Systematic and Applied Microbiology 33 (2010) 209–221
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Bacterial diversity in dry modern freshwater stromatolites from Ruidera Pools Natural Park, Spain ˜ a , Balbina Nogales b , Elena Soria-Soria a,1 , Fernando Santos a , Arantxa Pena a c M Ángeles García del Cura , Juan Antonio González-Martín d , Josefa Antón a,∗ a
División de Microbiología, Departamento de Fisiología, Genética y Microbiología, Universidad de Alicante, 03080 Alicante, Spain Grup de Microbiologia, Departament de Biologia, Universitat de les Illes Balears, 07122 Palma de Mallorca, Spain c Laboratorio de Petrología Aplicada, Unidad Asociada CSIC (Instituto Geología Económica – UA), Universidad de Alicante, 03080 Alicante, Spain d Departamento de Geografía, Universidad Autónoma de Madrid, Cantoblanco, 28049 Madrid, Spain b
a r t i c l e
i n f o
Article history: Received 11 November 2009 Keywords: Stromatolite Cyanobacteria Leptolyngbya Pseudoanabaenaceae
a b s t r a c t Ruidera Pools Natural Park, Spain, constitutes one of the most representative systems of carbonate precipitation in Europe. The prokaryotic community of a dry modern stromatolite recovered from the park has been analyzed by molecular techniques that included denaturing gradient gel electrophoresis (DGGE) and 16S rRNA gene clone library analysis, together with microscopic observations from the sample and cultures. Ribosomal RNA was directly extracted to study the putatively active part of the microbial community present in the sample. A total of 295 16S rRNA gene sequences were analyzed. Libraries were dominated by sequences related to Cyanobacteria, most frequently to the genus Leptolyngbya. A diverse and abundant assemblage of non-cyanobacterial sequences was also found, including members of Firmicutes, Bacteroidetes, Proteobacteria, Actinobacteria, Acidobacteria, Planctomycetes and Chloroflexi groups. No amplification was obtained when using archaeal primers. The results showed that at the time of sampling, when the pool was dry, the bacterial community of the stromatolites was dominated by groups of highly related Cyanobacteria, including new groups that had not been previously reported, although a high diversity outside this phylogenetic group was also found. The results indicated that part of the Cyanobacteria assemblage was metabolically active and could thus play a role in the mineralization processes inside the stromatolites. © 2010 Elsevier GmbH. All rights reserved.
Introduction Ruidera Pools Natural Park in Central Spain (38◦ 58 N 02◦ 52 W) is a fluvio-lacustrine system consisting of a series of successive lagoons stepped along the longitudinal profile of the High Guadiana Valley, connected by streams, waterfalls and springs. The lagoons are formed due to the impermeability of the geological substrate at the bottom of the valley and the presence of a natural tufa barrier, constituted by carbonate precipitation, which create natural dams separating the different pools. Ruidera Pools represents one of the most important sites of current carbonate precipitation in Europe ˜ (Ordónez et al. 1997) and contains well-preserved examples of present day fluvial and lacustrine tufas developed under semi-arid ˜ conditions (Ordónez et al. 2005). Tufas are formed by physico-
∗ Corresponding author at: División de Microbiología, Departamento de Fisiología, Genética y Microbiología, Universidad de Alicante, Apto. 99, 03080 Alicante, Spain. Tel.: +34 965903870; fax: +34 965909569. E-mail address:
[email protected] (J. Antón). 1 Present address: LABAQUA, Dpto. de Microbiología, C/Dracma, 16-18 Polígono Industrial Las Atalayas, 03114 Alicante, Spain. 0723-2020/$ – see front matter © 2010 Elsevier GmbH. All rights reserved. doi:10.1016/j.syapm.2010.02.006
chemical and biological precipitation, a process mediated by the presence of microbial biofilms generally dominated by diatoms, cyanobacteria and heterotrophic bacteria. These lithified, laminated, calcareous and benthic microbial deposits are frequent in Ruidera Pools and constitute the stromatolites that are organosedimentary layered structures accreted by sediment trapping, binding and in situ precipitation due to the growth and metabolic activities of microorganisms (Walter 1976). Although many naturally occurring microbial mat communities are known to trap and bind sediments, only a few ecosystems, such as the one described here, are conducive for the process of lithification and fossilization (Havemann and Foster 2008). In addition, living microbialites, such as the ones studied here, are only found within a few selected locations, as pointed out by Breitarbt and collaborators (Breitbart et al. 2009). Stromatolites present in Ruidera Pools have been studied ˜ morphologically, geologically and microscopically (Ordónez and ˜ García del Cura 1983; Ordónez et al. 1997; Souza-Egipsy et al. 2005), although very little is known about their microbial diversity. Microscopy analysis of submerged stromatolites shows abundant diatoms in the surface of calcite build-ups interspersed with a layer of EPS and unicellular and filamentous cyanobacteria and algae. It seems clear that microorganisms play an important role in the
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building of these organosedimentary structures and that microbial cell walls, sheaths and exopolymeric substances influence the process of calcite biomineralization (Souza-Egipsy et al. 2005). In fact, differences between cell walls and sheaths of cyanobacteria and algae, as well as the presence of exopolymeric substances, lead to differences in the process of biologically induced calcite mineralization between these two microbial groups (Souza-Egipsy et al. 2005). We chose a pool located in an erosional channel between two larger pools (Tomilla and Tinaja) that was generated in 1947 as a consequence of a flooding-related spillover (García del Cura et al. 1997). The water of Tomilla Pool, which feeds the small pool sampled, is almost saturated with calcite and dolomite. The pool is periodically dried due to drought and/or seasonal changes in the rain regime. The bottom of the pool is covered with stromatolites of different morphology which grow by the formation of mineralentrapping microbial layers that can be up to 2 mm thick. Below the surface of the stromatolite, a clear green band was observed (Fig. 1A), formed by the