, Volume 82, Issue 5, pp 693–716 | Cite as

The attachment of kinetochores to the pro-metaphase spindle in PtK1 cells

Recovery from low temperature treatment
  • Conly L. Rieder
  • Gary G. Borisy


When late prophase PtK1 cells are chilled to 6 ° C the nuclear envelope (NE) breaks down as in normal cells but the spindle is inhibited from forming. When these cells are subsequently warmed to 18 ° C the spindle slowly forms and pro-metaphase congression ensues. Using this approach we have been able to experimentally eliminate the influence of asynchronous NE breakdown on the formation and development of the spindle, and also to slow down (and thus increase the temporal separation of) the subsequent events which occur during the initial stages of spindle formation. Correlative light and high voltage electron microscopic studies on these cells, fixed after various times of recovery, reveal the following results: 1) the centrosomes generate microtubules (MTs) well before MTs are seen to be associated with the kinetochores; 2) as in untreated PtK1 cells (Roos, 1973a, 1976) the order in which chromosomes attach to the forming spindle is influenced by their proximity to a centrosome-kinetochores closest to a centrosome appear stretched towards the centrosome at a time during recovery when other kinetochores, more distal to the centrosome appear unstretched and unoriented; 3) as in untreated cells (Heneen, 1970; Roos, 1976) the predominant behavior during recovery is for a chromosome to initially mono-orient and associate with the near centrosome and only later to develop a bipolar association; and 4) MTs associated with early pro-metaphase kinetochores are almost always oriented towards a centrosome. — From our results we conclude that the proximity effect and the tendency of pro-metaphase chromosomes in PtK1 to initially mono-orient and associate with the near centrosome cannot be ascribed, as suggested by Roos (1976), to influences arising during the asynchronous breakdown of the NE. Rather, our data clearly demonstrate that a kinetochore-centrosome interaction occurs during spindle formation which cannot be attributed to transient influences. The proximity effect and the predominant tendency of PtK1 pro-metaphase chromosomes to mono-orient to the near pole are taken to signify the existance of a centrosomal influence on the attachment and orientation of chromosomes. Two possible mechanisms for this influence, both involving a structural interaction between the centrosome and the kinetochore, are outlined.


