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Photosynthetic characterization of two Nannochloropsis species and its relevance to outdoor cultivation

  • Avigad VonshakEmail author
  • Nurit Novoplansky
  • Ana M. Silva Benavides
  • Giuseppe Torzillo
  • John Beardall
  • Yussi M. Palacios
Article

Abstract

Despite the increased interest in exploring the potential of algal biomass production for food stock and renewable energy, very little work has been done in developing reliable screening protocols to enable the identification of species that are best suited to mass cultivation outdoors. Nannochloropsis is an algal genus identified as a potential source of lipids due to its ability to accumulate large quantities of these compounds, especially under nutrient-limiting conditions. The objective of the current work was to use two species of this genus, Nannochloropsis oceanica and N. oculata, as model organisms to develop a protocol that will allow the evaluation of their capacity to yield high biomass productivity under outdoor conditions. Growing the alga under different light intensities and measuring growth rate as well as a range of photosynthetic parameters based on light response curves and variable fluorescence highlighted significant differences between the two species. Our data show that N. oceanica cells have a better capacity to respond to higher light intensities, as reflected by growth measurements, photosynthetic electron transport rates, and oxygen evolution as well as their response to the very high photon flux densities expected in outdoor culture. On the other hand, N. oculata showed a higher tolerance to oxidative stress as reflected in its resistance to the reactive oxygen species generating compounds Rose Bengal (RB) and methyl viologen (MV). Based on the above evidence, we suggest that N. oceanica may perform better than N. oculata when grown under high light conditions typically found outdoors in summer, while N. oculata may perform better than N. oceanica under oxidative stress conditions usually found in outdoor cultures exposed to a combination of high light and low temperature commonly occurring in winter time.

Keywords

Photosynthetic characterization Nannochloropsis Outdoor cultivation Rose bengal Methyl viologen Light stress Oxidative stress 

Notes

Acknowledgements

The work by GT has been partially supported by the H2020 project “SABANA” funded from the EU Horizon 2020 research and innovation program under Grant Agreement No. 727874.