mineral-entrapping microbial community. These layers are formed only when the pool is covered by water and, therefore, stromatolite formation depends on the periodical flooding of the pool, as well as the development of an active microbial community that contributes to mineral precipitation. However, microorganisms have also been repeatedly observed by scanning electron microscopy inside dry lithified stromatolites from Ruidera Pools (A. de los Ríos, personal communication), although it is unknown whether these communities remain active when not covered by water, and which microbes compose them. To address these questions, culture-independent methods were chosen for examining the microbial diversity and composition of a dry stromatolite. The bacterial diversity was analyzed by using denaturing gradient gel electrophoresis (DGGE) and clone libraries from 16S rRNA genes amplified by PCR from total environmental DNA, and libraries from 16S rRNA, reversely transcribed and amplified from total environmental rRNA. Since RNA molecules are more labile than DNA, the detection of 16S rRNA in an environmental sample is an indication of the presence of microbial cells which contain ribosomes and, therefore, they are presumably metabolically active (Nogales et al. 2001). In addition, the microbial community in the dry stromatolite was analyzed by culture and microscopic methods. Materials and methods Samples A stromatolite sample (10 cm × 7 cm × 3.5 cm, 150 g, Fig. 1A) was taken from the bottom of the pool on September 15th, 2000 (the pool was dry at the time of sampling), and kept at 4 ◦ C for 3 h, during transportation to the laboratory. The stromatolite chosen for this study had a pinnate dome shape (h = 20 cm) and was located in the subcritical turbulent flow and in the deepest part of the pool. The main component of the pinnacle domes consists of layers of superposed light lamina of fibrous calcite macrocrystals with visible cyanobacterial trichomes in an intracrystalline position (Souza-Egipsy et al. 2005). The sample was stored at −70 ◦ C, and subsamples were taken for nucleic acid extraction by using a sterile spatula to scrape off material from the green (“algal”) layer (Fig. 1A), which was collected in a sterile Petri dish. Care was taken to avoid scraping detritus material deposited on the surface of the stromatolite. A piece of the original sample was kept at room temperature and used for culture inoculation. Microscopy Scraped material from the green layer of the stromatolite sample was used for phase contrast optical microscopy (DM 4000B,
LEICA) and confocal laser scanning microscopy (TCS SP2, LEICA) examinations after rehydrating in sterile water. Nucleic acid extraction and purification For nucleic acid extraction, two protocols were assayed. Protocol I, an adaptation of the protocol described previously (Cifuentes et al. 2000), was followed. A total of 5 g from the greenish layer of the stromatolite was mixed in a 15 mL polypropylene tube and washed twice with 7 mL cold AE buffer (20 mM sodium acetate pH 5.5, 1 mM EDTA) by centrifuging at 3900 × g for 5 min. Then, 10 mL of phenol–chloroform–isoamylalcohol 25:24:1 (PCI), pre-warmed at 60 ◦ C, and 300 L of 10% SDS were added. This mixture was incubated at 60 ◦ C for 5 min with vortexing every minute. After cooling on ice, the tubes were centrifuged at 3900 × g for 5 min at 4 ◦ C. The aqueous phase was transferred to a new tube to which 500 L of 2 M sodium acetate pH 5.2 and 10 mL of PCI were added, vortexed, and centrifuged as above. The removal of the aqueous phase, addition of sodium acetate and PCI and centrifugation were repeated until a clear interphase between the aqueous and the organic phases was observed (i.e. 2–3 times). Finally, nucleic acids were precipitated with ethanol and resuspended in 100 L of sterile deionized water. A second nucleic acid extraction protocol (protocol II) was also carried out for comparison (see Results). For this extraction, 5 g from the greenish layer of the stromatolite were mixed as described above with 7 mL of extraction buffer (100 mM Tris–HCl pH 8.0, 100 mM EDTA pH 8.0). Then, 80 L lysozyme (final concentration: 3 mg mL−1 ) were added and the sample was incubated for 15 min at 37 ◦ C (180 rpm). A total of 120 L proteinase K (final concentration: 150 g mL−1 ) and 800 L 10% SDS were added, followed by incubation for 30 min at 37 ◦ C (180 rpm). The sample was mixed with 1600 L 5 M NaCl and then incubated at 65 ◦ C for 10 min with 1200 L CTAB (10% CTAB, 0.7 M NaCl). Three freeze/thaw steps were carried out by using liquid nitrogen and a water bath at 65 ◦ C. One volume of PCI was added and the sample was centrifuged at 3900 × g for 5 min (4 ◦ C). After centrifugation, the aqueous phase was transferred into new tubes and the addition of PCI was repeated until a clear interphase between the aqueous and the organic phases was observed (i.e. 2 or 3 times). Nucleic acids were precipitated with a 0.1 volume of 3 M sodium acetate (pH 4.8), 0.01 volume of 1 M magnesium chloride and 0.6 volume of cold isopropanol and finally resuspended in 100 L sterile deionized water. Crude nucleic acid extracts were purified with GENECLEAN Kit II (Bio 101), according to the manufacturer’s protocol. To check the quality of the extracted nucleic acids, they were subjected to electrophoresis on 0.8% LE agarose gels (FMC Products, Rockland, ME, USA) in 1× Tris–acetic acid–EDTA (TAE) buffer, and visualized under UV light after ethidium bromide staining. All glassware was washed with 2N NaOH and rinsed with sterile deionized water to minimize RNA degradation. PCR amplification of 16S rRNA genes Forward primers 27f and 21F, and universal reverse primer 1492r were used for complete 16S rRNA gene amplification, as previously described (DeLong 1992). Gene fragments for denaturing gradient gel electrophoresis (DGGE) were amplified with forward primer 341f-GC for Bacteria, and 344F-GC for Archaea and universal reverse primer 907r, as previously described (Muyzer et al. 1993, 1998; Nagy and Johansen 2001), using different conditions for the touch down PCR (from 65 to 55 ◦ C, and from 60 to 50 ◦ C) that did not have an effect on the DGGE patterns. Primers CYA359GC and CYA781R (equimolar mixture between primers CYA781Ra and CYA781Rb) were used for PCR amplification of cyanobacterial 16S rRNA genes from environmental DNA, as previously described (Nübel et al. 1997).
