into seawater. This layer is often incomplete; the larger gaps may be produced during the dissection procedure. Each follicle cell is less than 0.5 pm thick except at its thicker perikaryon from which a slowly beating flagellum emerges. There appear to be about 25 to 75 follicle cells associated with each oocyte. They are roughly polygonal in shape and measure about 25 X 45 pm, according to superficial views by Nomarski microscope (Schroeder, 1981) and by SEM (Fig. 3a, inset). Follicle cells often do not exhibit broad areas of contact along their lateral margins; consequently, the underlying jelly coat is sometimes visible between the cells (Fig. 3a). As shown in Fig. 2a, the follicle cells are separated lo-15 pm from the oocyte surface by the intervening jelly layer. Each follicle cell is connected to the underlying oocyte by over 100 thin follicle cell processes which traverse the jelly layer (Runnstrom, 1944; Schroeder et ah, 1979; Schroeder, 1981). These can sometimes be seen by SEM (Fig. 3a). Insufficient washing in CaFSW causes some of these processes to remain embedded in the jelly coat, where they can be confused for spikes when seen in the light microscope; remnant processes are also sometimes seen by SEM (Fig. 3~). Although intercellular contacts between follicle cells and oocytes occur while the oocytes are still in the ovary (unpublished observations), the attenuated shape of the follicle cell processes probably results from the swelling of the jelly coat. During natural spawning, oocytes are released into seawater without investing follicle cells, which are retained in the ovary after follicle cell-oocyte contacts are separated. Jelly coat. The jelly coat swells to a thickness of lo15 pm upon contact with seawater. Most of the jelly coat appears fibrous in organization by Nomarski microscopy (Figs. 2a and b), although an extremely trans-
FIG. 2. Nomarski photomicrographs of starfish oocytes at 12°C after stepwise removal of investing coats (a-d) and progressively after lMA treatment (e-h). In each case only the animal hemisphere of the spherical oocyte is shown. The large germinal vesicle (gv) with its prominent nucleolus is visible (a-f), until later stages of maturation when these structures have disintegrated (g, h). (a) An oocyte dissected from the ovary into seawater with its fibrous jelly coat (j) and investing layer of follicle cells (fc). (b) In calcium-free seawater the follicle cells dissociate and the jelly coat swells but remains present. (c) Ionophore A23187 causes the vitelline layer (VI) to elevate. (d) Protease digestion in calcium-free seawater removes the vitelline layer and jelly coat, thus exposing the plasma membrane directly to the medium. (e) At 10 min after application of lo-’ Ml-MA, radiating projections called spikes (sp) are visible on the surface. (f) By 22 min after l-MA treatment, spikes at the surface have vanished. (g) By 65 min after l-MA treatment the germinal vesicle has broken down leaving in its place a somewhat yolk-free region (*). (h) The first polar body (pbl) forms at about 165 min after l-MA treatment; in this figure at 215 min the second meiotic apparatus is visible as a clear zone beneath the first polar body. The scale bar = 10 pm. Magnification = 335X.
parent component of the jelly gradually expands beyond the fibrous zone after oocytes remain in seawater or CaFSW for a few hours. In the SEM the jelly coat resembles a tangled mat of fine fibers (Figs. 3a and b). We do not know, however, how much of this organization is a product of specimen preparation. In our experience neither CaFSW or acidification to pH 4.15 dissolves the jelly coat, contrary to reports elsewhere (Jeffery, 19’77; Shoji et aZ., 19’78; Nakamura and Hiramoto, 19’78; Nemoto et aZ., 1980). Lowered pH does cause the jelly coat to dissipate temporarily, but it reappears when the pH is again raised. We find that it is necessary to remove the jelly coat along with the vitelline layer by digestion in protease, as described under Materials and Methods. Vitelline layer. The vitelline layer of Pisaster oocytes is about 0.25 pm thick, according to transmission electron micrographs (Schroeder, 1981), and thus is much thicker than its counterpart on sea urchin eggs (Schroeder, 1979). This layer was seldom observed by SEM, because we did not specifically endeavor to expose it by removing the jelly coat alone; nevertheless, in some
preparations the protease treatment digested only the jelly coat, while leaving the vitelline layer relatively intact (Fig. 3~). In such preparations, the vitelline layer appears quite smooth on its outer surface, although it has been shown that microvilli are embedded in the vitelline layer from below, before elevation occurs (Hirai et al., 1971; Cayer et ak, 1975; Schroeder, 1981). Sometimes we see small pores in the vitelline layer (Fig. 3~) through which follicle cell processes apparently project as they course toward the oocyte surface. These holes, however, are usually not visible. Thus, it is likely that the vitelline layer material can rearrange itself and obscure the holes left when the follicle cell-oocyte contacts are broken and the follicle cell processes withdraw. We have had little success using 1 M urea to remove the vitelline layer, in contrast with studies of oocytes from other species (Dan-Sohkawa, 1976). In our hands, urea only caused an occasional rupture of the vitelline layer, thereby revealing a portion of the oocyte surface. The microvilli in these cases appeared somewhat longer and more erect than after protease digestion.
