Microtubule cycles in oocytes of the surf clam, Spisula solidissima: An immunofluorescence study

DEVELOPMENTAL

BIOLOGY

114,151-160 (1986)

Microtubule Cycles in Oocytes of the Surf Clam, Spisula solidissima: An lmmunofluorescence Study RYOKO KURIYAMA,*~-~

GARY G. BORISY,$ AND YOSHIO MASUI?§

*Department of Anatomy, University of Minnesota, Minneapolis, Minnesota 55.455;tMarine Biological Laboratmy, Woods Hole, Massachusetts 025.4~;#Laboratory of Molecular Biology, University of Wisconsin, Madison, Wisconsin 53706; and #Department of Zoology, University of Toronto, Twmto, Ontario MESlAl, Canada Received June 28, 1985; accepted in revised

fwm September 24, 1985

Oocytes of the surf clam, Spisula solidissimu, underwent germinal vesicle breakdown and two meiotic divisions to give off polar bodies when they were fertilized or parthenogenetically activated with KCl. Fertilized eggs further proceeded to mitosis and cleaved, while parthenogenetically activated eggs remained uncleaved. We examined changes in microtubule-containing structures during meiotic divisions and subsequent mitotic processes by immunofluorescence. A monoclonal anti-tubulin antibody was applied to alcohol-fixed eggs from which the vitelline membrane had been removed by protease digestion. Up to the stage of second polar body formation, the pattern of microtubule organization in the first and second meiotic spindles was identical in both fertilized and parthenogenetically activated eggs. However, while fertilized eggs formed a sperm aster and mitotic spindles later, activated eggs formed only monaster- or ringshaped microtubule-containing structures which underwent cycles of alternating formation and breakdown. Lactoorcein staining of parthenogenetically activated eggs revealed that the chromosome cycle could occur in these eggs, in phase with this microtubule cycle. 0 1986 Academic Press, Inc. INTRODUCTION

In contrast to the great deal of information about the microtubules and their related structures from surf clam eggs in vitro, there have been very few reports on their Oocytes of the bivalve mollusc, Spisula solidissima, have been excellent material for biochemical studies of structural changes during meiotic and mitotic processes microtubules in meiotic and mitotic spindles. They are in intact cells. Over a quarter of a century ago, Rebhun easily obtained in a large quantity and develop syn- demonstrated movement and position changes of spinchronously after fertilization or parthenogenetic acti- dles in surf clam oocyte, visualized by staining of fervation. Rebhun and Sharpless (1964) first succeeded in tilized eggs with basic dyes (Rebhun, 1959). In a series of ultrastructural studies by thin-section electron mimass isolation of spindles from Spisulu eggs at different stages of development. This opened up a new way of croscopy, Longo and Anderson (1970a,b) showed the imstudying spindle microtubules in vitro (Keller and Re- portant role of microtubules in the formation of meiotic bhun, 1979; Suprenant and Rebhun, 1984) and made pos- and mitotic spindles in fertilized eggs. Most recently, sible the characterization of the proteins associated with Dan and Ito (1984) have isolated both nuclear and microtubule elements from Spisula eggs to show some of spindle microtubules (Murphy, 1980). Spindle microtubules are known to originate from their morphological changes from fertilization to second structures termed microtubule-organizing centers tiitotic cleavage. (MTOCs) (Pickett-Heaps, 1969). Using activated oocytes As seen above, the literature on microtubules and their of the surf clam, Weisenberg and his co-worker first related structures in intact eggs of the surf clam is rather demonstrated that the MTOC was a structure which re- scanty, and therefore it would be desirable to collect mained stable after isolation in solution (Weisenberg, more data from detailed structural studies of the entire 1973; Weisenberg and Rosenfeld, 1975). This stimulated microtubule system in whole eggs at various stages of a number of studies directed at the isolation and char- development. Since immunofluorescence staining has acterization of MTOCs from different kinds of organisms already proven to be an excellent, practical method for including cultured mammalian cells (Telzer et al, 1975; studying tubulin distribution in large cells such as sea Gould and Borisy, 1977), yeast (Hyams and Borisy, 1978; urchin eggs (Harris et al., 1980), we stained surf clam Byers et aL, 1978), quadriflagellata (Stearns et aL, 1976), eggs with an anti-tubulin antibody to obtain holistic imsea urchin eggs (Kuriyama and Borisy, 1983), and cel- ages of microtubules in a quite simple and effective way. lular (Kuriyama et ah, 1982) and plasmodial (Roobol et Comparing the distribution of microtubule arrays beal., 1982) slime molds. tween fertilized and parthenogenetically activated eggs, 151

0012-1606/86 $3.00 Copyright All rights

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

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DEVELOPMENTAL BIOLOGY

we found that a newly identified microtubule-containing structure was formed in parthenogenetically activated eggs after completion of maturation. Since in the activated eggs formation and breakdown of this microtubule-containing structure are tightly coupled with the cycle of nuclear-chromosome alteration, a possible explanation for the cycle of the microtubule structure is discussed in relation to cyclic changes in egg cytoplasmic activities. In reference to the origin and fate of the spindle poles, the difference in number and activity of centrioles between fertilized and artificially activated eggs is also discussed.

