Advertisement

UV-Visible Spectrophotometry-Based Metabolomic Analysis of Cedrela Fissilis Velozzo (Meliaceae) Calluses - A Screening Tool for Culture Medium Composition and Cell Metabolic Profiles

  • Fernanda Kokowicz PilattiEmail author
  • Christopher Costa
  • Miguel Rocha
  • Marcelo Maraschin
  • Ana Maria Viana
Conference paper
  • 634 Downloads
Part of the Advances in Intelligent Systems and Computing book series (AISC, volume 375)

Abstract

In plant cell cultures aiming at the production of secondary metabolites of industrial interest, the culture medium composition is a decisive step for obtaining cell growth and high yields of the target compound(s). A rapid and reliable methodology for screening metabolic responses to medium composition is fundamental for the development of this biotechnological field. Following this approach, UV-Vis scanning spectrophotometry of callus extracts and their spectra pre-processing, univariate and multivariate analysis were tested in the present work. The results obtained successfully discriminated the culture media investigated and shed light on what metabolic pathways might be responsible for the differences among the callus cultures’ metabolic profiles.

Keywords

Metabolic profiling Cedrela fissilis UV-Visible spectrophotometry R language Unsupervised methods 

Notes

Acknowledgements:

To CAPES (Coordination for the Improvement of Higher Education Personnel) for financial support, Post-Graduation Program in Cell and Developmental Biology (Federal University of Santa Catarina) and BIOSYSTEMS research group (University of Minho).

