Ca2+ as a Signal in the Induction of Callose Synthesis

  • H. Kauss
  • T. Waldmann
  • H. Quader
Part of the NATO ASI Series book series (volume 47)


In suspension-cultured cells and protoplasts, callose synthesis can be triggered by various biochemically unrelated substances. Laser scanning microscopy shows that the polycation chitosan is bound to the surface of protoplasts. Induction of callose synthesis by chitosan increases with the degree of chitosan polymerization up to several thousand, whereas N-acetylation of chitosan — statistically at every fourth to fifth glucosamine residue — decreases its potency. These results suggest that interaction of chitosan might occur over a large surface area with the phospholipid head groups on the plasma membrane. In contrast, the primary interaction of other callose elicitors (e.g. saponins, polyene antibiotics, acylated cyclic peptides) may occur with various constituents in the lipid phase of the plasma membrane. Taken together, signal perception for callose synthesis appears not to involve complementary receptors in the classical sense.

Common to all types of callose elicitors is the induction of a rapid K+ efflux, which is correlated with an external alkalinization and followed temporally by net Ca2+ uptake. As the plasma membrane-located 1,3-ß-glucan synthase has an absolute requirement for Ca2+ in the μM range, it has been suggested that Ca2+ uptake may lead to an increase in cytoplasmic [Ca2+] and thereby trigger callose synthesis. This idea is supported by the observations that external Ca2+ is essential for callose induction, and that inhibition of Ca2+ uptake by putative Ca2+-channel blockers decreases callose synthesis. Increasing Ca2+-uptake alone, however, appears not to be sufficient for the induction of callose formation as shown with the Ca2+-ionophore A 23187 and by low doses of Amphotericin B. Plasma membrane depolarization by itself appears not to represent an additional signal since callose synthesis is increased by the hyperpolarizing toxin fusicoccin and is decreased by protonophores.

Some of the substances capable of rapidly inducing callose synthesis have also been shown to elicit the slower production of phytoalexins, suggesting that the signal transduction mechanism involved in callose synthesis may also contribute to the regulation of other metabolic pathways.


Soybean Cell Polyene Antibiotic Callose Synthesis Callose Formation Channel Blocker Nifedipine 
These keywords were added by machine and not by the authors. This process is experimental and the keywords may be updated as the learning algorithm improves.


Unable to display preview. Download preview PDF.

Unable to display preview. Download preview PDF.