SCHROEDER AND STRICKER
Cell Surface
and Cortical
Actin
of Starfish
Oocytes
FIG. 3. Scanning electron micrographs of various surfaces exposed during the removal of investing coats surrounding Pim.steroocytes. The insets in the center illustrate the corresponding surfaces at low magnification. (a) Each follicle cell (fc) surrounding the oocyte possesses a single flagellum (fg); from the undersides of these cells, thin follicle cell processes (arrowheads) project toward the oocyte through the fibers of the jelly coat (jc) which is visible in the gaps between follicle cells. (b) In calcium-free seawater the jelly coat is entirely exposed and appears to be composed of fibers that form a thick mat. (c) In a few specimens, digestion in protease removed the jelly coat without removing the vitelline layer; wrinkles in the vitelline layer may be due to its elevation and expansion caused by ionophore A23187 treatment and to its subsequent collapse during preparation. Several long processes pass through small pores in the vitelline layer (arrowheads) are presumed to be fragmentary follicle cell processes still in contact with the oocyte; these are not usually seen and probably result from inadequate rinsing in CaFSW. Microvilli on the oocyte surface (mv) can be seen through a small tear in the vitelline layer. (d) After the complete digestion of the vitelline layer, the oocyte plasma membrane is entirely exposed; numerous short microvilli cover its surface. Scale bar = 1 pm (insets, 10 Mm). Magnification = 5400X (insets, 660X).
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Hormone Requirements and the Course of Maturation The threshold concentration of l-MA for Pisaster oocytes in vitro, i.e., the concentration at which 90+% undergo germinal vesicle breakdown (GVBD), is typitally 1 X lop6 M. It is temperature independent from 11 to 22°C and is the same in natural seawater or ar-
VOLUME98,1983
tificial CaFSW. The schedule for meiotic maturation, however, is distinctly temperature dependent. At -12°C GVBD begins at 55 min; the long-lasting first meiotic spindle is set up somewhat later; and, the first polar body is emitted at -165 min (Fig. 2). At -21°C GVBD begins 26 min after treatment with l-MA. The hormone-dependent period for Pisaster oocytes
FIG. 4. Changes in surface topography after application of l-MA at -12°C. (a) Before l-MA treatment the oocyte surface is uniformly covered with short microvilli. (b) Within 1 min after l-MA treatment, triangular projections -2 pm long emerge from the surface as microvilli begin to disappear. (c) By 10 min after l-MA the projections are transformed into definitive spikes measuring 3-5 pm in length (they are up to 10 pm long in living oocytes) and microvilli have been reduced to tiny bumps. (d) By 30 min after l-MA the formerly erect spikes have been replaced by thinner, longer, and flaccid processes which are interpreted as being the membranous remnants of the spikes without their rigid contents. Microvilli are virtually absent at this stage. Several visible depressions, which can also be seen before l-MA treatment, probably represent collapsed vacuoles that occupy the cortex of these oocytes. Scale bar = 1 pm. Magnification = 6790X.
SCHROEDER AND STRICKER
Cell Surface
the HDP is 11-12 at -12°C is 27-31 min. At -2l”C, min. These values suggest that the HDP is a constant fraction of the time to GVBD. At a given temperature, the HDP is the same for oocytes in seawater or CaFSW. The HDP is unchanged by the presence or absence of oocyte investing coats and is not affected by higher concentrations of l-MA.
