Advertisement

Analytical and Bioanalytical Chemistry

, Volume 405, Issue 10, pp 3359–3365 | Cite as

29Si{1H} CP-MAS NMR comparison and ATR-FTIR spectroscopic analysis of the diatoms Chaetoceros muelleri and Thalassiosira pseudonana grown at different salinities

  • Sian M. La Vars
  • Martin R. Johnston
  • John Hayles
  • Jason R. Gascooke
  • Melissa H. Brown
  • Sophie C. Leterme
  • Amanda V. EllisEmail author
Original Paper

Abstract

Diatoms are key indicators of marine environmental health. To further understand how diatoms respond to varying degrees of salinity, either due to climate change or brine waste discharge into marine environments, two different diatom species were studied. Thalassiosira pseudonana and Chaetoceros muelleri were cultured at three different salinities namely, 26 practical salinity units (PSU or parts per thousand), 36 PSU (standard salinity for culturing of seawater species) and 46 PSU. Changes in silica and organic content within the cultured diatoms were analysed using solid-state 29Si{1H} cross-polarization–magic angle spinning (CP-MAS) nuclear magnetic resonance (NMR) and attenuated total reflection–Fourier transform infrared (ATR-FTIR) spectroscopies coupled with analysis of variance. 29Si CP-MAS NMR showed that qualitatively the Q4:Q3 area ratios of C. muelleri, grown away from standard salinities, increased in response to the formation of more condensed (2 ≡SiOH → ≡Si–O–Si≡ + H2O) and/or an increase in closely associated organic matter to the Q4 component of the diatoms. This was not observed for T. pseudonana. However, both species showed the appearance of a new peak centered at 1575–1580 cm−1 in the ATR-FTIR spectra, designated as the C═N band of nitrogenous purine-type compounds. Further, the C. muelleri species was shown to produce more extracellular polymeric substances at non-standard salinities. On this basis, results suggest that there is a strong relationship between diatom composition and salinity and that C. muelleri is more sensitive to its environment than T. pseudonana.

Keywords

Diatoms Salinity 29Si solid-state nuclear magnetic resonance Fourier transform infrared spectroscopy 

Notes

Acknowledgments

The authors wish to acknowledge the financial support of an ARC Discovery Grant (DP110101679) and the National Centre of Excellence in Desalination Australia which is funded by the Australian Government through the Water for the Future initiative. We thank the Australian Research Council for a LIEF grant (LE0668489) for the purchase of the NMR spectrometer. We also thank Matthieu Bourgery for his assistance with the algal culture.

