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Journal of Applied Phycology

, Volume 28, Issue 1, pp 15–24 | Cite as

Optimization of environmental parameters for Nannochloropsis salina growth and lipid content using the response surface method and invading organisms

  • Meridith L. Bartley
  • Wiebke J. Boeing
  • David Daniel
  • Barry N. Dungan
  • Tanner Schaub
Article

Abstract

Algae biofuel has the potential to replace fossil fuels. However, cultivation and productivity of target algae need improvement, while controlling undesired organisms that can lower the efficiency of production systems. A central composite design and response surface model were utilized to predict cultivation optima of marine microalga, Nannochloropsis salina, under a suite of environmental parameters. The effects of salinity, pH, and temperature and their interactions were studied on maximum sustainable yield (MSY, a measure for biomass productivity), lipid content of N. salina, and invading organisms. Five different levels of each environmental predictor variable were tested. The environmental factors were kept within ranges that had previously been determined to allow positive N. salina growth (14.5–45.5 PSU; pH 6.3–9.7; 11–29 °C). The models created for this experiment showed that N. salina’s MSY and lipid content are not strongly affected over the broad range of salinity and temperature values. Calculated optima levels were 28 PSU/20 °C for MSY and 14.5 PSU/20 °C for lipid accumulation, but neither value significantly influenced the model. However, pH was the most important factor to influence algae productivity, and pH optimum was estimated around 8. Both MSY and lipid content were strongly reduced when pH deviated from the optimum. Occurrence of invading organisms seemed stochastic, and none of the environmental factors studied significantly influenced abundance. In conclusion, pH should be kept around 8 for maximum productivity of N. salina. Temperature and salinity should be kept around 20 °C and 28 PSU; however, moderate variations are not too much of a concern and might enhance lipid content of N. salina.

Keywords

Algae density Lipid productivity Environmental factors Biodiesel fuel Invasive organisms Response surface model 

Notes

Acknowledgments

We are grateful for the valuable work from the following undergraduate students: Herman Campos, Levi Chavez, Renee Pardee, Herberto Chaparro, and Zach Brecheisen. Neeshia Macanowicz was vital to the design and construction of the temperature control system. Darren James provided valuable help with statistical analyses for this research. This work is supported by the US Department of Energy under contract DE-EE0003046 awarded to the National Alliance for Advanced Biofuels and Bioproducts and by the Center for Animal Health, Food Safety and Biosecurity at New Mexico State University. This is a New Mexico Agricultural Experiment Station publication, supported by state funds and the US Hatch Act.

