, Volume 191, Issue 3–4, pp 178–190 | Cite as

Actin-microtubule interactions in the algaNitella: analysis of the mechanism by which microtubule depolymerization potentiates cytochalasin's effects on streaming

  • David A. Collings
  • Geoffrey O. Wasteneys
  • Richard E. Williamson


In the characean algaNitella, depolymerization of microtubules potentiates the inhibitory effects of cytochalasins on cytoplasmic streaming. Microtubule depolymerization lowers the cytochalasin B and D concentrations required to inhibit streaming, accelerates inhibition and delays streaming recovery. Because microtubule depolymerization does not significantly alter3H-cytochalasin B uptake and release, elevated intracellular cytochalasin concentrations are not the basis for potentiation. Instead, microtubule depolymerization causes actin to become more sensitive to cytochalasin. This increased sensitivity of actin is unlikely to be due to direct stabilization of actin by microtubules, however, because very few microtubules colocalize with the subcortical actin bundles that generate streaming. Furthermore, microtubule reassembly, but not recovery of former transverse alignment, is sufficient for restoring the normal cellular responses to cytochalasin D. We hypothesize that either tubulin or microtubule-associated proteins, released when microtubules depolymerize, interact with the actin cytoskeleton and sensitize it to cytochalasin.


Actin Cytoplasmic streaming Cytochalasin Microtubule depolymerization Nitella Oryzalin 



