Cyclic assembly-disassembly of cortical microtubules during maturation and early development of starfish oocytes

DEVELOPMENTAL

BIOLOGY

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(1984)

Cyclic Assembly-Disassembly of Cortical Microtubules during Maturation and Early Development of Starfish Oocytd THOMAS *F&g

E. SCHROEDER*

AND JOANN J. OTTOMAN

Harbor Lubomhh, Univemi~ of Washington, Friday Harbor, Washington 98350, and TDepartd qf Bidogical S’cienm, Purdue Univemit~, West L&a&te, Indianu 47997 Received Nuvember 18, 198s; accepted in revised form January 19, 1984

An extensive array of cortical microtubules in oocytes of the starfish Pisaste~ ochraeem undergoes multiple cycles of disappearance and reappearance during maturation and early development. These events were studied in isolated fragments of the oocyte cortex stained with antitubulin antibodies for indirect immunofluorescence. The meshwork of long microtubules is present in the cortex (a) of immature oocytes, i.e., before treatment with the maturationinducing hormone 1-methyladenine, (b) for 10-29 min after treatment with 1-methyladenine, (c) after formation of the second polar body (in reduced numbers in unfertilized oocytee), and (d) in the intermitotic period between first and second cleavage divisions. The array of cortical microtubules is absent in oocytes (a) undergoing germinal vesicle breakdown, (b) during the two meiotic divisions (polar body divisions), and (c) during mitosisof the first and, perhaps, subsequent cleavage divisions. The cycle of assembly-disassembly of cortical microtubules is synchronized to the cycle of nuclear envelope breakdown and reformation and to the mitotic cycle; specifically, cortical microtubules are present when a nucleus is intact (germinal vesicle, female pronucleus, zygote nucleus, blastomere nucleus) and are absent whenever a meiotic or mitotic spindle is present. These findings are discussed in terms of microtubule organizing centers in eggs, possible triggers for microtubule assembly and disassembly, the eccentric location of the germinal vesicle, and the regulation of oocyte maturation and cell division. INTRODUCTION

Fully grown oocytes of starfish are uniformly arrested at the germinal vesicle stage (meiotic prophase), but they can be stimulated to resume meiotic (maturation) divisions synchronously in vitro by treatment with the maturation-stimulating hormone 1-methyladenine (Kanatani, 1973). Their availability and ease of handling make starfish oocytes an important experimental system for studying the physiological mechanisms and biochemical pathways of maturation in animal oocytes. The reinitiation of maturation by 1-methyladenine involves radical changes in the nucleus, cytoplasm, cortex, and cell surface (Schroeder and Stricker, 1933). As a fortuitous finding while examining the actin-based organization of the oocyte cortex (Otto and Schroeder, 1934a), we discovered an unusual and extensive meshwork of long microtubules throughout the cortex of meiotically arrested immature starfish oocytes (Otto and Schroeder, 193413). Another array of long microtubules in the form of one or two premeiotic asters was also found in the subcortical cytoplasm between the eccentric germinal vesicle and the animal pole. At the time of those Andings, further examination of the microtubule systems was interrupted by the end of the reproductive season. We now wish to describe the dynamic behavior 1 Commemorating the fundamental insights into starfish reproduction by Prof. H. Kanatani (1939-1984) and Dr. S. Hirai (1943-1983). 493

of these microtubules during hormone-stimulated maturation and during the early developmental stages after fertilization. MATERIALS

AND METHODS

Oocytes of the starfish Ptimter ochraceus were obtained according to methods described previously (Schroeder and Stricker, 1933). Two different protocols were used in order to denude unfertilized oocytes and fertilized oocytes. For the former, immature oocytes were first defolliculated and enzymatically denuded in calcium-free seawater (CaFSW) and then treated with lo-’ M 1-methyladenine (l-MA) in CaFSW to initiate the resumption of maturation, precisely as previously described (Otto and Schroeder, 1934a,b). For denuding fertilized oocytes, immature oocytes with intact follicle cell coverings were separated from dissected ovaries, washed in natural seawater (NSW), and treated with 10-6M1-MA in NSW; they were allowed to develop until after germinal vesicle breakdown and were then inseminated with a dilute suspension of fresh sperm. At about 10 min after fertilization, when fertilization envelopes had elevated, oocytes were treated for 15 min with a solution of 1 mg/ml Pronase (Calbiochem-Behringer, La Jolla, Calif.) dissolved in NSW, washed, and then cultured in monolayers in NSW. Such treatment weakened and fragmented the fertilization envelopes but did not entirely remove them; nevertheless, the fragments 0912-16cW94 $3.99 Copyright All righta

Q 1984 by Aademic Press, Inc. of reproduction in any form rwxwd.