VOLUME114.1986

fixed with cold methanol in the presence of 50 mMEGTA (Harris et aL, 1980) as described elsewhere (Kuriyama and Borisy, 1985). After rehydration with phosphatebuffered saline (PBS), which consisted of 8.0 g NaCl, 0.2 g KCI, 0.2 g KH2POI, 0.99 g KzHPOl in 1 liter (pH 7.4), the coverslip was incubated with a monoclonal anti-tubulin antibody (YL1/2) which was raised against yeast tubulin (Kilmartin et al., 1982) for 1.5 hr at 37°C. This monoclonal antibody reacts specifically with the tyrosylated form of a-tubulin (Wehland et aL, 1983). The coverslip was then rinsed thoroughly with PBS and stained with the second antibody as described previously (Kuriyama and Borisy, 1985; Kuriyama et ab, 1984). In MATERIALS AND METHODS some cases, the whole cells on the coverslips were treated Animals. The surf clams were obtained from the sup- with 3% gelatin dissolved in PBS for 1 to 2 hr before ply department of the Marine Biological Laboratory incubating with the first antibody (Kuriyama and Borisy, (Woods Hole, Mass.), in June and July of 1983. 1985). Observations were made on a Zeiss PhotomicroOocytes. Oocytes were surgically released from ripe scope III (Carl Zeiss, Oberkochen, FRG) equipped with ovaries and washed with Ca-free artificial sea water epifluorescence optics, and photographs were taken with containing 1 mM ethylene glycol bis(@aminoethyl Kodak Tri-X films. ether)-N&V’-tetraacetic acid (EGTA). The Ca-free sea Observation of nuclei and chromosomes. Oocytes were water (CFSW) consisted of 27 g NaCl, 0.7 g KCI, 4.0 g pelleted and then resuspended in ethanol-acetic acid MgClz -6Hz0,ll g MgS04 7Hz0,0.4 g NaHC03 in 1 liter, mixture (3:l). After lo- to 60-min fixation in this mixture, being adjusted to pH 8.3 with 1 MNa&03. the cells were washed twice with distilled water and Removal of the vitelline membrane. In order to stain then resuspended in a small volume of lacto-orcein sothe microtubule-containing structures in whole surf lution which was prepared by mixing an equal volume clam eggs, it is necessary to remove the vitelline mem- of 2% orcein in acetic acid and 85% lactic acid in distilled brane to allow antibodies to penetrate into the cyto- water. Stained cells were observed with a bright-field plasm. Previously, Rebhun and his co-workers developed light microscope. the methods useful for removing the membrane by inRESULTS cubation of oocytes with alkaline-isotonic NaCl solution (Rebhun, 1962) or 1 Mglycerol (Rebhun and Sharpless, 1. Development of Fertilized and ParthenogeneticaUy 1964). These agents cause an expansion of the membrane Activated Oocytes up to the Stage of and its local dissolution. However, the methods require Second Polar Body Formation several centrifugation steps to wash the oocytes, thus Fully grown oocytes of the surf clam are about 55 pm causing some difficulties in collecting samples at exact in diameter. They contain a large nucleus of 30 pm in times following the initiation of development. In the present study, therefore, we adopted protease diameter, called the germinal vesicle, which is surtreatment to dissolve the vitelline membrane prior to rounded by a thin layer of cytoplasm. Although the apfertilization or parthenogenetic activation, which al- pearance of oocytes demembranated with protease lowed us to fix oocytes at desired times without delay. digestion shows no deterioration for many hours after The oocytes were treated with CFSW containing 1 mM the demembranation, they are so fragile that great care EGTA and 0.05 to 0.1% protease (Sigma, St. Louis, MO.) should be taken in subsequent handling of these oocytes. In the present study, oocytes were inseminated or acfor 5 to 10 min at room temperature. They were rinsed by treatment with KC1 several times with CFSW containing 1 mM EGTA and tivated parthenogenetically within 30 min postdemembranation. In both cases the then with filtered natural sea water. Fertilization and activation. To induce meiotic matu- oocytes developed in an almost identical fashion until ration, demembranated oocytes were fertilized or acti- they completed meiosis. The developmental timetables vated parthenogenetically by addition of 0.25 ml of 3 M for fertilized and parthenogenetically activated oocytes KC1 to 10 ml of an oocyte suspension in sea water ac- are compared in Fig. 1. There were no marked differences cording to the method of Allen (1953) with slight mod- in the schedule of early events such as germinal vesiele breakdown (GVBD) and the formation of the first ification. ImmunoJuorescent staining. Demembranated oocytes meiotic spindle between fertilized and parthenogenetiwere adhered to a polylysine-coated glass eoverslip and cally activated oocytes. However, differences became l

KURIYAMA, BORISY, AND MASUI

Time after Fertilization (min) 0

t

kNBD

T 20 T

1

r’”

1st Pb

\

1 2nd Pb \

60 ,

,

ist 1st miiatic spindle cleavage I

0

, T8O

I

20

Timzfter

Act%ion

~ 100

2nd cleavage

I

I 100

(min?

