This chapter reviews various aspects of changes in cytoskeletal organization which occur upon activating a highly motile cell phenotype. The first of these relates to the rapid formation of F-actin-rich ruffles on the apical cell surfaces following the addition of one of several motility factors. In a number of aspects these ruffles resemble leading edge lamellipodia. The ruffles form upon ligand binding and, at least for one cytokine, the receptors become associated with the ruffles. For several cytokines the ruffles are circular and in all cases they are associated with much increased pinocytosis. The possible significance is considered of these very early markers of a motile cell phenotype. A second topic covered is the role of microtubules (MTs) in maintaining cell polarity in some cell types but not others. The micro-injection of biotin-tubulin into cells has provided a valuable marker of MT turnover. Using this method it has been found that the microtubule network in secondary chick heart fibroblasts (2 degrees CHFs), where MTs are required to maintain a polarized motility, does not turn over significantly more slowly than in 1 degree CHFs which do not require an intact MT network for locomotion. In both cases the MT network turns over very quickly and no sub-population of longer-lived MTs has been found. In contrast, in motile epithelial (PtK2) cells, a sub-population of longer-lived microtubules has been identified and these appear to maintain the long cell processes.
ABSTRACT Changes in the distribution of myosin during the formation of the cellular blastoderm of Drosophila melanogaster were followed by staining sections of embryos with antibodies to myosin. These were visualized with indirect immunofluorescence. Prior to the start of cell membrane extension myosin is distributed between the nuclear caps as a thin sub-plasmalemma layer. There is also myosin present beneath the surface of the caps. When plasmalemma growth occurs, myosin is associated with the furrow canals, the tips of the advancing membranes. The fluorescence is distributed in an approximately hexagonal pattern around the growth points of each cell. The hexagons are joined up forming a network. It is suggested that this myosin is associated with bundles of microfilaments, orientated parallel to the surface, to form many interlocking contractile rings. The simultaneous contraction of these rings causes the cleavage of the blastoderm. During the first phase of membrane growth, myosin is also associated with the apical surfaces of the forming cells. At this stage these surfaces are rich in microvilli. However, by the time the furrow canals have reached the bases of the cells much of this myosin has disappeared. At about this time the apical surface becomes taut with a loss of the microvilli.
The F-actin distribution was studied during pole cell formation in Drosophila embryos using the phalloidin derivative rhodaminyl-lysine-phallotoxin. Nuclei were also stained with 4'-6 diamidine-2-phenylindole dihydrochloride to correlate the pattern seen with the nuclear cycle. The precursors of the pole cells, the polar surface caps, were found to have an F-actin-rich cortex distinct from that of the rest of the embryo surface and an interior cytoplasm that was less intensely stained but brighter than the cytoplasm deeper in the embryo. They were found to divide once without forming true cells and then a second time when cells formed as a result of a meridional and a basal cleavage. Three distinct distributions of the cortical F-actin have been identified during these cleavages. It is concluded that the first division, which cleaves the polar caps but does not separate them from the embryo, involves very different processes from those that lead to the formation of the pole cells. A contractile-ring type of F-actin organization may not be present during the first cleavage but is suggested to occur during the second.
By microinjecting rhodamine‐conjugated porcine tubulin into pea epidermis we recently showed how cortical microtubules reorientate from transverse to longitudinal in living cells (Yuan et al ., 1994, Proc. Nat. Acad. Sci, USA 91, 6050–6053). In the present paper we compare this reorientation with the contrary longitudinal to transverse realignment induced by adding gibberellic acid to pre‐injected cells on the microscope slide. Both kinds of reorientation are initiated by the appearance of ‘discordant’ microtubules which do not share the existing alignment but anticipate the new direction. These increase in number as the existing microtubules depolymerize, one alignment apparently replacing the other in a continuous process. By rotating stacks of confocal sections by computer methods we have previously shown that microtubules at the outer tangential cell wall do not necessarily have the same orientation as microtubules at the adjoining anticlinal walls of the same cell (Yuan et al ., 1995, Plant J. 7, 17–23). This suggests that microtubule reorientations in these epidermal cells occur mainly (or, at least, first) at the outer wall, indicating that the array may not reorientate as a whole. Collectively, these data emphasize the discontinuous nature of the realignment process, the importance of new microtubule polymerization, and the special property of the outer epidermal surface as a sensitive domain.
CO25 cells, a mouse myoblast line, contain multiple centrioles and primary cilia. A most unusual feature has been the finding of large numbers of separate structures in single cells-up to a maximum of nine centrioles, six primary cilia, and 12 of both organelles together. Aberrant multipolar spindles were occasionally seen containing variable numbers of centrioles. This strongly suggests that cells containing supernumerary centrioles and cilia are lost during mitosis, and that additional centriolar structures are generated during each interphase. No change in centriole or primary cilium frequency was detected after inducing the differentiation of myoblasts into myotubes. However, a significant migration of these structures occurred from a perinuclear to a supranuclear position prior to and during the phase of myoblast elongation. This shift was not maintained during cell fusion, when a net migration back to the periphery was observed, suggesting that it may have some function in relation to cell elongation and the change in the pattern of microtubule distribution which occurs as part of the process.
ABSTRACT The changes in F-actin organization during the cellularization of the Drosophila embryo have been studied with a confocal laser scanning microscope using fluorescein-phalloidin as a specific stain. Particular study has been made of the changes in the organization of the F-actin network associated with the leading edges of the growing membranes. The role of this actin network in the cellularization process is considered. Other actin-containing structures have also been examined, including the cortical actin layer and a conspicuous region of F-actin aggregates, present beneath the level of the forming cell membranes.
Rhodamine-labeled monoclonal antibodies, which react with tyrosinated alpha-tubulin (clone YL 1/2; Kilmartin, J. V., B. Wright, and C. Milstein, 1982, J. Cell Biol., 93:576-582) and label microtubules in vivo (Wehland, J., M. C. Willingham, and I. Sandoval, 1983, J. Cell Biol., 97:1467-1475) were microinjected into syncytial stage Drosophila embryos. At 1 mg/ml antibody concentration, the microtubule arrays of the surface caps became labeled by YL 1/2 but normal development was found to continue. The results are compared with the data from fixed material particularly with regard to interphase microtubules, centrosome separation, and spindle and midbody formation. At 5 mg/ml antibody concentration the microtubules took up larger quantities of antibodies and clumped around the nuclei. Nuclei with clumped microtubules lost their position in the surface layer and moved into the interior. As a result, the F-actin cap meshwork associated with such nuclei either failed to form or subsided. It is concluded that microtubule activity is required to maintain the nuclei in the surface layer and organize the F-actin meshwork of the caps.
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