Internal structure and photosynthetic performance of Nostoc sp. colonies in the high Arctic

• Autorzy: Kviderova J.
• Języki publikacji: EN

Abstrakty

EN

The physiological performance of Nostoc sp. colonies in the high Arctic was investigated based on their structure and function. To investigate the internal colony structure, a method based on vertical stacking of individual light microscopy images was tested under the conditions at a polar field station. The physiological state of sun-exposed and shaded surfaces of the colonies was assessed using variable chlorophyll fluorescence imaging under two distinct low- and high-light conditions. The 3D image of the internal structure of the colonies revealed a high number of cells in the central part of the colony. Two peaks of maximum cell density were observed, probably caused by two overlapping colony lobes or subcolonies. Light was the driving factor of photosynthetic activity, and the colony structure played a role in the rate of response to incoming light. Fluorescence imaging revealed heterogeneity of the photosynthetic activity in the colonies, with the maximum photosynthetic activity at the colony edge due to better access to nutrients. The differences between exposed and shaded surfaces were not as pronounced as was expected, either due to good photoacclimation to a broad range of light conditions, light distribution through translucent extracellular matrixes, or integration of fluorescence signals throughout the colonies. The slightly better photosynthetic performance under high light conditions may indicate photoacclimation of Nostoc sp. to a broad range of light conditions encountered in the field.

• Wydawca: -
• Czasopismo: Acta Societatis Botanicorum Poloniae
• Rocznik: 2018
• Tom: 87
• Numer: 4
• Opis fizyczny: Article 3602 [12p.],fig.,ref.

Twórcy

autor

Kviderova J.

• Center for Polar Ecology, Faculty of Science, University of South Bohemia in Ceske Budejovice, Na Zlate stoce 3, 370 05 Ceske Budejovice, Czech Republic

