, Volume 165, Issue 3, pp 311–321

Tagetitoxin affects plastid development in seedling leaves of wheat

  • J. H. Lukens
  • R. D. Durbin


Ultrastructural and biochemical approaches were used to investigate the mode of action of tagetitoxin, a nonhost-specific phytotoxin produced by Pseudomonas syringae pv. tagetis (Hellmers) Young, Dye and Wilkie, which causes chlorosis in developing — but not mature — leaves. Tagetitoxin has no effect on the growth rate or morphology of developing leaves of wheat (Triticum aestivum L.) seedlings. Its cytological effects are limited to plastid aberrations; in both light-and dark-grown leaves treated with toxin, internal plastid membranes fail to develop normally and plastid ribosomes are absent, whereas mitochondrial and cytoplasmic ribosomes are unaffected. The activity of a plastid stromal enzyme, ribulose-1,5-bisphosphate carboxylase (RuBPCase, EC, which is co-coded by nuclear and chloroplast genes, is markedly lower in extracts of both light-and dark-grown toxin-treated leaves, whereas the activity of another stromal enzyme, NADP-glyceraldehyde-3-phosphate dehydrogenase (NADP-G-3P-DH, EC, which is coded only by the nuclear genome, is significantly lower in extracts of light-grown, but not of dark-grown, treated leaves. The mitochondrial enzymes fumarase (EC and cytochrome-c oxidase (EC are unaffected by toxin in dark-grown leaves, but fumarase activity is reduced in light-grown ones. Four peroxisomal enzyme activities are lowered by toxin treatment in both light- and dark-grown leaves. Light- and dark-grown, toxintreated leaves contain about 50% and 75%, respectively, of the total protein of untreated leaves. There are threefold and twofold increases in free amino acids in light-grown and dark-grown treated leaves, respectively. In general, the effects of tagetitoxin are more extensive and exaggerated in light-grown than in dark-grown leaves. We conclude that tagetitoxin interferes primarily with a light-independent aspect of chloroplast-specific metabolism which is important in plastid biogenesis.

Key words

Chloroplast development Chloroplast ribosomes Etioplast (prolamellar body) Pseudomonas (toxin) Tagetitoxin Triticum (chloroplast development) 



NADP-glyceraldehyde-3-phosphate dehydrogenase


prolamellar body


ribulose-1,5-bisphosphate carboxylase


shikimic acid dehydrogenase


Unable to display preview. Download preview PDF.

Unable to display preview. Download preview PDF.