FIG. 1. Developmental timetables for fertilized and parthenogenetically activated oocytes. GVBD, germinal vesicle breakdown; spindle, first meiotic spindle formation in the center of the eggs; 1st and 2nd Pb, first and second polar body formation; 1st mitotic spindle, first mitotic spindle formation (fertilized eggs only); 1st and 2nd cleavage, first and second mitotic cleavage (fertilized eggs only).

Oocyte Microtubules

153

emanating from them become distinct (Fig. 2C), and they increase in length as well as in number. Eventually astral fibers originating from the opposite spots cross the nucleus to form bridges between the two spots (Figs. 2D, E, E’). At this stage, the nuclear envelope of the germinal vesicle has broken down almost completely. This is followed by the formation of the first meiotic spindle which is accompanied by a bright spot at each pole. By 12 to 15 min after activation, the meiotic spindle has grown to its maximum size. It is situated in the center of the oocyte and is associated with symmetrically elongated astral fibers (Figs. 2F, F’). Then, the spindle starts to move towards the periphery (Fig. 2G), and one of the poles becomes attached to the inner side of the oocyte surface (Fig. 2H). This asymmetric position of the spindle causes unequal cleavage of the oocyte resulting in the extrusion of the small first polar body (Fig. 21). The meiotic spindle pole remaining in the oocytes appears to divide immediately after first polar body formation. Two bright spots appear, which are connected to each other by fluorescent lines of microtubules (Fig. 25). The microtubules run perpendicularly to the axis of the first meiotic spindle and parallel to the oocyte surface. Since the axis of the second meiotic spindle is perpendicular to the oocyte surface (Fig. 2K), the two bright spots must be relocated to serve as poles for the second meiotic spindle. It is interesting to note that when the second meiotic spindle forms in the oocytes, a fully grown metaphase spindle also appears in the first polar body (Fig. 2K). Although we have not observed cleavage of the first polar body, as described in oocytes of the snail Crepidula (Conklin, 1901), an abortive cleavage furrow sometimes forms when the polar body is isolated (Kuriyama, unpublished). The distribution of tubulin in the oocyte cytoplasm also changes in the course of the meiotic divisions. The cytoplasm shows a very clean immunofluorescent background when oocytes have a fully grown spindle (Figs. 2F-H). After polar body formation, however, remnants of the spindle microtubules in both egg and polar body disintegrated and gradually faded away (Figs. W, J’), resulting in increased background fluorescence. These observations suggest an increase in the amount of nonpolymerized tubulin in the cytoplasm at the expense of the spindle tubulin.

apparent in later events as the oocytes approached the end of meiosis. For instance, activated oocytes were delayed in their second polar body release by 10 min at 20°C as compared with fertilized oocytes. Microtubule-containing structures in whole oocytes and their successive changes during meiotic maturation were clearly visualized by anti-tubulin immunofluorescence microscopy (Fig. 2). It should be noted here that the pattern of microtubule organization presented in this report was derived from staining of the tyrosinated form of tubulin and is not present when staining nontyrosinated tubulin. Since we have not isolated structures from the eggs or extracted eggs with a detergent-containing solution as done to improve the resolution of the fluorescence image (Balczon and Schatten, 1983; Otto and Schroeder, 1984), specially differentiated microtubule structures, if any, have not been detected in unfertilized/ unactivated eggs. As is evident in Figs. 2A and A’, in which phase-contrast and fluorescence images of unfertilized/unactivated eggs are shown, the thin layer of cytoplasm surrounding the germinal vesicle was faintly stained in these oocytes. In contrast, distinct microtubule structures eventually became visible by immunofluorescence in fertilized/activated oocytes without detergent extraction. At 2 to 5 min after insemination or KC1 treatment, oocytes develop two bright spots in the cytoplasm. These spots always appear in close proximity to the germinal vesicle, facing each other across the germinal vesicle. Their exact positions in a whole egg and the distance between them appear to be varied by the angle of observation. Soon after the appearance of the bright spots, the surface of 2. Development of Fertilized and Parthenogenetically the geminal vesicle became indented at the sites where Activated Oocytes a$ter the Second the spots were as if they are pushing their way into the Polar Body Formation nucleus (Fig. 2B). Following this initial stage of activation the oocytes Following the protrusion of the second polar body, develop astral fibers from each bright spot. As the flu- differences in the developmental pattern between parorescent spots become larger, individual microtubules thenogenetically activated and fertilized oocytes become