Early Surface Responses to l-MA The surface topography of Pisaster oocytes is rapidly and dramatically altered by treatment with l-MA. Qualitatively, these changes include the appearance of long, straight spikes, the simultaneous loss of microvilli, resorption of the spikes, and eventually a partial restoration of microvilli. Some of these events are discernible by light microscopy (Fig. 2), while others can only be appreciated by SEM (Figs. 4 and 5). Accurate quantification of these changes is complicated by the shrinkage of the specimens that typically
and Cortical Actin of Starjsh
Oocytes
379
occurs during preparation for SEM, as discussed fully elsewhere (Schroeder, 1979). Shrinkage reduces the dimensions of structures but presumably does not effect their numbers. In our SEM micrographs the overall oocyte diameter is -35% less than it is in life (100 pm vs 160 pm). We do not know, however, if this shrinkage is isometric at the level of the microvilli. Thus, we will simply cite measurements obtained from representative SEM micrographs, without correcting for shrinkage. Before treatment with l-MA the cell surface is uniformly covered with about 2.5 X lo5 short microvilli. In the SEM these microvilli measure -0.1 pm in diameter and -0.35 pm in length (Figs. 3d and 4a). Very soon after l-MA application, the oocyte surface begins to undergo changes that culminate in the transient appearance of spikes (Schroeder, 1981). The following description refers to the schedule of events at -12’C. Several minutes elapse before spikes can be seen by Nomarski microscopy (Fig. 2e), but at even earlier
FIG. 5. Fluorescence micrographs of l-MA-treated oocytes that have been fixed and stained with NBD-phallacidin, a stain that is specific for polymerized actin. The times after l-MA correspond to those illustrated in Fig. 3 by SEM. (a) An extremely low level of fluorescence is detectable before l-MA. (b) Within 1 min after l-MA application an extensive meshwork of thin polymerized actin bundles -2-pm-long begins to appear. (c) By 10 min after l-MA, when spikes are longest and most prominent, numerous bold actin bundles up to 10 pm long appear all over the oocyte surface. (d) By 30 min after l-MA, there is no evidence of polymerized actin by fluorescent staining with NBDphallacidin. Scale bar = 10 pm. Magnification = 1455~.
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DEVELOPMENTAL BIOLOGY
times there are topographical changes which are obvious by SEM. Within 1 min after treatment, the surface displays broad leaf-like projections (Fig. 4b); these are more or less triangular in shape and -2 pm on a side, bearing on their edges short projections that are reminiscent of microvilli. Microvilli are still visible between these projections (Fig. 4b), but they are much reduced in length. At about 10 min after l-MA application, the leaf-like projections are gone, and the oocyte surface is covered by about 6500 spikes. These structures are very straight, abruptly pointed, -0.3 pm in diameter, and 3-5 pm long when viewed by SEM (Fig. 4~). In living oocytes they are frequently 10 pm long. In Fig. 4c the density of fully formed spikes is about -0.3/pm2. By this stage, microvilli are further reduced to small, sparse bumps. Spikes can be seen to withdraw soon after they are fully formed. By 20 min after l-MA application very few spikes are visible by Nomarski (Fig. 2f), and by 30 min none are visible even by SEM (Fig. 4d). The stout, straight spikes are replaced by thin, curved, and seemingly flaccid projections that lie along the oocyte surface (Fig. 4d). These are not as numerous as spikes were at 10 min, and they soon disappear entirely. They undoubtedly represent membranous remnants of spikes that have lost their rigidity. Between these flaccid projections there are a few small bumps that probably represent shortened microvilli.
Cortical Actin aJter Treatment with l-MA The early-surface responses to l-MA, described above, are correlated with dramatic changes in actin of the oocyte cortex, as indicated by patterns of staining with NBD-phallacidin. Before l-MA is applied, the cortex displays only a very low level of fluorescence (Fig. 5d). Within 1 min after application of l-MA, an elaborate meshwork of very thin fluorescent lines is present throughout the cortex (Fig. 5b). These lines are -2 pm long and apparently represent bundles of polymerized actin in an early stage of formation. They do not occur in the deeper cytoplasm. By 10 min after treatment with l-MA, the putative actin bundles are fewer in number, but they are longer, thicker, and more boldly fluorescent (Fig. 5~) than at 1 min. These bright needles of fluorescence are 4-12 pm in length and occur only in the cortex as projections from the oocyte surface. Their location, density, and time of appearance correlate with those of the definitive spikes seen by Nomarski or SEM. By 30 min after lMA application, when spikes are completely withdrawn, no fluorescence can be detected in whole starfish oocytes stained with NBD-phallacidin (Fig. 5d). Thus, there is evidence of even less polymerized cortical actin at this stage than before l-MA is applied.
