Microbial community composition of Tirez lagoon (Spain), a highly sulfated athalassohaline environment

Physico-chemical characterization

Enumeration of cells in water and sediment

The total DAPI cell count in the water column from the rainy season was 2.6×10⁸ ± 0.8x10⁸ cells/ml. At high ionic strength cell numbers dropped one order of magnitude. In the sediments the maximum cell counts were detected at a depth of 10 cm and corresponded to 4×10⁷ ± 0.4×10⁷ cells/g of wet sediment. At deeper zones cell numbers decreased drastically reaching values of 8×10³ ± 0.8x10³ cells/g of wet sediment.

Microscopy observation of water samples

Isolation from enrichment cultures

Analysis of uncultivated microbes

16S rRNA gene cloning from water samples

More than half of the clones obtained from the bacterial library of 16S rRNA gene sequences from water column samples of the rainy season had sequences similar to Alphaproteobacteria (Figure 3). Most of the clones were affiliated with Loktanella vestfoldensis and aggregated in one OTU with 12 phylotypes (a posterior probability (pp) of 100% supported this clade). The remaining phylotypes, Rw_cb_4 and Rw_cb_6, formed a cluster with Roseobacteraceae sequences (100% pp) (Figure 3). The phylotype Rw_cb_9 was affiliated with the uncultured bacterium clone SN26 (JQ824918) in the Methylophaga clade of the Gammaproteobacteria (100% pp). The phylotype Rw_cb_46 did not clustered at species level within Gammaproteobacteria by phylogenetic analysis, this view was confirmed with BLASTN and Classifier tools since it presented a similarity of 91% with the uncultured bacterium clone SN26 (JQ824918) obtained from an haloalkaline soil (Table 1).

Also, four clones retrieved from the low salinity water column were affiliated with Bacteroidetes (genus Psychroflexus and uncultured bacterium clones) and Actinobacteria phyla (uncultured actinobacterium clone).

In the archaeal clone library from the 33% salinity brine a substantial proportion of clone sequences combined three OTUs clustering with the Halorubrum genus (Figure 4), with a pp of 100%. One exception was the Dw_ca_59 clone, which was affiliated to an Uncultured euryarchaeote clone SFH1E051 (FN391283) (Figure 4).

Although DNA was successfully extracted from water samples with low salinity, attempts to obtain amplified products with archaeal primers were unsuccessful. This suggests that there were undetectable levels of halophilic archaea in these conditions, probably due to their low tolerance to low salinity concentrations [28].

DGGE analysis of water samples

The genomic DNA from the rainy season water column samples was amplified with the bacterial primer pair 341fGC-907r and resolved by DGGE using a denaturing gradient of 30-70% (Figure 5). The band pattern was reproducible and the most prominent bands were sequenced, showing high similarity with members of Flavobacteriaceae and an uncultured Actinomycetales organism (Table 2).

DGGE analysis of sediment samples

Band patterns were obtained from a denaturing gradient of 40-70% for sediments from rainy and dry season (Figure 5). From a total of 15 identified bacterial bands from the rainy season, five phylotypes belonged to the Flavobacteriaceae. Also members of the Bacillaceae were detected. Of the Gammaproteobacteria the frequently observed members of the genus Halomonas, harbouring only halophiles, members of the sulfur oxidizing bacteria Thioalkalimicrobium and the heterotroph Marinobacter were identified. One band was identified by the tool Classifier as a Desulfohalobiaceae member a sulfate reducing halophilic group of the Deltaproteobacteria class (Table 2).

A total of seven bands were obtained from the dry season sediment sample. Their sequences indicated phylotypes closely related with members of the phylum Bacteroidetes (Muricauda flavescens, unidentified Bacteroidetes) and an unidentified Sphingobacteria. Of the Gammaproteobacteria, members of the halophilic genus Halomonas were identified and the band Rs_db_29 was related with the Clostridiaceae.

A DGGE pattern was obtained using a 40-70% denaturing gradient of the amplification product of sediment samples using the archaeal primers 344f and 915rGC (Figure 6). The partial 16S rRNA sequence of the main bands allowed us to identify members of the strict halophilic genera Halobacterium and Natronomonas in samples from both types of sediments, and a species of the genus Archaeoglobus in sediments from the wet season (Table 2).

