INTRODUCTION

Terrestrial ecosystems of Antarctica with sparse vegetation cover are mainly formed by microbial communities under the impact of extreme climatic conditions [13, 14, 57] and can be considered as the modern analogues of relict Precambrian biogeocenoses [6, 9, 14, 44–47]. The ice-free areas of Antarctica (oases) represent hot spots of biodiversity, where oxygenic phototrophic microorganisms (cyanobacteria and eukaryotic microalgae) often become the dominant component and the main producer of organic matter in aquatic and land algae–bacterial communities [4, 14, 49, 58]. They occur in the form of benthic and littoral mats in large lakes and shallow reservoirs of Antarctica, in the form of epi- and endolithic biofilms on rocky substrates in terrestrial ecotopes, and in the form of organic horizons in soils [10, 13, 14, 19, 48, 57, 59]. These horizons are often located under the layer of stones and gravels (stone pavement) in the so-called hypolithic (from Greek hypo—under and lithos—stone) ecological niches [14, 18, 40, 41]. Stone pavements are found in wet valleys of Antarctic oases or in rock basins. They protect microorganisms from unfavorable factors: strong wind, extreme temperature changes, aridity, and high levels of ultraviolet radiation. The landscapes of the East Antarctica oases were described in detail earlier [1, 13, 40]. They are assigned to snow-patch cryptogamic barrens [2], which are supplied with moisture during snow melting in spring and summer.

Some microbial communities in hypolithic horizons of Antarctic soils are formed with the significant participation of cyanobacteria [17, 22, 41, 43, 45, 55]. It is well-known that oxygenic phototrophic bacteria are the primary producers of organic matter, which is consumed by the heterotrophic part of microbial communities: chemoheterotrophic bacteria and micromycetes [13, 18, 44, 46, 47, 49]. According to published data, soil microbiota under extreme conditions of Antarctica is characterized by short food chains, which usually include representatives of Proteobacteria, Actinobacteria, Bacteroidetes, Acidobacteria, Gemmatimonadetes, Сyanobacteria, and Deinococcus-Thermus [4, 14, 19, 36, 45, 46, 49]. It has been revealed that their taxonomic composition depends not only on the physicochemical properties of soil, vegetation, geographical location, and climatic zone but also on the depth of the soil horizon [4, 6, 13]. The upper layers of Antarctic soils include representatives of chemoheterotrophs (Acidobacteria, Actinobacteria, Bacteroidetes, and Proteobacteria) together with phototrophic oxygenic (Cyanobacteria) and anoxygenic (Chloroflexi) bacteria [14, 33, 36]. At depths of 0.5–2 m, mainly Actinobacteria, Firmicutes, and Betaproteobacteria are found [4, 9, 14].

Through the activity of phototrophic bacteria, the hypolithic horizons of soils become an important source of available forms of organic carbon and nitrogen for the heterotrophic component of the microbiota, as well as for mosses, lichens, and some angiosperms occasionally found in Antarctic oases [10, 17]. The resulting organic matter is involved in biochemical weathering, formation of minerals, and aggregation of soil [13]. Cyanobacteria are a major contributor to soil-forming processes. However, the study of their biological diversity is not complete, as it is based either on the description of their morphological features in the natural environment [19, 44, 58–61], or on metabarcoding of the 16S rRNA gene [4, 33, 36, 46, 47]. No works focusing on the isolation of Antarctic cyanobacteria from soils into laboratory cultures and their description using a modern polyphasic approach have yet been published [15, 31]. The present paper is the first comprehensive study of Antarctic cyanobacteria in the hypolithic horizons of the Larsemann Hills, both in situ using the Rossi–Cholodny technique, light and confocal microscopy, or in vitro by the help of laboratory cultures isolated from soil samples.