VOLUME 98, 1983
Polar Body Formation Very little change in the topography of starfish oocytes occurs between elimination of the flaccid projections and formation of the first polar body (Fig. 6). Microvilli gradually reappear, although never to the density seen before application of l-MA (Figs. 3d and 4a). The microvilli become slightly polarized in distribution on the oocyte shortly before polar body formation. Microvilli are longer and slightly more dense within a circular zone about 75 pm in diameter around the animal pole where the polar body forms (Figs. 6a and b). Within this zone (comprising less than 10% of the oocyte surface), microvilli are 0.5 pm long and occur at a density of -3.5/pm2 (Fig. 6b). The polar body itself usually exhibits microvilli that are about the same length but are sparser (Fig. 6b). Outside this polar zone, microvilli are only -0.15 pm long and occur at a density of -3.0/pm2 (Fig. 6~). Staining with NBD-phallacidin at polar body stages yields a mottled pattern of weak fluorescence surrounding the polar body (Fig. 6d). A minor topographical feature that is also affected by l-MA is an array of surface depressions which are -2 brn in diameter and visible at all stages of maturation (Figs. 4a-d, and 6~). They presumably represent the clear vacuoles situated in the cortex and have collapsed during peparation for SEM. The surface overlying these depressions sometimes becomes perforated during specimen preparation, particularly at the time of germinal vesicle breakdown and at polar body stages (Fig. 6~). DISCUSSION
Surface Ultrastructure and the Control of Actin Polymerixation Using SEM to observe surface changes of experimentally denuded oocytes, we find that the immature oocyte is thoroughly covered with short microvilli. The hormone l-MA induces a rapid reorganization of the surface which culminates after 10 min in long spikes, previously described elsewhere (Rosenberg, 1978; Schroeder, 1981). Microvilli are quantitatively eliminated as spikes form. The loss or retraction of microvilli during oocyte maturation has been reported for other species of starfish (Hirai et aL, 1971; Cayer et aL, 1975; Rosenberg, 1978; Hirai and Shida, 1979). Based on the numbers and dimensions of microvilli and spikes (see Results), the total surface area of an oocyte is constant during spike formation, suggesting that spikes “borrow” plasma membrane from the microvilli. Our observations with NBD-phallacidin confirm an earlier hypothesis that spikes contain dense bundles of polymerized actin (Schroeder, 1981). Indeed, actin polymerization begins within a minute, and the forces which
SCHROEDER AND STRICKER
Cell ,%&ace and Cortical Actin of Star$sh
Ooc@es
FIG. 6. First polar body stage at 165 min after l-MA treatment. (a) Survey SEM view of the small first polar body on the surface of the oocyte. (b) A first polar body (pbl) as it is pinching off from the rest of the oocyte; this area corresponds to the trapezoidal box in a. Relatively long microvilli are present in this region of the oocyte surface. (c) At a location 90” away from the first polar body, in a region corresponding to the rectangular box in a, microvilli are much shorter and slightly less numerous; depressions similar to those in Figs. 4a and d sometimes appear to be perforated at this stage (arrowheads). (d) Staining with NBD-phallacidin reveals a weak fluorescence that outlines the polar body (pbl). Scale bars a and d = 10 pm; b and c = 1 Frn. Magnifications: a = 64 OX; b and c = 6790X; d = 1455X.