Supplementary material

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References

  1. Ben Sheleg A, Novoplansky N, Vonshak A (2019) Can Rose Bengal resilience be used as a marker for photosynthetic resilience of Nannochloropsis oceanica strains in excess light environments? Algal Res 41:101562CrossRefGoogle Scholar
  2. Bilger W, Bjӧrkman O (1990) Role of the xanthophyll cycle in photoprotection elucidated by measurements of light-induced absorbance changes, fluorescence and photosynthesis in leaves of Hedera canariensis. Photosynth Res 25:173–185CrossRefGoogle Scholar
  3. Borowitzka MA, Vonshak A (2017) Scaling up microalgal cultures to commercial scale. Eur J Phycol 52:407–418CrossRefGoogle Scholar
  4. Boussiba S, Vonshak A, Cohen Z, Avissar Y, Richmond A (1987) Lipid and biomass production by the halotolerant microalga Nannochloropsis salina. Biomass 12:37–48CrossRefGoogle Scholar
  5. Burnison BK (1980) Modified dimethyl sulfoxide (DMSO) extraction for chlorophyll analysis of phytoplankton. Can J Fish Aquat Sci 37:729–733CrossRefGoogle Scholar
  6. Cao S, Zhang X, Xu D, Fan A, Mou S, Wang Y, Ye N, Wang W (2013) A trans thylakoid proton gradient and inhibitors induce a non-photochemical fluorescence quenching in unicellular algae Nannochloropsis sp. FEBS Lett 587:1310–1315CrossRefGoogle Scholar
  7. Chukhutsina VU, Fristedt R, Morosinotto T, Croce R (2017) Photoprotection strategies of the alga Nannochloropsis gadidana. Biochem Biophys Acta 1858:544–522Google Scholar
  8. Dubinsky Z (1992) The functional and optical absorption cross-sections of phytoplankton photosynthesis. In: Fakowski PJ, Woodhead AD (eds) Primary productivity and biogeochemical cycles in the sea. Aquatic photosynthesis, Plenum Press, New York, p 31–45CrossRefGoogle Scholar
  9. Eilers PHC, Peeters JCH (1988) A model for the relationship between light intensity and the rate of photosynthesis in phytoplankton. Ecol Model 42:199–215CrossRefGoogle Scholar
  10. Fisher T, Minnaard J, Dubinsky Z (1996) Photoacclimation in the marine alga Nannochloropsis sp. (Eustigmatophyte): a kinetic study. J Plankton Res 18:1797–1818CrossRefGoogle Scholar
  11. Goss R, Jakob T (2010) Regulation and function of xanthophyll cycle-dependent photoprotection in algae. Photosynth Res 106:103–122CrossRefGoogle Scholar
  12. Hawkes TR (2014) Mechanisms of resistance to paraquat in plants. Pest Manag Sci 70:1316–1323CrossRefGoogle Scholar
  13. Iturbe-Ormaetxe I, Escuredo PR, Arrese-Igor C, Becana M (1998) Oxidative damage in pea plants exposed to water deficit or paraquat. Plant Physiol 116:173–181CrossRefGoogle Scholar
  14. Kandilian R, Lee E, Pilon L (2013) Radiation and optical properties of Nannochloropsis oculata grown under different irradiances and spectra. Bioresour Technol 137:63–73CrossRefGoogle Scholar
  15. Kochevar IE, Redmond RW (2000) Photosensitized production of singlet oxygen. Methods Enzymol 319:20–28CrossRefGoogle Scholar
  16. Kotabová E, Kaňa R, Jarešová J, Prášil O (2011) Non-photochemical quenching fluorescence in Chromera velia is enabled by fast violaxanthin de-epoxidation. FEBS Lett 585:1941–1945CrossRefGoogle Scholar
  17. Kovács L, Ayaydin F, Kálai T, Tandori J, Kós PB, Hideg É (2014) Assessing the applicability of singlet oxygen photosensitizers in leaf studies, photochemistry and photobiology. Photochem Photobiol 90:129–136CrossRefGoogle Scholar
  18. Kromkamp J, Forster RM (2003) The use of variable fluorescence measurements in aquatic ecosystems: differences between multiple and single turnover measuring protocols and suggested terminology. Eur J Phycol 38:103–112CrossRefGoogle Scholar
  19. Kromkamp J, Limbeek M (1993) Effect of short-term variation in irradiance on light harvesting and photosynthesis of the marine diatom Skeletonema costatum: a laboratory study simulating vertical mixing. J Gen Microbiol 139:2277–2284CrossRefGoogle Scholar
  20. Kromkamp JC, Beardall J, Sukenik A, Kopeck J, Masojidek J, van Bergeijk S, Gabai S, t Shaham E, Yamshon A (2009) Short-term variations in photosynthetic parameters of Nannochloropsis cultures grown in two types of outdoor mass cultivation systems. Aquat Microb Ecol 56:309–322CrossRefGoogle Scholar
  21. Ma X-N, Chen T-P, Yang B, Liu J, Chen F (2016) Lipid production from Nannochloropsis. Mar Drugs 14:61–79CrossRefGoogle Scholar