FIG. 2. Oocytes stained with monoclonal anti-tubulin antibody at various stages of maturation from the germinal vesicle stage to the second polar body formation. A: Phase-contrast. A’, B-K: Immunofluorescence. Fluorescence photographs in E’, F’, H’, and J’ represent the same images as E, F, H, and J at low magnification to show the uniformity of the staining reaction. The bright dot in the lower left of J’ is an air bubble. A, A’: Unactivated oocytes. B-J’: activated or fertilized eggs. Photographs were taken at 5.25 min (B), 6.3 min (C, D), 7.75 min (E’), 9.3 min (F’), 10 min (E), 13 min (F, G), 22 min (H’), 27.75 min (H, I, J’), 28 min (J), and 40 min (K) after either fertilization (J, J’) or parthenogenetic activation (B-I, K). A, A’, X220; B, X670; C, X620; D, X720; E, X760; F, G, X800, E’, X280; F’, H’, X260; H, I, X740; J, X770; K, X650; J’, X240.

154

FIG. 3. Fertilized eggs stained with monoclonal anti-tubulin antibody at various stages of development form the second polar body formation to the second mitotic metaphase. A-G: immunofluorescence (large blob in D is an air bubble). Photographs were taken at 39 min (A, A’), 55 min (B), 65 min (C, D), ‘71 min (E, F), and 96 min (G) after fertilization. A, A’, C, X720; B, D, X750, E, X730; F, X630; G, X620.

In contrast, after parthenogenetically activated eggs increasingly manifest. In parthenogenetically activated oocytes which are forming the second polar body, the have completed meiosis, they contain no mitotic spindles and fail to proceed to mitosis. Instead they form micromeiotic spindle is the only microtubule-containing structures different from mitotic structure (Fig. 2K). However, in the fertilized oocytes tubule-containing spindles. These sometimes resemble monasters (Figs. at the corresponding stage, another immunofluorescent structure could be seen in addition to the meiotic spindle 4A, B), or sometimes appear as brightly stained rings (Fig. 3A). This additional structure was identified as the or ovoidal structures with anti-tubulin antibody (Figs. sperm aster when the focal plane was adjusted to it (Fig. 4C, D). The number of the microtubules radiating from 3A’). Formation of sperm aster in anaphase eggs at the the center of monaster-like structure varies greatly, and second maturation has been reported in Crepidula it is rather difficult to detect microtubules radiating from the center of the ring-shaped structure. (Conklin, 1901). The monasterjring-shaped microtubule-containing Soon after the second polar body was extruded, fertilized oocytes begin their mitotic process. The first mi- structure goes through cyclic formation and breakdown totic spindle forms at the center of the egg and remains in the mature egg cytoplasm. In parallel wit@ the there for a while (Figs. 3B, C) but later moves to the changes in microtubule structures, nuclei were also found to change their morphology. Figure 5 illustrates periphery (Fig. 3D). This results in an unequal division of the egg at the first cleavage (Figs. 3E, F). In the ab- changing chromosomal morphology in eggs stained with sence of the vitelline membrane, which had already been lacto-orcein at different times after activation, repreremoved from the egg, blastomeres formed after first senting various degrees of chromosomal condensation cleavage were loosely attached to each other (Figs. 3F, from interphase (Fig. 5A), through intermediate stages G). The mitotic spindles for the second cleavage appear (Figs. 5B, C), and towards metaphase (Fig. 5D) and teloin the small and large blastomeres at the same time. phase stage (Fig. 5E) in mitosis. To compare the chromosome cycle with that of miThe spindle of the small blastomere remains in the center and divides the blastomere into equal halves, whereas crotubule-containing structure, we scored over 100 eggs, the spindle formed in the large blastomeres moves from stained either with anti-tubulin antibody or lacto-orcein the center to the periphery, thus cleaving the blastomere at various times after activation, and calculated perunequally (Fig. 3G). The larger blastomere thus cleaved centages of eggs with monaster/ring-like structures and with condensed chromosomes. The results are summain a similar manner to the whole egg in first cleavage.

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FIG. 4. Parthenogenetically nofluorescence. Photographs X700; E, X260; F, X250.