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of fertilization envelopes progressively disappeared during culturing and did not interfere with subsequent observations. All cultures were continuously maintained at 12-13°C and were periodically monitored with a dissecting microscope for major cytological landmarks of development. Alternatively, small aliquots of oocytes were spotchecked by differential interference-contrast microscopy (Nomarski). Neither the chronology or morphology of development was altered by enzymatic denuding or culturing in CaFSW instead of NSW. The time of germinal vesicle breakdown (GVBD) was recorded when disintegration of the nuclear envelope was observed in about 90% of the oocytes. The times of first and second polar body formation and first and second cleavage (PB I, PB II, Cl 1, and Cl 2 in Figs. 3 and 8) were recorded when signs of constriction were detectable in at least 50% of oocytes. At selected times, isolated cortices were prepared and examined by indirect immunofluorescence using antitubulin antibodies. The methods have been previously described (Otto and Schroeder, 1934a,b); briefly restated, one or two drops of oocytes in seawater were placed on a coverslip pretreated with polylysine, the oocytes were allowed to settle and adhere for 1 min, excessseawater was drained away, the oocytes were sheared with a stream of buffered detergent solution delivered from a squeeze bottle, and the coverslips with cortical fragments were fixed in formaldehyde and cold methanol. In most instances duplicate coverslips were prepared in order to compare different antibodies and minor variations in protocols. For indirect immunofluorescence detection of microtubules, four preparations of antitubulin antibodies were used: (a) ascites fluid containing a monoclonal antibody against cr-tubulin, donated by Dr. S. H. Blose (Cold Spring Harbor Lab, N. Y.); (b) a commercial antitubulin antiserum (Miles, Inc., Elkhart, Ind.); (c) an antiserum prepared against electrophoretically purified tubulin from chick brain, donated by Dr. W. E. Gordon (Cold Spring Harbor Lab, N. Y.); and (d) an affinity-purified antiserum against bovine tubulin, donated by Dr. B. R. Brinkley (Baylor College of Medicine, Houston, Tex.). When applied to isolated cortices, these antibodies were localized with fluorescein-conjugated second antibodies; anti-mouse for (a); anti-rabbit for (b) and (c); and antisheep for (d). Photobleaching of fluorescence in stained specimens was reduced to a negligible amount by dissolving 1.5% (w/v) n-propyl gallate (Giloh and Sedat, 1982) in a mounting medium composed of 50% glycerol in phosphate-buffered saline (pH 8.6). Continuous observation even for a few minutes did not perceptibly reduce the fluorescence intensity of the specimens; on the other

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hand, exposures of 5-20 set were sufficient for fluorescencephotomicrography. In nearly all instances, isolated cortices were too thin and too low in contrast to be examined profitably by phase-contrast or Nomarski microscopy. The array of cortical microtubules at various stages was evaluated semiquantitatively as follows. Assuming that a preparation of isolated cortices met several subjective standards of quality (uniformity of appearance among numerous cortices, individual fragments that were large yet undistorted, absenceof staining or drying artifacts, etc.), the whole population of cortices on the coverslips was categorized according to the pattern of cortical microtubules. The categories were: (1) a full complement of microtubules; (2) an intermediate array, distinctly less dense than the full complement; (3) no or virtually no microtubules. In Figs. 3, 5, and 6 these categories are indicated symbolically as “++,” “+,” and “0,” respectively. At a few transitional times, dual designations were assigned if subpopulations of cortices, each representing at least one-third of the total, could be recognized. In this study, we evaluated a total of 101 coverslips by these criteria, each of which contained 50100 isolated cortices. Aggregate lengths of microtubules were measured with an opisometer from representative photomicrographs of cortices in the three categories. The full complement (-I-+) contained >2 pm of lineal microtubules/ pm2 of cortex; the typical intermediate array (+) contained 0.2-l ~rn/~rn’; and the null array (0) contained <0.06 pm/fim2. RESULTS

Chrondogzl sfmajor cytological eventa The pattern of microtubules in the cortex changes in accord with the chronology of oocyte maturation, fertilization, and the mitotic cell divisions of cleavage. The schedule of these events was determined by microscopic examination of the living cultures at 12-13°C. Good synchrony of development was observed within and, usually, between batches of oocytes. Meiosis, as reinitiated by treatment with l-MA, includes germinal vesicle breakdown (GVBD) beginning at about 50 min, the progressive establishment of the first meiotic spindle (not easily visualized until late stages), pinching-off of the first polar body (PB I), rapid establishment of the second meiotic spindle, pinchingoff of PB II, and appearance of a female pronucleus. If an oocyte is unfertilized, PB I and PB II appear as protrusions at the animal pole (i.e., at meiotic telophase) at about 130 and 240 min after treatment with l-MA. If the oocyte is fertilized, these meiotic divisions occur precociously with PB I and PB II at 135 and 196 min,