F. Santos et al. / Systematic and Applied Microbiology 33 (2010) 209–221
All PCR products were electrophoresed onto 1% LE agarose gels (FMC Products, Rockland, ME, USA) in 1× TAE buffer. RT-PCR of 16S rRNAs A total of 20 L purified nucleic acid was digested with 3 U of RQ1 RNase-free DNase (Promega) in the buffer provided by the supplier, according to the manufacturer’s recommendations. An aliquot of the digestion product was analyzed by electrophoresis
211
to check for the absence of DNA. Bacterial 16S rRNAs positions 1055–1492, Escherichia coli numbering, were reversely transcribed and PCR amplified by using the SuperScript One-Step RT-PCR with Platinum Taq kit (Life Technologies), according to the manufacturer’s protocol, using primers B1055 (Amann et al. 1995) and 1492r, and the PCR protocol described above. The PCR products were electrophoresed as previously described. To check for the absence of contaminating DNA, a control PCR with RNA without previous reverse transcription was carried out.
Fig. 1. A piece of the stromatolite used in this study (A; the green surface is indicated with an arrow) and cyanobacterial filaments observed after rehydrating material scraped from it (B). Cyanobacterial filaments containing chlorophyll (C and D) developed after inoculation in enrichment media with (E and F) and without (G and H) a combined source of nitrogen.
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Cloning of PCR and RT-PCR products Prior to cloning, a low-cycle-number reamplification of a 10fold diluted mixed-template PCR product was performed with a fresh reaction mixture, as described to eliminate heteroduplexes from multitemplate PCR products (Thompson et al. 2002). Complete (PCR) and partial (RT-PCR) bacterial 16S rRNA products were then run on a 2% Metaphor agarose gel (FMC Products, Rockland, ME, USA). The bands of appropriate size (i.e. 1500 and 500 bp, respectively) were sliced from the gel using the GFXTM PCR DNA and gel band purification kit (Amersham, UK). Purified products were then run on a 1% LE agarose gel to check for their quality. Cloning of the products with the pCR2.1® vector, transformation of E. coli TOP10F’ cells and selection of transformants were carried out with the TOPO TA cloning kit (Invitrogen), according to the manufacturer’s protocol. Two bacterial 16S rRNA gene clone libraries (D libraries) from DNA samples obtained in two independent extractions were generated. Each library was generated with the pooled products of three independent PCR reactions. One bacterial 16S rRNA gene clone library from environmental RNA reversely transcribed to DNA (R library) was constructed. In addition, one small 16S rRNA gene clone library from DNA extracted from one of the cultures was also constructed (see below).
