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Cyano- = pigment color in the blue-green part of the visible spectrum due to the presence of light-capturing molecules including chlorophyll a (green) as well as water-soluble 'phycobilins' phycoerithrin (red), phycocyanin and allophycocyanin (blue), and other accessory yellow pigments, predominantly β-carotene and zeaxanthin (Hirschberg and Chamovitz 2004). Color varies from nearly black to colorless.

Old Names:  Myxophyceae, Schizophyceae (schizo- = split), Schizomycetes and Myxobacteria (myxo- = slime) are retired names (Geitler 1932), as is Cyanophyceae used by Geitler himself (1932, 1979).  The name ‘blue-green algae’ appeared as recently as 1971 by Stanier et al..  Other retirees include 'Cyanophycota' (Friedmann 1982) and 'Blue-greens' (Fay 1983). 'Cyanophyta' and 'blue-green algae' appeared again Recently (van den Hoek et al. 2005; Whitton 2011) as equivalent to Cyanobacteria while explicitly describing their bacterial structure. Old names retire slowly.

Bacteria: Because most cyanobacteria are photosynthetic and often share the same habitat as eucaryotic photosynthetic protists ("algae"), they were originally classed as cyanophyceae, where “‑phyceae” in botanical nomenclature signifies eukaryotic “algae”. Roger Stanier (1916 - 1982) stated clearly (ca. 1970) that these photosynthetic bacteria should be classified according to the rules of procaryote nomenclature, published in later papers ( Stanier et al. 1971, Stanier 1977), hence their currently accepted name.  In bacterial classification only living cultures are recognized as “holotypes” capable of being distinguished, in contrast to the “type specimen” (dead; not cultures) required in the Botanical Code.  Thus revisions are occurring both at the generic and species level (Ripka et al. 1978).

Polymorphism:  Without molecular information the identity of species and in some case genera is doubtful.  Ecads, or morphotypes modified by the microhabitat, modify sizes, shapes and both intensity and color of pigmentation.  The viscosity of extracellular glycans (polysaccharides with various structure) changes the shape and orientation of coccoid cells as well as filaments.  “In no class have there been so many records of polymorphism as in the Cyanophyceae” (West and Fritsch 1927).

Photopigments vary in dominance and ratio, but generally include Chl a, phycocyanin, allophycocyanin, and phycoerithrin. As a result the apparent color also varies from red to green to cyan to blue, as well as brown. Such variation may be influenced by the light available in any particular microhabitat, such as in the microstratified layers of a thermocline -- chromatic adaptation.

Photosynthetic pigment absorbance spectra of cyanobacteria (euryaerobic - from anaerobes to aerobes), purple bacteria (microaerobes), and green bacteria (strict anaerobes).  © 2007 Sinaur Associates posted as slide 19 online.

Chromatic adaptation is the ability of cyanobacteria, especially of similar strains, to vary their pigment ratio, especially the accessory phycobilins (allophycocyanin, phycocyanin, and phycoerithrin.  A clear example is the change in ratio of phycocyanin to phycoerithrin in Synechoccus in chemostats, both in uniculture and mixed culture, when confronted with either green or red light. the red strain (rich in phycoerithrin) outcompeted the green strain in green light, and vice versa (Stomp et al.  2004). Effectively this reduces inter-strain competition, and increases diversity of strains by complementarity – absorbing different light spectra when confronted with light shading by one of the strains.  Other more remotely related strains such as Tolypothrix can also compensate for changes in available light, as found in mixed cultures with the red or green strain of Synechococcus (Ibid.).

Another example: While a graduate student at the University of Minnesota I kept a 500 ml capped bottle of metalimnetic Oscillatoria agardhii (now Planktothrix agardhii) in the laboratory, with only the overhead fluorescent fixtures as a light source, and without any disturbance such as nutrient addition. Within a few weeks I was astounded to see that it had turned red, and it would have been called O. rubescens. To this day I wonder whether there were a few scarce red filaments in the lakewater, or whether the green trichomes turned red. I searched for some half-green half-red trichomes but found none...and never saw red trichomes in the source -- Deming Lake, Itasca State Park, Minnesota (47o 10' 13.66 N, 95o 10' 07.18 W). Confirmation of this 'duality' is clear: Although allozymes (a.k.a. alloenzymes -- variant forms of an enzyme that are coded by different alleles at the same locus are identical, absorbance spectra of Oscillatoria rubescens (1) and O. agardhii (O. agardhii) from Komárek (2000).