OBJECTS AND METHODS

Sampling sites. In the Larsemann Hills oasis, we investigated hypolithic organic horizons of Cryosols (Arenic) formed in small and large accumulative landforms: (1) shallow depressions on rock outcrops filled with fine earth and gravel (rock baths) and (2) an intermountain wet valley (Fig. 1). A brief description of the research objects is given in Table 1. Detailed characterization of the sites, where the fouling glasses were installed by Dr. N.S. Mergelov, and a description of the experiment and soils according to WRB [24] have been published previously [3, 41]. The taxonomic composition of cyanobacteria in rock baths (sites 1 and 2) was studied both in soil samples, from which laboratory cultures were isolated, and on fouling glasses—sterile glass slides 75 × 25 mm, which were placed vertically directly under the soil surface to a depth equal to the glass width and exposed in hypolithic horizons during four years. The taxonomic composition of cyanobacteria in the valley bottom (site 3) was studied only in laboratory cultures isolated from samples of hypolithic horizons. Compared to the rock baths, the valley bottom is a wetter habitat due to the intensive influx of meltwater from a large snow patch and hypolithic organic-accumulative horizons of microbial genesis have a greater thickness here. The material of stone pavements in the valley is partially formed through the input of desquamation plates from nearby rock outcrops (gneisses and granites). Such plates (site 4) often contain epilithic and endolithic algae-bacterial biofilms [26], which may replenish the pool of bacteria of the stone pavement and the underlying hypolithic horizons. This object was also used in the work.

Fig. 1.
figure 1

Study objects: (a, b) location and sampling sites; (c) shallow depressions on rock outcrops (rock baths) (sites 1 and 2); (d) inter-mountain wet valley (sites 3 and 4); (e, f) Cryosols (Arenic) with hypolithic organic-accumulative horizons at sites 1 and 3, respectively.

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Table 1.   Brief description of the study objects
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Methods. Soil samples, parts of desquamation plates, and fouling glasses were transported frozen and then studied by means of light and confocal microscopy, fluorescent in situ hybridization (FISH) [7], and microbiological and molecular-genetic methods [50].

The study of the taxonomic composition of soil cyanobacteria in situ. Microbial fouling patterns were studied by the methods of light and confocal microscopy. Shape, size, and linear parameters of cells were described with the help of phase contrast microscopes DM2500 (Leica). On the surface of some glasses, the cyanobacterial fouling was unevenly arranged and sometimes found only in the upper part. In order to make them available for microscopy, the material was scraped off the glasses with a disposable blade, and such preparations were studied individually.

In addition to light microscopy, the biodiversity of soil cyanobacteria was visualized by hybridization of 16S rRNA of bacteria on the glasses with specific fluorescent oligonucleotide probes [5, 20]. The material on the glasses was fixed by a 2% solution of paraformaldehyde in a phosphate-buffered saline (PBS) containing 137 mM NaCl, 2.7 mM KCl, 1.8 mM KN2PO4, and 10 mM Na2HPO4 (pH 7.4) at 10°C for 2 h. Then, the fixed material was washed with PBS, dried for 30 min at 37°C, and dehydrated in a series of ethyl alcohols with increasing concentration (70, 80, 96, and 100%). The dried material was hybridized with oligonucleotide sequences specific for 16S rRNA: EUB338I (for visualization of eubacteria) and СYA361/СYA762 (for cyanobacteria detection) probes conjugated with fluorochrome dyes AlexaFluor©488, AlexaFluor©533, and AlexaFluor©750 [30, 37]. The hybridization was performed at 46°C in buffer containing 25 mM tris-HCl (pH 8.0), 50 mM NaCl, 0.01% SDS, 5 mM Na2-EDTA, and 35% formamide. After this, slides were washed with PBS and dried. Bacterial DNA was additionally stained by 0.1% solution of 4',6-diamidino-2-phenylindole (DAPI) for 1 h at 37°C. Then, the slides were studied under a Leica TCS-SP5 laser scanning confocal microscope (Leica Microsystems, Germany) in the “Chromas” Research Centre of St. Petersburg State University.