  1. Beevers, H. (1979) Microbodies in higher plants. Annu. Rev. Plant. Physiol. 30, 159–193Google Scholar
  2. Bieleski, R.L., Turner, N.A. (1966) Separation and estimation of amino acids in crude plant extracts by thin-layer electrophoresis and chromatography. Anal. Biochem. 17, 278–293Google Scholar
  3. Blair, G.E., Ellis, R.J. (1973) Protein synthesis in chloroplasts. I. Light-driven synthesis of the large subunit of Fraction I protein by isolated pea chloroplasts. Biochim. Biophys. Acta 319, 223–234Google Scholar
  4. Boller, T., Kende, H. (1979) Hydrolytic enzymes in the central vacuole of plant cells. Plant Physiol. 63, 1123–1132Google Scholar
  5. Bradford, M.M. (1976) A rapid and sensitive method for quantitation of μg quantities of protein using the principle of protein-dye binding. Anal. Biochem. 72, 248–254Google Scholar
  6. Castelfranco, P.A., Beale, S.I. (1981) Chlorophyll biosynthesis. In: Stumpf, P.K., Conn, E.E., eds. The biochemistry of plants, vol. 8, pp. 375–421, Academic Press, New York LondonGoogle Scholar
  7. Dean, C., Leech, R.M. (1982) Genome expression during normal leaf development. I. Cellular and chloroplast numbers and DNA, RNA, and protein levels in tissues of different ages within a seven day old wheat leaf. Plant Physiol. 69, 904–910Google Scholar
  8. De Boer, J., Feierabend, J. (1974) Comparison of the effects of cytokinins on enzyme development in different cell compartments of the shoot organs of rye seedlings. Z. Pflanzenphysiol. 71, 261–270Google Scholar
  9. Ellis, R.J., Hartley, M.R. (1971) Sites of synthesis of chloroplast proteins. Nature New Biol. 233, 193–196Google Scholar
  10. Eyster, H.C. (1950) Catalase activity in pigment-deficient types of corn. Plant Physiol. 25, 630–638Google Scholar
  11. Fedtke, C. (1982) Biochemistry and physiology of herbicide action, Springer, New York Heidelberg BerlinGoogle Scholar
  12. Feierabend, J. (1975) Developmental studies on microbodies in wheat leaves. III. On the photocontrol of microbody development. Planta 123, 63–77Google Scholar
  13. Feierabend, J. (1977) Capacity for chlorophyll synthesis in heat-bleached 70S ribosome-deficient rye leaves. Planta 135 83–88Google Scholar
  14. Feierabend, J., Beevers, H. (1972a) Developmental studies on microbodies in wheat leaves. I. Conditions influencing enzyme development. Plant Physiol. 49, 28–32Google Scholar
  15. Feierabend, J., Beevers, H. (1972b) Developmental studies on microbodies in wheat leaves. II. Ontogeny of particulate enzyme associations. Plant Physiol. 49, 33–39Google Scholar
  16. Feierabend, J., Brassel, D. (1977) Subcellular localization of shikimate dehydrogenase in higher plants. Z. Pflanzenphysiol. 82, 334–346Google Scholar
  17. Feierabend, J., Kemmerich, P. (1983) Mode of interference of chlorosis-inducing herbicides with peroxisomal enzyme activities. Physiol. Plant. 57, 346–351Google Scholar
  18. Feierabend, J., Mikus, M. (1977) Occurrence of a high temperature sensitivity of chloroplast ribosome formation in several higher plants. Plant Physiol. 59, 863–867Google Scholar
  19. Feierabend, J., Schrader-Reichhardt, U. (1976) Biochemical differentiation of plastids and other organelles in rye leaves with a high temperature-induced deficiency of plastid ribosomes. Planta 129, 133–145Google Scholar
  20. Feierabend, J., Schubert, B (1978) Comparative investigation of the action of several chlorosis-inducing herbicides on the biogenesis of chloroplasts and leaf microbodies. Plant Physiol. 61, 1017–1022Google Scholar
  21. George, P. (1953) Intermediate compound formation with peroxidase and strong oxidizing agents. J. Biol. Chem. 201, 413–426Google Scholar
  22. Griffiths, W.T., Beer, N.S. (1982) Site of synthesis of NADPH protochlorophyllide oxidoreductase in rye (Secale cereale). Plant Physiol. 70, 1014–1018Google Scholar