cause spikes to protrude may derive from this polymerization and assembly into bundles. Prior to l-MA treatment, fluorescence due to stained polymerized actin is extremely low, though present; perhaps it is attributable to actin in microvilli, although our transmission EM observations have not yet revealed polymerized actin at this stage. Spikes are withdrawn by 30 min and no detectable sign of polymerized actin remains. When spikes collapse there appears to be a loss of cell surface (i.e., plasma membrane). The cell surface remains nearly smooth until microvilli partially reappear by the time of first polar
body formation. Even then, microvilli are still fairly sparse and are distributed in a polarized fashion, perhaps reflecting cortical movements (Hamaguchi and Hiramoto, 1978). Our rough calculations indicate that the total cell surface area at this stage is still 40% less than in the immature oocyte. The mechanisms by which l-MA causes actin polymerization or depolymerization are not known. There is evidence that an elevation of intracellular pH is at least part of the trigger for actin polymerization in cells (Tilney et ak, 19’78; Carron and Longo, 1982; Begg et al, 1982; Schroeder and Christen, 1982). In starfish oocytes,
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DEVELOPMENTALBIOLOGY
however, treatment with l-MA does not cause intracellular pH to rise (Johnson and Epel, 1982), yet it triggers the actin polymerization involved in spike formation. Furthermore, application of permeant weak bases which can elevate intracellular pH fails to induce the formation of spikes (Schroeder, unpublished observations); indeed, rather than mimicking l-MA, such weak bases actually inhibit l-MA-induced germinal vesicle breakdown (Cloud and Schuetz, 1979; Guerrier et al, 1978; Doree et ah, 1982). Thus, the triggering of actin polymerization in starfish oocytes may involve complex factors.
Identijcation
and Efects of Removing Investing Coats
In order to study the surface ultrastructure of oocytes, we devised methods for systematically removing investing layers (follicle cells, jelly coat, and vitelline layer) without altering the responses of oocytes to lMA. We have cleared up previous misinterpretations of these investments (Rosenberg et aL, 1977; Lee et al, 1977) which apparently arose through the unwarranted belief that CaFSW removes the jelly coat. On the other hand, we do confirm the findings of Cayer et al. (1975) and Cloud and Schuetz (1979) that CaFSW can completely remove the follicle cells and their processes, provided the oocytes are rinsed sufficiently. In the case of Pisaster oocytes, complete removal of the jelly coats and vitelline layers requires protease digestion after ionophore treatment. Responses of P&aster oocytes to l-MA proved to be identical before or after removing the investments and regardless of being cultured in natural seawater or CaFSW. These responses include (1) surface spike formation and withdrawal; (2) the time course, rate, and extent of maturation; (3) the threshold concentration of l-MA; and (4) the duration of the hormone-dependent period. We fail to confirm observations by Rosenberg and Lee (1981) that both the threshold concentration and the hormone-dependent period are specifically and significantly altered by the calcium content of the medium.
Relationship of Cortical Actin to Other Ph@ological Events Relating spike formation to other l-MA responses or to oocyte maturation is problematical. Spike formation is not necessary for germinal vesicle breakdown (Schroeder, 1981), yet both events share l-MA as their specific inducer; in fact, a single class of l-MA receptors may be involved in both responses, since the hormone specificities (using analogs of l-MA) and hormone sensitivities (using experimentally altered oocytes) of the two responses are indistinguishable (T. E. Schroeder and H. Ikadai, unpublished observations).
VOLUME 98. 1983 Ca2+ burst Actin polymerization
Actin depolymerization Phosphorylation of cortical proteins
x 4 $ kJ I
50-
End of hormone dependent period
60-
Membrane depolarization
k4
70-
w I I=
80-
OOJ
Cortical relaxation
Germinal vesicle breakdown
FIG. ‘7. A timetable of molecular and biophysical events that occur during maturation of starfish oocytes. The data are drawn from published reports of various species studied under a variety of conditions. The time scale is normalized (Time 0 corresponds to the application of l-MA and Time 100 represents the beginning of germinal vesicle breakdown) so that the timing of various events can be compared.