respectively. (The accelerating effect of fertilization on meiotic divisions has been noted previously (Chambers and Chambers, 1949)). The first and second cleavage furrows (i.e., at mitotic telophase), which only occur in fertilized oocytes, appear initially at about 299 and 355 min, respectively. The nucleus and its nuclear envelope periodically break down and reform in maturing and developing Piso&r oocytes. An intact nucleus (the germinal vesicle) is present until GVBD but during the two meiotic divisions there is none. A nucleus (the female pronucleus) reappears at telophase of the second meiotic division, i.e., when the PB II first appears as a protrusion. In unfertilized oocytes, no further development beyond the pronuclear stage takes place. In fertilized oocytes at

about 259 min, the male and female pronuclei fuse into a zygote nucleus which shortly thereafter breaks down when a mitotic apparatus appears. After mitosis, blastomere nuclei appear just before the onset of the cleavage furrow. During subsequent cleavage divisions, blastomere nuclei are intact for about one-half of each cell division cycle, breaking down during mitosis and reforming again when the cleavage furrow begins. Cortical rnicrot~~d po&z~~~ As previously described (Otto and Schroeder, 1984b), the unstimulated immature oocyte normally contains an extensive array of long microtubules that course throughout the cortex (Fig. la). As seen in isolated cortices, the pattern is remarkable for the apparent length of individual microtubules, the relative paucity of ends of microtubules,

FXO. 1. Fluorescence micrographs of portions of isolated cortices from unfertilii stsrhsh oocytes stained with antitubulin antibody. The sequence shows the loss and partial reappearance of cortical microtubules at times after 1-methyladenine (l-MA): (a) 0, (b) 25, (c) 2’70, and (d) 280 min after treatment with l-MA. The pattern of microtubules in (a) is typical of the immature oocyte. Virtually all cortical microtubules disappear by 26 min after l-MA treatment (b) and remain disassembled (c) until after the second polar body emerges (d), at which time some cortical microtubules reappear. Xl!ZOO. Scale bar = 10 pm.

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the randomness of microtubule orientation, the openness of the meshwork, and in general the lack of prominent focal centers of organization. Since extensive fields of numerous isolated cortices exhibit virtually identical patterns, we believe that all regions of the oocyte cortex possess essentially the same array of cortical microtubules. (As will be discussed separately below, there is a single exception to this generalization, in the form of a separate array of astral microtubules comprising a premeiotic aster that is situated near the animal pole.) In the present context the “cortex” of an oocyte is defined operationally by our method of isolation. By focusing at high power with the microscope, the thickness of isolated cortices was determined to be l-2 pm, although occasional irregularities projected 3 pm from the coverslip surface. “Cortical” microtubules, when

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present, course through these isolated cortices about 1 pm from the coverslip. Cortical microtubules in situ similarly occupy a zone about l-2 pm from the plasma membrane, based upon transmission electron microscopic analysis of thin-sectioned intact oocytes (Otto and Schroeder, 1934b; unpublished observations). Such microtubules do not make contact with the plasma membrane or extend deeper than about 3 pm into the cell. In order to follow the fate of this extensive array of cortical microtubules, we studied the time-course of changes in 5 batches of oocytes treated with l-MA. The results are recorded graphically in Fig. 3. The microtubule patterns from selected stages of unfertilized and fertilized oocytes are shown in Figs. 1 and 2. The full complement of cortical microtubules persists

FIG. 2. Isolated cortices stained with antitubulin antibody at (a) 135, (b) 220, (c) 240, and (d) 290 min after l-MA. In this series, oocytes were fertilized at 90 min when cortical microtubules are absent (see Fig. 1). Cortical microtubules reappear in large numbers after second polar body formation (b), then disappear again during mitosis of the first cleavage division (c). Some cortical microtubules reappear (d) in the intermitotic period before the next cleavage division. X1200. Scale bar = 10 pm.

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FIG. 3. The abundance of cortical microtubules in oocytes from five batches (A-E) as a function of time after application of l-MA. The vertical axis (applicable to each curve) identifies semiquantitative categories of the number and density of microtubule arrays (++, many microtubules; +, some microtubules; 0, no microtubules). Each data point represents the summary evaluation of 50-100 cortices per coverslipstained with antitubulin. In all, 89 coverslips were examined; for more than one-half of the time points, evaluations are based on replicate coverslips stained with different antibodies. At a few time