Biosystems) and vector primers. Nucleotide sequences of DGGE bands were obtained with primer 907r, after band excising and DNA reamplification. Sequences were compared initially with reference sequences at the NCBI (http://www.ncbi.nlm.nih.gov) using BLAST (Altschul et al. 1997) and they were subsequently aligned with 16S rRNA reference sequences in the ARB package (Ludwig et al. 2004). Phylogenetic trees were generated using nearly complete representative 16S rRNA gene sequences obtained from public databases, using the algorithms implemented in the ARB package. Partial sequences of the clones and DGGE bands obtained in this study were then added to the tree using the parsimony tool implemented in the ARB package. Neighbor-joining, parsimony and maximum likelihood analyses were performed in ARB with 1000 replicates, by using different data sets and, alternatively, by using masks that removed very frequent gaps. Once the different reconstructions obtained were compared, consensus trees were drawn and positions that could not be resolved were shown as multifurcations and thus did not show bootstrap values. Sequences were also checked for possible chimeric structures using the Mallard 1.02 and Pintail 1.0 softwares (Ashelford et al. 2005, 2006) and by analyzing the correlation between primary sequence and secondary structure using the ARB sequence editor. Nucleotide sequence accession numbers
Amplified ribosomal DNA restriction analysis (ARDRA) Inserts from the 16S rRNA gene clones were PCR amplified with vector-specific primers. The following conditions were used for amplification: a step of 94 ◦ C for 3 min, 30 cycles of 94 ◦ C for 15 s, 55 ◦ C for 30 s, and 72 ◦ C for 2 min; plus an extension step of 10 min at 72 ◦ C. Amplification products were electrophoresed onto 1% agarose gels, stained and visualized as above. Then, 10 L of the PCR-amplified inserts were incubated separately with 5 U of enzymes HinfI and MboI (New England Biolabs) in the corresponding enzyme buffers. Digestion products were electrophoresed onto 2% agarose gels in 0.5× Tris–boric acid–EDTA (TBE) buffer, stained and visualized as above. Clones representing different restriction patterns were selected for sequencing. Denaturing gradient gel electrophoresis (DGGE) DGGEs were carried out with a Dcode system [Bio-Rad, Hercules, CA, USA]. Electrophoresis conditions were 70 V for 16 h in a linear gradient of denaturing agents from 40% to 70% [where 100% denaturing agents were 7 M urea and 40% deionized formamide] in a 6% acrylamide–bis-acrylamide [37.5:1] gel, using TAE 1× as the running buffer. After electrophoresis, the gels were stained with SYBR-GREEN and visualized under UV light. Gel images were captured by using a Typhoon 9410 scanner. Cultures Small pieces of stromatolite containing the greenish upper layer were used for inoculating two cyanobacterial specific media (Rippka 1988) with and without a source of combined nitrogen [17.5 mM NaNO3 ]. The cultures were incubated at 30 ◦ C under white light until cyanobacterial filaments were released from the stromatolite piece used as inoculum [i.e. two months]. Aggregates from the cultures were observed microscopically. Sequencing of cloned PCR/RT-PCR products and DGGE bands The nucleotide sequences of the cloned products were determined from plasmid preparations (Wizard Plus SV Minipreps DNA Purification System, Promega) using the Big Dye Terminator Cycle Sequencing kit, an ABI PRISMTM 310 DNA sequencer (Applied
The sequences from this study are available in GenBank under accession numbers EU753607–EU753687, EU780449–EU780450 and AY566347–AY566320. Results Material scraped from the green layer of the stromatolite used for molecular analysis was observed microscopically and all the recognizable morphologies were attributed to filamentous Cyanobacteria. Most of the cyanobacterial filaments observed presented a width of about 2–3 m, with thin sheaths but lacking heterocysts and akinetes, thus resembling the morphologies of Pseudoanabaenaceae family members (Fig. 1B). Cell filaments also emitted bright red fluorescence when observed under the confocal laser scanning microscope with excitation at 633 nm (white fluorescence in Fig. 1C and D), indicating the presence of chlorophyll-containing filamentous cyanobacteria (Papineau et al. 2005) in the stromatolite sample. With the described extraction and purification protocols, DNA and rRNA suitable for PCR and RT-PCR amplification were obtained from the green layer of the dry, highly lithified stromatolite sample. Previously, different amounts of extraction buffers and sample were assayed (data not shown) and it was found that the amount of stromatolite per volume of extraction buffer was especially critical for obtaining enough good quality nucleic acid. In order to make sure that the DNA extraction procedure was not biasing the results, two subsamples were taken from opposite sides of the stromatolite sample and two different extraction protocols were used. Nucleic acids from each subsample were extracted with the two protocols and their bacterial communities were analyzed by denaturing gradient gel electrophoresis (Fig. 2A), as described above. No amplification with primers for Archaea was obtained in any case. This analysis revealed that the community in the upper layer of the stromatolite was rather homogeneous, independently of the subsample chosen. Eleven DGGE bands from the profile were successfully sequenced; however, a considerably higher number of bands were detected in the background of the gel, suggesting a high bacterial diversity in the sample, which was further analyzed by constructing and sequencing 16S rRNA gene libraries. DGGE sequences in the database matched members from the Bacteroidetes, Cyanobacteria, Firmicutes and Chloroflexi groups (Table 1). Except for DGGE bands
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Fig. 2. (A) DGGE profiles obtained with bacterial primers 341f-GC and 907r. Lanes 1–6 contain amplification products from DNA extracted with protocol I from one subsample; lanes 7–12 contain amplification products from DNA extracted with protocol II from a different subsample. PCR products in even lanes were obtained with a touch down from 65 to 55 ◦ C in the amplification cycles; PCR products in odd lanes were obtained with a touch down from 60 to 50 ◦ C in the amplification cycles. (B) DGGE profile obtained with cyanobacterial specific primers 359F-GC and 781R. Sequenced bands are marked with arrows.