Strain isolation. Soil cyanobacteria were isolated into laboratory cultures using standard microbiological techniques [24, 50]. For this, samples were incubated in a BG11 liquid medium [14], under white lamps illumination at an intensity of 10–15 µM of photons m–2 × s–1 and a temperature of 22–24°C for 2–3 months. The resulting laboratory cultures were used for inoculation of isolates on 1% agarized medium BG11 with cycloheximide (at a final concentration of 50 µg/mL). The taxonomic status of the obtained strains was determined within the framework of the conventional polyphasic approach for the description of cyanobacteria [15, 31].

Molecular-genetic analysis of cyanobacterial strains. DNA preparations were obtained using the DNAeasy Pro Plant (Qiagen) kit according to the manufacturer protocol. 16S rRNA gene were amplified with ∼30 ng of DNA samples and universal bacterial primers 27F/1492R using a ScreenMix-HS kit (Evrogen) following the standard protocol (95°C, 5 min; 30 cycles: 95°C, 1 min, 55°C, 30 s, 72°C, 1.5 min; 72°C, 10 min) on a Bio-Rad T100 thermal cycler. Internal transcribed spacers (ITS) of the ribosomal operon were amplified with primers 322F/340R [23] under the same conditions. The obtained PCR fragments were tested by electrophoresis in 1% agarose gel, from which the amplicons were cut out and purified using a Nucleospin Clean-Up kit (Macherey Nagel). Sequencing was performed according to Sanger with the help of the Resource Centre of St. Petersburg State University ‘Development of Molecular and Cell Technologies’. The sequenced 16S rRNAs of cyanobacteria were compared with those previously deposited in the NCBI GenBank database (https://blast.ncbi.nlm.nih.gov) using the BLASTn algorithm.

Phylogenetic analysis of 16S rRNAs was performed by the Neighbor-Joining method, using the two-parameter Kimura model in the MEGA X program [34, 51, 53]. The reliability of the tree topologies was assessed by a bootstrap test (10 000 replications), and the evolutionary distances were calculated using the Maximum Composite Likelihood method. The phylogenetic tree presented in this paper includes 102 sequences of the cyanobacterial 16S rRNA gene. The conservative and variable regions of the internal transcribed spacer (ITS) of the ribosomal operon were analyzed as described earlier [23, 25]. Secondary structures of ITS were analyzed using the RNAstructure program [11]. The sequences of the 16S rRNA gene and 16S–23S ITS of the studied strains were deposited in the GenBank database (OQ079417–OQ079431 and OQ079504–OQ079517, respectively).

RESULTS AND DISCUSSION

Taxonomic composition of cyanobacteria in soil biofilms. The fouling glasses revealed mainly filamentous heterocysts and non-heterocystic cyanobacterial morphotypes (Fig. 2). In addition to cyanobacteria, eukaryotic microalgae (green, diatom, and desmid), as well as micromycetes, were found in microbial communities [3]. Confocal microscopy detected a large number of bacteria hybridized with eubacterial probes and DAPI (Fig. 2j), but the identification of their taxa was not the aim of our research, because the biological diversity of the heterotrophic part of soil microbiota of the Larsemann Hills has been studied in detail [4, 6, 35, 36, 38].

Fig. 2.
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Morphotypes of cyanobacteria in microbial communities determined on fouling glasses: Nostoc (a, b, f, k, l, m), Scytonema (n); Hassallia (a, g, o); Сalothrix (c, q); Microcoleus (h); Phormidium (p); Phormidesmis (d, r); Leptolyngbya (e, p); Gloeocapsa (j, s, t); and cf. Gloeocapsopsis (i, m). Light (a–e) and laser-confocal (f–j) microscopy, and their joint application (k–t). Fluorescent detection of cyanobacteria (red or yellow), eubacteria (green), and staining of DNA material and capsules (blue). The scale is 100 µm (a, e, p), 10 µm (s, t), and 20 µm (b–o, q, r).