No flagella are present, but some "gliding” or “bending (flexing)” motility occurs in several filamentous genera lacking dense outer sheaths associated with individual trichomes. Good examples include Oscillatoria (some having been renamed "Planktothrix"), Arthrospira, and Microcoleus (with a common sheath containing multiple trichomes not firmly fixed in the sheath). The motility can take the form of reversible translocation along the axis of the trichome, generally with clockwise or counterclockwise rotation. In other cases the motility is pendulum-like bending or flexing of the axis of the trichome. In Lyngbya, hormogones (short fragments containing a few cells) can be seen travelling along the inside of an empty sheath, eventually escaping to form their own filament.

Motility mechanism:  Direct observation of mucilage secretion (Hoiczyk and Baumeister 1998) during gliding motility of unbranched trichomes, including mucilage-secreting pores and mucilage trails, along with electron micrographs of the pores at trichome cell junctions, enabled a model in which helical proteins in the outer cyanobacterial wall control secretion of mucilage to propel the trichome reversibly, with a reversing rotation and translocation. How all the cells in the trichome are communicating to act synergically (to propel reversibly) is not explained. Also, trichome bending may require a different model. How external stimuli (light, temperature, solutes) are sensed by the motility system is another series of questions.


Figure 8 from Hoiczyk and Baumeister (1998).

Origin and Age: The oldest known fossils are from the Archaean Era, ~3.5 Ga (billion years), in stromatolites found in Western Australia and South Africa. Cyanobacteria along with other bacteria and archaea likely were the first life forms to inhabit Earth. Their ultimate origin may well be extraterrestrial given their extraordinary ability (at least in spore form) to resist extremes of low temperature, low moisture, and capacity (at least some genera) for both anaerobic and aerobic metabolism. Thus as a group they may be older than Earth itself, currently estimated to have accreted 4.5 Ga. Given ca. 1 billion years of cooling to a temperature tolerable to terrestrial life, and the relative complexity both morphologically and physiologically, it is not unreasonable to expect extraterrestrial procaryotes to land on Earth, and perhaps distant hospitable planets outside the Solar System, and develop successful communities.

Importance in Carbon Fixation:  Cyanobacteria currently and collectively account for >30% of global photosynthesis primarily as oceanic picoplankton (Rae et al.), slowing (mitigating) both the rise in atmospheric CO2 and rate of ocean acidification.

Importance in Oxygen Fixation:  The cyanobacteria evolved chlorophyll a from bacteriochlorophyll (Burke et al. 1993) or possibly before bacteriochlorophyll (Lockhart et al. 1996), enabling them to generate molecular oxygen by oxidizing the oxygen in water (rather than sulfide in the earlier anoxygenic photosynthesis) and excluding O2. The resultant oxygen-rich environment (atmosphere, hydrosphere, biosphere) led to the evolution of all aerobes. Many or most cyanobacteria retain the capacity to switch between anoxygenic and oxygenic photosynthesis.

Importance in Nitrogen Fixation:  Along with several other photosynthetic and heterotrophic bacteria, the cyanobacteria can reduce elemental nitrogen (unavailable biologically except in this way) to ammonia - 'nitrogen fixation', the source of nitrogen for amino acids and proteins. Grula (2005) suggests that N-fixation was essential to early cyanobacteria, and evolved either before or at the same time as synthesis of chlorophyll.

Toxins:

Secondary metabolites (not required for growth) include nitrogen (N)-rich substances that are most likely a reserve for N but at the same time are toxic to humans, other mammals and likely other organisms.  Physiologically they are known to degrade either mammalian livers (hepatotoxins) or nerve networks (neurotoxins).

