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. 1980 Jun 1;85(3):853–865. doi: 10.1083/jcb.85.3.853

Functional implications of cold-stable microtubules in kinetochore fibers of insect spermatocytes during anaphase

PMCID: PMC2111453  PMID: 7391142

Abstract

In normal anaphase of crane fly spermatocytes, the autosomes traverse most of the distance to the poles at a constant, temperature-dependent velocity. Concurrently, the birefringent kinetochore fibers shorten while retaining a constant birefringent retardation (BR) and width over most of the fiber length as the autosomes approach the centrosome region. To test the dynamic equilibrium model of chromosome poleward movement, we abruptly cooled or heated primary spermatocytes of the crane fly Nephrotoma ferruginea (and the grasshopper Trimerotropis maritima) during early anaphase. According to this model, abrupt cooling should induce transient depolymerization of the kinetochore fiber microtubules, thus producing a transient acceleration in the poleward movement of the autosomal chromosomes, provided the poles remain separated. Abrupt changes in temperature from 22 degrees C to as low as 4 degrees C or as high as 31 degrees C in fact produced immediate changes in chromosome velocity to new constant velocities. No transient changes in velocity were observed. At 4 degrees C (10 degrees C for grasshopper cells), chromosome movement ceased. Although no nonkinetochore fiber BR remained at these low temperatures, kinetochore fiber BR had changed very little. The cold stability of the kinetochore fiber microtubules, the constant velocity character of chromosome movement, and the observed Arrhenius relationship between temperature and chromosome velocity indicate that a rate-limiting catalyzed process is involved in the normal anaphase depolymerization of the spindle fiber microtubules. On the basis of our birefringence observations, the kinetochore fiber microtubules appear to exist in a steady-state balance between comparatively irreversible, and probably different, physiological pathways of polymerization and depolymerization.

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Selected References

These references are in PubMed. This may not be the complete list of references from this article.

