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Confocal microscopy of F-actin distribution in Xenopus oocytes

Published online by Cambridge University Press:  26 September 2008

Amy D. Roeder  and
David L. Gard
Amy D. Roeder
Affiliation:
Department of Biology, University of Utah, Salt Lake City, Utah, USA.
David L. Gard*
Affiliation:
Department of Biology, University of Utah, Salt Lake City, Utah, USA.
*
David L. Gard, Department of Biology, University of Utah, Salt Lake City, UT 84112, USA. Tel: 801-581-7365. Fax: 801-581-4668.
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Summary

We have used rhodamine-conjugated phalloidin and confocal microscopy to examine the organisation of filamentous actin (F-actin) during oogenesis in Xenopus laevis. F-actin was restricted to a thin shell in the cortex of oogonia and post-mitotic oocytes less than 35 μm in diameter. In oocytes with diameters of 35–50 μm F-actin was observed in three cellular domains: in the cortex, in the germinal vesicle (GV) and in a network of cytoplasmic cables. Initially, actin cables were sparsely distributed in the cytoplasm, with no evidence of discrete organising centres. In larger stage I oocytes, a dense network of actin cables extended throughout the cytoplasm, linking the GV and mitochondrial mass to the cortical actin shell. Apart from the F-actin associated with the mitochondrial mass, no evidence of a polarised distribution of F-actin was apparent in stage I oocytes. F-actin was observed also in the cortex and the GV of stage VI oocytes, and a network of cytoplasmic cables surrounded the GV. Cytoplasmic actin cables extended from the GV to the animal cortex, and formed a three-dimensional network surrounding clusters of yolk platelets in the vegetal cytoplasm. Finally, disruption of F-actin in stage VI oocytes with cytochalasin resulted in distortion and apparent rotation of the GV in the animal hemisphere, suggesting that actin plays a role in maintaining the polarised organisation of amphibian oocytes.

Keywords

Amphibian oogenesisCytoskeletonMicrofilamentsPhalloidin
Type
Article
Information
Zygote , Volume 2 , Issue 2 , May 1994 , pp. 111 - 124
Copyright
Copyright © Cambridge University Press 1994