corresponding to Chloroflexi, the rest of the groups were well represented in the libraries (see below). Nucleic acid extracts obtained with protocol I were chosen for library construction, since this protocol yielded higher amounts of nucleic acids than protocol II. The screening of the 242 clones from two D libraries by ARDRA yielded a total of 94 different patterns. At least one clone per restriction pattern was selected for complete sequencing. Eighteen clones resulted in chimerical sequences (7.3% of analyzed clones) and these were then eliminated from further analyses, as well as two clones which corresponded to E. coli contaminating sequences. Two clones matched in the database with algal-plastid sequences (data not shown). The rest of the clones were related to members of the Cyanobacteria, Firmicutes, Alphaproteobacteria, Betaproteobacteria, Gammaproteobacteria, Bacteroidetes, Acidobacteria, Actinobacteria and Planctomycetes (Table 1). A total of 27 clones obtained from the amplification of environmental 16S rRNA reversely transcribed (labelled as R clones) were analyzed, and most of them were related to cyanobacterial 16S rRNA gene sequences (Table 1). However, clones related to Alphaproteobacteria, Gammaproteobacteria, Firmicutes, Actinobacteria, and Planctomycetes were also found. Shannon indexes (Hammer et al. 2001) for D and R libraries were 2.71 and 2.62, which were considerably higher than that found by Havemann and Foster (2008) for marine stromatolites (Alonso 2005; Vaulot et al. 2002). Cyanobacterial diversity A total of 139 (57.4% of the total D clones) corresponded to cyanobacterial sequences, most of them related to Leptolyngbya. Oscillatorial sequences were the most represented group in the D (110 clones) and R libraries (17 out of 27 R clones). All the oscillatorial-related R sequences clustered together with D sequences with identity values always above 97% (100% identity for clone R10 and sequences D1A07 and D1B02), indicating that all the “active” (i.e. ribosome containing) members of this assemblage had also been detected in the D libraries. The phylogenetic tree of the cyanobacterial sequences retrieved from the stromatolite (Fig. 3) was consistent with those calculated elsewhere (García-Pichel et al. 2001; Ishida et al., 2001; Liviatis 2002; Miller and Castenholz 2001; Nelissen et al. 1996) that also showed
Oscillatoriales as a paraphyletic group, in agreement with Rippka (Rippka 1988) who pointed out that “the classical oscillatorian taxonomy is rather confusing and often arbitrary”. A total of 65 of the Leptolyngbya-related clones matched in the database with uncultured cyanobacterial clones from endolithic microbial ecosystems, laminated Fe-formations on Sulawesi, Indonesia, and microbial mats from Antarctic freshwater, as well as terrestrial habitats (Taton et al. 2006b; Walker and Pace 2007; Yergeau et al. 2007). However, most of the sequences found in this work grouped separately from those retrieved from other environments (identities below 97%), as also found for cyanobacterial communities inhabiting desert crusts (García-Pichel et al. 2001) or the microbial mats of Lake Fryxell, McMurdo Dry Valleys, Antarctica (Taton et al. 2006a). Twenty-nine D clones and four R clones corresponded to cyanobacterial sequences belonging to the order Nostocales, constituted by filamentous, heterocyst-forming Cyanobacteria. They clustered together (Fig. 3) and close to Calothrix spp. and had their best hit in the database with different strains of Nostoc, Cyanospira, and Nodularia associated with benthic and planktonic ecosystems, as well as with cyanobacterial symbionts of the liverwort Blasia pusilla. Complete sequences from the D library had identities to each other of between 97% and 99%, and some R sequences exactly matched some D sequences, with identities of 100% (i.e. clones D1E07 and D3F11 with sequence R02; clones D1F06, D2H03 and D3B06 with sequences R03, R23 and R25, respectively). Except for sequence D1E09, the rest of the Nostocales-related sequences clustered together in the same branch and separately from previously reported sequences, thereby constituting a new group of putatively nitrogen-fixing cyanobacteria associated with the analyzed stromatolite. All DGGE bands obtained with primers 341f-GC and 907r assigned to Cyanobacteria clustered together with sequences obtained in the libraries. Since most of the clone sequences corresponded to Cyanobacteria, another DGGE analysis was performed with primers specific for this group (Fig. 2B) in order to obtain a better picture of cyanobacterial diversity. Except for Cya DGGE Band 01, that had a very low percentage identity with the databases, all the cyanobacterial DGGE bands (Table 2) were highly related to sequences obtained in the libraries. This probably indicated that these molecular approaches had unveiled most of the cyanobacterial diversity in the sample.
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Table 1 16S rRNA gene sequences obtained in this study. Phylogenetic group
Best GenBank hit
D librarya Number of clones
Identity
R libraryb Number of clones
Identity
DGGEb bands
Cyanobacteria; Oscillatoriales
Leptolyngbya sp. LLi18 (DQ786166)
43
94–96%
5
97%
n.d.c
–
Unclassified bacterium isolate MMB05 (EF126270) Unclassified cyanobacterium clone FQSS098 (EF522318) Unclassified cyanobacterium clone H-A02 (DQ181685) Leptolyngbya frigida ANT.LH70.1 (AY493574) Leptolyngbya antarctica ANT.LH67.1 (AY493572) Unclassified cyanobacterium clone RJ094 (DQ181682) Unclassified cyanobacterium clone H09 MO02 (EF220899) Unclassified bacterium clone GASP-0KB-616-H05 (EU043700) Cyanospira rippkae (AY038036) Unclassified bacterium clone JSC8-C11 (DQ532230) Nostoc sp. YK-01 (AB187507) Nostoc sp. SKS8 (EU022706) Nodularia harveyana BECID29 (AJ781156) Nostoc sp. PCC 9229 (AY742451) Unclassified bacterium clone Baqar.Water.Eubac.6 (AB355032) Nostoc sp. PCC 7120 (BA000019) Unclassified cyanobacterium clone 100M1 A6 (DQ513995) Unclassified bacterium clone JSC8-C11 Anabaena oscillarioides BO HINDAK 1984/43 (AJ630428) Calothrix sp. CAL3363 (AM230684)
39
95–96%
n.d.