VOLUME 114, 1986

activated oocytes stained with monoclonal anti-tubulin antibody at various times after activation. A-F: immuwere taken at 65 min (E), 115 min (A, B, D), and 176 min (F) after parthenogenic activation. A, B, D, X780; C,

rized in Fig. 6. In this figure, line A represents the percentage of cells that contain a monaster/ring-shaped structure. From this line, it is clearly seen that in a brief period following the second polar body formation, the microtubule structures disappear in all eggs. However, at 80 min after activation, microtubule-containing structures can be detected in more than 95% of the eggs, and thereafter percentages of the eggs that contain this structure fluctuate periodically several times until the eggs begin to deteriorate. Figs. 4E and F show low-magnification fluorescence micrographs of the egg samples taken at 65 and 176 min postactivation, representing the peaks of formation of the microtubule-containing structure. On the other hand, line B in Fig. 6 shows the percentages of eggs that contain condensed chromosomes. The line clearly shows that soon after second polar body formation (that is, 50 min postactivation), the interphase

FIG. 5. Parthenogenetically

activated

nucleus is formed in all eggs. Then, the nuclear envelope once again breaks down and chromosomes condense to metaphase in all eggs by 80 min when the formation of the microtubule-containing structure reaches its peak. Thereafter, the microtubule and chromosome cycles are both repeated with the same periodicity of 40 to 50 min. Clearly, the two cycles are well synchronized, suggesting that the nuclear cycle is coupled to the microtubule cycle. DISCUSSION

Oocytes once activated by insemination or KC1 treatment show dynamic rearrangements of microtubules during the course of maturation. A couple of minutes after activation, two bright fluorescent spots appear in the cytoplasm adjacent to the germinal vesicle. Microtubules continue to polymerize onto the spots, resulting in the formation of the first meiotic spindle in the center

oocytes stained with la&o-orcein.

A-E: bright

field. A, C, D, X300; B, X320; E, X350.

KURIYAMA, BORISY, AND MASUI

157

Oocyte Microtubules

1st 2nd

0

III,,

I 40

I 60

I

I 80

I

I 100

Time after FIG. 6. Monaster/ring-shaped microtubule-containing of oocytes with microtubule structure. B: Percentage body formation, respectively.

I

I I 120

Activation

I 140

I

11 160

I fi I 180 200

(min)

structure and chromosome cycles in parthenogenetically activated oocytes. A: Percentage of oocytes with condensed chromosomes. 1st and 2nd: times for the first and second polar

of eggs. Symmetric astral microtubules radiate from each pole of the spindle. To divide unequally, however, the spindle starts to move toward one side of the egg, and astral fibers are no longer symmetrical. This observation with immunofluorescence microscopy is in good agreement with the report recently made by Dan and Ito (1984), who demonstrated, by isolating the microtubule elements, that the metaphase spindle shifts from the cell center to a definite site on the cortex. This migration of the spindle can be inhibited by taxol, a microtubule-stabilizing drug, but not by cytochalasin B, a microfilament-disintegrating drug (R. Kuriyama, manuscript in preparation). This suggests that depolymerization of astral microtubules is required for the spindle movement. After extrusion of the first polar body, structures corresponding to the second meiotic spindles are formed not only in the egg but also in the first polar body. Here, it is interesting to note that the spindle forms simultaneously in those two cells whose cytoplasmic conditions are quite different in terms of quantity and quality (Long0 and Anderson, 19’70a).It may be suggested that the dynamic organization of microtubule systems in the oocyte and polar body has been programmed under the control of a common cytoplasmic clock defined in the ooplasm before the first meiotic division. After completion of the maturation process, parthenogenetically activated eggs form a monaster (Rebhun and Sharpless, 1964) or ring-shaped microtubule-containing structure.

In this stage, however, it is not clear how the two structures relate to each other. The microtubule structures undergo a series of periodic changes in which their formation alternates with breakdown and is coupled with chromosome cycles. The period of this cycle is 40 to 50 min, corresponding to the mitotic cycle in zygotes. Therefore, the oscillation of cytoplasmic activities responsible for monaster formation and chromosome condensation is a process independent of the presence of sperm factors. A similar case of monaster cycles has been reported in sea urchin zygotes treated with caffeine (Harris, 1983). It has been suggested that cytoplasmic activities related to meiosis or mitosis, which can be triggered by activation of oocytes, are autonomous and independent of nuclear functions. The evidence supporting this notion has been provided by the observations of cyclic changes in ooplasmic activities in enucleated eggs of various animal species. For example, the morphology of the hyaline layer and the cytoplasm of the sea urchin eggs (Kojima, 1960), the contractility of the frog egg cortex (Hara et aL, 1980), and the tension of the starfish oocyte (Yamamoto and Yoneda, 1983) undergo cyclic changes with the same periodicity as the division cycles of the zygote, even in the absence of normal meiotic and mitotic events such as spindle formation and cytokinesis. Therefore, it is probable that the microtuble cycle observed in parthenogenetically activated Spisulu oocytes, which follows completion of their maturation, is an expression of au-