Nearly all lakes have toxic cyanobacteria, even the most oligotrophic in New Hampshire USA also contain toxins such as microcystin, a liver toxin (Haney 19XX)

Evidence that the toxins have diminished production in extreme lakes with > 2000 µg total nitrogen (TN) is in Midwestern lakes in the USA (Graham et al. 2004).

                                

                                            From Graham et al. 2004, Figure 2a.

The lack of toxin production where N is in such excess reinforces the idea that they are primarily a N-storage mechanism.

   

Blooms:

A large concentration of biologically available phosphorus and limited concentrations of nitrate-N tend to favor high growth rates of N-fixing cyanobacteria with or without heterocysts, relative to other PS plankton.  Buoyant cyanobacteria accumulate at the water surface forming blooms.  Generally a period of rapid growth and population increase occurs well below the surface and beneath the impact of injurious UV radiation, and bloom formation can mark the end of the growth of a clone.  A series of several blooms may follow each other as subsequent clonal populations develop sequentially, ending their pulse by floating to the surface.

Wind can mix the blooms downward dispersing the cyanobacteria, or at lower velocity can concentrate the potentially toxic populations along the lee shores providing a danger to vertebrates.

Chemical control of cyanobacterial blooms in New Hampshire USA, at least until ~1980, included CuSO4 solution added to lake surfaces either by spraying or by dragging crystals in hemp sacks suspended from a power boat in transects across lakes.  Massive fish kills often followed copper treatments, largely from anoxia and toxin release as the cyanobacteria lysed.  Recognition that copper toxicity was not limited to the cyanobacteria, the temporary nature of the bloom control and high costs have made the application unpopular in the state.  An alternative chemical control is application of alum, a co-precipitate of Al2(SO4)3 and Na2Al2O4 (or related salts) that adsorbs phosphate from the water column and carries it to the sediments.  Expense, limited duration and potential aluminum toxicity limit its use.

Biological control may have at least a minimal impact on cyanobacterial growth prior to blooms.  Simocephalus vetulus, a large 2 – 4 mm long caldoceran, can grow on diets of some strains of Microcystis (but not all) and Oscillatoria (“Limnothrix”) sp. as well as on the green Scenedesmus acutus (Fernández et al. 2014).  Some ciliates and amoebae also consume cyanobacteria but are unlikely to prevent blooms.

Predators on cyanobacteria

The chrysophycean mixotrophic flagellate Ochromonas can feed on all Microcystis strains tested as well as on Pseudanabaena, and have a ‘strongly reduce’ Microcystis biomass and toxins”, thus potentially reducing severity of blooms, or at least reduce the frequency of small colonies (≤ 810 µm3) as demonstrated in culture.  Ochromonas and Microcystis co-occurred in 94% of 460 Norwegian lakes.  Apparently Ochromonas digests ingested cyanobacterial cells or even ingested dissolved microcystin (van Donk et al. 2009).  There is evidence that Ochromonas can be amoeboid, producing pseudopods to capture particles (Boenigk and Arndt  2000).

Natural populations of filamentous cyanobacteria such as Oscillatoria (a.k.a. Planktothrix) can also be decimated by ‘gulping’ ciliates such as Pseudomicrothorax dubius (Hausmann 2002).

Logically the best control is preventative:  Reduce phosphorus loading from the watershed.

 

References:

Boenigk, J. and H. Arndt 2000. Particle handling during interception feeding by four species of heterotrophic nanoflagellates. Journal of Eukaryotic Microbiology, 47, 350358.  

Burke, D. H., Hearst, J. E. & Sidow, A.  (1993).  Early evolution of photosynthesis:  clues from nitrogenase and chlorophyll iron proteins.  Proc. Natl. Acad. Sci. USA 90, 7134-7138.

Fay, P.  1983.  The Blue-greens. Edward Arnold, London.

Fernández, R., S. Nandini, S.S.S. Sarma and M.E. Castellanos-Paez  2014.  Effects of cyanobacteria, fish kairomones, and the presence of ostracods on the demography of Simocephalus vetulus (Cladocera).  Invertebrate Biology 133(4):371-380.

Friedman, I.  1982.  Cyanophycota. In: Synopsis and Classification of Living Organisms, Vol. I, Ed. S.P. Parker, pp. 45-52. McGraw-Hill, NY. [Imre Friedmann is the sole PhD student of Lothar Geitler.]