A normalized timetable of starfish oocytes is presented in Fig. 7. It summarizes the temporal relationship between actin polymerization and various other changes that occur during maturation. We have assumed (perhaps simplistically) that chronological events between l-MA application and germinal vesicle breakdown can be proportionalized and that comparisons between species are valid. The earliest known response to l-MA is a burst of intracellular free calcium ions that occurs within seconds (Moreau et ak, 1978). Putative actin polymerization and the associated topographical changes begin soon after the calcium burst (this study; Schroeder, 1981). When surface spikes reach their maximum extension, several events begin almost simultaneously: actin depolymerization starts (this report); protein kinase activity rises and phosphorylation of cortical proteins reaches a plateau (well before a steady state is achieved in the endoplasm) (Guerrier et aZ., 1977); the membrane potential declines from -70 to less than -20 mV by the time that germinal vesicle breakdown occurs (Miyazaki et al, 1975); and mechanical rigidity of the cortex rapidly declines (Nakamura and Hiramoto, 1978, Nemoto et aL, 1980). These changes appear to begin before the end of the hormone-dependent period, the time at which
SCHROEDER AND STRICKER
CeU Surface
the cell becomes irreversibly committed to undergo germinal vesicle breakdown. Very little is known about events between the end of the hormone-dependent period and the germinal vesicle breakdown; high activity of maturation-promoting factor appears only after germinal vesicle breakdown occurs (Kishimoto et al, 1981). The timetable presented in Fig. 7 provides a basis for exploring causal mechanisms in the cellular responses to l-MA, although we recognize that sequential events need not be causally linked. With regard to possible mechanisms of actin polymerization, the fact that a calcium burst is the only cytoplasmic event known to precede actin polymerization suggests that these two events may be linked; actin polymerization could be controlled by a calcium-sensitive regulator. On the other hand, spikes do not form merely by treating oocytes with ionophore A23187, so the potential role of calcium is not simple. Actin depolymerization and spike withdrawal coincide temporally with several known biochemical changes (Fig. 7). Phosphorylation of diverse proteins in the cortex seems to be a particularly important event during the hormone-dependent period (Guerrier et cd, 1977). It is possible that actin depolymerization results from phosphorylation of a key protein, for example, by activating a regulatory protein. Such a mechanism for regulating actin is not incompatible with the idea that protein phosphorylation is also involved in reinitiating meiosis. It is noteworthy that :Mabuchi (1981) has recently isolated from starfish oocytes a 17,000-dalton protein called “depactin” which causes rapid actin depolymerization in vitro. It is not yet known if depactin’s activity is altered by phosphorylation. Analyses of the phosphoproteins which appear soon after l-MA application (Preddie and Abastado, 1979; Doree et al., 1981b; Mazzei and Guerrier, 1982) sometimes reveal a protein about the size of depactin, but they are otherwise somewhat conflicting. Such studies set the stage for further work that could clarify the mechanisms and interrelations of spike formation, actin assembly-disassembly, and the processes leading to meiotic reinitiation in starfish oocytes. This study was supported by the U. S. Public Health Service in a suballocation of a Biomedical Research Support Grant RR 07096 to the University of Washington and by NIH Research Grant GM 19464. REFERENCES BARAK, L. S., YOCVM, R. R., NOTHNAGEL, E. A., and WEBB, W. W. (1980). Fluorescence staining of the actin cytoskeleton in living cells with 7-nitrobenz-2-oxa-3,3-diazole phallacidin. Proc Nat Acad. Sci USA 77.980-984. BEGG, D., REBHUN, L. I., and HYA’IT, H. (1982). Structural organization of actin in the sea urchin egg cortex: Microvillar elongation in the absence of actin filament bundle formation. J. CeU. Biol 93, 24-32.