  1. Aist JR, Gold RE, Bayles CJ, Morrison GH, Chandra S, Israel HW (1988) Evidence that molecular components of papillae may be involved in ml-o resistance to barley powdery mildew. Physiol Mol Plant Pathol 33: 17–32CrossRefGoogle Scholar
  2. Atkinson MM, Baker CJ, Collmer A (1986) Transient activation of plasmalemma K+ efflux and H+ influx in tobacco by a pectate lyase isozyme from Erwinia chrysanthemi. Plant Physiol 82: 142–146PubMedCrossRefGoogle Scholar
  3. Bolard J (1986) How do the polyene macrolide antibiotics affect the cellular membrane properties? Biochim Biophys Acta 864: 257–304PubMedGoogle Scholar
  4. Bonhoff A, Grisebach H (1988) Elicitor-induced accumulation of glyceollin and callose in soybean roots and localized resistance against Phytophthora megasperma f. sp. glycinea. Plant Sci 54: 203–209CrossRefGoogle Scholar
  5. Christensen O (1987) Mediation of cell volume regulation by Ca2+ influx through stretch-activated channels. Nature 330: 66–68PubMedCrossRefGoogle Scholar
  6. Clark HF, Shepard CC (1963) A dialysis technique for preparing fluorescent antibody. Virology 20: 642–644PubMedCrossRefGoogle Scholar
  7. Conrath U, Domard A, Kauss H (1989) Correlations in the chitosan-elicited synthesis of callose and of coumarin derivatives by parsley cell suspensions. Plant Cell Reports, in pressGoogle Scholar
  8. Delmer DP (1987) Cellulose biosynthesis. Annu Rev Plant Physiol 38: 259–290CrossRefGoogle Scholar
  9. Eilam Y, Grossowicz N (1982) Nystatin effects on cellular calcium in Saccharomyces cerevisiae. Biochim Biophys Acta 692: 238–423PubMedCrossRefGoogle Scholar
  10. Eschrich W 1956 Kallose. Protoplasma 47: 487–530CrossRefGoogle Scholar
  11. Falke LC, Edwards KL, Pickard BG, Misler S (1988) A stretch-activated anion channel in tobacco protoplasts. FEBS Lett 237: 141–144PubMedCrossRefGoogle Scholar
  12. Fink J, Jeblick W, Blaschek W, Kauss H (1987) Calcium ions and polyamines activate the plasma membrane-located 1,3-ß-glucan synthase. Planta 171: 130–135CrossRefGoogle Scholar
  13. Gögelein H, Hüby A (1984) Interaction of saponin and digitonin with black lipid membranes and lipid monolayers. Biochim Biophys Acta 773: 32–38PubMedCrossRefGoogle Scholar
  14. Gold RE, Aist JR, Hazen BE, Stolzenburg MC, Marshall MR, Israel HW (1986) Effects of calcium nitrate and chlortetracycline on papilla formation, ml-o resistance and susceptibility of barley toowdery mildew. Physiol Plant Pathol 29: 115–129CrossRefGoogle Scholar
  15. Hächler H, Hohl HR (1984) Temporal and spatial distribution patterns of collar and papillae wall appositions in resistant and susceptible tuber tissue of Solanum tuberosum infected by Phytophthora infestans. Physiol Plant Pathol 24: 107–118CrossRefGoogle Scholar
  16. Kauss H (1987) Some aspects of calcium-dependent regulation in plant metabolism. Annu Rev Plant Physiol 38: 47–72CrossRefGoogle Scholar
  17. Kauss H. (1990) Role of the plasma membrane in host/pathogen interactions. In: Larsson Ch, MRller IM (eds) The Plant Plasma Membrane — Structure, Function and Molecular Biology. Springer, in pressGoogle Scholar
  18. Kauss H, Jeblick W (1985) Activation by polyamines, polycations, and ruthenium red of the Ca2+-dependent glucan synthase from soybean cells. FEBS Lett 185: 226–230CrossRefGoogle Scholar
  19. Kauss H, Jeblick W (1986) Influence of free fatty acids, lysophosphatidylcholine, platelet-activating factor, acylcarnitine, and Echinocandin B on 1,3-ß-D-glucan synthase and callose synthesis. Plant Physiol 80: 7–13PubMedCrossRefGoogle Scholar
  20. Kauss H, Jeblick W, Domard A (1989) Degree of polymerization and N-acetylation of chitosan determine its ability to elicit callose formation in suspension cells and protoplasts. Planta, in pressGoogle Scholar
  21. Köhle H, Young DH, Kauss H (1984) Physiological changes in suspension-cultured soybean cells elicited by treatment with chitosan. Plant Sci Lett 33: 221–230CrossRefGoogle Scholar
  22. Köhle H, Jeblick W, Poten F, Blaschek W, Kauss H (1985) Chitosan-elicited callose synthesis in soybean cells as a Ca2+-dependent process. Plant Physiol 77: 544–551PubMedCrossRefGoogle Scholar
  23. Mayer MG, Ziegler E (1988) An elicitor from Phytophthora megasperma f. sp. glycinea influences the membrane potential of soybean cotyledonary cells. Physiol Mol Plant Pathol 33: 397–407CrossRefGoogle Scholar
  24. Ojalvo I, Rokem JS, Navon G, Goldberg I (1987) 31P NMR study of elicitor treated Phaseolus vulgaris cell suspension cultures. Plant Physiol 85:716–719PubMedCrossRefGoogle Scholar
  25. Osswald WF, Zieboll S, Elstner EF (1985) Comparison of pH changes and elicitor induced production of glyceollin isomers in soybean cotyledons. Z Naturforsch 40c: 477–481Google Scholar
  26. Pavlovkin J, Novacky A, Ullrich-Eberius CI (1986) Membrane potential changes during bacteria-induced hypersensitive reaction. Physiol Mol Plant Pathol 28: 125–135CrossRefGoogle Scholar
  27. Pelissier B, Thibaud JB, Grignon C, Esquerré-Tugayé MT (1986) Cell surfaces in plant-microorganism interactions. VII. Elicitor preparations from two fungal pathogens depolarize plant membranes. Plant Sci 46: 103–109CrossRefGoogle Scholar
  28. Ride JP, Drysdale RB (1972) A rapid method for the chemical estimation of filamentous fungi in plant tissue. Physiol Plant Pathol 2: 7–15CrossRefGoogle Scholar
  29. Storm DR, Rosenthal KS, Swanson PE (1977) Polymyxin B and related peptide antibiotics. Annu Rev Biochem 46: 723–763PubMedCrossRefGoogle Scholar
  30. Ullrich-Eberius CI, Pavlovkin J, Schindel J, Fischer K, Novacky A (1989) Changes in plasmalemma functions induced by phytopathogenic bacteria. In: Crane FL, Morré DJ, Lbw M (eds) Plasma membrane oxidoreductase in control of animal and plant growth. Plenum Press, New York, in pressGoogle Scholar
  31. Waldmann T, Jeblick W, Kauss H (1988) Induced net Ca2+ uptake and callose biosynthesis in suspension-cultured plant cells. Planta 173: 88–95CrossRefGoogle Scholar
  32. Wise BC, Glass DB, Jen Chou CH, Raynor RL, Katoh N, Schatzman RC, Turner RS, Kibler RF, Kuo JF (1982) Phospholipid-sensitive Ca2+-dependent protein kinase from heart. J Biol Chem 257: 8489–8495PubMedGoogle Scholar
  33. Young DH, Kauss H (1983) Release of calcium from suspension-cultured Glycine max cells by chitosan, other polycations, and polyamines in relation to effects on membrane permeability. Plant Physiol 73: 698–702PubMedCrossRefGoogle Scholar

Copyright information

© Springer-Verlag Berlin Heidelberg 1990

Authors and Affiliations

  • H. Kauss
    • 1
  • T. Waldmann
    • 1
  • H. Quader
    • 2
  1. 1.Fachbereich BiologieUniversität KaiserslauternKaiserslauternBundesrepublik Deutschland
  2. 2.ZellenlehreUniversität HeidelbergHeidelbergBundesrepublik Deutschland

Personalised recommendations