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Protoplasma

, Volume 212, Issue 3–4, pp 174–185 | Cite as

A plasmolytic cycle: The fate of cytoskeletal elements

  • I. Lang-Pauluzzi
  • B. E. S. Gunning
Article

Summary

In most plant cells, transfer to hypertonic solutions causes osmotic loss of water from the vacuole and detachment of the living protoplast from the cell wall (plasmolysis). This process is reversible and after removal of the plasmolytic solution, protoplasts can re-expand to their original size (deplasmolysis). We have investigated this phenomenon with special reference to cytoskeletal elements in onion inner epidermal cells. The main processes of plasmolysis seem to be membrane dependent because destabilization of cytoskeletal elements had only minor effects on plasmolysis speed and form. In most cells, the array of cortical microtubules is similar to that found in nonplasmolyzed states except that longitudinal patterns seen in some control cells were never observed in plasmolyzed protoplasts of onion inner epidermis. As soon as deplasmolysis starts, cortical microtubules become disrupted and only slowly regenerate to form an oblique array, similar to most nontreated cells. Actin microfilaments responded rapidly to the plasmolysis-induced deformation of the protoplast and adapted to its new form without marked changes in organization and structure. Both actin microfilaments and microtubules can be present in Hechtian strands, which, in plasmolyzed cells, connect the cell wall to the protoplast. Anticytoskeletal drugs did not affect the formation of Hechtian strands.

Keywords

Cytoskeleton Hechtian strands Latrunculin B Onion inner epidermis Oryzalin Plasmolysis 

Abbreviations

DIC

differential interference contrast

DiOC6(3)

