Role of specialized microvilli and the fertilization envelope in the spatial positioning of blastomeres in early development of embryos of the starfish Astropecten scoparius.
In mammals, "compaction" of the embryo occurs at the 8-cell stage, leading to binding of the blastomeres together (Tarkowski and Wroblewska, 1967; Hillman et al., 1972; Fleming et al., 1989). The presumptive fate of the blastomeres is first determined at this stage. In these embryos there is accordingly a close relationship between the formation of a binding device for the blastomeres and their differentiation.
In most invertebrate embryos, however, the presumptive fate of each blastomere is determined at a much earlier stage than the formation of intercellular binding devices such as desmosomes (cf. Kuraishi and Osanai, 1989; Kiyomoto and Shirai, 1993a, 1993b). It is known that a number of morphogenetic determinants, which play a role in the determination of the blastomeres' presumptive fate, are located unevenly within the egg of these animals (Boveri, 1910; Horstadius, 1935; Anderson and Nusslein-Volhard, 1984). In those embryos, cell division occurs not just to increase cell numbers, but also to distribute the various determinants to the respective blastomeres and induce cell specialization. If the distribution of the determinants is upset, or some of the blastomeres are removed, and the spatial positions of blastomeres are changed, normal development of the embryo is disturbed (Chabry, 1887; Horstadius, 1935; Gilbert, 1994). Animal development provides examples of many ways to maintain the relative positions am ong blastomeres in the period before the formation of intercellular binding devices. For example, in the embryos of jellyfish (Dan and Dan, 1947) and hydra (Martin et al., 1997), species that lack an investing layer, spinning processes or ladderlike membrane structures are formed, respectively, between blastomeres. In sea urchins, a hyaline layer is formed on the surface of the egg immediately after fertilization (Endo, 1961), and this layer holds the outer surface of the embryo in place by means of microvilli (Dan and Ono, 1952; Endo, 1961). If the embryo is exposed to antibodies against the hyaline layer, normal morphogenesis is inhibited (Adelson and Humphreys, 1988). In sea urchin embryos, therefore, the relative positions among the blastomeres are maintained by the hyaline layer and microvilli (Burgess and Schroeder, 1977; Adelson and Humphreys, 1988). In the embryos of starfish, which belong to the same class of Echinodermata as sea urchins, a hyaline layer is not formed in the early cleavage stages (Ma tsunaga et al., 2000), so some other means must exist to unite the blastomeres. In the embryos of the starfish Astropecten scoparius, we observed the individual blastomeres lining up along the inner surface of the fertilization envelope and found threadlike structures (fixing processes) connecting the blastomeres to the fertilization envelope. Normal morphogenesis was prevented if the connection between the blastomeres and the fertilization envelope was severed, suggesting that the fertilization envelope and the fixing process play an important role in maintaining the relative positions of the blastomeres in A. scoparius embryos. In this study, we examined the shape, character, and function of these fixing processes.
Materials and Methods
Handling of gametes
The starfish Astropecten scoparius was collected from Tateyama Bay in Chiba Prefecture, Japan. Isolated ovaries were treated with calcium-free seawater for 20-30 min, and then ovulation was induced by returning them to filtered normal seawater (NSW), yielding immature oocytes without the follicular envelope (Nemoto et al., 1980). "Dry sperm" taken from isolated testes were diluted with NSW just before use. Maturation division of oocytes was induced with 2 [micro]M of 1-methyladenine (1-MeAde; Sigma-Aldrich, St. Louis, MO) dissolved in NSW (Kanatani, 1969). At 20 [degrees]C, germinal vesicle breakdown (GVBD) and extrusion of the first and second polar bodies occurred, respectively, at about 25, 60, and 90 min after treatment with 1-MeAde. The maturing oocytes were fertilized 40 min after treatment with 1-MeAde. The fertilized oocytes and embryos were rinsed once with NSW and then cultured at 20 [degrees]C. The average diameters of the oocytes and the fertilization envelope were 210 and 300 [micro]m, respective ly.
Oocytes, eggs, and embryos were flattened to a 180-[micro]m thickness and examined under a differential interference-contrast microscope (Axiophoto; Zeiss, Oberkochen, Germany). Microphotographs were taken with Neopan 400 Presto film (Fuji Photo Film, Tokyo, Japan).
Transmission electron microscopy
Two fixing methods were used in this study. A quick-freeze and freeze-substitution method was mainly used for examination of cytoskeletons, and chemical fixation was used for examination of other structures.