–
n.d.
–
8
93–95%
n.d.
–
n.d.
–
18
92–93%
n.d.
–
n.d.
–
1
94%
n.d.
–
n.d.
–
1
93%
n.d.
–
n.d.
–
–
n.d.
–
1
Sporosarcina sp. S11-2 (DQ514313) Unclassified bacterium clone AKIW836 (DQ129270) Bacillus sp. YY (AF414443) Bacillus sp. JAMB-204 (AB180249) Bacillus sp. JL-38 (AY745843) Bacillus sp. CNJ904 PL04 (DQ448797) Unclassified clostridiales bacterium clone SRB16 (DQ069224) Unclassified eubacterium clone AKAU3868 (DQ125755) Unclassified bacterium clone RB506 (AB240379) Clostridium thermocellum ATCC 27405 (CP000568) Unclassified bacterium clone AKAU3928 (DQ125791) Unclassified bacterium clone D8A 3 (AY768823) Bacillus sp. HM06-04 (EU004566) Glacial ice bacterium SB100-9-5-1 (AF479372) Bacteroidetes bacterium MWH-CFBk5 (AJ565431) Unclassified bacterium clone 21BSF11 (AJ863254) Unclassified bacteroidetes clone MSB-5A10 (DQ811911) Unclassified bacteroidetes clone S1-4-CL9 (AY728066) Unclassified bacterium clone oc11 (AY491561) Flexibacter flexilis, strain IFO 16028 (AB078054) Unclassified bacterium clone 21BSF16 (AJ863255) Unclassified bacteroidetes clone PS17 (DQ004377) Unclassified bacterium clone B60 (AM162442)
6 2
98% 94%
n.d. n.d.
– –
n.d. n.d.
– –
13 4 9 1 1
97–99% 98% 98% 97% 91%
n.d. n.d. n.d. n.d. n.d.
– – – – –
n.d. 1 n.d. n.d. n.d.
– 99% – – –
1
94%
n.d.
–
n.d.
–
1
93%
n.d.
–
n.d.
–
1
93%
n.d.
–
n.d.
–
2
96%
n.d.
–
n.d.
– –
Cyanobacteria; Nostocales
Firmicutes
Bacteroidetes
Proteobacteria; Alphaproteobacteria
Identity
n.d.
–
3
95%
n.d.
–
n.d.
–
5
95%
n.d.
–
n.d.
–
–
–
1
93%
2 9
93% 95%
n.d. n.d.
– –
n.d. n.d.
– –
5 5 6 1 1
93% 92% 92% 92% 92%
n.d. n.d. n.d. n.d. n.d.
– – – – –
n.d. n.d. n.d. n.d. n.d.
– – – – –
1 n.d.
93% –
n.d. 1
– 93%
n.d. n.d.
– –
n.d. n.d.
– –
1 2
93% 94%
n.d. n.d.
– –
n.d.
94%
n.d.
–
2
90%
n.d.
n.d. n.d.
– –
n.d. n.d.
– –
1 1
5
91%
n.d.
–
n.d.
–
2
97%
n.d.
–
n.d.
–
1
92%
n.d.
–
n.d.
–
1
95%
n.d.
–
n.d.
–
1
97%
n.d.
–
n.d.
–
n.d.
–
n.d.
–
1
89%
n.d.
–
n.d.
–
2
98%
n.d.
–
n.d.
–
1
99%
92%
n.d.
–
n.d.
4
97% 99%
–
F. Santos et al. / Systematic and Applied Microbiology 33 (2010) 209–221
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Table 1 (Continued ) Phylogenetic group
Proteobacteria; Betaproteobacteria
Proteobacteria; Gammaproteobacteria
Planctomycetes
Actinobacteria
Acidobacteria
Chloroflexi
Best GenBank hit
D librarya Number of clones
Identity
R libraryb Number of clones
Identity
DGGEb bands
Unclassified bacterium isolate BF0002C082 (AM697385) Brevundimonas subvibrioides (AJ227784) Unclassified bacterium clone M1-34 (EU015114) Unclassified bacterium clone KSC2-93 (DQ532296) Paracoccus sp. NPO-JL-65 (AY745834) Brevundimonas variabilis ATCC (AJ227783) Blastochloris viridis strain UN (AF084496) Unclassified bacterium clone IYF06 (DQ984584) Unclassified alphaproteobacterium clone FI-2F D05 (EF220410) Unclassified bacterium clone CV108 (DQ499328) Unclassified soil bacterium clone 119 (AY493943) Unclassified bacterium DSSD75 (AY328773) Unclassified gammaproteobacterium clone 478 (AB252882) Unclassified bacterium clone HB33 (EF648043) Unclassified bacterium clone PeM12 (AJ576382) Unclassified planctomycete clone A12 MO02 (EF220755) Unclassified actinobacterium clone F15 1F FL (EF683003) Unclassified actinobacterium clone EB1077 (AY395396) Knoellia sp. O-008 (DQ812538) Unclassified bacterium clone Par-s-4 (EF632908) Unclassified bacterium clone MIZ18 (AB179509) Unclassified bacterium clone AKIW799 (DQ129639) Unclassified Antarctic bacterium clone LB3-100 (AF173817) Unclassified bacterium clone FFCH12062 (EU134097)
2
98%
n.d.