artificial pond water


cytoplasraic free calcium concentration


dimethyl sulfoxide






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  1. Asai DJ, Thompson WC, Wilson L, Dresden CF, Schulman H, Purich DL (1985) Microtubule-associated proteins (MAPs): a monoclonal antibody to MAPI decorates microtubules in vitro but stains stress fibers and not microtubules in vivo. Proc Natl Acad Sci USA 82: 1434–1438PubMedGoogle Scholar
  2. Baines AJ, Bennett V (1986) Synapsin I is a microtubule-bundling protein. Nature 319: 345–347Google Scholar
  3. Bech-Hansen NT, Till JE, Ling V (1976) Pleiotropic phenotype of colchicine-resistant CHO cells; cross-resistance and collateral sensitivity. J Cell Physiol 88: 23–32PubMedGoogle Scholar
  4. Bradley MO (1973) Microfilaments and cytoplasmic streaming: inhibition of streaming with cytochalasin. J Cell Sci 12: 327–343PubMedGoogle Scholar
  5. Chu B, Kerr GP, Carter JV (1993) Stabilizing microtubules with taxol increases microfilament stability during freezing in rye root tips. Plant Cell Environ 16: 883–889Google Scholar
  6. Collings DA (1994) The organisation, functions and interactions of the actin cytoskeleton in plants. PhD thesis, The Australian National University, Canberra, ACTGoogle Scholar
  7. —, Wasteneys GO, Miyazaki M, Williamson RE (1994) Elongation factor lα is a component of the subcortical actin bundles of characean algae. Cell Biol Int 18: 1019–1024PubMedGoogle Scholar
  8. — —, Williamson RE (1995) Cytochalasin rearranges cortical actin of the algaNitella into short, stable rods. Plant Cell Physiol 36: 765–772Google Scholar
  9. Cross D, Vial C, Maccioni RB (1993) A tau-like protein interacts with stress fibres and microtubules in human and rodent cultured cell lines. J Cell Sci 105: 51–60PubMedGoogle Scholar
  10. Danowski BA (1989) Fibroblast contractility and actin organization are stimulated by microtubule inhibitors. J Cell Sci 93: 255–266PubMedGoogle Scholar
  11. Dentler WL, Adams C (1992) Flagellar microtubule dynamics inChlamydomonas: cytochalasin D induces periods of microtubule shortening and elongation: and colchicine induces disassembly of the distal, but not proximal, half of the flagellum. J Cell Biol 117: 1289–1298PubMedGoogle Scholar
  12. Eleftheriou EP, Palevitz BA (1992) The effect of cytochalasin D on preprophase band organization in root tip cells ofAllium. J Cell Sci 103: 989–998Google Scholar
  13. Hertel C, Marmé D (1983) Herbicides and fungicides inhibit Ca2+ uptake by plant mitochondria: a possible mechanism of action. Pestle Biochem Physiol 19: 282–290Google Scholar
  14. Hope AB (1971) Ion transport and membranes. Butterworths, LondonGoogle Scholar
  15. Ishikawa M, Murofushi H, Sakai H (1983) Bundling of microtubules in vitro by fodrin. J Biochem 94: 1209–1217PubMedGoogle Scholar
  16. —, Kagami O, Hayashi C, Kohama K (1992a) Characterization of smooth muscle caldesmon as a microtubule-associated protein. Cell Motil Cytoskeleton 23: 244–251PubMedGoogle Scholar
  17. — — — — (1992b) The binding of nonmuscle caldesmon from brain to microtubules. FEBS Lett 299: 54–56PubMedGoogle Scholar
  18. Kadota A, Wada M (1992) The circular arrangement of cortical microtubules around the subapex of tip-growing fern protonemata is sensitive to cytochalasin B. Plant Cell Physiol 33: 99–102Google Scholar
  19. Kajstura J, Bereiter-Hahn J (1993) Disruption of microtubules induces formation of actin fibrils in density-inhibited 3T3 cells. Cell Biol Int 17: 1023–1031PubMedGoogle Scholar
  20. Keifer AQ, Callaham DA, Hepler PK (1992) Inhibitors of cell division and protoplastic streaming fail to cause a detectable effect on intracellular calcium levels in stamen-hair cells ofTradescantia virginiana L. Planta 186: 361–366Google Scholar
  21. Kilmartin JV, Wright B, Milstein C (1982) Rat monoclonal antitubulin antibodies derived by using a new nonsecreting rat cell line. J Cell Biol 93: 576–582Google Scholar
  22. Lessard JL (1989) Two monoclonal antibodies to actin: one muscle selective and one generally reactive. Cell Motil Cytoskeleton 10: 349–362Google Scholar
  23. MacRobbie EAC, Dainty J (1958) Ion transport inNitellopsis obtusa. J Gen Physiol 42: 335–353PubMedGoogle Scholar
  24. Marchesi VT, Ngo N (1993) In vitro assembly of multiprotein complexes containing alpha, beta and gamma tubulins, heat shock protein HSP70, and elongation factor la. Proc Natl Acad Sci USA 90: 3028–3032PubMedGoogle Scholar
  25. Menzel D, Schliwa M (1986) Motility in the siphonous green algaBryopsis. II. Chloroplast movement requires organized arrays of both microtubules and actin filaments. Eur J Cell Biol 40: 286–295Google Scholar
  26. Mitsui H, Yamaguchi-Shinozaki K, Shinozaki K, Nishikawa K, Takahashi H (1993) Identification of a gene family (kat) encoding kinesin-like proteins inArabidopsis thaliana and the characterization of secondary structure ofkatA. Mol Gen Genet 238: 362–368PubMedGoogle Scholar
  27. —, Nakatan K, Yamaguchi-Shinozaki K, Shinozaki K, Nishikawa K, Takahashi H (1994) Sequencing and characterization of the kinesin-related geneskatB andkatC ofArabidopsis thaliana. Plant Mol Biol 25: 865–876PubMedGoogle Scholar
  28. Okuhara K, Murofushi H, Sakai H (1989) Binding of kinesin to stress fibers in fibroblasts under condition of microtubule depolymerization. Cell Motil Cytoskeleton 12: 71–77PubMedGoogle Scholar
  29. Plieth C, Hansen U-P (1992) Light dependence of protoplasmic streaming inNitella flexilis L. as measured by means of laservelocimetry. Planta 188: 332–339Google Scholar
  30. Schellenbaum P, Vantard M, Lambert A (1992) Higher plant microtubule-associated proteins (MAPs): a survey. Biol Cell 76: 359–364Google Scholar
  31. Seagull RW (1990) The effects of microtubule and microfilament disrupting agents on cytoskeletal arrays and wall deposition in developing cotton fibres. Protoplasma 159: 44–59Google Scholar
  32. Suzuki N, Mihashi K (1991) Binding mode of cytochalasin B to F-actin is altered by lateral binding of regulatory proteins. J Biochem 109: 19–23PubMedGoogle Scholar
  33. Tanaka I, Wakabayashi T (1992) Organization of the actin and microtubule cytoskeleton preceding pollen germination. Planta 186: 473–482Google Scholar
  34. Tiezzi A, Moscatelli A, Bartalesi A, Cresti M (1992) An immunoreactive homolog of mammalian kinesin inNicotiana tabacum pollen tubes. Cell Motil Cytoskeleton 21: 132–137PubMedGoogle Scholar
  35. Vantard M, Schellenbaum P, Fellous A, Lambert A (1991) Characterization of maize microtubule-associated proteins, one of which is immunologically related to tau. Biochemistry 30: 9334–9340PubMedGoogle Scholar
  36. Verkhovsky AB, Surgucheva IG, Gelfland VI, Rosenblat VA (1981) G-actin-tubulin interaction. FEES Lett 135: 290–294Google Scholar
  37. Wasteneys GO, Williamson RE (1987) Microtubule orientation in developing internodal cells ofNitella: a quantitative analysis. Eur J Cell Biol 43: 14–22Google Scholar
  38. — — (1989) Reassembly of microtubules inNitella tasmanica: quantitative analysis of assembly and orientation. Eur J Cell Biol 50: 76–83Google Scholar
  39. — — (1991) Endoplasmic microtubules and nucleus-associated actin rings inNitella internodal cells. Protoplasma 162: 86–98Google Scholar
  40. —, Jablonsky PP, Williamson RE (1989) Assembly of purified brain tubulin at cortical and endoplasmic sites in perfused internodal cells of the algaNitella tasmanica. Cell Biol Int Rep 13: 513–528Google Scholar
  41. —, Gunning BES, Hepler PK (1993) Microinjection of fluorescent brain tubulin reveals dynamic properties of cortical microtubules in living plant cells. Cell Motil Cytoskeleton 24: 205–213Google Scholar
  42. —, Collings DA, Gunning BES, Hepler PK, Menzel D (1996) Actin in living and fixed characean internodal cells: identification of a cortical array of fine actin strands and chloroplast actin rings. Protoplasma 190: 25–38Google Scholar
  43. Williamson RE, Ashley CC (1982) Free Ca2+ and cytoplasmic streaming in the algaChara. Nature 296: 647–651PubMedGoogle Scholar

Copyright information

© Springer-Verlag 1996

Authors and Affiliations

  • David A. Collings
    • 1
  • Geoffrey O. Wasteneys
    • 1
  • Richard E. Williamson
    • 1
  1. 1.Plant Cell Biology Group, Research School of Biological SciencesThe Australian National UniversityCanberraAustralia

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