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without modification for at least 10 min after application of l-MA then rapidly disappears well before GVBD (Figs. lb and 3). When these long microtubules disappear, a small number of microtubule fragments often remains (Figs. lb and c). Except for these few fragments, no microtubules again appear in the isolated cortex until formation of PB II (Figs. Id, 2b, and 3). Isolated cortices lacking organized microtubules were often very difficult to detect because of the low levels of fluorescent staining, especially when highly specific antitubulin antibodies such as antibodies (a) and (c) were used (see Materials and Methods). Antibody (b), however, faintly stained some nonmicrotubule component of the isolated cortex in addition to microtubules and thus outlined the cortex even when most microtubules were absent (e.g., Figs. lb, c, 2a, and c). Cortical microtubules reappear after PB II in all oocytes, whether unfertilized (Figs. Id and 3a) or fertilized (Figs. 2b and 3b), regardless of the schedule of development. The array of microtubules that reassemble after PB II in unfertilized oocytes is consistently less dense than the array in fertilized oocytes at the same stage (+ vs ++, Fig. 3). Microtubules at this stage in unfertilized oocytes frequently are about 15 pm long, forming a very loose meshwork (Fig. Id). In fertilized oocytes the reassembled microtubules form an array that appears as dense as that of immature oocytes (compare Figs. 2b and la), although many more microtubule ends are evident as if the constituent elements are shorter. After the reappearance of a reduced meshwork of microtubules at PB II in unfertilized oocytes, no further developmental change is observed and the loose array of microtubules persists (Fig. 3a). In fertilized oocytes, however, the extensive array persists for only about 30 min before it disappears again at the time that mitosis begins (Figs. 2c and 3b). This second episode of disassembly of cortical microtubules is as complete and as reproducible as the first episode that occurs soon after treatment with l-MA, it appears to coincide with the onset of mitosis of the first cleavage division-that is, with breakdown of the zygote nucleus.

points (bB and bE), presumably coinciding with times of rapid transition, two microtubule densities representing two clear subpopulations of isolated cortices were observed. Curves connecting data points represent interpretations. (a) Cortices treated with l-MA but not fertilized. (b) Gocytes treated with l-MA and subsequently fertilized (at times indicated by small arrows). Major cytological landmarks (for all cultures except D, for which times were not obtained and which appears to be on a different schedule) are indicated by the broad arrows, whose widths depict variability within and between batches of oocytes (GVBD, germinal vesicle breakdown; PB I and PB II, appearance of first and second polar bodies as protrusions at the animal pole; CL 1 and 2, appearance of cleavage furrows of the first and second cleavage divisions).

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Remarkably, cortical microtubules reappear a second time in all fertilized oocytes coincidently with the onset of cleavage furrow formation (Figs. 2d and 3b). This occurrence completes two cycles of microtubule disassembly and reassembly. At first cleavage, however, the density of the array of microtubules is quite low (Fig. 2d), even lower than the density after maturation of unfertilized oocytes. Individual microtubules are again 15 pm or more in length but they are so sparse that they hardly ever intersect. Following this late, but partial, reappearance of microtubules during first cleavage, the sequence of events becomes unclear. Most oocytes continue to display low densities of microtubules, but some begin to lack them

FIG. 4. Phase-contra& (left) and matching fluorescence micrographs (right) of cortices isolated soon after l-MA application, Cortical microtuhules are abundant at 0 min and are still evident at 10 min when a&in-filled surface spikes (Schroeder and Stricker, 1933) are prominent (arrows). There is no obvious relationship between the positions of spikes and microtubules. By 25 min virtually all surface spikes and cortical microtubules are gone. X1200. Scale bar = 10 pm.

VOLUME 103,1934

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FIG. 6. The abundance of cortical microtubules (vertical axis) as a function of time after l-MA application during the initial events. ++, Many microtubules; +, intermediate densities of microtubules; 0, no microtubules. In all, 12 coverslips (including some replicates) were prepared from a single batch of oocytes and examined. At 10 min cortical microtubules have begun to disappear in about one-third of the cortices; by 25 min, but not earlier, they are entirely gone.

as time progresses. Since the synchrony between oocytes becomes increasingly imperfect with time in culture, it remains unresolved whether the cyclic assembly-disassembly of microtubules persists during cleavage divisions or whether cortical microtubules just gradually fade away altogether. Rates of microtubule disassembly and reassembly. We specifically examined the rate of the initial disassembly of cortical microtubules in a single batch of oocytes (Figs. 4 and 5). As in other experiments, denuded oocytes were cultured in CaFSW; GVBD was observed at 44 min. Specimens of isolated cortices were prepared every 5 min after l-MA treatment. The array of cortical microtubules remained unchanged until 10 min after lMA when about one-third of the oocytes began to exhibit a slightly diminished network. By 15 and 20 min no cortices contained the full array of microtubules, although most contained intermediate arrays. By 25 min virtually all microtubules had disappeared (Fig. 4), as has already been mentioned (Fig. la). These changes are summarized graphically in Fig. 5 and suggest that, once perceptible microtubule disassembly begins, the array completely disappears within a 15-min interval. During the first 30 min after l-MA treatment, the oocyte cortex undergoes many changes in the actin-based cytoskeleton (Otto and Schroeder, 1984a). As shown in Fig. 4, the cortical microtubules bear no discernible positional relation to the actin-filled spikes that appear soon after l-MA treatment (Schroeder, 1981; Schroeder and Stricker, 1983). On the other hand, since it is known that spikes reach a maximum length at about 10 min and begin to withdraw thereafter, there is a temporal coincidence between spike withdrawal and the disas-