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tonomous cell cycle activities of the cytoplasm. In this connection, it would be of particular importance to know whether or not chromosome replication also cycles with the same period as the monaster cycle as observed in NHIOH-activated sea urchin eggs (Mazia and Ruby, 1974; Mazia, 1974). The origin and fate of the centrioles pose intriguing problems, and centriolar behavior has already been described in Crepidula eggs from as early as 1901 (Conklin, 1901). The first question is how centrioles appear in parthenogenetically activated oocytes. It has already been shown that while the astral microtubules assembled in oocyte homogenates prepared at 2.5 min postactivation were associated only with a “procentriole,” a dense central cylinder, those formed in the homogenates prepared at 4.5 min postactivation contained a distinct centriole at each focal point (Weisenberg and Rosenfeld, 1975). In this stage, astral microtubules were found to radiate from a large number of cytoplasmic granules surrounding the centriole. These structures most likely correspond to the spots identified immunofluorescently in this paper. Therefore, it may be assumed that de nova formation of centrioles can occur in the oocyte cytoplasm shortly after activation. This is illustrated in Fig. 7, which shows a putative scheme of centriole behavior in developing Spisula oocytes. The second question concerns the number of centrioles at each pole of meiotic spindles. In oocytes of the mussel, Mytilus edulis, which are spawned at the germinal vesicle stage and later proceed to metaphase I, but remain suspended at this stage until fertilization, each pole of the first meiotic spindle contained a pair of centrioles (Long0 and Anderson, 1969). In Sp$sula, however, no evidence has been provided for the existence of paired centrioles at the pole of the first meiotic spindle in intact oocytes.

VOLUME 114, 1986

Rather, Weisenberg and Rosenfeld (1975) indicated that each aster assembled in vitro from oocyte homogenates contained only a single centriole. Apparently, there exists a discrepancy between the observations on the oocytes of two different molluscan species. Therefore, it would be important to examine carefully the number of centrioles at meiotic spindle poles in intact Sfisula oocytes to reconcile this discrepancy. If two centrioles are found at each pole of this first meiotic spindle, as shown in Fig. 7, then we must ask how centriole duplication can take place at an early phase of the first meiosis such as step 2. In Spisula, the second meiotic spindle is formed within 10 to 15 min after the first polar body extrusion. It would be interesting to know how many centrioles reside at each pole of the spindle at this stage. Although we have no direct evidence to answer this question, the formation of the second meiotic spindle could not be inhibited by taxol, a drug shown to be a potent inhibitor of centriole duplication in cultured mammalian cells (R. Kuriyama et al., manuscript in preparation). Therefore, it may be that centriole duplication does not occur at step 4 as illustrated in Fig. 7, thus bringing only one centriole to each pole of the second meiotic spindle. However, it is also possible that the duplication, if it occurs, is not a prerequisite for completing the second meiosis, or that the duplication has been programmed earlier so that it remains unaffected by taxol treatment. Interestingly, Longo and Anderson (1969) mentioned that each aster of the second meiotic spindle in Myths oocytes contained only one centriole, supporting our scheme in Fig. 7. The behavior of centrioles in fertilized eggs is expected to be more complicated than that in parthenogenetically activated eggs, since the sperm introduces centrioles of

ACTIVATION

FERTILIZATION

FIG. 7. Putative scheme of centriolar behavior in parthenogenetically activated and fertilized oocytes during maturation rectangle, maternal centriole; hatched rectangle, paternal centriole. The numbers refer to steps of centriole development.

and cleavage. Open

KURIYAMA, BORISY, AND MASUI

paternal origin into the egg. This creates complex relationships between centrioles of two parental origins. In Spisula, according to Longo and Anderson (19’70b), the sperm carries a pair of centrioles at the base of its flagellum and brings them into the oocyte together with the male nucleus. However, these centrioles are located in close proximity to the nucleus, without being associated with polymerized microtubules, until the sperm chromatin has completely dispersed and the nuclear envelope is formed. The results of light (Conklin, 1901) and thin-section electron microscopy (Long0 and Anderson, 1970b) indicate that no sperm centriole becomes active in assembling astral microtubules until the oocyte chromosomes completed meiosis. Similarly, our immunofluorescence study reveals that the sperm aster can develop only after the second polar body begins to form. Taken together, these results lead us to the conclusion that the centrioles, which serve to organize microtubules for the first meiotic spindle at step 7, are totally of maternal origin. We are puzzled again by the question of why only the maternal centrioles become active to organize microtubules almost immediately after oocytes are activated (step ‘7),while the paternal centrioles remain silent until oocytes complete meiosis (step lo), despite the fact that the same nine triplet microtubules of both origins reside in a common ooplasmic environment. To answer this question, we may need information about the pericentriolar material and its distribution among centrioles of different origins, since it is not the centriole itself, but the amorphous cloud of pericentriolar material surrounding the centrioles located at spindle pole, that is directly responsible for organizing microtubules (Gould and Borisy, 1977; Robbins et al, 1968). The last question concerns the fate of the maternal centrioles in oocytes and polar bodies, that is, whether they disintegrate in the cytoplasm or are retained throughout development. It would be interesting to know whether or not the centrioles reappear in association with cycling monaster after step 6 in parthenogenetically activated eggs. In fertilized eggs, the classical notion, which has been well-accepted as Boveri’s theory, states that only the sperm centrioles play an essential role in organizing mitotic spindles (Wilson, 1928). However, the fate of maternal centrioles as well as the mechanism for the organization of spindle poles during successive mitotic cycles remain as questions to be answered by future investigations. We thank Dr. J. V. Kilmartin of the MRC Laboratory of Molecular Biology, Cambridge, England, for providing monoclonal antitubulin antibody; Mr. H. Thorsen of the University of Wisconsin for printing photographs; Dr. R. P. Elinson, Ms. E. K. Shibuya, and Mr. H. J. Clarke of the University of Toronto for reading the manuscript. One of us (R.K.) expresses her thanks to Dr. J. L.‘Rosenbaum of the Yale Uni-

