Cells of many kinds adhere firmly to glass or plastic surfaces which have been pretreated with polylysine. The attachment takes place as soon as the cells make contact with the surfaces, and the flattening of the cells against the surfaces is quite rapid. Cells which do not normally adhere to solid surfaces, such as sea urchin eggs, attach as well as cells which normally do so, such as amebas or mammalian cells in culture. The adhesion is interpreted simply as the interaction between the polyanionic cell surfaces and the polycationic layer of adsorbed polylysine. The attachment of cells to the polylysine-treated surfaces can be exploited for a variety of experimental manipulations. In the preparation of samples for scanning or transmission electron microscopy, the living material may first be attached to a polylysine-coated plate or grid, subjected to some experimental treatment (fertilization of an egg, for example), then transferred rapidly to fixative and further passed through processing for observation; each step involves only the transfer of the plate or grid from one container to the next. The cells are not detached. The adhesion of the cell may be so firm that the body of the cell may be sheared away, leaving attached a patch of cell surface, face up, for observation of its inner aspect. For example, one may observe secretory vesicles on the inner face of the surface (3) or may study the association of filaments with the inner surface (Fig. 1). Subcellular structures may attach to the polylysine-coated surfaces. So far, we have found this to be the case for nuclei isolated from sea urchin embryos and for the microtubules of flagella, which are well displayed after the membrane has been disrupted by Triton X-100 (Fig. 2).

Various techniques have heen adapted from protein chemistry to the cytological demonstration of proteins (e.g., ninhydrin reaction, Mazia and Jaeger, 1939; Millon reaction, Pollister and Ris, 1947; Sakaguchi reaction, Thomas, 1946;Serra, 1946) but none has found wide use for both the resolution of morphological detail in terms of protein distribution and the measurement of relative protein concentra-

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The forms and locations of centrosomes in mouse oocytes and in sea urchin eggs were followed through the whole course of fertilization and first cleavage by immunofluorescence microscopy. Centrosomes were identified with an autoimmune antiserum to centrosomal material. Staining of the same preparations with tubulin antibody and with the DNA dye Hoechst 33258 allowed the correlation of the forms of the centrosomes with the microtubule structures that they generate and with the stages of meiosis, syngamy, and mitosis. The results with sea urchin eggs conform to Boveri's view on the paternal origin of the functional centrosomes. Centrosomes are seen in spermatozoa and enter the egg at fertilization. Initially, the centrosomes are compact, but as the eggs enter the mitotic cycle the forms of the centrosomes go through a cycle in which they spread during interphase, apparently divide, and condense into two compact poles by metaphase. In anaphase, they spread to form flat poles. In telophase and during reconstitution of the daughter nuclei, the centrosomal material is dposed as hemispherical caps around the poleward surfaces of the nuclei. Mouse sperm lack centrosomal antigen. In the unfertilized mouse oocyte, the meiotic spindle poles are displayed as broad-beaded centrosomes. In addition, centrosomal material is detected in the cytoplasm as particles, about 16 in number, which are foci of small aster-like arrays of microtubules. The length and number of astral microtubules correlate with the size of the centrosomal foci. After sperm incorporation, as the pronuclei develop and more cytoplasmic microtubules assemble, a few of the foci associate with the peripheries of the nuclei. The number of foci multiplies during the first cell cycle. At the end of interphase, all of the centrosomal foci have concentrated on the nuclear peripheries and the cytoplasmic microtubules have disappeared. At prophase, the centrosomes are seen as two irregular clusters, marking the poles which, at metaphase and anaphase, appear as rough bands with foci, and the spindle is typically barrel-shaped. At telophase, the centrosomes are seen as arcs that lie on the nuclear peripheries after cleavage. The ordering of microtubules in all the stages reflects the shapes of the centrosomes. The findings on the sea urchin confirm the classical theory of the paternal origin of centrosomes and contrast with observations tracing the mitotic poles of the mouse egg to maternal centrosomal material. This evidence strengthens the conclusion that mouse centrosomes derive from the oocyte. Mouse and sea urchin fertilization was as described (6). Sea urchin eggs were extracted in a microtubule-stabilization buffer (7), and mouse egg cytoskeletons were stabilized with a similar mixture (4). The cells were affixed to polylysinecoated coverslips (8). Sea urchin eggs were fixed in methanol at -10TC and mouse eggs were fixed in 10 mM ethylene glycol bis(succinimidyl)succinate (9). Autoimmune centrosomal antiserum 5051 was derived from a patient with scleroder...