Normal Cell Developmental Biology Nuclear Envelope Structural Interaction Subsequent Event 
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  1. Aubin, J.E., Osborn, M., Weber, K.: Variations in the distribution and migration of centriolar duplexes in mitotic PtK2 cells studied by immunofluorescence microscopy. J. Cell Sci. 43, 177–194 (1980)Google Scholar
  2. Bajer, A.S., Molè-Bajer, J.: Spindle dynamics and chromosome movement. Int. Rev. Cytol., Suppl. 2 (1972)Google Scholar
  3. Barber, H., Callan, H.: The effects of cold and colchicine on mitosis in the newt. Proc. roy. Soc., Lond., B. 131, 258–274 (1942)Google Scholar
  4. Behnke, O., Rostgaard, J.: Your “third hand” in mounting serial sections on grids for electron microscopy. Stain Technol. 39, 205–208 (1964)Google Scholar
  5. Brinkley, B.R., Stubblefield, E., Hsu, T.C.: The effects of colcemid inhibition and reversal on the fine structure of the mitotic apparatus of Chinese hamster cells in vitro. J. Ultrastr. Res. 19, 1–18 (1967)Google Scholar
  6. DeBrabander, M., Geuens, G., DeMey, J., Joniau, M.: Light microscopic and ultrastructural distribution of immunoreactive tubulin in mitotic mammalian cells. Biol. cellulaire 34, 213–226 (1979a)Google Scholar
  7. DeBrabander, M., DeMey, J., Geuens, G., Joniau, M.: The distribution, function and regulation of microtubules in cultured cells studied with a new “complete” immunocytochemical approach. 37th Ann. Proc. Electron Microsc. Soc. Amer. (G.W. Bailey, ed), pp. 10–13 (1979b)Google Scholar
  8. DeBrabander, M., Geuens, G., DeMey, J., Joniau, M.: The organized assembly and function of the microtubule system throughout the cell cycle. In: Cell movement and neoplasia (M. DeBrabander, M. Mareel and L. DeRidder, eds.), pp 19–40. New York: Pergamon Press (1980a)Google Scholar
  9. DeBrabander, M., Geuens, G., Nuydens, R., Willebrords, R., DeMey, J.: The microtubule nucleating and organizing activity of kinetochores and centrosomes in living PtK2 cells. In: Microtubules and microtubule inhibitors (M. DeBrabander and J. DeMey, eds.), pp. 255–268. New York: North Holland 1980bGoogle Scholar
  10. DeRobertis, E.M., Longthorne, R.R., Gurdon, J.B.: Intracellular migration of nuclear proteins in Xenopus oocytes. Nature (Lond) 272, 254–256 (1978)Google Scholar
  11. Goode, D.: Kinetics of microtubule formation after cold disaggregation of the mitotic apparatus. J. molec. Biol. 80, 531–538 (1973)Google Scholar
  12. Heneen, W.K.: In situ analysis of normal and abnormal patterns of the mitotic apparatus in cultured rat-kangaroo cells. Chromosoma (Berl.) 29, 88–117 (1970)Google Scholar
  13. Heneen, W.K.: Ultrastructure of the prophase kinetochore in cultured cells of rat kangaroo (Potorous tridactylis). Hereditas (Lond) 79, 209–220 (1975)Google Scholar
  14. Inoué, S.: Effect of temperature on the birefringence of the mitotic spindle. Biol. Bull 103, 316a (1952)Google Scholar
  15. Inoué, S.: Organization and function of the mitotic spindle. In: Primitivve motile systems in cell biology (R.D. Allen and N. Kamiya, eds.). New York. Academic Press 1964Google Scholar
  16. Izutsu, J., Sato, H., Nakabayashi, H., Aoki, N.: The behavior of spindle fibers and movement of chromosomes in dividing grasshopper spermatocytes. Cell Structure and Function 2, 119–133 (1977)Google Scholar
  17. Jensen, C., Bajer, A.S.: Effects of dehydration on the microtubules of the mitotic spindle. Studies in vitro and with the electron microscope. J. Ultrastr. Res. 26, 367–386 (1969)Google Scholar
  18. Jensen, C., Jensen, L., Rieder, C.L.: The occurrence and the structure of primary cilia in a subline of Potorous tridactylus (PtK1). Exp. Cell Res. 123, 444–449 (1979)Google Scholar
  19. Jokelainen, P.T.: The differentiation of sister kinetochores during metakinesis. J. Cell Biol 27 (2 pt 2), 48a (1965)Google Scholar
  20. Jokelainen, P.T.: The ultrastructure and spatial organization of the metaphase kinetochore in mitotic rat cells. J. Ultrastr. Res. 19, 19–44 (1967)Google Scholar
  21. Lambert, A.M.: The role of chromosomes in anaphase trigger and nuclear envelope activity in spindle formation. Chromosoma (Berl.) 76, 295–308 (1980)Google Scholar
  22. Lambert, A.M., Bajer, A.S.: Microtubule distribution and reversible arrest of chromosome movements induced by low temperature. Cytobiologie 15, 1–23 (1977)Google Scholar
  23. Leibovitz, A.: The growth and maintenance of tissue-cell cultures in free gas exchange with the atmosphere. Amer. J. Hygene 78, 173–180 (1963)Google Scholar
  24. Mazia, D.: Mitosis and the physiology of cell division. In: The cell (J. Brachet and A.E. Mirsky, eds), pp 77–412. New York: Academic Press 1961Google Scholar