  22. MacIntyre HL, Kana TM, Anning T, Geider RJ (2002) Photoacclimation of photosynthesis irradiance response curves and photosynthetic pigments in microalgae and cyanobacteria. J Phycol 38:17–38CrossRefGoogle Scholar
  23. Murgia I, Tarantino D, Vannini C, Bracale M, Carravieri S, Soave C (2004) Arabidopsis thaliana plants overexpressing thylakoidal ascorbate peroxidase show increased resistance to Paraquat-induced photooxidative stress and to nitric oxide-induced cell death. Plant J 38:940–953CrossRefGoogle Scholar
  24. Padmasree K, Padmavathi L, Raghavendra AS (2002) Essentiality of mitochondrial oxidative metabolism for photosynthesis: optimization of carbon assimilation and protection against photoinhibition. Crit Rev Biochem Mol Biol 37:71–119CrossRefGoogle Scholar
  25. Palacios YM, Vonshak A, Beardall J (2018) Photosynthetic and growth responses of Nannochloropsis oculata (Eustigmatophyceae) during batch cultures in relation to light intensity. Phycologia 57:492–502CrossRefGoogle Scholar
  26. Ruban AV (2014) Evolution under the sun: optimizing light harvesting in photosynthesis. J Exp Bot 66:7–23CrossRefGoogle Scholar
  27. Sandnes JM, Källqvist T, Wenner D, Gislerød HR (2005) Combined influence of light and temperature on growth rates of Nannochloropsis oceanica: linking cellular responses to large-scale biomass production. J Appl Phycol 17:515–525CrossRefGoogle Scholar
  28. Schreiber U, Klughammer C, Kolbowski J (2012) Assessment of wavelength-dependent parameters of photosynthetic electron transport with a new type of multi-color PAM chlorophyll fluorometer. Photosynth Res 113:127–144CrossRefGoogle Scholar
  29. Seely GR, Duncan MJ, Vidaver WE (1972) Preparative and analytical extraction of pigments from brown algae with dimethyl sulfoxide. Mar Biol 12:184–188CrossRefGoogle Scholar
  30. Simionato D, Sforza E, Corteggiani-Carpinelli E, Bertucco A, Giacometti GM, Morosinotto T (2011) Acclimation of Nannochloropsis gaditana to different illumination regimes: effects on lipids accumulation. Bioresour Technol 102:6026–6032CrossRefGoogle Scholar
  31. Simionato D, Block MA, La Rocca N, Jouhet J, Maréchal E, Finazzi G, Morosinotto T (2013) The response of Nannochloropsis gaditana to nitrogen starvation includes de novo biosynthesis of triacylglycerols, a decrease of chloroplast galactolipids, and reorganization of the photosynthetic apparatus. Eukaryot Cell 12:665–676CrossRefGoogle Scholar
  32. Solovchenko A, Lukyanov A, Solovchenko O, Didi-Cohen S, Boussiba S, Khozin-Goldberg I (2014) Interactive effects of salinity, high light, and nitrogen starvation on fatty acid and carotenoid profiles in Nannochloropsis oceanica CCALA 804. Eur J Lipid Sci Technol 116:635–644CrossRefGoogle Scholar
  33. Sukenik A, Beardall J, Kromkamp JC, Kopecky J, Masojídek J, van Bergeijk S, Gabai S, Shaham E, Yamshon A (2009) Photosynthetic performance of outdoor Nannochloropsis mass cultures under a wide range of environmental conditions. Aquat Microb Ecol 56:297–308CrossRefGoogle Scholar
  34. Szabó M, Parker K, Guruprasad S, Kuzhiumparambil U, Lilley RM, Tamburic B, Schliep M, Larkum AW, Schreiber U, Raven JA, Ralph PJ (2014) Photosynthetic acclimation of Nannochloropsis oculata investigated by multi-wavelength chlorophyll fluorescence analysis. Bioresour Technol 167:521–529CrossRefGoogle Scholar
  35. Vonshak A, Laorawat S, Bunnag B, Tanticharoen M (2014) The effect of light availability on the photosynthetic activity and productivity of outdoor cultures of Arthrospira platensis (Spirulina). J Appl Phycol 26:1309–1315CrossRefGoogle Scholar
  36. Weis E, Berry JA (1987) Quantum efficiency of Photosystem II in relation to “energy”-dependent quenching of chlorophyll fluorescence. Biochim Biophys Acta 894:198–208CrossRefGoogle Scholar

Copyright information

© Springer Nature B.V. 2020

Authors and Affiliations

  1. 1.The Jacob Blaustein Institutes for Desert ResearchBen-Gurion University of the NegevMidreshet Ben-GurionIsrael
  2. 2.Escuela de Biología, Centro de Investigación en Ciencias del Mar y Limnología (CIMAR)Universidad de Costa RicaSan PedroCosta Rica
  3. 3.CNR – Institute of BioeconomyFlorenceItaly
  4. 4.School of Biological SciencesMonash UniversityClaytonAustralia

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