References

  1. Abu-Rezq TS, Al-Musallam L, Al-Shimmari J, Dias P (1999) Optimum production conditions for different high-quality marine algae. Hydrobiologia 403:97–107CrossRefGoogle Scholar
  2. Bartley ML, Boeing WJ, Corcoran AA, Holguin FO, Schaub T (2013) Effects of salinity on growth and lipid accumulation of biofuel microalga Nannochloropsis salina and invading organisms. Biomass Bioenergy 54:83–88CrossRefGoogle Scholar
  3. Bartley ML, Boeing WJ, Dugan BN, Holguin FO, Schaub T (2014) pH effects on growth and lipid accumulation of the biofuel microalgae Nannochloropsis salina and invading organisms. J Appl Phycol 26:1431–1437CrossRefGoogle Scholar
  4. Becker EW (1994) Microalgae biotechnology and microbiology. Cambridge University Press, Cambridge, pp 128–142Google Scholar
  5. Bigelow NW, Hardin WR, Barker JP, Ryken SA, MacRae AC, Cattolico RA (2011) A comprehensive GC-MS sub-microscale assay for fatty acids and its applications. J Am Oil Chem Soc 88:1329–1338CrossRefPubMedPubMedCentralGoogle Scholar
  6. Borowitzka MA (1998) Limits to growth. In: Wong YS, Tam NFY (eds) Wastewater treatment with algae. Springer, Berlin, pp 203–218CrossRefGoogle Scholar
  7. Brennan L, Owende P (2010) Biofuels from microalgae—a review of technologies for production, processing, and extractions of biofuels and co-products. Renew Sust Energ Rev 14:557–577CrossRefGoogle Scholar
  8. Brown MR, Garland CD, Jeffrey SW, Jameson ID, Leroi JM (1993) The gross and amino acid compositions of batch and semi-continuous cultures of Isochrysis sp. (clone T. ISO), Pavlova lutheri and Nannochloropsis oculata. J Appl Phycol 5:285–296CrossRefGoogle Scholar
  9. Campos H, Boeing WJ, Dungan BN (2014) Cultivating the marine microalgae Nannochloropsis salina under various nitrogen sources: effect on biovolume yields, lipid content and composition, and invasive organisms. Biomass Bioenergy 66:301–307CrossRefGoogle Scholar
  10. Chen CY, Durbin EG (1994) Effects of pH on the growth and carbon uptake of marine phytoplankton. Mar Ecol Prog Ser 109:83–94CrossRefGoogle Scholar
  11. Chen M, Tang H, Ma H, Holland TC, Ng KS, Salley SO (2011) Effect of nutrients on growth and lipid accumulation in the green algae Dunaliella tertiolecta. Bioresource Technol 102:1649–1655CrossRefGoogle Scholar
  12. Cheng-Wu Z, Zmora O, Kopel R, Richmond A (2001) An industrial-size flat plate glass reactor for mass production of Nannochloropsis sp. (Eustigmatophyceae). Aquaculture 195:35–49CrossRefGoogle Scholar
  13. Chi Z, Lui Y, Frear C, Shulin C (2009) Study of a two-stage growth of DHA-producing marine algae Schizochytrium limacinum SR21 with shifting dissolved oxygen level. Appl Microbiol Biotechnol 81:1141–1148CrossRefPubMedGoogle Scholar
  14. Chisti Y (2007) Biodiesel from microalgae. BiotechnolAdv 25:294–306Google Scholar
  15. Clavero E, Hernández-Mariné M, Grimalt JO, Garcia-Pichel F (2008) Salinity tolerance of diatoms from thalassic hypersaline environments. J Phycol 36:1021–1034CrossRefGoogle Scholar
  16. Doan TTY, Sivaloganathan B, Obbard JP (2011) Screening of marine microalgae for biodiesel feedstock. Biomass Bioenergy 35:2534–2544CrossRefGoogle Scholar
  17. Griffiths MJ, van Hille RP, Harrison ST (2010) Selection of direct transesterification as the preferred method for assays of fatty acid content of microalgae. Lipids 45:1053–1060CrossRefPubMedGoogle Scholar
  18. Guillard RRL, Rhyther JH (1962) Studies of marine planktonic diatoms: I. Cyclotella nana Hustedt, and Detonula confervacea (Cleve) Gran. Can J Microbiol 8:229–239CrossRefPubMedGoogle Scholar
  19. Hill PS, Tripati AK, Schauble EA (2014) Theoretical constraints on the effects of pH, salinity, and temperature on clumped isotope signatures of inorganic carbon species and precipitating carbonate minerals. Geochim Cosmochim Acta 125:610–652CrossRefGoogle Scholar
  20. Hu H, Gao K (2006) Response of growth and fatty acid compositions of Nannochloropsis sp. to environmental factors under elevated CO2 concentration. Biotechnol Lett 28:987–992CrossRefPubMedGoogle Scholar
  21. Inouye BD (2001) Response surface experimental design for investigating interspecific competition. Ecology 82:2696–2706CrossRefGoogle Scholar
  22. Laurens LML, Quinn M, Van Wychen S, Templeton DW, Wolfrum EJ (2012) Accurate and reliable quantification of total microalgae fuel potential as fatty acid methyl esters by in situ transesterification. Anal Bioanal Chem 403:167–178CrossRefPubMedPubMedCentralGoogle Scholar
  23. Ma J, Lu N, Qin W, Xu R, Wang Y, Chen X (2006) Differential responses of eight cyanobacterial and green algal species, to carbamate insecticides. Ecotox Environ Safe 62:268–274CrossRefGoogle Scholar