Chemical fixation. Oocytes and embryos were first fixed in 2% glutaraldehyde (TAAB, Berkshire, England) dissolved in NSW for 2 h at room temperature, then in Karnovsky's fixative (Karnovsky, 1965) for 2 h at room temperature, followed by 10 h at 4 [degrees]C. After being rinsed several times with 0.1 M sodium-cacodylate buffer (pH 7.2), they were then fixed in 1% osmium tetroxide (Polyscience, Warrington, PA) dissolved in 0.1 M sodium cacodylate for 60 min on ice (Steinbeck et al., 1993). The fixed specimens were rinsed twice with distilled water (DW) and then dehydrated with ethanol.
Quick-freeze and freeze-substitution fixation. Oocytes were first treated with 15% butanediol (Wako Pure Chemical, Tokyo, Japan) diluted with NSW for 20 min on ice (Reimer and Crawford, 1997), and then they were collected with a hand-driven centrifuge. A 20-[micro]1 aliquot of the specimen was placed on an electron microscopy grid. Excess NSW was drawn off with a piece of filter paper, and the specimens were frozen by immersion for 10 s in liquid propane that had itself been cooled with liquid nitrogen. The frozen materials were placed in 4% osmium tetroxide/acetone cooled to -90 [degrees]C. Five days later, the specimens were first brought up to -20 [degrees]C and then to 4 [degrees]C for 2 h each. They were finally placed at room temperature (Burke et al., 1998) and dehydrated using acetone.
Embedding and examination. The dehydrated specimens were embedded in epoxy resin (Kushida, 1980), and ultrathin sections (90 min in thickness) were produced using an ultramicrotome (MT-XL, RMC Inc., Tucson, AZ). The sections were stained with uranyl nitrate and lead acetate for 2 min each and examined under a JEOL 1010 transmission electron microscope (JEOL, Tokyo, Japan).
Ultrathin sections made from the chemically fixed specimens underwent 5 min of etching with 3% sodium metaperiodate. After being rinsed with DW, they were treated with an antibody raised against sea urchin hyaline layer (Yazaki, 1968) diluted in 0.2% Tween 80 for 5 min at room temperature. After another rinse with DW, the sections were treated with an anti-rabbit IgG antibody (Sigma-Aldrich) bound to protein A-colloidal gold (20 nm in diameter), diluted 1/40 in 0.2% Tween 80, for 5 mm (Morris and Ciraolo, 1997). These sections were not stained with lead acetate or uranyl nitrate.
Cleavage and development of fertilized eggs
Oocytes fertilized immediately following germinal vesicle breakdown (GVBD) extruded a second polar body 1 [1/2] h after treatment with 1-MeAde, completing their maturation division (Fig. 1A). As with the embryos of other starfish species, elevation of the fertilization envelope continued for about 2 h after fertilization. The first cleavage occurred some 40 mm after the formation of the second polar body. After the first cleavage, the two blastomeres moved in a direction perpendicular to the cleavage plane, as if to separate from each other. Some 15-20 mm after the first cleavage, the two blastomeres impressed themselves into the fertilization envelope at the surface of their mitotic poles (Fig. 1B). As the area of contact between the blastomeres and the fertilization envelope became progressively larger, the blastomeres changed their shape to ellipsoids with the long axis in the animal-vegetal pole direction, producing space between them. The second cleavage occurred some 30 min after the first. After this c leavage, as after the first, space was produced between the blastomeres (Fig. 1C). The cleavage plane for the third cleavage was formed at right angles to the first and second cleavage planes. After this division also, the blastomeres did not separate from the fertilization envelope (Fig. 1D). The blastomeres at this stage adhered to the fertilization envelope over a wide area and had become flattened. Adjacent blastomeres had become widely separated from each other. A hyaline layer could not be distinguished. At the 16-cell stage and beyond, blastomeres continued to undergo cell division without separating from the fertilization envelope, and they lined up along its inside surface (Fig. lE). As a result, the embryo did not go through the cell-mass, "morula" shape, but became a blastula consisting of a single cell layer (Fig. 1F). When hatching commenced, a space developed between the fertilization envelope and the embryo, and the embryo began to spin around (Fig. 1G). Gastrulation began about 3 h after hatch ing. The embryos developed to mid-gastrulae 10 h after hatching (Fig. 1H). Development beyond this stage was described by Oguro et al. (1976).