–
n.d.
–
1 1
99% 96%
n.d. n.d.
– –
n.d. n.d.
– –
1
98%
n.d.
–
n.d.
–
1 1 n.d. n.d.
99% 98% – –
n.d. n.d. 1 2
– – 97% 98%
n.d. n.d. n.d. n.d.
– – – –
n.d.
–
1
96%
n.d.
–
Total a b c
Identity
8
92–94%
n.d.
–
n.d.
–
1
98%
n.d.
–
n.d.
–
1
99%
n.d.
–
n.d.
–
2
99%
n.d.
–
n.d.
–
1
90%
n.d.
–
n.d.
–
1
90%
n.d.
n.d.
n.d.
–
1
96%
n.d.
–
n.d.
–
1
99%
n.d.
–
n.d.
–
1
94%
n.d.
–
n.d.
–
1 n.d.
99% –
n.d. 1
– 97%
n.d. n.d.
– –
2
93%
n.d.
–
n.d.
–
2
93%
n.d.
–
n.d.
–
n.d.
–
n.d.
–
1
90%
n.d.
–
n.d.
–
1
89%
220
27
11
Nearly complete sequences. Partial sequences. Not detected.
Table 2 16S rRNA gene sequences from cyanobacterial DGGE bands and clones from the culture. DGGE bands/clones
Length
Best hit
Hit with DNA library
Cya DGGE Band 01 Cya DGGE Band 02 Cya DGGE Band 03 Cya DGGE Band 04 Cya DGGE Band 05 Cya DGGE Band 06 Cya DGGE Band 07
301 346 353 365 270 279 349
Unclassified bacterium clone QB81 88% Unclassified cyanobacterium clone D2D07 98% (N) Unclassified cyanobacterium clone D3B06 97% (N) Unclassified cyanobacterium clone D1D12 99% (O) Unclassified cyanobacterium clone D1H05 98% (N) Unclassified cyanobacterium clone D1G12 98% (O) Unclassified cyanobacterium clone D1G12 98% (O)
Unclassified cyanobacterium clone D1G07 88% (O)
Culture Clone N01 Culture Clone N02 Culture Clone N03 Culture Clone N04 Culture Clone N06 Culture Clone N08 Culture Clone N14 Culture Clone N16
424 425 429 426 425 426 427 426
Unclassified cyanobacterium clone D3B01 96% (O) Leptolyngbya badia CRS-1 94% Leptolyngbya frigida ANT.LH52.2 96% Unclassified bacterium clone 2N6-23R 96% Unclassified cyanobacterium clone 100M2 H9 95% Unclassified cyanobacterium clone D3B01 96% (O) Leptolyngbya frigida ANT.LH52.2 97% Unclassified bacterium clone 2N6-23R 96%
(O): oscillatoriales; (N): nostocales.
Unclassified cyanobacterium clone D1A07 94% (O) Unclassified cyanobacterium clone D1A07 94% (O) Unclassified cyanobacterium clone D1A07 95% (O) Unclassified cyanobacterium clone 3D02 93% (O) Unclassified cyanobacterium clone D1A07 96% (O) Unclassified cyanobacterium clone D1A07 95% (O)
216
F. Santos et al. / Systematic and Applied Microbiology 33 (2010) 209–221
Fig. 3. Phylogenetic inference based on 16S rRNA gene sequences from clones and DGGE bands belonging to the Cyanobacteria group. The sequence of Arthrobacter globiformis was used as the outgroup. Sequences from the R library are enclosed in boxes; sequences from the D library and DGGE are indicated by boldface type. A sequence from every ARDRA profile has been included in the tree and the number of clones per profile is indicated in parentheses. The tree is based on the results of neighbor-joining inferences for only complete or nearly complete sequences. The tree topology was corrected based on the results of additional maximum parsimony and maximum likelihood analyses. For branches without bootstrap values, the values were less than 60%. Bar = 10% estimated sequence divergence.
F. Santos et al. / Systematic and Applied Microbiology 33 (2010) 209–221
217
Fig. 4. Phylogenetic inference based on 16S rRNA gene sequences from clones and DGGE bands belonging to the Firmicutes group. The sequence of Arthrobacter ramosus was used as the outgroup. Sequences from the R library are enclosed in boxes; sequences from the D library and DGGE are indicated by boldface type. A sequence from every ARDRA profile has been included in the tree and the number of clones per profile is indicated in parentheses. The tree is based on the results of maximum likelihood inferences for only complete or nearly complete sequences. The tree topology was corrected based on the results of additional neighbor-joining and maximum parsimony analyses. For branches without bootstrap values, the values were less than 60%. Bar = 10% estimated sequence divergence.