Bibliografia

• 1. Broady PA. Diversity, distribution and dispersal of Antarctic algae. Biodivers Conserv. 1996;5:1307–1335. https://doi.org/10.1007/BF00051981
• 2. Elster J. Ecological classification of terrestrial algal communities in polar environments. In: Beyer L, Bötler M, editors. Geoecology of Antarctic ice-free coastal landscapes. Springer; 2002. p. 303–326. (Ecological Studies; vol 154). https://doi.org/10.1007/978-3-642-56318-8_17
• 3. Elster J, Benson EE. Life in the polar terrestrial environment with a focus on algae and cyanobacteria. In: Fuller BJ, Lane N, Benson EE, editors. Life in the frozen state. Boca Raton, FL: CRC Press; 2004. p. 111–150. https://doi.org/10.1201/9780203647073.ch3
• 4. Vincent WF. Cyanobacterial dominance in the Polar regions. In: Whitton BA, Potts M, editors. The ecology of cyanobacteria. Dordrecht: Kluwer Academic Publishers; 2000. p. 321–340. https://doi.org/10.1007/0-306-46855-7_12
• 5. Zakhia F, Jungblut AD, Taton A, Vincent WF, Wilmotte A. Cyanobacteria in cold ecosystems. In: Margesin R, Schinner F, Marx JC, Gerday C, editors. Psychrophiles: from biodiversity to biotechnology. Berlin: Springer; 2008. p. 121–135. https://doi.org/10.1007/978-3-540-74335-4_8
• 6. Jungblut AD, Vincent WF. Cyanobacteria in the polar and alpine ecosystems. In: Margesin R, editor. Psychrophiles: from biodiversity to biotechnology. Cham: Springer; 2017. p. 181–206. https://doi.org/10.1007/978-3-319-57057-0_9
• 7. Morgan-Kiss RM, Priscu JC, Pocock T, Gudynaite-Savitch L, Huner NPA. Adaptation and acclimation of photosynthetic microorganisms to permanently cold environments. Microbiol Mol Biol Rev. 2006;70(1):222–252. https://doi.org/10.1128/MMBR.70.1.222-252.2006
• 8. Schulze ED, Beck E, Müller-Hohenstein K. Plant ecology. Berlin: Springer; 2005.
• 9. Gao K, Ai H. Relationship of growth and photosynthesis with colony size in an edible cyanobacterium, Ge-Xian-Mi Nostoc (Cyanophyceae). J Phycol. 2004;40:523–526. https://doi.org/10.1111/j.1529-8817.2004.03155.x
• 10. Li Y, Gao K. Photosynthetic physiology and growth as a function of colony size in the cyanobacterium Nostoc sphaeroides. Eur J Phycol. 2004;39:9–15. https://doi.org/10.1080/0967026032000157147
• 11. Paerl HW, Pinckney JL, Steppe TF. Cyanobacterial-bacterial mat consortia: examining the functional unit of microbial survival and growth in extreme environments. Environ Microbiol. 2000;2(1):11–26. https://doi.org/10.1046/j.1462-2920.2000.00071.x
• 12. Belnap J, Lange OL. Structure and functioning of biological soil crusts: a synthesis. In: Belnap J, Lange OL, editors. Biological soil crusts: structure, function, and management. Berlin: Springer; 2001. p. 471–479. https://doi.org/10.1007/978-3-642-56475-8_33
• 13. Pushkareva E, Johansen JR, Elster J. A review of the ecology, ecophysiology and biodiversity of microalgae in Arctic soil crusts. Polar Biol. 2016;39(12):2227–2240. https://doi.org/10.1007/s00300-016-1902-5
• 14. Schneider T, Schmid E, de Castro JV, Cardinale M, Eberl L, Grube M, et al. Structure and function of the symbiosis partners of the lung lichen (Lobaria pulmonaria L. Hoffm.) analyzed by metaproteomics. Proteomics. 2011;11(13):2752–2756. https://doi.org/10.1002/pmic.201000679
• 15. Tashyreva D, Elster J. Production of dormant stages and stress resistance of polar cyanobacteria. In: Hanslmeier A, Kempe S, Seckbach J, editors. Life on Earth and other planetary bodies. Dordrecht: Springer; 2012. p. 367–386. https://doi.org/10.1007/978-94-007-4966-5_21
• 16. Aguilera A, Souza-Egipsy V, Gomez F, Amils R. Development and structure of eukaryotic biofilms in an extreme acidic environment, Rio Tinto (SW, Spain). Microb Ecol. 2007;53(2):294–305. https://doi.org/10.1007/s00248-006-9092-2
• 17. Flemming HC, Wingender J. The biofilm matrix. Nat Rev Microbiol. 2010;8(9):623–633. https://doi.org/10.1038/nrmicro2415
• 18. Kvíderová J. Biofilm. In: Amils R, Gargaud M, Cernicharo Quintanilla J, Cleaves HJ, Irvine WM, Pinti D, et al., editors. Encyclopedia of astrobiology. Berlin: Springer; 2015. p. 1–3. https://doi.org/10.1007/978-3-642-27833-4_170-3
• 19. Tamaru Y, Takani Y, Yoshida T, Sakamoto T. Crucial role of extracellular polysaccharides in desiccation and freezing tolerance in the terrestrial cyanobacterium Nostoc commune. Appl Environ Microb. 2005;71(11):7327–7333. https://doi.org/10.1128/aem.71.11.7327-7333.2005