Morphogenesis of embryos that have lost attachment between blastomeres and the fertilization envelope
When embryos at the 8-cell stage (3 h postfertilization) were subjected to a 1-mm pulse treatment with acidified seawater adjusted to pH 4 with HC1, blastomeres changed from a flattened shape to a globular one, and the area of the cell surface in contact with the fertilization envelope became smaller (compare Fig. 2A with Fig. 2B). Blastomeres also became globular when the fertilization envelope was removed. When embryos treated with acidified seawater were returned to NSW and then gently spun with a hand-driven centrifuge, blastomeres were gathered at the centrifugal end inside the fertilization envelope (Fig. 2C). In untreated embryos, however, the arrangement of the blastomeres was not disturbed even by such centrifugation. These results indicate that the attachments between the fertilization envelope and the blastomeres are severed by treatment with acidified seawater.
Even after embryos pulse-treated with acidified seawater were returned to NSW, the blastomeres did not restore their connection with the fertilization envelope but remained globular, not returning to their normal flattened shape. At 24 h postfertilization, at which time control embryos had formed gastrulae, the embryos treated with acidified seawater alone were morphologically abnormal, many with distorted shapes or abnormal archenterons, and some embryos had multiple archenterons (Fig. 3A, B). The effects of treatment with acidified seawater were especially marked if conducted between 3 and 4.5 h postfertilization (8- to 64-cell stage). No morphological abnormalities were seen in embryos treated with acidified seawater 6.5 h postfertilization ([2.sup.9]-cell stage) (Fig. 3C), and normal gastrulae were formed 24 h postfertilization (Fig. 3D). The effect of acidified seawater treatment was no longer observed when it was done with embryos 6.5 h postfertilization and beyond. These results indicate that in the ea rly stages of cleavage, blastomeres in A. scoparius embryos maintain their spatial positions relative to each other by means of the fertilization envelope.
The mechanism of anchoring blastomeres to the fertilization envelope
The above observations suggest the existence of some means that attaches blastomeres to the fertilization envelope. Within a minute postinsemination, when the fertilization envelope could be identified, a number of particles were observed in the perivitelline space (perivitelline-space particles) (Fig. 4A). A few minutes after insemination, many thin processes connecting the fertilization envelope and the egg proper could be seen (Fig. 4B, arrow). These processes will hereafter be referred to as fixing processes. These fixing processes elongated rapidly with elevation of the fertilization envelope and were located over the entire surface of the egg. The perivitelline-space particles moved along the fixing processes towards the fertilization envelope (Fig. 4B, arrowheads), reaching it by several minutes after insemination, by which time very few could be seen in the penvitelline space (Fig. 4C). The perivitelline-space particles adhering to the fertilization envelope eventually disappeared, leaving a smooth in ner surface (Fig. 4C). Around this time, the basal part of the process assumed the shape of a cone (Fig. 4C, white arrowheads). After the first cleavage, fixing processes could still be seen between the blastomeres and the fertilization envelope but not on the surface of the cleavage furrow (Fig. 4D; cf. Fig. 5A). The length of the fixing processes varied with the distance between the egg surface and the fertilization envelope; the longest, some 60 [micro]m, was seen in the vicinity of the cleavage furrow (Fig. 4D). Near the boundary between the areas of the blastomeres touching the fertilization envelope, fixing processes were short and straight, with no bending being seen. Short fixing processes were also observed in embryos at later stages of development (Fig. 5A, white arrowheads).
In embryos at the 8-cell stage, blastomeres still adhered to the fertilization envelope over a wide area and were ellipsoid (Fig. 5A; cf. Fig. 2A). Fixing processes (white arrowheads), near the boundary between the areas of the blastomeres touching the fertilization envelope, were thin and straight. After a 1-min pulse treatment with acidified seawater (Fig. 5B), fixing processes were badly damaged and became deformed and thicker (arrowheads) than those of embryos before treatment; they looked as if they had lost their rigidity. Blastomeres separated from the fertilization envelope and became roughly spherical. These observations suggest that the fixing processes are the way that blastomeres are anchored to the fertilization envelope.
Transmission electron microscopic examination of fixing processes
Microvilli, some 700 nm in length, were identified embedded in the vitelline coat on the surface of both immature (Fig. 6A) and maturing unfertilized oocytes (Fig. 6B). At 30 s postfertilization, the tips of the microvilli were connected to the vitelline coat that was transforming into the fertilization envelope (Fig. 6C, arrows). Subsequently, the microvilli elongated with the elevating of the fertilization envelope (Fig. 6D, E). The fixing processes seen under light microscopy (Figs. 4 and 5) are therefore considered to be specialized microvilli. These specialized microvilli sprout from all over the surface of the egg. At early cleavage stages, short fixing processes some 500 nm in length were seen on the surface of the blastomeres adhering to the fertilization envelope (Fig. 6F), and the tips of those processes were connected with the fertilization envelope (arrows). No extracellular matrix surrounding the embryo, such as a hyaline layer, was observed (Fig. 6D-F). Some membranous structures (white asterisk ) were sometimes seen in the space between the blastomeres and the fertilization envelope.
