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Ooplasm segregation in the squid embryo, loligo pealeii.

After fertilization, squid egg ooplasm streams toward the animal pole to create a clear lens-shaped blastodisc cap where meroblastic cleavage occurs (1). Exposing the embryo to cold (4 [degrees]C) after fertilization inhibits blastodisc formation (2), suggesting that microtubules may be associated with this ordered movement of cytoplasm (3). Although microtubules have been correlated with cytoplasmic movements that follow fertilization in amphibians (4), ascidians (5), and annelids (6), these embryos do not form blastodisc caps and, unlike the squid, they undergo complete or holoblastic cleavage. Interestingly, ooplasmic segregation, blastodisc cap formation, and meroblastic cleavage all take place in the zebrafish embryo, but here microfilaments and not microtubules have been shown to direct the segregation of ooplasm from the yolk (7).

To clarify the role of the cytoskeleton during early development in squid, embryos cultured at 20 [degrees]C in petri dishes lined with 0.2% agarose (Sigma, Type II) and filled with Millipore-filtered seawater (MFSW), were treated 30 min after in vitro fertilization (8) with either 0.01-20 [micro]g/ml of the microfilament inhibitor cytochalasin D (Sigma) (3 trials, 50 to 75 embryos per dish) or 0.5-10 [micro]g/ml of the microtubule inhibitor colchicine (Sigma) (6 trials, 50 to 75 embryos per dish). Stock solutions of each inhibitor were prepared in dimethyl sulfoxide (DMSO) (Sigma), and DMSO (0.1%) was therefore added to MFSW as a control. Embryos were observed for at least 4 h after fertilization during blastodisc formation and after incubation overnight at 17 [degrees]C.

Embryos cultured in MFSW (Fig. 1a) or MFSW and DMSO (Fig. 1d) formed normal 40-50 [micro]m thick blastodiscs by 4 h and underwent normal cleavage and early development. Blastodisc cap formation occurred in all embryos treated with 0.01, 0.05, 0.1, 0.5, 1.0, 2.0, 4.0, and 10.0 [micro]g/ml cytochalasin D; however, at all but the lowest concentration of microfilament inhibitor, the entire cortical yolk cell membrane appeared disrupted by the presence of small and large vesicles of cytoplasm (Fig. 1c). This result was similar to what had previously been reported in squid (9). Normal yolk cell membranes were observed in 45% of the embryos treated with 0.05 [micro]g/ml cytochalasin D, while all yolk membranes appeared similar to the control in the 0.01 [micro]g/ml group. It is important to note that embryos from these different treatment groups failed to undergo normal development. In contrast, colchicine prevented ooplasm segregation and blastodisc formation in all embryos cultured at concentrations of 10.0, 7.5, and 5.0 [micro]g/ml, although the thin layer of cytoplasm present prior to fertilization was maintained (Fig. 1b). Embryos cultured in 2.5 [micro]g/ml all formed thinner, <20 [micro]m, blastodisc caps by 4 h but failed to retain them after culture overnight. While blastodisc caps (30-35 [micro]m) formed in the cultures treated with 1.0 and 0.5 [micro]g/ml colchicine, after overnight culture these caps were either lost or reduced to small abnormal shaped discs or sacs of cytoplasm at the animal pole. Normal cleavage and development were never observed in any embryos treated with cytoskeletal inhibitors. Normal blastodisc cap formation and cleavage occurred in all embryos treated with DMSO (0.1%) (Fig. 1d).

These results suggest that a microtubule-associated mechanism is responsible for ooplasm segregation and blastodisc formation in the squid. Although microfilaments do not seem to be required for ooplasm movement to the blastodisc, the disruption of the cortical yolk cell membrane by cytochalasin D suggests that they may have a role in stabilizing the yolk cell membrane or regulating cortical cytoplasm flow toward the animal cap. In contrast, microtubules do not seem to be involved in ooplasm segregation and blastodisc formation in the zebrafish embryo (10), where microfilaments have been shown to direct ooplasm flow along streamers of cytoplasm within the central region of the yolk cell. Interestingly, in embryos of another fish, medaka, both microtubules and microfilaments have been shown to be involved in ooplasm segregation. In this fish, not only did cytochalasin D inhibit blastodisc formation, but colchicine treatment also resulted in less directed ooplasm movements (11). Thus it seems, in medaka, microf ilaments and microtubule networks may function in concert during ooplasm segregation.

One possible clue to understanding the subtle similarities and differences observed in embryos of these fish and the squid may be found when the location of ooplasm within the unfertilized egg and the route of cytoplasmic flow are considered. In contrast to the central flow of ooplasm in zebrafish, ooplasm flow in medaka occurs cortically along meridonal pathways (11). Similarly, in the squid embryo, where microtubular arrays can be visualized by antibody labeling, the ooplasm flow to the cortex of the yolk cell surface is restricted to the outermost cortical layer of cytoplasm (unpubl. results). With antibodies to [beta]-tubulin, unfertilized eggs were observed to possess circular swirls of tubulin staining within the cortex. Yolk just below this thin layer did not label for [beta]-tubulin. After fertilization, these patterns change and [beta]-tubulinrich streams oriented toward the animal pole are formed along the outer cortex of the embryo. Perhaps the reliance on two supporting cytoskeletal mechanisms wit hin the cortex to move ooplasm to the blastodisc, as shown in medaka, is characteristic of eggs that possess a dense central yolk and cortical ooplasm, and may underly this process in the squid embryo. With this possibility in mind, it will be important to reexamine these elements in other embryos where microtubules alone, or in concert with microfilaments, have been linked to ooplasm segregation and movement following fertilization. In addition, further analysis of microtubule and microfilament arrays during ooplasm segregation and in the presence of cytoskeletal inhibitors will further extend our understanding of the mechanism of blastodisc formation and early development in the squid embryo.

This work was supported by a Faculty Development Grant from St. Mary's College of Maryland to Karen Crawford and would not have been possible without Dr. Robert Baker and his laboratory group at the Marine Biological Laboratory.

Literature Cited

(1.) Arnold, J. M. 1968. Dev. Biol. 18: 180-197.

(2.) Crawford, K. 2000. Biol. Bull. 199: 207-208.

(3.) Yahara, I., and F. Kakimoto-Sameshima. 1978. Cell 15: 251-259.

(4.) Houliston, E., and R. P. Elinson. 1991. Development 112: 107-117.

(5.) Sawada, T., and G. Shatten. 1989. Dev. Biol. 132: 331-342.

(6.) Eckberg, W. R. 1981. Differentiation 19: 55-58.

(7.) Leung, C. F., S. E. Webb, and A. L. Miller. 1998. Develop. Growth Differ. 40: 313-326.

(8.) Klein, K. C., and L. A. Jaffe. 1984. Biol. Bull. 167: 518.

(9.) Arnold, J. M., and L. D. Williams-Arnold. 1974. J. Embryol. Exp. Morphol. 31: 1-25.

(10.) Leung, C. F., S. E. Webb, and A. L. Miller. 2000. Dev. Growth Differ. 42: 29-40.

(11.) Abraham, V. C., S. Gupta, and R. A. Fluck. 1993. Biol. Bull. 184: 115-124.

[Figure 1 omitted]
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Author:Crawford, Karen
Publication:The Biological Bulletin
Geographic Code:1USA
Date:Oct 1, 2001
Words:1155
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