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Actin
of Star&sh
Oocytes
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BRYAN, J., and SATO, H. (1970). The isolation of the meiosis I spindle from the mature oocyte of Pisaster ochraceus. Exp. Cell Res. 59, 371-378. CARRON, C. P., and LONGO, F. J. (1982). Relation of cytoplasmic alkalinization to microvillar elongation and microfilament formation in the sea urchin embryo. Dev. Biol 89, 129-137. CAYER, M. L., KISHIMOTO, T., and KANATANI, H. (1975). Formation of the fertilization membrane by insemination of immature starfish oocytes pretreated with calcium-free seawater. Deuelop. Growth D$ feren 17,119-B%. CLOUD, J. G., and SCHUETZ, A. W. (1979). 1-Methyladenine induction of oocyte (starfish) maturation: Inhibition by procaine and its pH dependency. J. Exp. Zool. 210,11-16. DAN-S• HKAWA, M. (1976). A “normal” development of denuded eggs of the starfish, Asterina pectin$ero. Den Grwth D&?ren. 18,439445. DOREE, M., KISHIMOTO, T., LE PEUCH, C. J., DEMAILLE, J. G., and KANATANI, H. (1981a). Calcium-mediated transduction of the hormonal message in meiosis reinitiation of starfish oocytes. Exp. Cell Res. 135, 237-249. DOREE, M., CAVADORE, J.-C., LE PEUCH, C. J., and DEMAILLE, J. G. (1981b). Hormone-induced phosphorylation of a 16,000 dalton polypeptide following meiosis reinitiation in starfish oocytes. Exp. Cell Res. 135, 251-257. DOREE, M., SANO, K., and KANATANI, H. (1982). Ammonia and other weak bases applied at any time of the hormone-dependent period inhibit 1-methyladenine-induced meiosis reinitiation of starfish oocytes. Dev. Biol SO, 13-17. GUERRIER, P., and DOREE, M. (1975). Hormonal control of reinitiation of meiosis in starfish: The requirement of 1-methyladenine during nuclear maturation. Dev. Biol 47. 341-348. GUERRIER, P., MOREAU, M., and DOREE, M. (1977). Hormonal control of meiosis in starfish: Stimulation of protein phosphorylation induced by 1-methyladenine. Mol. Cell Endocrinol. 7, 137-150. GUERRIER, P., MOREAU, M., and DOREE, M. (1978). Control of meiosis reinitiation in starfish: Calcium ion as the primary effective trigger. Ann. Biol. Anim. Bid Biophys. 15,441-452. HAMAGUCHI, M. S., and HIRAMOTO, Y. (1978). Protoplasmic movement during polar-body formation in starfish oocytes. Exp. Cell Res. 112, 55-62. HIRAI, S., KUBOTA, J., and KANATANI, H. (1971). Induction of cytoplasmic maturation by 1-methyladenine in starfish oocytes after removal of the germinal vesicle. Exp. Cell Res. 68, 137-143. HIRAI, S., and SHIDA, H. (1979). Shortening of microvilli during the maturation of starfish oocyte from which vitelline coat was removed. Bull Mar. Biol Sta Asamushi, Tohoku Univ. 16, 161-167. IKADAI, H., and KANATANI, H. (1982). Regeneration of oocyte-surface factor responsible for 1-methyladenine-induced oocyte maturation in starfish. Biomed Res. 3, 378-383. JEFFERY, W. R. (1977). Hormonal action of 1-methyladenine: The effect of enzymatic digestion of intact starfish oocytes on the induction of meiosis. Gen Comp. Endocrinol 31, 259-269. JOHNSON, C. H., and EPEL, D. (1982). Starfish oocyte maturation and fertilization: Intracellular pH is not involved in activation. De-v. Biol 92,461-469. KANATANI, H. (1973). Maturation-inducing substance in starfishes. Int. Rev. Cytd 35, 253-298. KANATANI, H., and HIRAMOTO, Y. (1970). Site of action of l-methyladenine in inducing oocyte maturation in starfishes. Exp. Cell Res. 61, 280-284. KANATANI, H., and SHIRAI, H. (1969). Mechanisms of starfish spawning. II. Some aspects of action of a neural substance obtained from radial nerve. Biol. BuU 137, 297-311. KISHIMOTO, T., HIRAI, S., and KANATANI, H. (1981). Role of germinal