References

  1. 1.
    Falciatore A, Bowler C (2002) Revealing the molecular secrets of marine diatoms. Annu Rev Plant Biol 53(1):109–130CrossRefGoogle Scholar
  2. 2.
    Gröger C, Sumper M, Brunner E (2008) Silicon uptake and metabolism of the marine diatom Thalassiosira pseudonana: solid-state 29Si NMR and fluorescence microscopic studies. J Struc Biol 161(1):55–63CrossRefGoogle Scholar
  3. 3.
    Kröger N, Lorenz S, Brunner E, Sumper M (2002) Self-assembly of highly phosphorylated silaffins and their function in biosilica morphogenesis. Science 298:584–586CrossRefGoogle Scholar
  4. 4.
    Sumper M (2002) A phase separation model for the nanopatterning of diatom biosilica. Science 295:2430–2433CrossRefGoogle Scholar
  5. 5.
    Del Amo Y, Brzezinski MA (1999) The chemical form of dissolved Si taken up by diatoms. J Phycol 35:1162–1170CrossRefGoogle Scholar
  6. 6.
    Vrieling EG, Beelen TPM, van Santen RA, Gieskes WWC (1999) Silicon deposition in diatoms: control by the pH inside the silicon deposition vesicle. J Biotechnol 70:39–51CrossRefGoogle Scholar
  7. 7.
    Kröger N, Poulsen N (2008) Diatoms—from cell wall biogenesis to nanotechnology. Annu Rev Genet 42:83–107CrossRefGoogle Scholar
  8. 8.
    Brunner E, Gröger C, Lutz K, Richthammer P, Spinde K, Sumper M (2009) Analytical studies of silica biomineralization: towards an understanding of silica processing by diatoms. Appl Microbiol Biotechnol 84(4):607–616CrossRefGoogle Scholar
  9. 9.
    Hedges JI, Baldock JA, Gélinas Y, Lee C, Peterson ML, Wakeham SG (2002) The biochemical and elemental compositions of marine plankton: a NMR perspective. Mar Chem 78(1):47–63CrossRefGoogle Scholar
  10. 10.
    Tesson B, Gaillard C, Martin-Jézéquel V (2008) Brucite formation mediated by the diatom Phaeodactylum tricornutum. Mar Chem 109(1-2):60–76CrossRefGoogle Scholar
  11. 11.
    Beal CM, Webber ME, Ruoff RS, Hebner RE (2010) Lipid analysis of Neochloris oleoabundans by liquid state NMR. Biotechnol Bioeng 106(4):573–583CrossRefGoogle Scholar
  12. 12.
    Chauton MS, Størseth TR, Johnsen G (2003) High-resolution magic angle spinning 1H NMR analysis of whole cells of Thalassiosira pseudonana (Bacillariophyceae): broad range analysis of metabolic composition and nutritional value. J Appl Phycol 15(6):533–542CrossRefGoogle Scholar
  13. 13.
    Chauton MS, Størseth TR, Krane J (2004) HR MAS DEPT 13C NMR analysis of whole cells of Chaetoceros muelleri (Bacillariophyceae), and comparison with 13C NMR and DEPT 13C NMR analysis of lipophilic extracts. J Phycol 40(3):611–618CrossRefGoogle Scholar
  14. 14.
    Gillam AH, Wilson MA (1985) Pyrolysis-GC–MS and NMR studies of dissolved seawater humic substances and isolates of a marine diatom. Org Geochem 8(1):15–25CrossRefGoogle Scholar
  15. 15.
    Kinrade SD, Gillson A-ME, Knight CTG (2002) Silicon-29 NMR evidence of a transient hexavalent silicon complex in the diatom Navicula pelliculosa. J Chem Soc Dalton Trans, 307–309Google Scholar
  16. 16.
    Størseth TR, Hansen K, Skjermo J, Krane J (2004) Characterization of a β-d-(1→3)-glucan from the marine diatom Chaetoceros mulleri by high-resolution magic-angle spinning NMR spectroscopy on whole algal cells. Carbohydrate Res 339(2):421–424CrossRefGoogle Scholar
  17. 17.
    Størseth TR, Kirkvold S, Skjermo J, Reitan KI (2006) A branched β-D-(1→3,1→6)-glucan from the marine diatom Chaetoceros debilis (Bacillariophyceae) characterized by NMR. Carbohydrate Res 341(12):2108–2114CrossRefGoogle Scholar
  18. 18.
    Gélabert A, Pokrovsky OS, Schott J, Boudou A, Feurtet-Mazel A, Mielczarski J, Mielczarski E, Mesmer-Dudons N, Spalla O (2004) Study of diatoms/aqueous solution interface. I. Acid–base equilibria and spectroscopic observation of freshwater and marine species. Geochim Cosmochim Acta 68(20):4039–4058CrossRefGoogle Scholar
  19. 19.
    Giordano M, Kansiz M, Heraud P, Beardall J, Wood B, McNaughton D (2001) Fourier Transform Infrared spectroscopy as a novel tool to investigate changes in intracellular macromolecular pools in the marine microalga Chaetoceros muellerii (bacillariophyceae). J Phycol 37(2):271–279CrossRefGoogle Scholar
  20. 20.
    Heredia A, van der Strate HJ, Delgadillo I, Basiuk VA, Vrieling EG (2008) Analysis of organo–silica interactions during valve formation in synchronously growing cells of the diatom Navicula pelliculosa. Chem BioChem 9(4):573–584Google Scholar
  21. 21.
    Hirschmugl CJ, Bayarri Z-E, Bunta M, Holt JB, Giordano M (2006) Analysis of the nutritional status of algae by Fourier transform infrared chemical imaging. Infrared Phys Tech 49(1-2):57–63CrossRefGoogle Scholar