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sembly of cortical microtubules. In the present experiment (Fig. 4), both structures have virtually disappeared by 26 min. In addition to this evidence for rapid disassembly of cortical microtubules, Fig. 3 provides data that microtubule reas8smbly can also be very rapid. In our best example (Fig. 3b, batch E) no cortical microtubules were detected at 185 min after l-MA, yet by 203 min oocytes from the same culture displayed an extensive array. Complete disassembly and complete reassembly of the cortical array, therefore, both occur at comparable rates and can take place within about 15 min.

Pqwwiotti asters, nwiotic spindles, and polur bodies. In a relatively small, yet significant number of isolated cortices from oocytes at different stages, we have observed a radially arranged “aster” of microtubules. Since we do not see more than one such array per cortex, we infer that it represents a true centrosome-aster complex or premeiotic aster, as previously described near the animal pole in whole oocytes (Otto and Schroeder, 1934b). Because it is a localized structure, presumably we observe it only when the relevant part of the cortex is isolated, At stages when the cortical array of microtubules is present, microtubules of premeiotic asters seem to be superimposed upon and intermingled with cortical microtubules (Fig. 6a). Radiating microtubules extend about 66 pm from one or two adjacent tubulin-containing focal centers, which may be centrosomes. At stages when the cortical array is normally absent, for example during meiosis, solitary asters of microtubules still persist (Fig. 6b) indicating that the two systems of microtubules behave differently during maturation and development. Microtubules of the premeiotic asters at these latter stages are somewhat sparser and shorter. Although the meiotic spindles of maturing starfish oocytes are also situated at the extreme cell periphery (attached to the cortex in some sense), we have not unambiguously observed a spindle belonging to an oocyte during the early stages of the first meiotic division, i.e., before polar body formation. On the other hand, as illustrated in Fig. 7, we have frequently observed meiotic spindles at later stages, when they are specifically contained within and belonging to polar bodies. Judging by the configurations of microtubules and negative images of chromosomes or nuclei some of these spindles are at meiotic metaphase (Figs. 7s and b), telophase (Figs. 7ce), and interphase (Figs. 7f and g). Late-stage spindles often exhibit midbodies, identifiable as nonstaining bands across bundles of microtubules (Figs. 7c and d, arrows). Microtubule patterns in polar bodies provide additional indications of a cyclic alternation between meiotic microtubules and a putative intermeiotic array. At

FIG. 6. Antitubulin staining patterns in isolated cortices illustrating putative premeiotic asters and centrosomes (bright dots where microtubules converge). Before l-MA treatment (a) asters with very long microtubule rays are superimposed upon a more random array of cortical microtubules. It is presumed that such asters are associated with the cortex of the animal pole. After l-MA treatment (b), when nonastral cortical microtubules have disappeared, asters with shorter rays are still seen. X1266. Scale bar x 10 pm.

meiotic metaphase, spindle microtubules predominate (Fig, 7b), whereas at interphase polar bodies contain no obvious spindle but exhibit prominent “basket-like” arrays emanating from a single focal center (Figs. 7f and g). These “basket-like” arrays resemble the cortical microtubules of the oocyte less than they resemble asters. Unfortunately, it is usually not possible to correlate precise stage-dependent changes in a polar body with microtubule configurations in the parent oocyte since the two cells are often separated during the procedure for isolating cortices. DISCUSSION

Phase relations of the microtub& assembly-disassembly cycle. After discovering the extensive array of cortical microtubules in starfish oocytes (Otto and Schroeder, 1934b), we suspected they would sooner or later disassemble in response to 1-methyladenine treatment. In

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FIG. 7. Antitubulin staining patterns in meiotic apparatuses and polar bodies found in maturing oocytes during meiosis. A variety of microtubule configurations can be seen. (a and b) Phase-contrast and fluorescence micrographs of a polar body at the edge of an isolated cortex; it contains a meiotic spindle with nonstaining images of metaphase chromosomes on an equatorial plate. (c-e) Meiotic apparatuses in which polar bodies (upper portions) are attached to arrays of microtubules which extend into oocytes; these two portions are separated by constrictions and/or nonstaining midbodies (arrows). (f and g) Fully formed polar bodies filled with basket-like arrays of microtubules. X1200. Scale bar = 10 pm.