• 20. Deng Z, Hu Q, Lu F, Liu G, Hu Z. Colony development and physiological characterization of the edible blue-green alga, Nostoc sphaeroides (Nostocaceae, Cyanophyta). Prog Nat Sci. 2008;18(12):1475–1483. https://doi.org/https://doi.org/10.1016/j.pnsc.2008.03.031
• 21. Armstrong R, Bradwell T. Growth of crustose lichens: a review. Geografiska Annaler: Series A, Physical Geography. 2010;92(1):3–17. https://doi.org/10.1111/j.1468-0459.2010.00374.x
• 22. Wharton RA, Parker BC, Simmons GM. Distribution, species composition and morphology of algal mats in Antarctic dry valley lakes. Phycologia. 1983;22(4):355–365. https://doi.org/10.2216/i0031-8884-22-4-355.1
• 23. MacIntyre HL, Geider RJ, Miller DC. Microphytobenthos: the ecological role of the “secret garden” of unvegetated, shallow-water marine habitats. I. Distribution, abundance and primary production. Estuaries. 1996;19(2):186–201. https://doi.org/10.2307/1352224
• 24. de los Ríos A, Ascaso C, Wierzchos J, Fernández-Valiente E, Quesada A. Microstructural characterization of cyanobacterial mats from the McMurdo Ice Shelf, Antarctica. Appl Environ Microb. 2004;70(1):569–580. https://doi.org/10.1128/aem.70.1.569-580.2004
• 25. Komárek O, Komárek J. Diversity and ecology of cyanobacterial microflora of Antarctic seepage habitats: comparison of King George Island, Shetland Islands, and James Ross Island, NW Weddell Sea, Antarctica. In: Seckbach J, Oren A, editors. Microbial mats: modern and ancient microorganisms in stratified systems. Dordrecht: Springer; 2010. p. 515–539. https://doi.org/10.1007/978-90-481-3799-2_27
• 26. Taton A, Grubisic S, Brambilla E, de Wit R, Wilmotte A. Cyanobacterial diversity in natural and artificial microbial mats of Lake Fryxell (McMurdo Dry Valleys, Antarctica): a morphological and molecular approach. Appl Environ Microb. 2003;69(9):5157–5169. https://doi.org/10.1128/aem.69.9.5157-5169.2003
• 27. Revsbech NP, Jorgensen BB, Blackburn TH, Cohen Y. Microelectrode studies of the photosynthesis and O2, H2S, and pH profiles of a microbial mat. Limnol Oceanogr. 1983;28(6):1062–1074. https://doi.org/10.4319/lo.1983.28.6.1062
• 28. Glud RN, Kühl M, Kohls O, Ramsing NB. Heterogeneity of oxygen production and consumption in a photosynthetic microbial mat as studied by planar optodes. J Phycol. 1999;35(2):270–279. https://doi.org/10.1046/j.1529-8817.1999.3520270.x
• 29. Vopel K, Hawes I. Photosynthetic performance of benthic microbial mats in Lake Hoare, Antarctica. Limnol Oceanogr. 2006;51(4):1801–1812. https://doi.org/10.4319/lo.2006.51.4.1801
• 30. Secker NH., Chua JPS, Laurie RE, McNoe L, Guy PL, Orlovich DA, et al. Characterization of the cyanobacteria and associated bacterial community from an ephemeral wetland in New Zealand. J Phycol. 2016;52(5):761–773. https://doi.org/10.1111/jpy.12434
• 31. Komárek J, Kováčik L, Elster J, Komárek O. Cyanobacterial diversity of Petuniabukta, Billefjorden, central Spitzbergen. Pol Polar Res. 2012;33(4):347–368. https://doi.org/10.2478/v10183-012-0024-1
• 32. Nedbal L, Soukupová J, Kaftan D, Whitmarsh J, Trtílek M. Kinetic imaging of chlorophyll fluorescence using modulated light. Photosynth Res. 2000;66:3–12. https://doi.org/10.1023/A:1010729821876
• 33. Nedbal L, Whitmarsh J. Chlorophyll fluorescence imaging of leaves and fruits. In: Papageorgiou GC, Govindjee, editors. Chlorophyll a fluorescence. A signature of photosynthesis. Dordrecht: Springer; 2004. p. 389–407. (Advances in Photosynthesis and Respiration; vol 19). https://doi.org/10.1007/978-1-4020-3218-9_14
• 34. Schneider CA, Rasband WS, Eliceiri KW. NIH Image to ImageJ: 25 years of image analysis. Nat Method. 2012;9:671. https://doi.org/10.1038/nmeth.2089
• 35. Roháček K, Soukupová J, Barták M. Chlorophyll fluorescence: a wonderful tool to study plant physiology and plant stress. In: Schoefs B, editor. Plant cell compartments-selected topics. Kerala: Research Signpost; 2008. p. 41–104.
• 36. Oxborough K, Baker NR. Resolving chlorophyll a fluorescence images of photosynthetic efficiency into photochemical and non-photochemical components – calculation of qP and Fv'/Fm'; without measuring Fo'. Photosynth Res. 1997;54(2):135–142. https://doi.org/10.1023/A:1005936823310