  24. Mata TM, Martins A, Caetano NS (2010) Microalgae for biodiesel production and other applications: a review. Renew Sust Energ Rev 14:217–232CrossRefGoogle Scholar
  25. Moazami N, Ashori A, Ranjbar R, Tangestani M, Eghtesadi R, Nejad AS (2012) Large-scale biodiesel production using microalgae biomass of Nannochloropsis. Biomass Bioenergy 39:449–453CrossRefGoogle Scholar
  26. Moheimani NR, Borowitzka MA (2011) Increased CO2 and the effect of pH on growth and calcification of Pleurochrysis carterae and Emiliania huxleyi (Haptophyta) in semicontinuous cultures. Appl Microbiol Biot 90:1399–1407CrossRefGoogle Scholar
  27. Neter J, Kutner M, Nachtsheim C, Wasserman W (2004) Applied linear regression models—4th edition. McGraw-HillGoogle Scholar
  28. Patil P, Reddy H, Muppaneni T, Mannarswamy A, Holguin O, Schaub T, Nirmalakhandan N, Cooke P, Deng S (2012) Power dissipation in microwave-enhanced in-situ transesterification of algal biomass to biodiesel. Green Chem 14:809–818CrossRefGoogle Scholar
  29. R Core Team (2014) R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria. http://www.R-project.org/
  30. Rebolloso-Fuentes MM, Navarro-Perez A, Garćia-Camacho F, Ramos-Miras JJ, Guil-Guerrero JL (2001) Biomass nutrient profiles of the microalga Nannochloropsis. J Agric Food Chem 49:2966–2972CrossRefPubMedGoogle Scholar
  31. Renaud SM, Parry DL (1994) Microalgae for use in tropical aquaculture II: Effect of salinity on growth, gross chemical composition and fatty acid composition of three species of marine microalgae. J Appl Phycol 6:347–356CrossRefGoogle Scholar
  32. Richmond A, Cheng-Wu Z (2001) Optimization of a flat plate glass reactor for mass production of Nannochloropsis sp. outdoors. J Biotechnol 85:259–269CrossRefPubMedGoogle Scholar
  33. Rocha J, Garcia JEC, Henriques MHF (2003) Growth aspects of the marine microalga Nannochloropsis gaditana. Biomol Eng 20:237–242CrossRefPubMedGoogle Scholar
  34. Rodolfi L, Zittelli GC, Bassi N, Padovani G, Biondi N, Bonini G, Tredici MR (2009) Microalgae for oil: strain selection, induction of lipid synthesis and outdoor mass cultivation in a low-cost photobioreactor. Biotechnol Bioeng 102:100–112CrossRefPubMedGoogle Scholar
  35. Roessler PG (1990) Environmental control of glycerolipid metabolism in microalgae: commercial implications and future research directions. J Phycol 26:393–399CrossRefGoogle Scholar
  36. Sforza E, Bertucco A, Morosinotto T, Giacometti GM (2012) Photobioreactors for microalgal growth and oil production with Nannochloropsis salina: from lab-scale experiments to large-scale design. Chem Eng Res Des 90:1151–1158CrossRefGoogle Scholar
  37. Søgaard DH, Hansen PJ, Rysgaard S, Glud RN (2011) Growth limitation and three Arctic sea ice algal species: effects of salinity, pH, and inorganic carbon availability. Polar Biol 34:1157–1165CrossRefGoogle Scholar
  38. Sommer U, Gliwicz ZM, Lampert W, Duncan A (1986) The PEG model of a seasonal succession of planktonic events in fresh waters. Arch Hydrobiol 106:433–471Google Scholar
  39. Sporalore P, Joannis-Cassan C, Duran E, Isambert A (2006) Optimization of Nannochloropsis oculata growth using the response surface method. J Chem Technol Biotechnol 81:1049–1056CrossRefGoogle Scholar
  40. Sukenik A, Zmora O, Carmeli Y (1993) Biochemical quality of marine unicellular algae with special emphasis on lipid composition. II Nannochloropsis sp. Aquaculture 117:313–326CrossRefGoogle Scholar
  41. Van Wagenen J, Miller TW, Hobbs S, Hook P, Crowe B, Huesemann M (2012) Effects of light and temperature on fatty acid production in Nannochloropsis salina. Energies 5:731–740CrossRefGoogle Scholar
  42. Xu Y, Boeing WJ (2014) Modeling maximum lipid productivity of microalgae: review and next step. Renew Sust Energ Rev 32:29–39CrossRefGoogle Scholar
  43. Zittelli GC, Pastorelli R, Tredici MR (2000) A modular flat panel photobioreactor (MFPP) for indoor mass cultivation of Nannochloropsis sp. under artificial illumination. J Appl Phycol 12:521–526Google Scholar
  44. Zittelli GC, Rodolfi L, Tredici MR (2003) Mass cultivation of Nannochloropsis sp. in annular reactors. J Appl Phycol 15:107–114Google Scholar

Copyright information

© Springer Science+Business Media Dordrecht 2015

Authors and Affiliations

  • Meridith L. Bartley
    • 1
  • Wiebke J. Boeing
    • 1
  • David Daniel
    • 2
  • Barry N. Dungan
    • 3
  • Tanner Schaub
    • 3
  1. 1.Department of Fish, Wildlife and Conservation EcologyNew Mexico State UniversityLas CrucesUSA
  2. 2.Department of Applied StatisticsNew Mexico State UniversityLas CrucesUSA
  3. 3.Chemical Analysis and Instrumentation LaboratoryNew Mexico State UniversityLas CrucesUSA

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