fact, we find that the cortical microtubules not only rapidly and synchronously disassemble but they later reassemble in a stage-dependent manner. Indeed, we have documented that cortical microtubules undergo two complete cycles of disassembly and reassembly between meiotic prophase (germinal vesicle stage) and first cleavage, as summarized in Fig. 8. Their subsequent fate remains to be determined but we suspect that the cycle persists into the early cleavage divisions. The presence and absence of cortical microtubules correlate with the phases of two other cellular cycles: the cycle of nuclear envelope dissolution and reformation

VOLUME 103,1934

and the cycle of formation and breakdown of the division apparatus (meiotic or mitotic spindle). In general, the presence of an intact nucleus and the absence of a division apparatus correlates with the presence of cortical microtubules; the single, and perhaps minor, exception to this correlation is the 20-min period just before germinal vesicle breakdown when the nuclear envelope appears intact but cortical microtubules have already disassembled. Conversely, the absence of an intact nucleus and the presence of a meiotic or mitotic apparatus correlate closely with intervals when cortical microtubules are absent. Thus, the existence of cortical microtubules exhibits a parallel correlation with intact nuclei and an inverse correlation with spindle microtubules. As Chambers and Chambers (1949) first demonstrated, meiotic divisions in fertilized starfish oocytes occur precociously compared to meiotic divisions in urlfertilized oocytes. In fertilized Pisaeter oocytes, meiosis and the first reappearance of cortical microtubules in fertilized oocytes are coordinately accelerated by about 45 min, relative to the same events in unfertilized oocytes (Fig. 3). This represents a natural experiment demonstrating that the cycles of cortical microtubules are independent of elapsed time per se and are indeed phenomenologically linked to other cellular events. Starfish oocytes are not alone in exhibiting inverse cycles of cytoplasmic and spindle microtubules. A similar phenomenon occurs during the cell cycle in cultured mammalian cells (Brinkley et uL, 1975; Aubin et ah, 1980; Brenner and Brinkley, 1981) in which a “cytoplasmic microtubule complex” during interphase alternates with mitotic microtubules. Such cytoplasmic microtubules are organized by the centrosomes and are often thought to be cytoskeletal in function; disappearance of the complex allows the cultured cells to round-up during cell division. The mechanism responsible for the reciprocal regulation of the assembly-disassembly of these two microtubules systems, however, remains unresolved. We have interpreted the disappearance and reappearance of microtubules in isolated cortices as evidence of disassembly and reassembly, respectively. In the future, we intend to analyze these events by indirect immunofluorescence of whole mounts and in thin sections of intact oocytes. Functional sign~cance of cortical microtubules. The observation that the first disappearance of cortical microtubules corresponds to the end of the hormone-dependent period (Schroeder and Stricker, 1983) raises thd possibility that they are essential for the action of lMA. This seems unlikely, however, since colchicine has no effect upon the timing of GVBD or the hormonedependent period (unpublished observations), even though cortical microtubules are known to be eliminated by this treatment (Otto and Schroeder, 198413).

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FIG. 8. Summary and interpretation of the cyclic change in occurrence of cortical microtubules in starfish oocytes during early stages of normal development, including hormone-induced germinal vesicle breakdown, fertilization (sperm symbol at 90 min), maturation, and early cleavage divisions. Vertical axis: ++, many microtubules; +, intermediate density of microtubules; 0, no microtubules. An extensive cortical array of microtubules is a natural feature of immature oocytes stored in the ovary (Otto and Schroeder, 1994b). It persists unchanged for about 10 min after l-MA application, then disappears before germinal vesicle breakdown. No cortical microtubules are found during fertilization or while the two meiotic divisions are under way. Microtubules reappear in large numbers at the end of maturation when the second polar body (polar body II) forms; at this time both male and female pronuclei are present (male and female symbols). Cortical microtubules then disappear again as mitosis of the first cleavage division ensues and then reappear (although only partially) when the first cleavage furrow begins (cleavage 1). The final two disappearances of microtubules, coinciding with the mitotic phases of the second and third divisions, are postulated events and have not yet been documented concretely.

Are cortical microtubules in starfish oocytes basically cytoskeletal in function? Cortical microtubules probably exist during the long period that oocytes are stored in the ovary when cytoskeletal support of the cortex might be advantageous. Immature oocytes exhibit a very high level of mechanical stiffness (resistance to deformation) which declines after the application of l-MA at roughly the same time that cortical microtubules disassemble (Nakamura and Hiramoto, 1978; Nemoto et oL, 1980, Yamamoto and Yoneda, 1933), and it is conceivable that the transitory stiffness is due to the transitory ,meshwork of microtubules. To test this possibility, Professor M. Yoneda visiting from Kyoto University, Japan compared the stiffness of immature Pisaster oocytes before and after treatment with sufficient colchicine to depolymerize cortical microtubules. Using his calibrated compression method (Yoneda, 1973), he determined that the stiffness is equally high for both cases: 3.54 f 0.48 dyne/cm (SD; n = 5) for denuded control oocytes in CaFSW and 3.79 f 0.47 dyne/cm (SD, n = 5) for oocytes incubated for 2 hr in lo-’ M colchicine in CaFSW (personal communication). Thus, neither the presence or the cyclic behavior of cortical microtubules in starfish oocytes explain the initial high level of stiffness in oocytes or its decline after l-MA. To explain such major changes in the mechanical properties of maturing starfish oo-