In the specimens prepared by the quick-freeze and freeze-substitution method, bundles of fibers could be identified within microvilli in immature oocytes (Fig. 6A). After oocyte maturation, these fiber bundles extended in the direction of the cytoplasm (Fig. 6B, white arrow), reaching a length of 2 [micro]m. The diameter of these fibers was 5-6 nm, so they can be considered microfilaments. Crossing the microfilament bundle, repeated bands were observed (Fig. 6B, black arrowheads).
We could not examine microfilaments in the fixing processes by transmission electron microscopy, because the fixing processes were not preserved by the quick-freeze and freeze-substitution fixation that effectively preserves microfilaments. To get information on these microfilaments, we examined the effects of cytochalasin D on the fixing processes. When the cytochalasin was applied to fertilized eggs, intact fixing processes disappeared within several minutes of treatment, and only fragmented fixing processes were observed in the perivitelline space (Fig. 7A). In embryos pulse-treated with the cytochalasin at early cleavage stages, the fixation of blastomeres on the fertilization envelope were severely damaged, and the blastomeres changed shape from flattened to globular (compare Fig. 7B with Figs. 1D and 2A). These results suggest the presence of microfilaments in the fixing processes.
Immuno-electron microscopic examination of the perivitelline-space particles
In unfertilized maturing oocytes, granular structures with fairly high electron density could be seen along the cortex (Fig. 8A, asterisks). On fertilization, these granules opened onto the cell surface and discharged the contents into the perivitelline space (Fig. 8B-D; cf. Fig. 6D, E), resulting in their disappearance from the cytoplasm (Fig. 6F). These granules are therefore considered to be cortical granules. Following exocytosis of the cortical granules, electrondense particles appeared in the perivitelline space, indicating that the particles correspond to the perivitelline-space particles observed under light microscopy (Fig. 4A, B) and are derived from the cortical granules. To confirm this, immuno-electron microscopy was performed using an antibody against the sea urchin egg's hyaline layer (Yazaki, 1968), which is known to derive from the cortical granules (Endo, 1961). As shown in Figure 8A, gold particles are present inside the cortical granules (asterisks) in unfertilized oocytes. After fertiliza tion, gold particles could be seen specifically in the cortical granule contents that were just being discharged outside the egg (Fig. 8B, asterisk). After their discharge into the perivitelline space (Fig. 8C, asterisk), the cortical granule contents, recognizable by the gold particles, reached the fertilization envelope (Fig. 8D, asterisk). As can be clearly seen in Figures 6 and 8, there is no hyaline layer over the egg surface, and gold particles cannot be identified outside the cell surface (Fig. 8C, D). As the perivitelline-space particles began to arrive at the fertilization envelope, a layer with fairly high electron density appeared on the inner part of the fertilization envelope, to which the tips of the fixing processes were joined (Fig. 6D, G). Near the cleavage furrows (Fig. 9), the inner layer was occasionally seen detached from the fertilization envelope and was linked with a number of fixing processes (arrows).
In the early cleavage stages before the establishment of some mechanism, such as desmosomes, to directly bind blastomeres, animal embryos devise various means to keep the blastomeres together and maintain their spatial positions relative to each other. In sea urchin eggs, a hyaline layer is formed by the exocytosis of cortical granules immediately after fertilization (Endo, 1961). Microvilli are embedded into the hyaline layer, thus fixing the surface of fertilized eggs to the hyaline layer. Each blastomere produced by cleavage is therefore held in place by the hyaline layer (Dan and Ono, 1952) by means of microvilli, thereby maintaining the three-dimensional spatial relationship among the blastomeres. In sea urchin embryos, even if the fertilization envelope is removed, normal morphogenesis continues as long as the hyaline layer remains (Showman and Foerder, 1979).
In this study, we found that in Astropecten scoparius embryos, which lack a hyaline layer, the relative positions among blastomeres are maintained by the fertilization envelope and the cytoplasmic threads (fixing processes) attached to it. As can be seen in Figure 6, fixing processes are altered microvilli. On fertilization, the tips of the microvilli buried in the vitelline coat are attached to the fertilization envelope, and the microvilli rapidly lengthen with elevation of the fertilization envelope. Because fixing processes are located all over the cell surface, the balance of their tensions may keep the egg placed in the center of the fertilization envelope at the one-cell stage before the first cleavage. Hiramoto (1954, 1955) predicted the presence of a certain structure that connects the egg proper and the fertilization envelope in sea urchin zygotes, but its true nature is unknown.