  1. Begg D. A., Ellis G. W. Micromanipulation studies of chromosome movement. I. Chromosome-spindle attachment and the mechanical properties of chromosomal spindle fibers. J Cell Biol. 1979 Aug;82(2):528–541. doi: 10.1083/jcb.82.2.528. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Begg D. A., Ellis G. W. Micromanipulation studies of chromosome movement. II. Birefringent chromosomal fibers and the mechanical attachment of chromosomes to the spindle. J Cell Biol. 1979 Aug;82(2):542–554. doi: 10.1083/jcb.82.2.542. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Bibring T., Baxandall J., Denslow S., Walker B. Heterogeneity of the alpha subunit of tubulin and the variability of tubulin within a single organism. J Cell Biol. 1976 May;69(2):301–312. doi: 10.1083/jcb.69.2.301. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Brinkley B. R., Cartwright J., Jr Cold-labile and cold-stable microtubules in the mitotic spindle of mammalian cells. Ann N Y Acad Sci. 1975 Jun 30;253:428–439. doi: 10.1111/j.1749-6632.1975.tb19218.x. [DOI] [PubMed] [Google Scholar]
  5. Cande W. Z., Lazarides E., McIntosh J. R. A comparison of the distribution of actin and tubulin in the mammalian mitotic spindle as seen by indirect immunofluorescence. J Cell Biol. 1977 Mar;72(3):552–567. doi: 10.1083/jcb.72.3.552. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Cande W. Z., Snyder J., Smith D., Summers K., McIntosh J. R. A functional mitotic spindle prepared from mammalian cells in culture. Proc Natl Acad Sci U S A. 1974 Apr;71(4):1559–1563. doi: 10.1073/pnas.71.4.1559. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Connolly J. A., Kalnins V. I., Cleveland D. W., Kirschner M. W. Intracellular localization of the high molecular weight microtubule accessory protein by indirect immunofluorescence. J Cell Biol. 1978 Mar;76(3):781–786. doi: 10.1083/jcb.76.3.781. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. FORER A. LOCAL REDUCTION OF SPINDLE FIBER BIREFRINGENCE IN LIVING NEPHROTOMA SUTURALIS (LOEW) SPERMATOCYTES INDUCED BY ULTRAVIOLET MICROBEAM IRRADIATION. J Cell Biol. 1965 Apr;25:SUPPL–SUPPL117. doi: 10.1083/jcb.25.1.95. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Forer A. Characterization of the mitotic traction system, and evidence that birefringent spindle fibers neither produce nor transmit force for chromosome movement. Chromosoma. 1966;19(1):44–98. doi: 10.1007/BF00332793. [DOI] [PubMed] [Google Scholar]
  10. Fuseler J. W. Temperature dependence of anaphase chromosome velocity and microtubule depolymerization. J Cell Biol. 1975 Dec;67(3):789–800. doi: 10.1083/jcb.67.3.789. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Griffin J. D., Light S., Livingston D. M. Measurements of the molecular size of the simian virus 40 large T antigen. J Virol. 1978 Jul;27(1):218–226. doi: 10.1128/jvi.27.1.218-226.1978. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Harris P. The role of membranes in the ogranization of the mitotic apparatus. Exp Cell Res. 1975 Sep;94(2):409–425. doi: 10.1016/0014-4827(75)90507-8. [DOI] [PubMed] [Google Scholar]
  13. Herman I. M., Pollard T. D. Comparison of purified anti-actin and fluorescent-heavy meromyosin staining patterns in dividing cells. J Cell Biol. 1979 Mar;80(3):509–520. doi: 10.1083/jcb.80.3.509. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Inoué S., Fuseler J., Salmon E. D., Ellis G. W. Functional organization of mitotic microtubules. Physical chemistry of the in vivo equilibrium system. Biophys J. 1975 Jul;15(7):725–744. doi: 10.1016/S0006-3495(75)85850-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Inoué S., Ritter H., Jr Dynamics of mitotic spindle organization and function. Soc Gen Physiol Ser. 1975;30:3–30. [PubMed] [Google Scholar]
  16. Inoué S., Sato H. Cell motility by labile association of molecules. The nature of mitotic spindle fibers and their role in chromosome movement. J Gen Physiol. 1967 Jul;50(6 Suppl):259–292. [PMC free article] [PubMed] [Google Scholar]
  17. LaFountain J. R., Jr Analysis of birefringence and ultrastructure of spindles in primary spermatocytes of Nephrotoma suturalis during anaphase. J Ultrastruct Res. 1976 Mar;54(3):333–346. doi: 10.1016/s0022-5320(76)80020-2. [DOI] [PubMed] [Google Scholar]
  18. LaFountain J. R., Jr Birefringence and fine structure of spindles in spermatocytes of Nephrotoma suturalis at metaphase of first meiotic division. J Ultrastruct Res. 1974 Feb;46(2):268–278. doi: 10.1016/s0022-5320(74)80061-4. [DOI] [PubMed] [Google Scholar]
  19. LaFountain J. R., Jr Spindle shape changes as an indicator of force production in crane-fly spermatocytes. J Cell Sci. 1972 Jan;10(1):79–93. doi: 10.1242/jcs.10.1.79. [DOI] [PubMed] [Google Scholar]
  20. Margolis R. L., Wilson L., Keifer B. I. Mitotic mechanism based on intrinsic microtubule behaviour. Nature. 1978 Mar 30;272(5652):450–452. doi: 10.1038/272450a0. [DOI] [PubMed] [Google Scholar]
  21. Margolis R. L., Wilson L. Opposite end assembly and disassembly of microtubules at steady state in vitro. Cell. 1978 Jan;13(1):1–8. doi: 10.1016/0092-8674(78)90132-0. [DOI] [PubMed] [Google Scholar]
  22. McIntosh J. R., Cande W. Z., Snyder J. A. Structure and physiology of the mammalian mitotic spindle. Soc Gen Physiol Ser. 1975;30:31–76. [PubMed] [Google Scholar]
  23. Nicklas R. B. Mitosis. Adv Cell Biol. 1971;2:225–297. doi: 10.1007/978-1-4615-9588-5_5. [DOI] [PubMed] [Google Scholar]
  24. SWANN M. M., MITCHISON J. M. Refinements in polarized light microscopy. J Exp Biol. 1950 Sep;27(2):226–237. doi: 10.1242/jeb.27.2.226. [DOI] [PubMed] [Google Scholar]
  25. Salmon E. D., Ellis G. W. Compensator transducer increases ease, accuracy, and rapidity of measuring changes in specimen birefringence with polarization microscopy. J Microsc. 1976 Jan;106(1):63–69. doi: 10.1111/j.1365-2818.1976.tb02384.x. [DOI] [PubMed] [Google Scholar]
  26. Salmon E. D., Goode D., Maugel T. K., Bonar D. B. Pressure-induced depolymerization of spindle microtubules. III. Differential stability in HeLa cells. J Cell Biol. 1976 May;69(2):443–454. doi: 10.1083/jcb.69.2.443. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Salmon E. D. Pressure-induced depolymerization of spindle microtubules. I. Changes in birefringence and spindle length. J Cell Biol. 1975 Jun;65(3):603–614. doi: 10.1083/jcb.65.3.603. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Salmon E. D. Pressure-induced depolymerization of spindle microtubules. II. Thermodynamics of in vivo spindle assembly. J Cell Biol. 1975 Jul;66(1):114–127. doi: 10.1083/jcb.66.1.114. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Sato H., Ellis G. W., Inoué S. Microtubular origin of mitotic spindle form birefringence. Demonstration of the applicability of Wiener's equation. J Cell Biol. 1975 Dec;67(3):501–517. doi: 10.1083/jcb.67.3.501. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Sherline P., Schiavone K. High molecular weight MAPs are part of the mitotic spindle. J Cell Biol. 1978 Apr;77(1):R9–12. doi: 10.1083/jcb.77.1.r9. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Stephens R. E. A thermodynamic analysis of mitotic spindle equilibrium at active metaphase. J Cell Biol. 1973 Apr;57(1):133–147. doi: 10.1083/jcb.57.1.133. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Stephens R. E. Structural chemistry of the axoneme: evidence for chemically and functionally unique tubulin dimers in outer fibers. Soc Gen Physiol Ser. 1975;30:181–206. [PubMed] [Google Scholar]

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