cytes, it may be more relevant to focus attention on the actin-based cortical cytoskeleton (Otto and Schroeder, 1934a) or on the organizational and consistency changes involving the entire cytoplasm (Hiramoto, 1976; Shoji et cd, 1978; Coffe et ah, 1982). Do the cortical microtubules provide a store of tubulin for other structures in the cell? The inverse correlation of assembly-disassembly cycles of cortical and spindle microtubules may suggest a priori that tubulin subunits are exchanged between these mutually exclusive microtubule arrays. Alternatively, it is frequently assumed that cells contain a sufficient excess of tubulin subunits to sustain several competing requirements, thereby obviating a need for storage forms. These ideas cannot be critically evaluated without further data on tubulin pool size and physiological factors involved in microtubule assembly. In this connection, we have already demonstrated that cortical microtubules account for about 50 pg of tubulin per oocyte and that an additional (and presumably much larger) pool of unpolymerized tubulin exists in immature oocytes since treatment with taxol causes a marked augmentation of the array of cortical microtubules (Otto and Schroeder, 1934b). How may subpopulutions of microtubules? The behavior of the premeiotic asters (Fig. 6) obviously differs from the behavior of cortical microtubules and meiotic

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apparatus microtubules. These asters do not appear to undergo comparable cycles of assembly and disassembly since they have been observed before and after the cortical array assembles. Similarly, these asters are found before and during stages when the meiotic apparatus is also present. Therefore, the meiotic asters may represent a third subpopulation of microtubules whose distinctive properties include stability during early aspects of the meiotic cycle. A fourth subpopulation of microtubules may be represented by the sperm aster of fertilized starflsh oocytes. Although not described here, this array of microtubules is organized by MTOCs (centrosomes) introduced by the fertilizing sperm. Sperm asters exhibit phases of growth that are distinct from those of other microtubule subpopulations (Hirai et al, 1981). Does the premeiotic aster anchor the germinal vesicle? A stable premeiotic aster situated at the animal pole may be important as a cytoskeletal anchor that maintains the polar eccentricity of the germinal vesicle. The centrosomes of this aster, being located next to the germinal vesicle (Otto and Schroeder, 1984b), may actually be attached to the nuclear envelope, just as centrioles are attached to nuclear envelopes in other cells (Bornens, 19’77; Maro and Bornens, 1980). Microtubules of the premeiotic aster could fasten the centrosome-germinal vesicle complex to the cortex of the animal pole, where it is maintained during the long period of storage in the ovary and during ovulation. Similarly, since the premeiotic aster persists at the animal pole even after GVBD, it may also anchor the peripheral pole of the meiotic apparatus to the cortex of the animal pole of the maturing oocyte; some such cortical anchorage would appear to be necessary in all instances of polar body formation, where cell division must be highly asymmetric and eccentric. Indeed, a premeiotic aster may actually be a precursor of the peripheral aster, so the continuous association between its microtubules and the cortex could guarantee that the meiotic apparatus forms with its peripheral centrosome next to the animal pole, as previously suggested by Shimizu (1981) from his studies of Z’~e~ eggs. Regulation of microtubule assembly-disassembly. It is generally believed that immature oocytes contain two centrosomes which serve as the microtubule organizing centers (MTOCs) for constructing meiotic spindles and asters (Wilson and Mathews, 1895). This idea is supported by our previous observations of fluorescent dots at the center of premeiotic asters (Otto and Schroeder, 1984b). It seems doubtful, however, that cortical microtubules in star&h oocytes are also assembled at centrosomes, for the following circumstantial reasons: they are not radially organized around one or two central foci so, if they do arise from centrosomes, they must