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On the fouling glasses, filamentous cyanobacteria from the order Nostocales Cavalier-Smith predominated. In addition to vegetative cells, they had specialized cells—heterocysts (for nitrogen fixation) and akinetes (resting stages). Members of Nostocales often formed typical blue-green globular colonies, in which isopolar trichomes were loosely or densely packed. The trichomes consist of cylindrical or barrel-shaped vegetative cells contained intercalary or terminal rounded heterocysts and large barrel-shaped akinetes [30]. Morphotypes generally corresponding to the genus Nostoc Vaucher (family Nostocaceae Eichler) were detected on the fouling glasses (Figs. 2a, 2b, 2f, 2k–2m) [21, 30]. Among them, microscopic colonies of N. сf. сommune Vaucher and N. сf. punctiforme Hariot were regularly found. Macroscopic colonies typical for N. сf. pruniforme Agardh and N. сf. sphaericum Vaucher were somewhat less common [21, 30]. It is important to note that Nostoc species identification currently takes into account molecular-genetic features and life cycle observations in culture [30–32].

Apart from the Nostoc species, morphotypes of members Tolypotrichaceae Hauer et al. (some defined as Hassallia Berkeley) were also found on the glasses (Figs. 2a, 2g, 2o, 2p). Their false-branched heteropolar trichomes consisted of barrel-shaped or disc-shaped vegetative cells enclosed in dark brown-colored sheaths. Along with them, there were species of the genus Сalothrix Agardh ex Bornet & Flahault (family Сalothrichaceae Bennet & Murray) characterized by heteropolar trichomes with dense dark colored sheaths, basal heterocysts, and discoid vegetative cells in the trichome (Figs. 2c and 2q). Representatives of the genera Hassallia, Rexia Casamatta et al., and Coleodesmium Borzě ex Geitler were also encountered by microscopy, but they were difficult to distinguish on the basis of morphological characters observed in nature alone and had to be identified by molecular-phylogenetic analysis [21, 30, 31]. In addition to the aforementioned cyanobacteria, members of the genus Scytonema Agardh (family Scytonemataceae Rabenhorst, order Nostocales) [30] with false-branched heteropolar trichomes in dense blue-black sheaths were occasionally revealed on fouling glasses (Fig. 2n).

Non-heterocystous cyanobacteria with thin trichomes—Leptolyngbyа Anagnostidis & Komárek (Figs. 2e, 2p) and Phormidesmis Turicchia et al. (Figs. 2d, 2r) were often found on the fouling glasses together with Nostocales. Both mentioned taxa of filamentous cyanobacteria belong to the family Leptolyngbyаceae Komárek et al. (order Synechococcales Hoffmann et al.). They differ in the shape and size of cells and in the thickness of the sheaths (often dark-colored, as in Phormidesmis nigrescens Raabová et al.) [29, 31]. Molecular-genetic data are also required to clarify their taxonomic position [54, 56].

Images of biofilms on fouling glasses also revealed large trichomes of cyanobacteria from the order Oscillatoriales Schaffner: Microcoleus Desmaziéres (family Microcoleaceae Strunecký et al.), Phormidium Kützing and Lyngbya Agardh (family Oscillatoriaceae Engler) (Figs. 2h, 2p). They all differ in cell size and shape (discoid or isodiametric), presence/absence of calyptras in terminal cells, and the type of trichomes organization (single or in bundles) [21, 29, 31].

Unicellular morphotypes assigned to the order Сhroococcales von Wettstein were also found frequently on fouling glasses [26, 28]: Gloeocapsa Kützing (2–4 rounded cells surrounded by multilayered mucous sheaths) (Figs. 1j, 2s, 2t) and сf. Gloeocapsopsis Geitler (groups of rounded or polygonal cells assembled in irregular aggregates) (Figs. 2i, 2m).

To summarize, light and laser-confocal microscopy methods enabled us to detect in hypolithic horizons of the Larsemann Hills predominantly filamentous cyanobacteria of the genera Nostoc, Hassallia, Сalothrix, Scytonema, Microcoleus, Phormidium, Leptolyngbya, Phormidesmis and unicellular species—Gloeocapsа or сf. Gloeocapsopsis. As it is difficult to classify closely related cyanobacterial taxa on the basis of morphological characters alone, methods of molecular-genetic identification of laboratory strains were used [15, 31].