  25. McGill, M., Brinkley, B.R.: Human chromosomes and centrioles as nucleating sites for the in vitro assembly of microtubules from bovine brain tubulin. J. Cell Biol. 67, 189–199 (1975)Google Scholar
  26. McIntosh, J.R., Cande, W.Z., Snyder, J.D.: Structure and physiology of the mammalian mitotic spindle. In: Molecules and cell movement (S. Inoué and R.E. Stephens, eds.), pp 31–74. New York: Raven Press 1975Google Scholar
  27. Molè-Bajer, J.: The role of centrioles in the development of the astral spindle (newt). Cytobios 13, 17–40 (1975)Google Scholar
  28. Molè-Bajer, J., Bajer, A.S.: Studies of selected endosperm cells with the light and electron microscope. The technique. Cellule 67, 257–265 (1968)Google Scholar
  29. Molè-Bajer, J., Bajer, A., Owczarzak, A.: Chromosome movements in pro-metaphase and aster transport in the newt. Cytobios 13, 45–65 (1975)Google Scholar
  30. Moore, M.J.: Removal of glass coverslips from cultures flat embedded in epoxy resins using hydroflouric acid. J. Microsc. 104, 205–207 (1975)Google Scholar
  31. Nicklas, R.B., Brinkley, B.R., Pepper, D.A., Kubai, D., Rickards, G.K.: Electron microscopy of spermatocytes previously studied in life: Methods and some observations on micromanipulated chromosomes. J. Cell Science 35, 87–104 (1979)Google Scholar
  32. Paweletz, N.: Elektronenmikroskopische Untersuchungen am frühen Stadien der Mitose bei HeLa Zellen. Cytobiologie 9, 368–390 (1974)Google Scholar
  33. Rattner, J.B., Branch, A., Hamakalo, B.A.: Electron microscopy of whole mount metaphase chromosomes. Chromosoma (Berl.) 52, 329–338 (1975)Google Scholar
  34. Rattner, J.B., Berns, M.W.: Distribution of microtubules during centriole separation in rat kangaroo (Potorous) cells. Cytobios 15, 37–43 (1976a)Google Scholar
  35. Rattner J.B., Berns, M.W.: Centriole behavior in early mitosis of rat kangaroo cells (PtK2). Chromosoma (Berl.) 54, 387–395 (1976b)Google Scholar
  36. Rieder, C.L.: Localization of ribonucleoprotein in the trilaminar kinetochore of PtK1. J. Ultrastr. Res. 66, 109–119 (1979)Google Scholar
  37. Rieder, C.L.: The structure of the mammalian kinetochore and of the centrohelidian centroplast as revealed by Bernhard's staining of thick sections. In: Microtubules and microtubule inhibitors (M. DeBrabander and J. DeMey, eds.), pp. 311–324. New York: North Holland 1980Google Scholar
  38. Rieder, C.L., Bajer, A.S.: Effect of elevated temperatures on spindle microtubules and chromosome movements in cultured newt lung cells. Cytobios 18, 201–234 (1977)Google Scholar
  39. Ris, H.: The anaphase movement of chromosomes in spermatocytes of the grashopper. Biol. Bull. 96, 90–106 (1949)Google Scholar
  40. Roos, U-P.: Light and electron microscopy of rat kangaroo cells in mitosis. I. Formation and breakdown of the mitotic apparatus. Chromosoma (Berl.) 40, 43–82 (1973a)Google Scholar
  41. Roos, U-P.: Light and electron microscopy of rat kangaroo cells in mitosis. II. Kinetochore structure and function. Chromosoma (Berl.) 41, 195–220 (1973b)Google Scholar
  42. Roos, U-P.: Light and electron microscopy of rat kangaroo cells in mitosis. III. Patterns of chromosome behavior during prometaphase. Chromosoma (Berl.) 54, 363–385 (1976)Google Scholar
  43. Roos, U-P.: The fibrillar organization of the kinetochore and the kinetochore region of mammalian chromosomes. Cytobiologie 16, 82–90 (1977)Google Scholar
  44. Roth, L.E.: Electron microscopy of mitosis in amebae. III. Cold and urea treatments: a basis for tests of direct effects of mitotic inhibitors on microtubule formation. J. Cell Biol. 34, 47–59 (1967)Google Scholar
  45. Stephens, R.E.: Studies on the development of the sea urchin Strongylocentrotus droebachiensis. II. Regulation of mitotic spindle equilibrium by environmental temperature. Biol. Bull 142, 145–159 (1972)Google Scholar
  46. Tippit, D.H., Pickett-Heaps, J.D., Leslie, R.: Cell division in two large pennate diatoms Hantzschia and Nitzschia. III. A new proposal for kinetochore function during prometaphase. J. Cell Biol. 86, 402–416 (1980)Google Scholar
  47. Wilson, E.B.: The Cell in Development and Heredity. 3rd edit. New York: MacMillian 1925Google Scholar
  48. Witt, P.L., Ris, H., Borisy, G.G.: Origin of kinetochore microtubules in Chinese hamster ovary cells. Chromosoma (Berl.) 81, 483–505 (1980)Google Scholar

Copyright information

© Springer-Verlag 1981

Authors and Affiliations

  • Conly L. Rieder
    • 1
  • Gary G. Borisy
    • 1
  1. 1.Laboratory of Molecular BiologyUniversity of WisconsinMadisonUSA

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