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secondarily become disconnected and dispersed-a phenomenon that has been reported elsewhere (Vorobjev and Chentsov, 1983); furthermore, microtubules reappear in mature oocytes after formation of the second polar body, a stage of maturity at which sea urchin eggs lack centrosomes (Kuriyama and Borisy, 1983). Thus, cortical microtubules may not emanate from centrosomes. It seems more likely that cortical microtubules are assembled at multiple sites distributed throughout the cortex. Differential activity of dispersed assembly sites could explain the partial reassembly observed at certain stages (intermediate arrays in Figs. 3 and 8). For comparison, evidence for the existence of subcortical sites for the assembly of microtubules in sea urchin eggs can be seen in cases of parthenogenetic activation (Mar, 1980), putative partial activation (Harris, 1979), and treatment with taxol (Schatten et d, 1982). Compared to MTOC-associated microtubules, such microtubules could be designated “free” microtubules (De Brabander,, 1982). The diversity of assembly-disassembly cycles for var-’ ious subpopulations of microtubules in starfish oocytes points out the complexity of the question of regulation. Perhaps each set of microtubules (the cortical array, the spindle microtubuies, the meiotic and mitotic asters, and the sperm aster) has its own MTOCs and specific regulators for microtubule assembly-disassembly. Various physiological regulators have been suggested as important for assembly or disassembly of microtubules. Based on both in vitro and in viva experiments on various cell types, pH, and the availability of highenergy phosphates are thought to be important (reviewed by De Brabander, 1981; De Brabander et al, 1982). On the other hand, it is too early to generalize, as indicated by contradictory experimental data: in certain cases elevated intracellular pH causes microtubules to disassemble (Maro and Bornens, 1982) yet the opposite occurs in sea urchin eggs where microtubules assemble when the intracellular pH is experimentally elevated (Coffe et d, 1982; Hamaguchi, 1982). We do not yet know enough about the physiological regulators of microtubule assembly-disassembly, about subpopulations of microtubules with diverse stabilities, or about different kinds of MTOCs with varied responsiveness to intracellular changes. In conclusion, although the function and regulation of cortical microtubules in starfish oocytes are presently elusive, the cyclic nature of their assembly and disassembly provides striking evidence of dramatic underlying changes during oocyte maturation and postfertilization development. Further study of these cycles may offer insights into the mechanisms by which development is initiated in starfish.

SCHROEDER AND Orro

C+es

We gratefully acknowledge the gifts of antitubulin antibodies from Dr. S. H. Blose, Dr. B. R. Brinkley, and Dr. W. E. Gordon. We also appreciate the key contribution of Dr. M. Yoneda in measuring the mechanical properties of oocytes. Our work was supported by research Grants PCM-8291866 to T.E.S. and PCM-8929984 to J.J.O. from the National Science Foundation. REFERENCES Aum~, J. E., OSBORN, M.. and WEBER, K. (1989). Variations in the distribution and migration of centriole duplexes in mitotic PtK2 cells studied by immunofluorescence microscopy. J. CeU s%i 43,1’77194. BORNENS,M. (1977). Is the centriole bound to the nuclear membrane? Nature (London) 370,89-82. BRENNER, S. L., and BRINKLEY, B. R. (1981). Tubulin assembly sites and the organization of microtubule arrays in mammalian cells. Cold Spring Harbor Synpr Qwmt Bid 46.241264. BRINKLEY, B. R., FULLER, G. M., and HIGHFIELD, D. P. (1975). Cytoplasmic microtubules in normal and transformed cell in culture: Analysis by tubulin antibody immunofluorescence. Proc Nat1 Acad Sci USA 72,4981-4985. CIUMB~RS, R., and CIUMBERS, E. L. (1949). Nuclear and cytoplasmic interrelations in the fertilization of the Aster& egg. BioL Bull 96, 270-282. COFFE, G., ROLA, F. H., SOYER, M. O., and PUDLES, J. (1982). Parthenogeuetic activation of sea urchin egg induces a,cyclic variation of the cytoplasmio resistance to hexylene glycol-Triton X-199 treatment. Exp. CeU Ra 137,63-72. DE BRABANDER, M. (1982). A model for the microtubule organizing activity of the centrosomes and kinetochores in mammalian cells. CM.?Bid Int Rsp 6,991-916. DEBRABANDER, M., GEUENS, G., NTJYDENS,R., WJLLENBRORDS,R., and DE MEY, J. (1981). Microtubule stability and assembly in living cells: The influence of metabolic inhibitors, tax01 and pH. Cold spring Harbor Sgmp Quant Bid 44,227-%&O. GILOH, H., and SEDAT, J. H. (1982). Fluorescence microscopy: Reduced photobleaching of rhodamine and fluorescein protein conjugates by n-propyl gallate. soience 217,1262-1265. . HAMAGUIXI, M. S. (1982). The role of intracellular pH in fertilization of sand dollar eggs analyzed by microinjection methods. Deu. Growth D@r. 24,443-461. HARRIS, P. (1979). A spiral cortical fiber system in fertilized sea urchin eggs. De-v. Bid 68,526-632. HIRAI, S., NAGAHAMA, Y., KISHIMOTQ T., and KANATANI, H. (1981). Cytoplasmic maturity revealed by the structural changes in incorporated spermatozoon during the course of starAsh oocyte maturation. Dev. Gmwth D&W. 23,466-478.

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