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versity for his kind support and advice. The authors are thankful to Mrs. Rossana Soo for her assistance in typing the manuscript. This work was supported by Step Toward Independence Fellowship from Marine Biological Laboratory at Woods Hole, Minnesota Medical Foundation CRF-69-85, ACS Institutional Grant IN-13-X-8 (R.K.), NIH Grant GM 30385 (G.G.B.), NIH Training Grant 5-T35HD07098 awarded to the Embryology Course, Marine Biological Laboratory, Woods Hole, and NSERC Canada Grant A5855 (Y.M.). REFERENCES ALLEN, R. D. (1953). Fertilization and artificial activation in the egg of the surf clam Spisula solidissima Biol. Bull (Woo& Hole, Mass.) 105,213-239. BALCZON, R., and SCHATTEN, G. (1983). Microtubule-containing detergent-extracted eytoskeletons in sea urchin eggs from fertilization through cell division: Antitubulin immunofluorescence microscopy. Cell Motil. 3, 213-226. BYERS, B., SHRIVER, K., and GOETSCH, L. (1978). The role of spindle pole bodies and modified microtubule ends in the initiation of microtubule assembly in Saccharomyces cerevisiae. J. Cell Sci 30,331352. CONKLIN, E. G. (1901). Centrosome and sphere in the maturation, fertilization and cleavage of Crepidula. And. Anz. 19,280-287. DAN, K., and ITO, S. (1984). Studies of unequal cleavage in Molluscs: I. Nuclear behavior and anchorage of a spindle pole to cortex as revealed by isolation technique. Dev. Growth Difler. 26,249-262. GOULD, R. R., and BORISY, G. G. (1977). The pericentriolar material in Chinese hamster ovary cells nucleates microtubule formation. J. Cell Biol. 73, 601-615. HARA, K., TYDEMAN, P., and KIRSCHNER, M. (1980). A cytoplasmic clock with the same period as the division cycle in Xenopus eggs. Proc. Natl. Acad. Sci. USA 77,462-466. HARRIS, P. (1983). Caffeine-induced monaster cycling in fertilized eggs of the sea urchin Strangylocentrotus prpuratus. Dev. Biol. 96,277284. HARRIS, P., OSBORN,M., and WEBER, K. (1980). Distribution of tubulincontaining structures in the egg of the sea urchin Strongylocentrotus purpuratus from fertilization through first cleavage. J. Cell Biol 84, 668-679. HYAMS, J. S., and BORISY, G. G. (1978). Nucleation of microtubles in vitro by isolated spindle pole bodies of the yeast Saccharcmzyces cerevisiae. J. Cell Biol. 78.401-414. KELLER, T. C. S., and REBHUN, L. I. (1979). Properties of polymerizable tubulin from isolated Spisula spindles. Biol Bull. (Woods Hole, Mass.) 157,374-375. KILMARTIN, J. V., WRIGHT, B., and MILSTEIN, C. (1982). Rat monoclonal antitubulin antibodies derived by using a new nonsecreting rat cell line. J. Cell Biol. 93, 576-582. KOJIMA, M. K. (1960). Cyclic changes of the cortex and the cytoplasm of the fertilized and the activated sea urchin eggs. I. Changes in the thickness of the hyaline layer. Embryologia 5.1-7. KURIYAMA, R., and BORISY, G. G. (1983). Cytasters induced within unfertilized sea urchin eggs. J. Cell Sci. 61, 175-189. KURIYAMA, R., and BORISY, G. G. (1985). Identification of molecular components of the centrosphere in the mitotic spindle of sea urchin eggs. J. Cell Biol. 101,524-530. KURIYAMA, R., KERYER, G., and BORISY, G. G. (1984). The mitotic spindle of Chinese hamster ovary cells isolated in taxol-containing medium. J. Cell Sci. 66, 265-275. KURIYAMA, R., SATO, C., FUKUI, Y., and NISHIBAYASHI, S. (1982). In vitro nucleation of microtubules from microtubule-organizing center prepared from cellular slime mold. CeU Motil. 2,257-272. LONGO, F. J., and ANDERSON, E. (1969). Cytological aspects of fertil-