Physico-chemical conditions

The circumneutral pH of the Tirez lagoon could be the result of the low concentration of carbonates in relation to the concentration of calcium. The difference in pH between the water column and the sediments is probably due to the difference in the ionic concentrations between both phases. In summer, the relative abundance of the most numerous cations of water was as follows: Na+K>Mg>Ca therefore, a low quantity of CO₃^2- is sequestered by carbonate precipitation, which leads to a slightly alkaline pH (Table 1). On the contrary, in the sediments the bivalent cations dominate precipitating the CO₃^2- and resulting in a neutral pH.

The ratio of Cl^- to SO₄^2- underlines the athalassohaline characteristic of the Tirez lagoon. The relationship between chloride and sulfate increases from the rainy to the dry season as a consequence of the sequestration of sulfate in different salts (i.e. epsomite, mirabilite, thenardite, hexahydrite and bloedite) that precipitate before halite does.

Methanogens make a living in habitats where electron acceptors such as O₂, NO₃^-, Fe³⁺, and SO₄^2- are limiting [30]. Under the sulfated conditions prevailing in Tirez methanogens might be excluded. On the other hand, the negative redox potential values detected in the sediments (Table 1), especially in the rainy season, are not low enough to allow methanogenesis to proceed, being necessary a redox potential lower than -330 mV [31]. Such conditions are enough to justify the lack of evidence of methanogenic archaea by 16S rRNA gene sequencing. However, hydrogenotrophic methanogens have been reported in Tirez lagoon [29].

The negative redox potential values detected in the sediments (Table 1), especially in the rainy season, are not low enough to allow methanogenesis to proceed, being necessary a redox potential lower than -330 mV [31]. This result strongly suggests that measured redox potentials might be a gross estimate of the sediment potentials and that microniches with appropriate physico-chemical conditions must develop in those sediments to facilitate the growth of these strict anaerobic microorganisms.

Microbial diversity

After comparing the class richness among three domains it was evident that Bacteria was the richest domain given that bacteria classes represented 50% of all classes detected. Proteobacteria was the most common phylum and accounted for the largest OTU fraction (31%) in all the phases analyzed. Thus, in Tirez, members of the Proteobacteria class were the best-represented group of Bacteria in the range of salinities studied, diverging from other characterized athalassohaline systems where the dominant role was played by members of the Bacteroidetes group [15, 32]. Frequently, Salinibacter ruber is recognized as a common and abundant bacterium inhabitant of hypersaline environments, including the athalassohaline lake Chaka [32]. However, this microorganism has not been detected in Tirez lagoon in the different seasons and phases analyzed, which suggests a more complex distribution of this species [33].

Water

Most of the oxygenic photosynthetic organisms were detected and identified by morphotype analysis. The Chlorophyta and the Cyanobacteria detected are frequently found in hypersaline environments. These microorganisms play an important role in the global carbon and mineral cycles of hypersaline environments [34]. During the dry season, water evaporates promoting massive salt precipitation. However, a thin 3 mm layer of cyanobacteria could be observed between the sediment and the salt crust. Although competition with macrophytes is practically nonexistent at these hypersaline conditions, the cyanobacterial layer becomes thin and ephemeral. One possible explanation proposed by Guerrero and de Wit is that thick cohesive mats are usually associated with permanently covered sediments, in contrast to the thin mats that are temporally submerged, as in our case [35]. Another possible explanation is the Tirez fine sediment granulometry, dominated by particles of ≤0.002 mm, which have been described as the possible cause for the generation of thinner mats [36].

Though a prevalence of Loktanella vestfoldensis sequences among clone libraries has not been reported in other hypersaline systems, Loktanella sp. has been found in cold saline groundwater springs (Axel Heiberg Island, Canada) rich in sulfate [37]. It is remarkably that the genera Marinobacter and Halomonas have not been reported in other hypersaline systems rich in sulfate.

The presence of Woloszynskia cincta in the wet season seems to be a peculiarity of this environment since there is no report of this species in hypersaline systems. Indeed, there are few reports of dinoflagellates in hypersaline environments [38]. W. sincta is absent at high osmolarity conditions, it is possible to explain its survival until the next season by cyst formation [39]. On the other hand, the algae Dunaliella sp. is present in dry and flooded seasons. It has been shown that Dunaliella tertiolecta growth is inhibited by Mg²⁺ salts when compared with Na⁺ salts [40]. This response is of interest because Mg²⁺ is present in high concentrations in Tirez (Figure 1). However, the proliferation of Dunaliella sp. in Tirez lagoon is not entirely surprising since this genus has been reported in the Dead Sea [41] under high concentrations of Mg²⁺. Fuji [40] argues that the ability of D. tertiolecta to grow in a MgSO₄ medium may be related to the high intracellular concentration of SO₄^2-. Therefore, the high sulfate concentration characteristic of Tirez might have positive consequences on Dunaliella sp. growth despite the high concentrations of bivalent cations.