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In this study, the reproduction of the mitotic centers in the eggs of a sea urchin, Strongylocentrotus purpuratus and a sand dollar Dendraster excentricus has been studied by means of experimental designs that do not depend on the actual visualization of centrioles. The centers are defined in operational terms as potential poles. Blockage of mitosis by mercaptoethanol, it was found, inhibits the duplication of the centers, but does not inhibit the splitting and separation of centers that have already duplicated and thus potential poles could be realized as actual poles in multipolar divisions. At all times, the center is at least a duplex structure; that is, it contains two potential poles. The actual duplication process is the earliest event in a given mitotic cycle, taking place at very early interphase or in late telophase of the previous division. The splitting of the centers following duplication is a distinct process, dissociable from the duplication as such. Duplication and splitting normally occur at about the same time in the mitotic cycle, with a precession of the former. That is, as the two members of a pair of "old" centers split, each one gives rise to a new one, which remains associated with it until the next phase of splitting and duplication occurs. The results are consistent with what is termed a "generative" model of the self-reproduction of an intracellular body. According to this, the body does not immediately produce a full-fledged copy of itself, with simultaneous fission, but the primary duplication event involves only a part of the parent structure. This gives rise to a "germ" or "seed" which then grows to be equivalent to the parent body, and finally splits from it.

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Amoebae of Dictyostelium discoideum were attached to a surface coated with polylysine, and the upper portion of the cells was sheared off with a stream of buffer. Scanning and transmission electron microscopy showed that the cytoplasmic surface of the exposed membrane was covered with fibers consisting of actin-containing filaments. The actin was identified by its solubility properties and its ability to interact with muscle myosin.In preceding work (1-3) proteins from amoebae of Dictyostelium discoideum that closely resemble the muscle proteins actin and myosin were l)urified and characterized, and it was shown biochemically that some of the Dictyostelium actin is associated with the cell membrane (2). In the present study, membrane-associated actin filaments were visualized by electron microscopy, utilizing a new technique in which cells were attached to surfaces coated with polylysine, and then disrupted to expose the cytoplasmic face of the cell membrane. The use of polylysine as an adhesive for electron microscopy was recently described by Mazia and coworkers (4); protamille sulfate was similarly employed by Vacquier (5) in his examination of the inner membrane surfaces in sea urchin eggs. MATERIALS AND METHODSBuffers and Reagents. The phosphate buffer (Buffer P) used to resuspend D. discoideurn cells contained 20 mM potassium phosphate, pH 6.4, 20 mM KCl, 5 mM MgCl2, and 0.5 mg/ml of streptomycin sulfate (6). The buffer used for cell disrultion (Buffer D) contained 10 mM Tris-HCl, pH 7.5, 0.1 M KCl, and 1 mM MgCl2. Poly(Llysine) (molecular weight 80,000-100,000) was obtained from New England Nuclear, Boston.Growth Scanning Electron Microscopy. Clean glass slides were broken into 1 cm2 plates, and were coated with an aqueous solution of polylysine (1 mg/ml). They were rinsed with distilled H20 and then with Buffer P, and the sample was applied. Samples were fixed by immersing the plates in a solution containing 2.5% glutaraldehyde and 1% acrolein made up in Buffer 1), and were post-fixedl in osmium tetroxide. They were then prel)ared for microscopy by freon 13 critical point drying andlplatinum shadowing, and were examined in a Coates & Welter field emission scanning electron microscope.Transmission Electron Microscopy. Grids were coated with a Parlodion film over which a layer of carbon was evaporated. Polylysine (1 mg/ml in H20) was applied to this surface, which was then rinsed with distilled H20 and Buffer P. The cell suspension was applied, and the cells were disrupted as described above, then negatively stained with 1% uranyl acetate. In some cases, the disrupted cells were treated with the myosin subfragment 1 (S5) according to the procedures described by Moore et al. (7) Other Materials. Actin was purified from amoebae of D. discoideum by column chromatography, as previously described (2). The myosin subfragment SI was prepared from rabbit muscle according to the procedure of Cooke (8). RESULTSWhen a Dictyostelium discoideun amoeba adhered to a polylvsine-coated surface, its attachiment was...