  23. Grob, K., Matile P (1980) Compartmentation of ascorbic acid in vacuoles of horseradish root cells. Z. Pflanzenphysiol. 98, 235–243Google Scholar
  24. Gruber, P.J., Becker, W.M., Newcomb, E. (1972) The occurrence of microbodies and peroxisomal enzymes in achlorophyllous leaves. Planta 105, 114–138Google Scholar
  25. Jutte, S.M., Durbin, R.D. (1979) Ultrastructural effects in zinnia leaves of a chlorosis-inducing toxin from Pseudomonas tagetis. Phytopathology 69, 839–843Google Scholar
  26. Lee, T.T. (1973) On extraction and quantitation of plant peroxidase isoenzymes. Physiol. Plant. 29, 198–203Google Scholar
  27. Lück, H. (1965) Catalase. In: Bergmeyer, H.U., ed. Methods of enzymatic analysis, pp. 885–894, Academic Press, New York LondonGoogle Scholar
  28. Lukens, J.H. (1983) Investigations into the mode of action of tagetitoxin in plants. Ph.D. thesis, University of Wisconsin, Madison, USAGoogle Scholar
  29. Mitchell, R.E., Bieleski, R.L. (1977) Involvement of phaseolotoxin in the halo blight of beans. Plant Physiol. 60, 723–729Google Scholar
  30. Mitchell, R.E., Durbin, R.E. (1981) Tagetitoxin, a toxin produced by Pseudomonas syringae pv. tagetis: purification and partial characterization. Physiol. Plant Pathol 18, 157–168Google Scholar
  31. Mitchell, R.E., Hart, P.A. (1983) The structure of tagetitoxin, a phytotoxin of Pseudomonas syringae pv. tagetis. Phytochemistry 22, 1425–1428Google Scholar
  32. Müller, B., Ziegler, I., Ziegler, H. (1969) Lichtinduzierte, reversible Aktivitätssteigerung der NADP-abhängigen Glycerinaldehyd-3-Phosphatdehydrogenase in Chloroplasten. Zum Mechanismus der Reaktion. Eur. J. Biochem. 9, 101–106Google Scholar
  33. Nadolny, L. (1978) The involvement of peroxidase in protection of tobacco against the hypersensitive reaction induced by the Pseudomonas solanacearum B1 isolate. M.S. thesis, University of Wisconsin, Madison, USAGoogle Scholar
  34. Nilsson, G. (1978) Effects of glyphosate on the amino acid content in spring wheat plants. Weed Abstr. 27, 407Google Scholar
  35. Racker, E. (1962) Ribulose diphosphate carboxylase from spinach leaves. Methods Enzymol. 5, 226–270Google Scholar
  36. Reiss, T., Bergfeld, R., Link, G., Thien, W., Mohr, H. (1983) Photooxidative destructive of chloroplasts and its consequences for cytosolic enzyme levels and plant development. Planta 159, 518–528Google Scholar
  37. Steinrücken, H.C., Amrhein, N. (1980) The herbicide glyphosate is a potent inhibitor of 5-enolpyruvyl-shikimic acid-3-phosphate synthase. Biochem. Biophys. Res. Commun. 94, 1207Google Scholar
  38. Styer, D.J., Durbin, R.D. (1982a) Common ragweed: a new host for Pseudomonas syringae pv. tagetis. Plant Dis. 66, 71Google Scholar
  39. Styer, D.J., Durbin, R.D. (1982b) Isolation of Pseudomonas synringae pv. tagetis from sunflower in Wisconsin. Plant Dis. 66, 701Google Scholar
  40. Tolbert, N.E., Oeser, A., Kisacki, T., Hageman, R.H., Yamazaki, R.K. (1968) Peroxisomes from spinach leaves containing enzymes related to glycolate metabolism. J. Biol. Chem. 243, 5179–5184Google Scholar
  41. Trimboli, D., Fahy, P.C., Baker, K.F. (1978) Apical chlorosis and leaf spot of Tagetes spp. caused by Pseudomonas tagetis Hellmers. Aust. J. Agric. Res. 29, 831–839Google Scholar
  42. Wintermans, J.F.G.M., De Mots, A. (1965) Spectrophotometric characteristics of chlorophyll a and b and their pheophytins in ethanol. Biochim. Biophys. Acta 109, 448–453Google Scholar

Copyright information

© Springer-Verlag 1985

Authors and Affiliations

  • J. H. Lukens
    • 1
    • 2
    • 3
  • R. D. Durbin
    • 1
    • 2
  1. 1.Department of Plant PathologyUniversity of WisconsinMadisonUSA
  2. 2.Agricultural Research ServiceU.S. Department of AgricultureMadisonUSA
  3. 3.Biological LaboratoriesHarvard UniversityCambridgeUSA

Personalised recommendations