Sediments

Most ecological studies on hypersaline ecosystems are focused on the aqueous phase. Therefore, the following data about bacteria and archaea present in sediments is remarkable in terms of diversity of halophiles.

Firstly, 16S rRNA gene sequence techniques showed a predominance of Haloarchaea, particularly members of the Halobacteriaceae, which can grow well heterotrophically in the dark [42]. Most of the identified bacteria are described as heterotrophs, with Gammaproteobacteria being the dominant taxa followed by members of the Flavobacteria class.

The thermophilic sulfate reducing Archaea Archaeoglobus was detected in Tirez sediments. Interestingly, there are previous evidences of thermophilic genus (Thermoplasmatales) described in other hypersaline environments [14, 43, 44]. To understand this ecological singularity, a contingent adaptation of thermophilic microorganisms to the high osmolarity conditions founded in Tirez should be considered. Thermophilic microorganisms succeeded in stabilizing intracellular macromolecules by the synthesis and/or accumulation of compatible solutes [45, 46]. This physiological adaptation is an absolute requirement for organisms under osmotic stress [47]. In fact, the synthesis of compatible solutes has been reported in species of Archaeoglobus[48].

Another interesting case was Desulfohalobium, a halophilic sulfate reducing bacterium that was identified by PCR-DGGE. Although sulfate reducing bacteria have been detected previously in thalassohaline environments [49], this is the first report of this genus in an athalassohaline sulfate-rich ecosystem.

Microorganisms of the functional groups methanogens, sulfur oxidizers and sulfate reducers have been detected in sediment and by molecular biology techniques and enrichment [29]. Interestingly, methanogenic archaea and sulfur oxidizing bacteria were undetected by PCR-cloning of the gene 16S rRNA. Moreover, the sulfate reducing bacteria here detected by 16S rRNA (Desulfohalobium sp.) does not coincide with those encountered with the functional gene marker Apr, i.e. Desulfonema and Desulfonatronovibrio[29]. This inconsistence between gene markers has been reported previously [50].

Sampling

Samples were taken in triplicate on February and July 2005. Water and sediments were obtained from the same area of the lake. Water samples were taken aprox. 10 cm above sediments using sterile 50 mL Falcon tubes and kept at 4°C (for 4 h) until processed. In the dry season 50 mL sterile syringes were used for brine collection. For core extraction a Ring Kit core-sampler (Eijkelkamp Agrisearch equipment, The Netherlands) for soft soil was used. The sampler was inserted down to 40 cm, the core was kept at 4°C until further processing. Samples were used to inoculate cultures in triplicate and for molecular analysis

Physico-chemical characterization

For water samples, in situ temperature, pH, Eh and dissolved oxygen were measured using a multi-parametric probe (YSI 556 MPS, YSI Environmental). Eh and pH along the sediment cores were measured with a probe connected to a potentiometer Orion Model 290A+Thermo Orion (Thermo Fisher Scientific) calibrated at high ionic strength using equivalent Na₂SO₄ solutions, and dissolved oxygen and temperature with a Syland TM model Simplair. Elemental analysis was performed by TXRF (Extra-II) and ICP-MS (ELAN-6000 PE-Sciex) instruments and ionic chromatography with an IC Dionex DX-600 apparatus. Ion data from Tirez lagoon and other saline systems were used to build a ternary diagram using the software ProSim Ternary Diagram (ProSim, France).

Microscopic examination and cell enumeration

Samples were fixed with formaldehyde at a final concentration of 4% (v/v). The identification of algae, Cyanobacteria, Arthropoda, Rotifera and Fungi was carried out by microscopic observation of fixed samples using a Zeiss Axiovert 200M microscope coupled to a CCD camera.

To quantify cell numbers, preparations were stained with 4′, 6′-diamino-2-phenylindole (DAPI), Molecular Probes (Invitrogen), as previously described [51] and counted under a Zeiss Axiovert 200M microscope.