Analysis of the taxonomic position of cyanobacterial strains. Fifteen strains of Antarctic cyanobacteria isolated from soil samples are shown in Table 2. The species of filamentous cyanobacteria identified as Nostoc, Coleodesmium, Leptolyngbya, Plectolyngbya, and Phormidesmis (Fig. 3) comprise the core of our collection. The morphological characters and results of genetic identification of the strains are given in Table 2, and their phylogenetic positions are shown in Fig. 4.

Table 2.  Morphological description and the results of molecular-genetic identification of strains
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Fig. 3.
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Micrographs of strains of Antarctic soil cyanobacteria: (a) Nostoc sp. 15CТ-3.1; (b) Nostoc sp. PS27-2.2, PS27-1.2; (c) Halotia sp. PS27-3.2, P52-1.2, PS33-1, P52-3.2; (d) Coleodesmium sp. PS30-1.2; (e) strain S121; (f) Plectolyngbya hodgsonii PS24-1; (g) Leptolyngbya sp. 15ST-6, PS27-2.1; (h) Phormidesmis sp 62T-1; (i) strain P52 (I); and (j) strain PS30-1.1. The scale is 10 µm.

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Fig. 4.
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Phylogenetic position of soil strains of Antarctic cyanobacteria.

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According to the BLASTn 16S rRNA sequences comparison results, strains 15СT-6, PS27-2.1, 62T-1, and PS24-1 with thin non-heterocystous trichomes (morphotypes Leptolyngbya and Phormidesmis found on fouling glasses) are similar (99.6–99.8%) to Antarctic strains previously isolated from benthic mats of oasis lakes [54–56]. Analysis of the secondary D1-D1' structures (Fig. 5a) also confirms these findings: strains 15СT-6 and PS27-2.1 (Fig. 3g) are identical to Leptolyngbya sp. ANT.L52.1 [54]; strain 62T-1 (Fig. 3h) is similar to Phormidesmis priestleyi ANT.L61.2 [54]; and strain PS24-1 (Fig. 3f), to Plectolyngbya hodgsonii ANT.LPR2.2 [56].

Fig. 5.
figure 5

Secondary structures of 16S–23S ITS: (a) domain D1–D1' and (b) B-box.

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The primary 16S rRNA sequences of strains PS30-1.2 (Fig. 3d) and Coleodesmium sp. ANT.L52B.5 show 99.8% similarity. The latter strain was previously found in benthic mats of the Larsemann Hills oasis [54]. The analysis of secondary ITS structures (loop D1–D1' and the B-box anti-termination region) also confirms their identity (Figs. 5a, 5b). The phylogenetic position and molecular-genetic features of strains PS30-1.2 and Coleodesmium sp. ANT.L52B.5 indicate that they do not belong to the genus Coleodesmium (because their secondary ITS structures differ from those of the type species Coleodesmium wrangelii Borzì ex Geitler; Figs. 5a, 5b) or to the genera Rexia and Hassallia (Fig. 4). This suggests that strains PS30-1.2 and Coleodesmium sp. ANT.L52B.5 are new and apparently endemic species of cyanobacteria, which occur both in soils and in benthic mats of the Larsemann Hills.

The 16S rRNA gene of strain PS30-1.1 (Fig. 3j) was found to be more than 98% homologous to sequences of uncultured cyanobacteria, clustered to the genus Stenomitos Miscoe & Johansen (Leptolyngbyaceae) (Fig. 4). However, the similarity of strain PS30-1.1 with Stenomitos species is only 96%. Taking into account that the accepted taxonomic similarity threshold for 16S rRNA is 98.65% (for a new species) and 97.5% (for a new genus) [27], strain PS30-1.1 can be considered as a potentially endemic species of filamentous cyanobacteria, which was isolated in laboratory culture for the first time.