After maturation division begins in A. scoparius oocyres, microfilament bundles within the microvilli lengthen, protruding some distance into the cytoplasm from the bases of the microvilli in unfertilized oocytes. Such lengthening following maturation division can be thought to be preparation for rapid elongation of microvilli postfertilization. Repeated bands were observed crossing the microfilament bundles (Fig. 6B, black arrowheads). The interval between the strips is some 60 [micro]m. These bands may be formed of a kind of fascins, a family of actin-bundling proteins, although the band interval is wider than that reported in other types of cells (11 nm) (see Edwards and Bryan, 1995, for review).
After the first cleavage, the fixing processes near the cleavage furrow can be as long as 60 [micro]m This rivals the length of the acrosomes of the sea cucumber Dendrochirotida thyone (Tilney and Inoue, 1982)--a well-known example of the long actin fibers in gametes. In embryos after the second cleavage, fixing processes are short around the contact region between the blastomeres and the fertilization envelope (Fig. 6F). On the surface of the blastomeres in contact with the fertilization envelope, fixing processes are so short that they are difficult to detect by light microscopy (see Figs. 4D and 5A). Such short processes are thought to be the product of reduction of the long fixing processes seen before the first cleavage, although the mechanism of the reduction is now known.
Before the first cleavage, perivitelline-space particles reach the fertilization envelope and form its inner layer (Fig. 8D). As can be seen in Figures 6 and 8, this inner layer is derived from substances contained in the cortical granules, and the tips of the fixing processes are attached to the layer. Near the cleavage furrows, the inner layer, with fixing processes attached, could sometimes be seen detached from the fertilization envelope (Fig. 9), suggesting that the inner layer, derived from cortical granule contents, reinforces the attachment of the fixing processes to the fertilization envelope.
Treatment with acidified seawater breaks the attachments between the blastomeres and the fertilization envelope. The effects of acidified seawater are not seen on embryos more than 6.5 h postfertilization, indicating that after that time, the fixing processes play a lesser role in holding the blastomeres in place. Intercellular binding devices such as desmosomes are probably fulfilling this role at this stage (cf. Dan-Sohkawa et al., 1986). It is not known when the connections between fixing processes and the fertilization envelope are broken, but around the time of hatching the embryo is able to spin around inside the fertilization envelope, so the attachments have been severed by this time at the latest.
The spatial relationship between blastomeres is maintained by the hyaline layer and microvilli in sea urchin embryos, and in A. scoparius embryos, which lack a hyaline layer, by the fertilization envelope and microvilli. The morphology and location of the hyaline layer and the fertilization envelope are different, but each contains substances derived from cortical granules. It is of great interest that, as shown in the immuno-electron microscopic findings (Fig. 8), they contain some common elements. This way of using the fertilization envelope as a scaffold for morphogenesis, which has been seen in A. scoparius embryos, may be a common mechanism for starfish embryos that lack a hyaline layer in early cleavage stages.
We thank Dr. I. Yazaki, Tokyo Metropolitan University, for the kind gift of an antibody against the hyaline layer of sea urchin eggs, and Dr. Y. Nakajima, Keio University, for valuable advice and kind help in preparing specimens fixed by the quick-freeze and freeze-substitution method. Our thanks are also due to Mr. M. Yamaguchi and Dr. M. Kiyomoto, Tateyama Marine Laboratory, Ochanomizu University, for technical assistance and providing facilities.
Received 25 June 2001; accepted 4 March 2002.
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MAKI MATSUNAGA (1)
ISAO UEMURA (2)
MIWA TAMURA (1)
SHIN-ICHI NEMOTO (1), *
* To whom correspondence should be addressed. E-mail: firstname.lastname@example.org
(1.) Department of Biology Faculty of Science, Ochanomizu University, Otsuka, Bunkyo, Tokyo 112-8610, Japan;
(2.) Department of Biology, Faculty of Science, Tokyo Metropolitan University, Minami-Oosawa, Hachioji, Tokyo 192-0397, Japan
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|Author:||Matsunaga, Maki; Uemura, Isao; Tamura, Miwa; Nemoto, Shin-Ichi|
|Publication:||The Biological Bulletin|
|Date:||Jun 1, 2002|
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