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vesicle material in producing maturation-promoting factor in starfish oocyte. Dev. BioL 81, 177-181. LEE, H. H., ROSENBERG, M. P., and HOESCH, R. (1977). The starfish (Patiria and Pisaster) oocyte: A model system for the study of hormone-surface interaction in oogenesis. Lab. Anim Sci 27,860-865. MABUCHI, I. (1981). Purification from starfish eggs of a protein that depolymerizes actin. J. B&hem 89, 1341-1344. MASER, M. D., and TRIMBLE, J. J. (1977). Rapid chemical dehydration of biologic samples for scanning electron microscopy using 2,2-dimethoxypropane. J. Histochem. Cytochewz. 25, 247-251. MAZZEI, G., MEIJER, L., MOREAU, M., and GUERRIER, P. (1981). Role of calcium and cyclic nucleotides during meiosis reinitiation in starfish oocytes. Cell mere% 10, 139-145. MAZZEI, G., and GUERRIER, P. (1982). Changes in the pattern of protein phosphorylation during meiosis reinitiation in starfish oocytes. Dev. BioL 91, 246-256. MEIJER, L., and GUERRIER, P. (1981). Calmodulin in starfish oocytes. I. Calmodulin antagonists inhibit meiosis reinitiation. Dev. BioL 88, 318-324. MIYAZAKI, S.-I., OHMORI, H., and SASAKI, S. (1975). Potassium rectifications of the starfish oocyte membrane and their changes during oocyte maturation. J PhysioL 246, 55-78. MOREAU, M., and GUERRIER, P. (1980). In vitro interactions between membrane, hormone, and cyclic nucleotides as revealed with aequorin. Dev. BioL 79, 488-492. MOREAU, M., GUERRIER, P., DOREE, M., and ASHLEY, C. C. (1978). Hormone-induced release of intracellular Ca’+ triggers meiosis in starfish oocytes. Nature (London) 272, 251-253. MORISAWA, M., and KANATANI, H. (1978). Oocyte-surface factor responsible for 1-methyladenine-induced oocyte maturation in starfish. Gam. Res. 1, 157-164. NAKAMURA, S., and HIRAMOTO, Y. (1978). Mechanical properties of the cell surface in starfish eggs. Dev. Growth Di&ren. 20, 317-327. NEMOTO, S-I., YONEDA, M., and UEMURA, I. (1980). Marked decrease in the rigidity of starfish oocytes induced by 1-methyladenine. Dev. Growth Di$krm 22, 315-325. NOTHNAGEL, E. A., BARAK, L. S., SANGER, J. W., and WEBB, W. W. (1981). Fluorescence studies on modes of cytochalasin B and phallotoxin effects on cytoplasmic streaming in Chara. J. Cell BioL 88, 364-372.
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PREDDIE, E., and ABASTADO, R. (1979). A phosphorylated protein in KCl-Triton extracts of 1-methyladenine treated cortices from oocytes of the starfish Marthasterias glacialis. Dev. BioL 73, 345-352. ROSENBERG,M. P. (1978). Relationship of changes in the cortical layer to the resumption of meiosis in starfish oocytes: Hormonal and cation requirements. Ph.D. dissertation. Univ. Toledo, Ohio. ROSENBERG,M. P., HOESCH, R., and LEE, H. H. (1977). The relationship between 1-methyladenine induced surface changes and fertilization in starfish oocytes. Exp. Cell Res. 107, 239-245. ROSENBERG, M. P., and LEE, H. H. (1981). The roles of Ca and Mg in starfish oocyte maturation induced by 1-methyladenine. J. Exp. ZooL 217,389-397. RUNNSTROM, J. (1944). Notes on the formation of the fertilization membrane and some features of the early development of the Aster&s egg. Acta ZooL 25, 159-167. SCHROEDER, P. C., LARSEN, J. H., and WALDO, A. E. (1979). Oocytefollicle cell relationships in a starfish. Cell. Tissue Res. 203, 249256. SCHROEDER, T. E. (1979). Surface area change at fertilization: Resorption of the mosaic membrane. Dev. BioL 70, 306-326. SCHROEDER, T. E. (1981). Microfilament-mediated surface change in starfish oocytes in response to I-methyladenine: Implications for identifying the pathways and receptors for maturation-inducing hormones. .I Cell BioL SO, 362-371. SCHROEDER,T. E. (1982). Novel surface specializations on a sea anemone egg: “Spires” of actin-filled microvilli. J. Morphol. 174, 207-216. SCHROEDER, T. E., and CHRISTEN, R. (1982). Polymerization of actin without acrosomal exocytosis in starfish sperm. Exp. Cell Res. 140, 363-371. SCHUETZ, A. W. (1969). Induction of oocyte shedding and meiotic maturation in Pismter ochraceus: Kinetic aspects of radial nerve factor and ovarian factor induced changes. BioL Bull. 137, 524-534. SHOJI, Y., HAMAGUCHI, M. S., and HIRAMOTO, Y. (1978). Mechanical properties of the endoplasm in starfish oocytes. Exp. Cell Res. 117, 79-87. TILNEY, L. G., KIEHART, D. P., SARDET, C., and TILNEY, M. (1978). Polymerization of actin. IV. Role of Ca++ and H+ in the assembly of actin and in membrane fusion in the acrosomal reaction of echinoderms. .J. Cell BioL 77, 536-550.