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References

Blose, S.H., Meltzer, D.I. & Feramisco, J.R. (1984). 10-nm filaments are induced to collapse in living cells microinjected with monoclonal and polyclonal antibodies against tubulin. J. cell Biol. 98, 847–58.CrossRefGoogle ScholarPubMed
Brenner, S.L. & Korn, E.D. (1980). The effects of cytochalasins on actin polymerization and actin ATPase provide insights into the mechanism of polymerization. J. Biol. Chem. 255, 841–4.CrossRefGoogle ScholarPubMed
Campanella, C., Chaponnier, C., Quaglia, L., Gualtieri, R. & Gabbiani, G. (1990). Different cytoskeletal organization in maturation stages of Discoglossus pictus (Anura) oocytes: thickness and stability of actin microfilaments and tropomyosin localization. Mol. Reprod. Dev. 25, 130–9.CrossRefGoogle Scholar
Clark, T.G. & Merriam, R.W. (1977). Diffusible and bound actin in nuclei of Xenopus laevis oocytes. Cell 12, 883–91.CrossRefGoogle ScholarPubMed
Clark, T.G. & Merriam, R.W. (1978). Actin in Xenopus oocytes. I. Polymerization and gelation in vitro. J. Cell Biol. 77, 427–38.CrossRefGoogle Scholar
Clark, T.G. & Rosenbaum, J.L. (1979). An actin filament matrix in hand-isolated nuclei of X. laevis oocytes. Cell 18, 1101–8.CrossRefGoogle ScholarPubMed
Coggins, L.W. (1973). An ultrastructural and radioautograpic study of early oogenesis in the toad, Xenopus laevis. J. Cell Sci. 12, 71–93.CrossRefGoogle ScholarPubMed
Coleman, A., Morser, J., Lane, C., Besley, J., Wylie, C. & Valle, G. (1981). Fate of secretory proteins trapped in oocytes of Xenopus laevis by disruption of the cytoskeleton or by imbalanced subunit synthesis. J. Cell Biol. 91, 770.–8.CrossRefGoogle Scholar
Colombo, R., Benedusi, P. & Valle, G. (1981). Actin in Xenopus development: indirect immunofluorescence study of actin localization. Differentiation. 20, 45–51.CrossRefGoogle ScholarPubMed
Cooper, J.A. (1987). Effects of cytochalasin and phalloidin on actin. J. Cell Biol. 105, 1473–8.CrossRefGoogle ScholarPubMed
Dictus, W.J.A.G., van Zoelen, E.J.J., Tetteroo, P.A.T., Tertoolen, L.G.J., De Laat, S.W. & Bluemink, J.G. (1984). Lateral mobility of plasma membrane lipids in Xenopus eggs: regional differences related to animal/vegetal polarity become extreme upon fertilization. Dev. Biol. 101, 201–11.CrossRefGoogle ScholarPubMed
Dumont, J.N. (1972). Oogenesis in Xenopus laevis (Daudin) I. Stages of oocyte development in laboratory maintained animals. J. Morphol. 136, 153–80.CrossRefGoogle ScholarPubMed
Franke, W.W., Rathke, P.C., Seib, E., Trendelenburg, M.F., Osborn, M. & Weber, K. (1976). Distribution and mode of arrangement of microfilamentous structures and actin in the cortex of the amphibian oocyte. Cytobiologie 14, 111–30.Google ScholarPubMed
Franz, J.K., Gall, L., Williams, M.A., Picheral, B. & Franke, W.W. (1983). Intermediate-sized filaments in a germ cell: expression of cytokeratins in oocytes and eggs of the frog Xenopus. Proc. Natl. Acad. Sci. USA 80, 6254–8.CrossRefGoogle Scholar
Gard, D.L. (1991). Organization, nucleation, and acetylation of microtubules in Xenopus laevis oocytes: a study by confocal immunofluorescence microscopy. Dev. Biol. 143, 346–62.CrossRefGoogle ScholarPubMed
Gard, D.L. (1993). Ectopic spindle assembly during maturation of Xenopus oocytes: evidence for functional polarization of the oocyte cortex. Dev. Biol. (in press).CrossRefGoogle ScholarPubMed
Gerhart, J.C. (1980). Mechanisms regulating pattern formation in the amphibian egg and early embryo. In Biological Regulation and Development, ed. Goldberger, R.F., pp. 133–316. New York: Plenum Press.CrossRefGoogle Scholar
Gerhart, J., Black, S., Gimlich, R. & Scharf, S. (1983). Control of polarity in the amphibian egg. In: Time, Space, and Pattern in Embryonic Development. ed, Jeffery, W.R. & Raff, R.A., pp. 261–86. New York: Alan R. Liss.Google Scholar
Giloh, H. & Sedat, J.W., (1982). Fluorescence microscopy: reduced photobleaching of rhodamine and fluorescein protein conjugates by n-propylgallate. Science 217, 1252–5.CrossRefGoogle Scholar
Godsave, S.F., Wylie, C.C., Lane, E.B. & Anderton, B.H. (1984). Intermediate filaments in the Xenopus oocyte: the appearence and distribution of cytokeration-containing filaments. J. Embryol. Exp. Morphol. 83, 157–67.Google ScholarPubMed
Heasman, J., Quarmby, J. & Wylie, C.C. (1984).The mito-chondrial cloud of Xenopus oocytes: the source of germinal granule material. Dev. Biol. 105, 458–69.CrossRefGoogle Scholar
Huchon, D., Jessus, C., Thibier, C. & Ozon, R. (1988). Presence of microtubules in isolated cortices of prophase I and metaphase II oocytes in Xenopus laevis. Cell Tissue Res. 154, 415–20.Google Scholar
Karsenti, E., Gounon, P. & Bornens, M. (1978). Immunocytochemical study of lampbrush chromosomes: presence of tubulin and actin. Biol. Cell. 31 219–24.Google Scholar
Klymkowsky, M.W., Manyell, L.A. & Polson, A.G. (1987). Polar asymmetry in the organization of the cortical cyrokertin system of Xenopus laevis oocytes and embryos. Development 100 543–57.CrossRefGoogle ScholarPubMed
Klymkowsky, M.W., Maynell, L.A. & Nislow, C. (1991). Cytokeratin phosphorylation, cytokeratin filament severing, and the solubilization of the maternal mRNA Vg1 J. Cell Biol. 114, 787–97.CrossRefGoogle ScholarPubMed
Lessman, C.A. (1987). Germinal vesicle migration and dissolution in Rana pipiens oocytes: effects of steroids and microtubule poisons. Cell Differ. 20, 238–51.CrossRefGoogle ScholarPubMed
Merriam, R.W. & Clark, T.G. (1978). Actin in Xenopus oocytes II. Intracellular distribution and polymerizability. J. Cell Biol. 77, 439–47.CrossRefGoogle ScholarPubMed
Palecek, J., Habrova, V., Nedvidek, J. & Romanovsky, A. (1985). Dynamics of tubulin structures in Xenopus laevis oogenesis. J. Embryol. Exp. Morphol. 87, 75–86.Google ScholarPubMed
Scheer, U., Hinssen, H., Franke, W.W. & Jockusch, B.M. (1984). Microinjection of actin-binding proteins and actin antibodies demonstrates involvement of nuclear actin in transcription of lampbrush chromosomes. Cell 39, 111–22.CrossRefGoogle ScholarPubMed
Theurkauf, W.E., Smiley, S., Wong, M.L. & Alberts, B.M. (1992). Reorganization of the cytoskeleton during Drosophila oogenesis: implications for axis specification and intracellular transport. Development 115, 923–36.CrossRefGoogle Scholar
Torpey, N.P., Heasman, J. & Wylie, C.C. (1992) Distinct distribution of vimentin and cytokeratin in Xenopus oocytes and early embryos. J.Cell Sci. 101, 151–60.CrossRefGoogle ScholarPubMed
Warn, R.M., Gutzeit, H.O., Smith, L. & Warn, A. (1985). F-actin rings are associated with the ring canals of the Drosophila egg Chamber. Exp. Cell Res. 157, 355–63.CrossRefGoogle ScholarPubMed
Weeks, D.L. & Melton, D.A. (1987). A maternal mRNA localized to the vegetal hemisphere in Xenopus eggs codes for a growth factor related to TGF-a. Cell 51, 861–7.CrossRefGoogle Scholar
Yisraeli, J.K., Sokol, S. & Melton, D.A. (1990). A two-step model for the localization of maternal mRNA in Xenopus oocytes: involvemnt of microtubules and microfilaments in the translocation and anchoring of Vg1 mRNA. Development 108, 289–98.CrossRefGoogle Scholar