References

  1. 1.
    Goulart, H.R., Kimura, E.A., Peres, V.J., Couto, A.S., Aquino Duarte, F.A., Katzin, A.M.: Terpenes arrest parasite development and inhibit biosynthesis of isoprenoids in Plasmodium falciparum. Antimicrob. Agents Chemother. 48, 2502–2509 (2004)CrossRefGoogle Scholar
  2. 2.
    Leite, A.C., Bueno, F.C., Oliveira, C.G., Fernandes, J.B., Da Vieira, P.C., Silva, M.F.G.F., Bueno, O.C., Pagnocca, F.C., Hebling, M.J.A., Bacci Jr, M.: Limonoids from Cipadessa fruticosa and Cedrela fissilis and their insecticidal activity. J. Braz. Chem. Soc. 16, 1391–1395 (2005)CrossRefGoogle Scholar
  3. 3.
    Ambrozin, A.R.P., Leite, A.C., Bueno, F.C.: Limonoids from andiroba oil and Cedrela fissilis and their insecticidal activity. J. Braz. Chem. Soc. 3, 542–547 (2006)Google Scholar
  4. 4.
    Ramos, D.F., Leitão, G.G., Costa, F.N., Abreu, L., Villarreal, J.V., Leitão, S.G., Fernández, S.L.S., Da Silva, P.E.A.: Investigation of the antimycobacterial activity of 36 plant extracts from the Brazilian Atlantic Forest. Braz. J. Pharm. Sci. 44, 669–674 (2008)Google Scholar
  5. 5.
    Leite, A.C., Ambrozin, A.R.P., Castilho, M.S., Vieira, P.C., Fernandes, J.B., Oliva, G., Silva, M.F.G.F., Thiemann, O.H., Lima, M.I., Pirani, J.R.: Screening of Trypanosoma cruzi glycosomal glyceraldehyde-3-phosphate dehydrogenase enzyme inhibitors. Rev. Bras. Farmacogn. 19, 1–6 (2009)Google Scholar
  6. 6.
    Nunes, E.C., Castilho, C.V., Moreno, F.N., Viana, A.M.: In vitro culture of Cedrela fissilis Vellozo (Meliaceae). Plant Cell Tiss. Org. 70, 259–268 (2002)CrossRefGoogle Scholar
  7. 7.
    Nunes, E.C., Benson, E.E., Oltramari, A.C., Araujo, P.S., Moser, J.R., Viana, A.M.: In vitro conservation of Cedrela fissilis Vellozo (Meliaceae), a native tree of the Brazilian Atlantic Forest. Biodivers. Conserv. 12, 837–848 (2003)CrossRefGoogle Scholar
  8. 8.
    Nunes, E.C., Laudano, W.L.S., Moreno, F.N., Castilho, C.V., Mioto, P., Sampaio, F.L., Bortoluzi, J.H., Benson, E.E., Pizolatti, M.G., Carasek, E., Viana, A.M.: Micropropagation of Cedrela fissilis Vell. (Meliaceae). In: Jain, S.M., Häggman, H. (ed.) – Protocols for Micropropagation of Woody Trees and Fruits, pp. 221–235. Springer, The Netherlands (2007)Google Scholar
  9. 9.
    Nielsen, J., Jewett, M.C.: Metabolomics - A powerful pool in systems biology. Springer-Verlag, Berlin, Heidelberg (2007)Google Scholar
  10. 10.
    Villas-Bôas, S.G., Roessner, U., Hansen, M.A.E., Smedsgaard, J., Nielsen, J.: Metabolome analysis: An introduction. Wiley, New Jersey (2007)CrossRefGoogle Scholar
  11. 11.
    Putri, S.P., Nakayama, Y., Matsuda, F., Uchikata, T., Kobayashi, S., Matsubara, A., Fukusaki, E.: Current metabolomics: practical applications. J. Biosci. Bioeng. 115, 579–589 (2013)CrossRefGoogle Scholar
  12. 12.
    Fiehn, O.: Combining genomics, metabolome analysis, and biochemical modelling to understand metabolic networks. Comp. Funct. Genomics 2, 155–168 (2001)CrossRefGoogle Scholar
  13. 13.
    Dunn, W.B., Ellis, D.I.: Metabolomics: current analytical platforms and methodologies. Trend. Anal. Chem. 24, 285–294 (2005)CrossRefGoogle Scholar
  14. 14.
    Harborne, J.B.: Phytochemical Methods, 3rd edn. Chapman & Hall, London (1998)Google Scholar
  15. 15.
    Sumner, L.W., Mendes, P., Dixon, R.A.: Plant metabolomics: large-scale phytochemistry in the functional genomics era. Phytochemistry 62, 817–836 (2003)CrossRefGoogle Scholar
  16. 16.
    Xia, J., Mandal, R., Sinelnikov, I.V., Broadhurst, D., Wishart, D.S.: MetaboAnalyst 2.0 - a comprehensive server for metabolomic data analysis. Nucl. Acids Res. 37, W652–W660 (2009)Google Scholar
  17. 17.
    Murashige, T., Skoog, F.: A revised medium for rapid growth and bioassays with tobacco tissue cultures. Physiol. Plant. 15, 473–497 (1962)CrossRefGoogle Scholar
  18. 18.
    Wickham, H., Chang, W., RStudio, R Core team: Tools to Make Developing R Packages Easier (2015)Google Scholar
  19. 19.
    Beleites, C.: Import and Export of Spectra Files. Vignette for the R package hyperSpec (2011)Google Scholar
  20. 20.
    Bosch Ojeda, C., Sanchez Rojas, F.: Recent developments in derivative ultraviolet/visible absorption spectrophotometry. Anal. Chim. Acta 518, 1–24 (2004)CrossRefGoogle Scholar
  21. 21.
    Gamborg, O.L., Murashige, T., Thorpe, T.A., Vasil, I.K.: Plant tissue culture media. In vitro. 12, 473–478 (1976)CrossRefGoogle Scholar
  22. 22.
    Rolland, F., Baena-Gonzalez, E., Sheen, J.: Sugar sensing and signaling in plants: conserved and novel mechanisms. Annu. Rev. Plant Biol. 57, 675–709 (2006)CrossRefGoogle Scholar
  23. 23.
    Miflin, B.J., Habash, D.Z.: The role of glutamine synthetase and glutamate dehydrogenase in nitrogen assimilation and possibilities for improvement in the nitrogen utilization of crops. J. Exp. Bot. 53, 979–987 (2002)CrossRefGoogle Scholar
  24. 24.
    Forde, B.G., Lea, P.J.: Glutamate in plants: metabolism, regulation, and signalling. J. Exp. Bot. 58, 2339–2358 (2007)CrossRefGoogle Scholar
  25. 25.
    Flores-Samaniego, B., Olivera, H., Gonzalez, A.: Glutamine synthesis is a regulatory signal controlling glucose catabolism in Saccharomyces cerevisiae. J. Bacteriol. 175, 7705–7706 (1993)Google Scholar
  26. 26.
    DeBerardinis, R.J., Mancuso, A., Daikhin, E., Nissim, I., Yudkoff, M., Wehrli, S., Thompson, C.B.: Beyond aerobic glycolysis: transformed cells can engage in glutamine metabolism that exceeds the requirement for protein and nucleotide synthesis. PNAS 104, 19345–19350 (2007)CrossRefGoogle Scholar

Copyright information

© Springer International Publishing Switzerland 2015

Authors and Affiliations

  • Fernanda Kokowicz Pilatti
    • 1
    Email author
  • Christopher Costa
    • 2
  • Miguel Rocha
    • 2
  • Marcelo Maraschin
    • 1
  • Ana Maria Viana
    • 3
  1. 1.Plant Morphogenesis and Biochemistry LaboratoryFederal University of Santa CatarinaFlorianópolisBrazil
  2. 2.Centre Biological Engineering School of EngineeringUniversity of MinhoBragaPortugal
  3. 3.Botany DepartmentFederal University of Santa CatarinaFlorianópolisBrazil

Personalised recommendations