  22. 22.
    Guillard RRL, Ryther JH (1962) Studies of marine planktonic diatoms. I. Cvclotella nana Hustedt and Detonula confervacea (Cleve). Gran Can J Microbiol 8:229–239CrossRefGoogle Scholar
  23. 23.
    Bevington PR, Robinson KD (2003) Data reduction and error analysis for the physical sciences, 3rd edn. McGraw-Hill, USAGoogle Scholar
  24. 24.
    G’elabert A, Pokrovsky OS, Schott J, Boudou A, Feurtet-Mazel A, Mielczarski J, Mielczarski E, Mesmer-Dudons N, Spalla O (2004) Study of diatoms/aqueous solution interface. I. Acid–base equilibria and spectroscopic observation of freshwater and marine species. Geochim Cosmochim Acta 68(20):4039–4058CrossRefGoogle Scholar
  25. 25.
    Kammer M, Hedrich R, Ehrlich H, Popp J, Brunner E, Krafft C (2010) Spatially resolved determination of the structure and composition of diatom cell walls by Raman and FTIR imaging. Anal Bioanal Chem 398(1):509–517CrossRefGoogle Scholar
  26. 26.
    Stehfest K, Toepel J, Wilhelm C (2005) The application of Microscopy FTIR spectroscopy to analyse nutrient-stress related changes in biomass composition of phytoplankton algae. Plant Physiol Biochem 43(7):717–726CrossRefGoogle Scholar
  27. 27.
    Jiang W, Saxena A, Song B, Ward BB, Beveridge TJ, Myneni SCB (2004) Elucidation of functional groups on gram-positive and gram-negative bacterial surfaces using infrared spectroscopy. Langmuir 20(26):11433–11442CrossRefGoogle Scholar
  28. 28.
    Yuan P, Wu DQ, He HP, Lin ZY (2004) The hydroxyl species and acid sites on diatomite surface: a combined IR and Raman study. Appl Surf Sci 227(1–4):30–39CrossRefGoogle Scholar
  29. 29.
    Tesson B, Masse S, Laurent G, Maquet J, Livage J, Martin-Jézéquel V, Coradin T (2008) Contribution of multi-nuclear solid state NMR to the characterization of the Thalassiosira pseudonana diatom cell wall. Anal Bioanal Chem 390:1889–1898CrossRefGoogle Scholar
  30. 30.
    Christiansen SC, Hedin N, Epping JD, Janicke MT, del Amo Y, Demarest M, Brzezinski MA, Chmelka BF (2006) Sensitivity considerations in polarization transfer and filtering using dipole–dipole couplings: implications for biomineral systems. Solid State Nucl Mag 29:170–182CrossRefGoogle Scholar
  31. 31.
    Vrieling EG, Sun Q, Tlan M, Kooyman PJ, Gleskes WWC, van Santen RA, Sommerdijk NAJM (2007) Salinity-dependent diatom biosilicification implies an important role of external ionic strength. Proc Nat Acad Sci USA 104(25):10441–10446CrossRefGoogle Scholar
  32. 32.
    Fujii S, Nishimoto N, Notoya A, Hellebust JA (1995) Growth and osmoregulation of Chaetoceros muelleri in Relation to Salinity. Plant Cell Physiol 36(5):759–764Google Scholar
  33. 33.
    Tsuboi M, Takahashi S, Harada I (1973) In: Duchesne J (ed) Infrared and Raman spectra of nucleic acids-vibrations in the base residues. In physico-chemical properties of nucleic acids, 2nd edn. Academic, New York, pp 91–145Google Scholar
  34. 34.
    Movasaghi Z, Rehman S, Rehman IU (2008) Fourier transform infrared (FTIR) spectroscopy of biological tissues. Appl Spectrosc Rev 43:134–179CrossRefGoogle Scholar
  35. 35.
    Dovbeshkoa GI, Gridinab NY, Kruglovac EB, Pashchuka OP (2000) FTIR spectroscopy studies of nucleic acid damage. Talanta 53(1):233–246CrossRefGoogle Scholar
  36. 36.
    Hecky RE, Mopper K, Kilham P, Degens ET (1973) The amino acid and sugar composition of diatom cell walls. Mar Biol 19:323–331CrossRefGoogle Scholar
  37. 37.
    Smith DJ, Underwood GJ (1998) Exopolymer production by intertidal epipelic diatoms. Limnol Oceanogr 43(7):1578–1591CrossRefGoogle Scholar
  38. 38.
    Krembs C, Eicken H, Junge K, Deming JW (2002) High concentrations of exopolymeric substances in Arctic winter sea ice: implications for the polar ocean carbon cycle and cryoprotection of diatoms. Deep-Sea Res 1 Oceanogr Res Pap 49:2163–2218CrossRefGoogle Scholar

Copyright information

© Springer-Verlag Berlin Heidelberg 2013

Authors and Affiliations

  • Sian M. La Vars
    • 1
  • Martin R. Johnston
    • 1
  • John Hayles
    • 2
  • Jason R. Gascooke
    • 1
  • Melissa H. Brown
    • 2
  • Sophie C. Leterme
    • 2
    • 3
  • Amanda V. Ellis
    • 1
    Email author
  1. 1.Flinders Centre for Nanoscale Science and Technology, School of Chemical and Physical SciencesFlinders UniversityBedford ParkAustralia
  2. 2.School of Biological SciencesFlinders UniversityBedford ParkAustralia
  3. 3.SARDI Aquatic SciencesWest BeachAustralia

Personalised recommendations