3,3-dihexyloxacarbocyanine iodide

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References

  1. Bachewich CL, Heath IB (1997) Differential cytoplasm-plasma membrane-cell wall adhesion patterns and their relationships to hyphal tip growth and organeile motility. Protoplasma 200: 71–86Google Scholar
  2. Baluška F, Vitha S, Barlow PW, Volkmann D (1997) Rearrangements of F-actin arrays in growing cells of intact maize root apex tissues: a major developmental switch occurs in the postmitotic transition region. Eur J Biol Chem 72: 113–121Google Scholar
  3. Burridge K, Chrzanowska-Wodnicka M, Zhong C (1997) Focal adhesion assembly. Trends Cell Biol 7: 342–347Google Scholar
  4. Canut H, Carrasco A, Galaud J-P, Cassan C, Bouyssou H, Vita N, Ferrara P, Pont-Lezica R (1998) High affinity RGD-binding sites at the plasma membrane ofArabidopsis thaliana links the cell wall. Plant J 16: 63–71PubMedGoogle Scholar
  5. Cleary A (1995) F-actin redistributions at the division site in livingTradescantia stomatal complexes as revealed by microinjection of rhodamine-phalloidin. Protoplasma 185: 152–165Google Scholar
  6. —, Mathesius U (1996) Rearrangements of F-actin during stomatogenesis visualized by confocal microscopy in fixed and permabilizedTradescantia leaf epidermis. Bot Acta 109: 15–24Google Scholar
  7. Epstein DL, Rowlette LL, Roberts BC (1999) Actomyosin drug effects and aqueous outflow function. Invest Ophthalmol Vis Sci 40: 74–81PubMedGoogle Scholar
  8. Erwee MG, Goodwin PB (1984) Characterisation of theEgeria densa leaf symplast: response to plasmolysis, deplasmolysis and to aromatic amino acids. Protoplasma 122: 162–168Google Scholar
  9. Eschrich W (1957) Kallosebildung in plasmolysiertenAllium cepa-Epidermen. Planta 48: 578–586Google Scholar
  10. Galway ME, Hardham AR (1986) Microtubule reorganisation, cell wall synthesis and establishment of the axis of elongation in regenerating protoplasts of the algaMougeotia. Protoplasma 135: 130–143Google Scholar
  11. — — (1989) Oryzalin-indueed microtubule disassembly and recovery in regenerating protoplasts of the algaMougeotia. J Plant Physiol 135: 337–345Google Scholar
  12. Gordon-Kamm WJ, Steponkus PL (1984) The behavior of the plasma membrane following osmotic contraction of isolated protoplasts: implications in freezing injury. Protoplasma 123: 83–94Google Scholar
  13. Gunning BES, Steer MW (1996) Bildatlas zur Biologie der Pflanzenzelle: Struktur und Funktion, 4th edn. Gustav Fischer, StuttgartGoogle Scholar
  14. Gupta GD, Heath IB (1997) Actin disruption by latrunculin B causes turgor-related changes in tip growth ofSaprolegnia ferax hyphae. Fung Genet Biol 21: 64–75Google Scholar
  15. Hecht K (1912) Studien über den Vorgang der Plasmolyse. Beitr Biol Pflanzen 11: 133–189Google Scholar
  16. Hepler PK, Palevitz BA, Lancelle SA, McCauley MM, Lichtscheidl I (1990) Cortical endoplasmic reticulum in plants. J Cell Sci 96: 355–373Google Scholar
  17. Hush JM, Overall RL (1996) Cortical microtubule reorientation in higher plants: dynamics and regulation. J Microsc 181: 129–139Google Scholar
  18. Iwata K (1995) Regulation of the orientation of cortical microtubules inSpirogyra cells. J Plant Res 108: 531–534Google Scholar
  19. Kjellbom P, Snogerup L, Stöhr C, Reuzeau C, McCabe PF, Pennell RI (1997) Oxidative cross-linking of plasma membrane arabinogalactan proteins. Plant J 12: 1189–1196PubMedGoogle Scholar
  20. Kropf DL, Williamson RE, Wasteneys GO (1997) Microtubule orientation and dynamics in elongating characean internodal cells following cytosolic acidification, induction of pH bands, or premature growth arrest. Protoplasma 197: 188–198Google Scholar
  21. Lichtscheidl IK (1995) Organization and dynamics of endoplasmic reticulum:Allium cepa L. inner epidermis. Wiss Film 47: 95–109Google Scholar
  22. —, Url WG (1987) Investigation of the protoplasm ofAllium cepa inner epidermal cells using ultraviolet microscopy. Eur J Cell Biol 43: 93–97Google Scholar
  23. — — (1990) Organization and dynamics of cortical endoplasmic reticulum in inner epidermal cells of onion bulb scales. Protoplasma 157: 203–215Google Scholar
  24. —, Lancelle SA, Hepler PK (1990) Actin-endoplasmic reticulum complexes inDrosera: their structural relationship with the plasmalemma, nucleus, and organelles in cells prepared by high pressure freezing. Protoplasma 155: 116–126Google Scholar
  25. McCulloch SR, Beilby MJ (1997) The electrophysiology pf plasmolysed cells ofChara australis. J Exp Bot 312: 1383–1392Google Scholar