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ization in the lamellibranch, Mytilus edulis. I. Polar body formation and development of the female pronucleus. J. Exp. ZooL 172,69-96. LONGO,F. J., and ANDERSON,E. (19’70a).An ultrastructural analysis of fertilization in the surf clam, Spisula solidissima: Polar body formation and development of the female pronucleus. J. Ultrastruct. Res. 33, 495-514. LONGO,F. J., and ANDERSON,E. (1970b). An ultrastructural analysis of fertilization in the surf clam, Spisulu solidissima: Development of the male pronucleus and the association of the maternally and paternally derived chromosomes. J. Ultrastract. Res. 33.515-527. MAZIA, D. (1974). Chromosome cycles turned on in unfertilized sea urchin eggs exposed to NH,OH. Proo. NatL Acad Sci. USA 71,690693. MAZIA, D., and RUBY,A. (1974).DNA synthesis turned on in unfertilized sea urchin eggs by treatment with NHIOH. Exp. Cell Res. 85,167172. MURPHY,D. B. (1980). Identification of microtubule-associated proteins in the meiotic spindle of surf clam oocytes. J. Cell BioL 84.235-245. OSBORN,M., and WEBER,K. (1976). Cytoplasmic microtubules in tissue culture cells appear to grow from an organizing structure towards the plasma membrane. Proc. NatL Acad Sci. USA 73,867-871. Oreo, J. J., and SCHROEDER,T. E. (1984). Microtubule arrays in the cortex and near the germinal vesicle of immature starfish oocytes. Den BioL 101.274-281. PICKETT-HEAPS,J. D. (1969). The evolution of the mitotic apparatus: An attempt at comparative ultrastructure cytology in dividing plant cells. Cytobios 3, 257-280. REBHUN,L. I. (1959). Studies of early cleavage in the surf clam, Spisula solidissima, using methylene blue and toluidine blue as vital stains. BioL Bull. (Woods Hole, Mass.) 117, 518-545.

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REBHUN,L. I. (1962). Dispersal of the vitelline membrane of the eggs of Spisula solidissima by alkaline, isotonic NaCl. J. Ultrastract. Res. 6,123-134. REBHUN,L. I., and SHARPLESS,T. K. (1964). Isolation of spindles from the surf clam Spisula solidissima. J. Cell Biol. 22, 488-492. ROBBINS,E. L., JENTZSCH,G., and MICALI, A. (1968). The centriole cycle in synchronized HeLa cell. J. Cell BioL 36,329-339. ROOBOL,A., HAVERCROFT,J. C., and GULL, K. (1982). Microtubule nucleation by the isolated microtubule-organizing center of Physarum polycephalum myxamoebae. J. Cell Sci 55,365-381. STEARNS,M. E., CONNOLLY,J. A., and BROWN,D. L. (1976). Cytoplasmic microtubule organizing centers isolated from Polytomella agilis. Science (Washington, D. C!) 191,188-191. SUPRENANT,K. A., and REBHUN,L. I. (1984). Purification and characterization of oocyte cytoplasmic tubulin and meiotic spindle tubulin of the surf clam Spisula solidissima. J. Cell BioL 98,253-266. TELZER,B. R., MOSES,M. J., and ROSENBAUM,J. L. (1975). Assembly of microtubules onto kinetochores of isolated mitotic chromosomes of HeLa cells. Proc. NatL Acad. Sci USA 72,4023-4027. WEHLAND,J., WILLINGHAM,M. C., and SANDOVAL,I. V. (1983). A rat monoclonal antibody reacting specifically with the tyrosylated form of a-tubulin. J. Cell BioL 97,1467-1475. WEISENBERG,R. C. (1973). Regulation of tubulin organization during meiosis. Amer. ZooL 13, 981-987. WEISENBERG,R. C., and ROSENFELD,A. C. (1975). In vitro polymerization of microtubules into asters and spindles in homogenates of surf clam eggs. J. Cell BioL 64, 146-158. WILSON,E. B. (1928). “The Cell in Development and Heredity,” 3rd ed. Macmillan Co., New York. YAMAMOTO,K., and YONEDA,M. (1983). Cytoplasmic cycle in meiotic division of starfish oocytes. Dev. BioL 96,166-172.