Microorganism isolation and culture

Sediment samples were dispersed in 1× PBS (0.1 M NaCl, 2 mM KCl, 4 mM Na₂HPO₄, pH 7.4) and the suspension used to inoculate different media plates. Each plate was inoculated with 100 μl of sediment slurry. For isolation of Cyanobacteria, BG11 medium plates [52] were inoculated and incubated at 20°C for up to 3–4 weeks under a 16:8 light:dark cycle at 150 mmol photons m²s^-1 irradiance and a temperature of 19°C. Fungi were isolated in PDA medium (potato-dextrose-agar) containing 0.050 mg/mL streptomycin and 0.1 mg/mL ampicillin and incubated at 30°C. Heterotrophic microorganisms were isolated on marine agar (Difco, Marine Broth 2216) and media for halophilic strains prepared with salts obtained from water of Tirez lagoon (rainy season) and crystallized with vacuum at room temperature. Salt composition was determined by ionic chromatography with an IC Dionex DX-600 apparatus. Ionic composition was as follows (ppm): Na⁺-K⁺ (8140), Ca²⁺ (1091), Mg²⁺ (4602), NH4⁺ (53.6), Cl^- (258) and SO₄^2- (6695). The crystallized salts were dissolved at a final concentration of 10, 20 or 30% (w/v). Dissolved salts were enriched with yeast extract (<0.5 g/L) and glycerol (<0.5 g/L) as reported by Bolhuis et al. [53]. Representative individual colonies from each medium were reinoculated in the same growth condition. All plate isolates were transferred to liquid media to obtain enough biomass to allow DNA extraction for molecular analysis. Sulfate-reducing bacteria were grown in anaerobic SRB medium modified from Raskin et al. [54] and supplemented with 500 mg/L L-cysteine, as reductive agent, and the following organic substrates: 250 mg/L yeast extract, 770 mg/L glutamic acid, 15 mg/L glycine, 250 mg/L peptone, 14 mM methanol and 27 mM methylamine.

Molecular methods

PCR conditions for rRNA gene amplification

Amplifications were performed using a Thermal Cycler 2720 (Applied Biosystems) in a final volume of 50 μL, each containing: 1 mM of dNTP, 3 mM MgCl₂, 1 mM of each primer, 1× PCR buffer and 0.025 u/μL Taq DNA Polimerase (AmpliTaq DNA Polymerase, Roche Molecular Systems). DNA was added in a volume of 3 μL, containing about 1–5 ng of template. Bacterial 16S rDNA was amplified using primers 27f [56] and 1492mr (5′-TACGGYTACCTTGTTACGACTT-3′) modified from [57] (annealing 57°C; 30 cycles). The 25f [58] and 1492mr primers were used for Archaea domain (52°C; 27 cycles). Both 16S rDNA amplification procedures consisted of initial denaturation (94°C for 10 min) followed by the above-indicated number of cycles of denaturation (94°C for 1 min), annealing (at the temperatures indicated above for 1 min) and extension (72°C for 3 min) followed by a final cycle of extension (72°C for 10 min). PCR amplification of 18S rDNAs were performed using the primer pair Euk1Af-Euka516 as described in [59].

For dinoflagellates an 18S rDNA fragment was amplified using dinoflagellate-specific primers Dino18SF1m (5′-AAGGGTTGTGTTTATTAGNTACAGAAC-3′) modified from [60] and 18ScomR1 [61]. The reaction was performed with an initial denaturation (94°C for 5 min), followed by 30 cycles of denaturation at 94°C for 1 min, annealing at 56°C for 1 min, and extension at 72°C for 3 min.

PCR amplifications for DGGE analysis of rRNA gene fragments were performed for Bacteria using the primers 341fGC [62] and 907r [56] while for Archaea the region between the primers 344fGC [62] and 915r [63] was used. Amplification conditions for Bacteria were the following: 94°C for 7 min, 35 cycles of 94°C for 45 s, 49°C for 45 s and 72°C for 1.5 min, and a final extension of 72°C for 10 min. For Archaea primers: 94°C for 5 min, 32 cycles of 94°C for 30 s, 54°C for 1 min and 72°C for 1 min, and a final extension of 72°C for 10 min. A 16S rRNA Deltaproteobacteria-specific region was amplified as described in [50] using the 385fGC-907r primer pair. Functional gene primers used for detection of sulfate reducing and methanogenic activities are described in [29]. In PCR reactions GC was equivalent to a 40 bp GC clamp at the 5′ end to prevent complete melting of the DNA fragments.