Strain P52 is also assigned to filamentous cyanobacteria of the family Leptolyngbyaceae by the morphotype (Fig. 3i). According to 16S rRNA analysis, it is 97% similar to sequences from the cyanobacteria of the genus Myxacoris Pietrasiak & Johansen [39]. However, strain P52 is located in the sister clade to the genus Myxacoris on the phylogenetic tree (Fig. 4). Moreover, P52 differs from species of the genus Myxacoris in its secondary structures of ITS (Figs. 5a, 5b). Our results allow us to consider strain P52 as a new and probably endemic taxon of Antarctic soil cyanobacteria.

Three strains (15ST-3.1, PS27-2.2, and 15ST-1.2) of Nostoc spp. were isolated in laboratory cultures (Figs. 3a and 3b). They are more than 98% similar to 16S rRNA of Nostoc commune, which is widespread in soils around the world [8]. At the same time, the 16S rRNAs of heterocystous strains PS27-3.2, PS33-1, P52-1.2 and P52-3.2 (Fig. 3c) were up to 97–98% similar to the recently described cyanobacterium Halotia [Genuário et al., 2015]. However, these strains are located on the phylogenetic tree separately from the sequences of other Halotia spp. (Fig. 4). The comparative analysis of ITS regions also confirms their difference (Figs. 5a and 5b). It can be assumed that our strains are a new species of the genus Halotia.

Endolithic unicellular cyanobacteria were found under the surface of the desquamation plate as blue-green patches (Table 1, site 4). According to 16S rRNA analysis, strain S121 was the most similar to uncultured cyanobacteria found in the Atacama Desert [35] and in another arid habitats. Morphologically, strain S121 is represented by 4–8 cells grouped in irregular aggregates (Fig. 3e). The closest sequenced homologues are unicellular cyanobacteria Aphanocapsa sp CCNUW2 and Pseudoacaryochloris sahariense PLM 132 (a lithobiont found in the Sahara Desert) [39], but they have only ~93% similarity. Strain S121 is phylogenetically distant from them and differs greatly in its morphology. On the phylogenetic tree, strain S121 primarily clusters with uncultured cyanobacteria (Fig. 4). The combination of morphological and molecular-genetic features indicates that strain S121 is a new taxon firstly described as endolithic unicellular Antarctic cyanobacteria.

CONCLUSIONS

Microbial complexes with cyanobacteria are present on almost all fouling glasses exposed in the soil [3], but their development and occurrence vary depending on the location of glass exposure (ecotope), material visualization (type of microscopy applied), and the study approach (microscopic or culture techniques). Standard methods of light microscopy without differentiated coloration and/or visualization of bacterial material with the use of confocal microscopy and FISH (with additional staining by DAPI) do not always detect all taxa of soil microorganisms, including cyanobacteria, in the samples. Therefore, a study of fouling glasses by various microscopic methods, as well as of soil samples, from which strains were isolated and analyzed, has been performed in order to find out the true taxonomic composition of cyanobacteria in the hypolithic horizons of the Larsemann Hills. The obtained data on the biodiversity of Antarctic cyanobacteria detected on fouling glasses are supported by the results of identification of strains in our collection. Clean mineral surfaces are obviously colonized by microorganisms (sterile fouling glasses model) in all previously studied ecotopes [3] that had sufficient moisture supply and clear evidence of soil formation [61].