Geitler, L.  1932.  Cyanophyceae.  Johnson Reprint Corporation, NY, London.  [Cyanophyceae von Europe, a part of Rabenhorst’s Kryptogamen-Flora von Deutschland, Österreich und der Schweiz.]

Geitler, L.  1979.  Einige kritische Bemerkungen zur neuen zusammenfassenden Darstellung der Morphologie und Systematik der Cyanophyceen.  Plant Syst. Evol. 132:153-60.

Fernández, R., S. Nandini, S.S.S. Sarma and M.E. Castellanos-Paez  2014.  Effects of cyanobacteria, fish kairomones, and the presence of ostracods on the demography of Simocephalus vetulus (Cladocera).  Invertebrate Biology 133(4):371-380.

Graham, J.L., J.R. Jones, S.B. Jones, J.A. Downing and T.E. Clevenger  2004.  Environmental factors influencing microcystin distribution and concentration in the Midwestern United States.  Water Research 38:4395-4404. (online)

Grula, J.W. 2005. Evolution of photosynthesis and biospheric oxygenation contingent upon nigroten fixation? Internat. J. of Astrobiology 4:251-257. (online)

Hausmann, K. 2002.  Food acquisition, food ingestion and food digestion by protists.  Japanese Journal of Protozoology 35(2): 85 – 95.

Hirschberg, J., and D. Chamovitz  2004.  Carotenoids in cyanobacteria. Advances in Photosynthesis and Respiration 1:559-579.

Hoiczyk, E., and W. Baumeister  1998.  The junctional pore complex, a prokaryotic secretion organelle, is the molecula motor underlying gliding motility in cyanobacteria. Current Biology 8:1161-1168

Lockhart, P.J., A.W.D. Larkum, M.A. Steel, P.J. Waddell, and D. Penny  1996.  Evolution of chlorophyll and bacteriochlorophyll:  The problem of invariant sites in sequence analysis.  Proc. Natl. Acad. Sci. USA 93:1930-1934.

Matthews, R.A.  2016.  Volume 1, Cyanobacteria. Books and Monographs.  Book 6.

       http://cedar.wwu.edu/cedarbooks/6

Rae, B.D., B.M. Long, L.F. Whitehead, B. Förster, M.R. Badger, G.D.  2013.  Cyanobacterial carboxysomes: Microcompartments that facilitate CO2 fixation.  J. Mol. Microbiol. Biotechnol. 23: 300 - 307.

Rippka, R., J. Deruelles, J.B. Waterbury, M. Herdman and R.Y. Stanier  1978. Generic assignments, strain histories and properties of pure cultures of cyanobacteria. Journal of General Microbiology 111:1 - 61.

Stanier, R.Y., R. Kunisawa, M. Mandel, and G. Cohen-Bazire 1971.  Purification and properties of unicellular blue-green algae (Order Chroococcales).  Bacterial Rev. 35:171-205.

Stanier, R.Y. 1977. The position of the Cyanobacteria in the world of phototrophs. Carlsberg Res. Comm. 42:77-98.

Stomp, M., J. Huisman, F. de Jongh, A.J. Veraart, D. Gerla, M. Rijkeboer, B.W. Ibellings, U.I.A. Wollenzien, and L.J. Stal.  2004.  Adaptive divergence in pigment composition promotes phytoplankton biodiversity. J. Gen. Microbiol. 111:1 – 61

van den Hoek, C., D.G. Mann, and H.M. Jahns 1995. Algae: an introduction to phycology. Cambridge University Press (623 pp).

van Donk, E., S. Cerbin, S. Wilken, N.R. Helmsing, R. Ptacnik and A.M. Verschoor   2009.  The effect of a mixotrophic chrysophyte on toxic and colony-forming cyanobacteria.  Freshwater Biology 54:1843-1855.

Whitton, B.A. 2011. Phylum Cyanobacteria (Cyanophyta). In: John, D.M., B.A. Whitton, and A.J. Brook (Eds.) The Freshwater Algal Flora of the British Isles. (878 pp)

 

 

 

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