Cloning of 16S rRNA

PCR amplified products were cloned using the TOPO TA Cloning kit (Invitrogen Corporation, California) according to the manufacturer’s indications. From each clone library putative positive transformants were randomly sampled to perform minipreps according to standard alkaline lysis protocols.

Denaturing gradient gel electrophoresis

DGGE analysis of PCR-amplified 16S rRNA gene fragments using a 30-70%, 40-70%, 40-70% and 30-50% gradients was performed as described by Muyzer et al. [64] using a D-Code Universal Detection System (BioRad Laboratories). PCR samples were loaded onto 8% (w/v) polyacrylamide gels in 1× TAE buffer (20 mM Tris, 10 mM acetate, 0.5 mM Na-EDTA, pH 7.4). Electrophoresis was carried out at 60°C, at a constant voltage of 200 V for 4.5 h. After electrophoresis, the gel was stained for 15 min with ethidium bromide (0.5 μg/mL), rinsed in distilled water for 30 min and photographed with a Polaroid Kodak digital 16 camera. DGGE bands were excised from the gel under UV light and eluted in 50 μl of milliQ water overnight at 4°C. An aliquot of 3 μL was taken from each eluted sample and re-amplified by PCR in the conditions described above. The primers used for re-amplifications were the corresponding ones used in the first amplification but without the tailing sequence.

Sequence analysis

PCR products from DGGE gel bands and plasmid DNAs containing inserts were sequenced with the primers used for amplification and the pair M13F/M13R, respectively using an ABI PRISM Big Dye Terminator Cycle Sequencing Ready Reaction Kit (ABI) and an Applied Biosystem ABI 310 (PE Applied Biosystems, Foster City, California, USA) automated sequencer. Chromatograms were transformed into contiguous sequences combining FinchTV (http://www.geospiza.com/finchtv) and GeneDoc (http://www.nrbsc.org) tools. Chimeric sequences were identified by using Mallard[65]. The 16S and 18S rRNA sequences obtained from DGGE bands, culture isolates and clones were collapsed into OTUs by similarity analysis using FastGroup II (http://fastgroup.sdsu.edu). OTUs were compared with those available in GenBank (NCBI) and Ribosomal Database Project (RDBP) to identify them using the Basic Local Aligment Search Tool Nucleotide (BLASTN) and Classifier algorithms, respectively. Similarity analysis was performed with FastGroup II (http://fastgroup.sdsu.edu)

Phylogenetic analysis

The 16S rRNA gene sequences with a length ≥ 1300 bp were aligned with representative ones published in Bergey’s Manual [66] using ClustalX[67] using default parameters. Alignments were optimized manually using BioEdit version 7.0.5.3 [68]. A similarity matrix was calculated by using the similarity matrix tool located at the Ribosomal Database Project homepage (http://rdp.cme.msu.edu/cgis/phylip.cgi). Operational Taxonomic Units (OTUs) were defined as sequences obtained from the same technique that showed a similarity more than 97% with each other. Similarity analysis was performed with FastGroup II (http://fastgroup.sdsu.edu). Alignments of OTUs obtained from cloning were exported to test different nucleotide substitution models using Phylip available in http://phylemon.bioinfo.cipf.es. GTR was consequently the optimal model. Posterior probability and topology of the phylogenetic trees were obtained with Mr. Bayes version 3.1.2 [69] defining the parameters GTR+I+G. Tree analysis was a consensus of 5x10⁵ generations (SD=0.02) in Archaea and 2.5×10⁵ generations in Bacteria (SD=0.04), in both cases it was performed a “burnin” of 50%.

GenBank sequence accession numbers

The SSU rRNA fragment gene sequences were deposited in the GenBank database under accession numbers, EU734574, EU725589-EU725602, EU722643-EU722714, FJ172052-FJ172100 and FJ236710-FJ236714. Prefixes of sequences describe: community sampled (Rw for rainy water, Rs for rainy sediment, Dw for dry water and Ds for dry sediment), technique used (c for clone, i for isolate and d for DGGE-band) and domain (b for Bacteria and a for Archaea) and identification number.