A comprehensive description of the biological diversity of cyanobacteria in the hypolithic soil horizons of the Larsemann Hills is given for the first time. The dominant taxa in them have been studied on the basis of the described strains. As a result, a unique collection of Antarctic soil cyanobacteria has been formed in Russia. The cyanobacteria are characterized by a set of morphological and molecular genetic features, and their taxonomic status generally confirms the results of in situ detection on fouling glasses: filamentous representatives of the order Nostocales (strains Nostoc and Halotia) and the order Synechococcales (strains Phormidesmis, Plectolyngbya, and Leptolyngbya) predominate in the hypolithic horizons of the Larsemann Hills. The results of the analysis of the primary sequences of 16S rRNA gene and the features of ITS secondary structures indicate that strains PS30-1.1, S121, PS33-1, P52-3.2, PS27-3.2, P52-1.2, and P52 are isolated in cultures for the first time and are obviously new, previously not described, and potentially endemic taxa of Antarctic cyanobacteria. According to available data, from 21 to 57% of the 16S rRNA gene sequences of cyanobacteria found in Antarctica and deposited in GenBank are endemic [33, 59]. These are Aphanocapsa cf. holastica, Aphanocapsa cf. hyalina and Arthronema sp., Geitlerinema deflexum, Leptolyngbya antarctica, Oscillatoria subproboscidea, Plectolyngbya hodgsonii, Phormidesmis priestleyi, and Phormidium pseudopriestleyi. Endemic taxa of cyanobacteria are found in Antarctic habitats along with cosmopolitan species (for example, Nostoc commune or Oscillatoria sp.) [8, 22, 32, 54–60].

The ability of cyanobacteria to spread beyond water spaces has been repeatedly described earlier [43, 44, 46, 48, 52, 54, 56, 60, 61]. An important factor in this case is the level of wetness of their habitats. For instance, in the Dry Valleys of Antarctica, the soils of the Beacon Valley with no substantial water reservoirs are generally poorer in biodiversity of cyanobacteria than soils of the Miers Valley that has lake and ponds [43, 60]. In the present study, the taxonomic composition of cyanobacteria in Antarctic soil microbial communities also changes qualitatively and becomes depleted in the drier ecotopes [3]. High level of similarity of 16S rRNA gene sequences of some strains in our collection (62T-1, PS24-1, PS27-2.1, 15ST-6, and PS30-1.2) with the strains previously found in benthic mats of water reservoirs in the Larsemann Hills (Phormidesmis priestleyi ANT.L61.2, Plectolyngbya hodgsonii ANT.LPR2.2, Coleodesmium sp. ANT.L52B.5, and Leptolyngbya sp. ANT.L52.1) enables us to consider them as amphibious species of cyanobacteria [46, 47, 49, 54–60].

According to observations, many biotopes in the oasis feed on meltwater and may pass through the subaqueous stage in addition to the subaerial one, i.e., they are characterized by features of amphibious landscapes. The duration and intensity of the meltwater influx are very different. For example, a small snow patch in a rock bath (site 2) melts quickly at the beginning of the warm season, which causes a rapid development of cyanobacteria in microbial community of small puddles or upper soil horizons, which subsequently dry up. On the contrary, a large snowfield in the valley bottom (site 3) is able to provide stable influx and even flooding by meltwater during the entire period with temperatures above zero. Strong winds, which redistribute fine earth particles covered by bacterial biofilms within the oasis and beyond it, may also make the taxonomic composition of adjacent biotopes more uniform (and thereby provide a wide representation of amphibious species). A significant reduction (to 1–2 species) of the cyanobacterial taxonomic diversity is observed in the driest (site 1) and endolithic (site 4) habitats.

Thus, the greatest taxonomic diversity of cyanobacteria has been found in benthic and littoral mats of Antarctic water bodies with representatives of the genera Oscillatoria, Phormidium, Leptolyngbya, Nostoc, Geitlerinema, Lyngbya, Pseudanabaena, Hydrocoryne, Nodularia, Schizothrix, Microcoleus, Tychonema, Aphanocapsa, and Gloeocapsa, as well as species Lyngbya antarctica, Plectolyngbya hodgsonii, and Leptolyngbya antarctica [3, 43, 45, 56–58]. Most of them, however, are also found in soil communities [3, 32, 44, 46, 48, 58–61]. Our results indicate the ability of Antarctic cyanobacteria from aquatic habitats to inhabit both edaphic and lithic niches due to the metabolism plasticity, which helps to adapt to conditions of limited available water and to environmental extremes in general.