  26. Mizuta S, Tsuji T, Tsurumi S (1995) Role of plasma membrane proteins and cell wall in meridional arrangement of cortical microtubules inBoodlea coacta. Protoplasma 189: 123–131Google Scholar
  27. Oparka KJ (1994) Plasmolysis: new insights into an old process. New Phytol 126: 571–591Google Scholar
  28. —, Prior DAM, Harris N (1990) Osmotic induction of fluid-phase endocytosis in onion epidermal cells. Planta 180: 555–561Google Scholar
  29. — —, Crawford JW (1994) Behavior of plasma membrane, cortical endoplasmic reticulum and plasmodesmata during plasmolysis of onion epidermal cells. Plant Cell Environ 17: 163–171Google Scholar
  30. — — — (1996) Membrane conservation during plasmolysis. In: Small-wood M, Knox JP, Bowles DJ (eds) Membranes: specialized functions in plants. Bios Scientific Publishers, Oxford, pp 39–56Google Scholar
  31. Palta JP, Lee-Stadelmann OY (1983) Vacuolated plant cells as ideal osmometer: reversibility and limits of plasmolysis, and estimation of protoplasm volume in control and water-stress-tolerant cells. Plant Cell Environ 6: 601–610Google Scholar
  32. Pont-Lezica RF, McNally JG, Pickard BG (1993) Wall-to-membrane linkers in onion epidermis: some hypotheses. Plant Cell Environ 16: 111–123Google Scholar
  33. Porter JC, Hogg N (1998) Integrins take partners: cross-talk between integrins and other membrane receptors. Trends Cell Biol 8: 390–396PubMedGoogle Scholar
  34. Reuzeau C, Doolittle KW, McNally JG, Pickard BG (1997a) Covisualisation in living onion cells of putative integrin, putative spectrin, actin, putative intermediate filaments and other proteins at the cell membrane and in an endomembranous sheath. Protoplasma 199: 173–197PubMedGoogle Scholar
  35. —, McNally JG, Pickard BG (1997b) The endomembrane sheath: a key structure for understanding the plant cell? Protoplasma 200: 1–9PubMedGoogle Scholar
  36. Schnepf E, Deichgräber G, Bopp M (1986) Growth, cell wall formation and differentiation in the protonema of the moss,Funaria hygrometrica: effects of plasmolysis on the developmental program and its expression. Protoplasma 133: 50–65Google Scholar
  37. Sheetz MP, Felsenfeld DP, Galbraith CG (1998) Cell migration: regulation of force on extracellular matrix-integrin complexes. Trends Cell Biol 8: 51–54PubMedGoogle Scholar
  38. Sitte P (1963) Zellfeinbau bei Plasmolyse. Protoplasma 57: 304–333Google Scholar
  39. Smith DL (1972) Staining and osmotic properties of young gametophytes ofPolypodium ulgare L. and their bearing on rhizoid function. Protoplasma 74: 465–479Google Scholar
  40. Sonobe S, Shibaoka H (1989) Cortical fine actin filaments in higher plant cells visualized by rhodamine-phalloidin after pretreatment with m-maleimidobenzoyl N-hydroxysuccinimide ester. Protoplasma 148: 80–86Google Scholar
  41. Stadelmann E (1964) Zu Plasmolyse und Deplasmolyse vonAllium-Epidermen. Protoplasma 59: 14–68Google Scholar
  42. Traas JA, Doonan JH, Rawlings DJ, Shaw PJ, Watts J, Lloyd CW (1987) An actin network is present in the cytoplasm throughout the cell cycle of carrot cells and associates with the dividing nucleus. J Cell Biol 105: 387–395Google Scholar
  43. Ueda K, Matsuyama T, Hashimoto T (1999) Visualization of microtubules in living cells of transgenicArabidopsis thaliana. Protoplasma 206: 201–206Google Scholar
  44. Url W (1960) Rosettensystrophe in Thioharnstoff-Zucker-Mischlösungen. Protoplasma 52: 260–273Google Scholar
  45. — (1971) The site of penetration resistance to water in plant proto plasts. Protoplasma 72: 427–447Google Scholar
  46. — (1974) Plasmolyse und Zytorrhyse. Institute for Scientific Films, Göttingen, Film C 1144, pp 3–14Google Scholar
  47. Wasteneys GO, 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
  48. Wymer C, Lloyd C (1996) Dynamic microtubules: implications for cell wall patterns. Trends Plant Sci 1: 222–228Google Scholar
  49. Zandomeni K, Schopfer P (1994) Mechanosensory microtubule reorientation in the epidermis of maize coleptiles subjected to bending stress. Protoplasma 182: 96–101PubMedGoogle Scholar

Copyright information

© Springer-Verlag 2000

Authors and Affiliations

  • I. Lang-Pauluzzi
    • 1
  • B. E. S. Gunning
    • 2
  1. 1.Institute of Plant PhysiologyUniversity of ViennaViennaAustria
  2. 2.Plant Cell Biology Group, Research School of Biological SciencesAustralian National UniversityCanberra

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