• 37. Statsoft. Statistica, version 13. 2015 [cited 2018 Dec 29]. Available from: http://www.statsoft.com/Products/STATISTICA-Features
• 38. Vincent WF, Castenholz RW, Downes MT, Howard-Williams C. Antarctic cyanobacteria: light, nutrients, and photosynthesis in the microbial mat environment. J Phycol. 1993;29(6):745–755. https://doi.org/10.1111/j.0022-3646.1993.00745.x
• 39. Vincent WF, Downes MT, Castenholz RW, Howard-Williams C. Community structure and pigment organisation of cyanobacteria-dominated microbial mats in Antarctica. Eur J Phycol. 1993;28(4):213–221. https://doi.org/10.1080/09670269300650321
• 40. Hawes I, Howard-Williams C, Vincent W. Desiccation and recovery of antarctic cyanobacterial mats. Polar Biol. 1992;12(6):587–594. https://doi.org/10.1007/BF00236981
• 41. Potts M. Desiccation tolerance of prokaryotes. Microbiol Mol Biol Rev. 1994;58(4):755– 805.
• 42. Potts M. Mechanisms of desiccation tolerance in cyanobacteria. Eur J Phycol. 1999;34:319–328. https://doi.org/10.1080/09670269910001736382
• 43. Tashyreva D, Elster J. Annual cycles of two cyanobacterial mat communities in hydro-terrestrial habitats of the high Arctic. Microb Ecol. 2016;71(4):887–900. https://doi.org/10.1007/s00248-016-0732-x
• 44. Makhalanyane TP, Valverde A, Velázquez D, Gunnigle E, van Goethem MW, Quesada A, et al. Ecology and biogeochemistry of cyanobacteria in soils, permafrost, aquatic and cryptic polar habitats. Biodivers Conserv. 2015;24(4):819–840. https://doi.org/10.1007/s10531-015-0902-z
• 45. Deming JW, Young JN. The role of exopolysaccharides in microbial adaptation to cold habitats. In: Margesin R, editor. Psychrophiles: from biodiversity to biotechnology. Cham: Springer; 2017. p. 259–284. https://doi.org/10.1007/978-3-319-57057-0_12
• 46. Cockell CS, Knowland J. Ultraviolet radiation screening compounds. Biol Rev. 1999;79:311–345. https://doi.org/10.1017/S0006323199005356
• 47. Garcia-Pichel F, Castenholz RW. Occurrence of UV-absorbing, mycosporine-like compounds among cyanobacterial isolates and an estimate of their screening capacity. Appl Environ Microb. 1993;59(1):163–169.
• 48. Sinha RP, Klisch M, Gröniger A, Häder DP. Ultraviolet-absorbing/screening substances in cyanobacteria, phytoplankton and macroalgae. J Photochem Photobiol B. 1998;47(2– 3):83–94. https://doi.org/https://doi.org/10.1016/S1011-1344(98)00198-5
• 49. Dodds WK. Photosynthesis of two morphologies of Nostoc parmelioides (Cyanobacteria) as related to current velocities and diffusion patterns. J Phycol. 1989;25(2):258–262. https://doi.org/10.1111/j.1529-8817.1989.tb00121.x
• 50. Komárek J. Süßwasserflora von Mitteleuropa 19/3. Cyanoprokaryota. 3. Teil: Heterocytous genera. Heildelberg: Springer; 2013. https://doi.org/10.1007/978-3-8274-2737-3
• 51. Hrouzek P, Ventura S, Lukešová A, Mugnai MA, Turicchia S, Komárek J. Diversity of soil Nostoc strains: phylogenetic and phenotypic variability. Algol Stud. 2005;117(1):251–264. https://doi.org/10.1127/1864-1318/2005/0117-0251
• 52. Lukavský J. Controlled cultivation of algae on agar plates. Algol Stud. 1974;10:90–104.
• 53. Lukavský J. Analysis of growth rate of algae by cultivation on solid media. Algol Stud. 1975;14:105–136.
• 54. Kvíderová J, Elster J, Šimek M. In situ response of Nostoc commune s. l. colonies to desiccation in central Svalbard, Norwegian high Arctic. Fottea. 2011;11(1):87–97. https://doi.org/10.5507/fot.2011.009
• 55. Gao K, Qiu B, Xia J, Yu A, Li Y. Effect of wind speed on loss of water from Nostoc flagelliforme colonies. J Appl Phycol. 1998;10(1):55–58. https://doi.org/10.1023/A:1008020914486
• 56. Dodds WK, Gudder DA, Mollenhauer D. The ecology of Nostoc. J Phycol. 1995;31(1):2– 18. https://doi.org/10.1111/j.0022-3646.1995.00002.x
• 57. Fukuda S, Yamakawa R, Hirai M, Kashino Y, Koike H, Satoh K. Mechanisms to avoid photoinhibition in a desiccation-tolerant cyanobacterium, Nostoc commune. Plant Cell Physiol. 2008;49(3):488–492. https://doi.org/10.1093/pcp/pcn018
• 58. Poza-Carrión C, Fernández-Valiente E, Piñas FF, Leganés F. Acclimation of photosynthetic pigments and photosynthesis of the cyanobacterium Nostoc sp. strain UAM206 to combined fluctuations of irradiance, pH, and inorganic carbon availability. J Plant Physiol. 2001;158(11):1455–1461. https://doi.org/10.1078/0176-1617-00555

Identyfikatory

DOI

10.5586/asbp.3602