In an attempt to cultivate the cyanobacteria detected by molecular methods, as well as to demonstrate the presence of viable cyanobacteria in the stromatolite (as suggested by the results of microscopy and rRNA analysis), we inoculated two cyanobacterial enrichment media with pieces of the stromatolite. In the medium with a source of combined nitrogen, several filamentous cyanobacteria were observed (Fig. 1E and F). Some filamentous cyanobacteria with thin sheaths and a width of about 2–3 m (Fig. 1F) resembled those observed directly when the scraped material from the stromatolite was viewed under optical and confocal microscopy. Other filaments, wider and with thicker sheaths, were also observed (Fig. 1E and F). Different cyanobacterial morphologies were observed in the culture without the nitrogen source (Fig. 1G and H). A 16S rRNA gene library was constructed for the culture grown with combined nitrogen, and eight clones were partially sequenced. Their sequences (labelled as “N” sequences in Table 2) matched in the databases with Pseudoanabaenaceaerelated strains and sequences obtained from the stromatolite, but the identity percentages were always below 97%. This indicated that the enrichment culture was not selecting for the cyanobacteria detected with the culture-independent approach, which was not surprising since our enrichment medium, although very commonly used for cyanobacterial culture, could have been selecting for the faster growing cyanobacteria present in the sample. However, it is also noteworthy that the eight clones all had their closest BLAST
matches with sequences from the stromatolite libraries, thus indicating that the cyanobacterial diversity analyzed by all the applied approaches seemed to be quite homogeneous in terms of phylogenetic assemblages. Other bacterial diversity A total of 95 non-cyanobacterial 16S rRNA gene sequences (from libraries and DGGE bands) were also retrieved from the stromatolite (Table 1), and they displayed a considerably higher diversity than the cyanobacteria-related sequences. From the D libraries, 41 clones (16.9% of the total D clones) corresponded to the Firmicutes group (Fig. 4). Most of these sequences, as well as DGGE Bands 06 and 07, were highly related to each other and with marine isolates from the genera Bacillus and Sporosarcina. The rest of the sequences were more distantly related to members of the Clostridiales. Only two Firmicutes-related sequences were obtained from the R library (clones R05 and R14), and they were rather different from the rest, since they had their best match to a clone from a 5 km deep terrestrial fault (Acc. no. AY768823). Sequences affiliated to other groups (Proteobacteria, Bacteroidetes, Actinobacteria, Acidobacteria and Planctomycetales) accounted for 16.5% of the total clones in the D library (Table 1 and Fig. 5). Some sequences matched in the database with identities from 97% to 99% with cultivated Alphaproteobacteria (Brevundi-
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F. Santos et al. / Systematic and Applied Microbiology 33 (2010) 209–221
Fig. 5. Phylogenetic inference based on 16S rRNA gene sequences from clones and DGGE bands belonging to the domain Bacteria (excluding the Cyanobacteria and Firmicutes groups). The sequence of Leptolyngbya sp. Lli18 was used as the outgroup. Sequences from the R library are enclosed in boxes; sequences from the D library and DGGE are indicated by boldface type. A sequence from every ARDRA profile has been included in the tree and the number of clones per profile is indicated in parentheses. The tree is based on the results of maximum parsimony inferences for only complete or nearly complete sequences. The tree topology was corrected based on the results of additional neighbor-joining and maximum likelihood analyses. For branches without bootstrap values, the values were less than 60%. Bar = 10% estimated sequence divergence.
F. Santos et al. / Systematic and Applied Microbiology 33 (2010) 209–221
monas subvibrioides CB81, Brevundimonas variabilis ATCC 15255T and Paracoccus sp. NPO-JL-65) isolated from waters, and cultivated Betaproteobacteria (Duganella sp. BD-a14) from a hydrocarboncontaminated soil. Two clones were related (99% identity BLASTn value) to Knoellia sp. O-0008 and Arthrobacter sp. Tibet-IX22 (within the Actinobacteria group), isolated from the Baltic Sea and a permafrost in the Qinghai-Tibet Plateau, respectively. Although both kinds of libraries and DGGE indicated the presence of the same bacterial groups in the stromatolite (with the exception of Chloroflexi that was only detected by DGGE), there was no complete overlap of the bacterial assemblages recovered between the two techniques. This is a relatively frequent phenomenon when different rRNA-based techniques are used to describe the diversity in a given environment. For instance, Díez et al. (2001) when analyzing the diversity of marine picoeukaryotic assemblages found groups of microorganisms that could be detected or not depending on whether they analyzed clone libraries or DGGE patterns. In this second case, the microbial diversity retrieved also depended on the primer set used for PCR amplification prior to DGGE separation.
Discussion As far as we know, this is the first study in which ribosomal RNA has been directly analyzed to study presumably metabolicallyactive microorganisms in lithified stromatolites. Ribosomal RNA content (Amann et al. 1995; Klappenbach et al. 2000), together with membrane integrity, respiratory activity, DNA condensed in nucleoids, cell growth, detection of specific mRNA, and substrate uptake, are properties that can be analyzed for the interpretation of metabolic activity (for a review, see Alonso 2005). For this reason, libraries constructed from rRNA contain clones from presumably metabolically-active cells (Nogales et al. 2001) or, at least, from cells with recent activity (Miskin et al. 1999), although some authors have found inactive bacteria containing high levels of ribosomes (Wagner et al. 1995). Thus, the comparison of what we have called the D and R libraries can provide information not only on the composition of the microbial community associated with the stromatolite but also on the putatively active part of this community. Although the number of clones analyzed in the R library was low, a coverage value of 71.4% was obtained, calculated as described in Miskin et al. (1999). This fact could also be related to a lower diversity of presumably active microbiota associated with the dry stromatolite, which might be considered an extreme environment at the time of sampling. It is known that the relative abundance of rRNA gene sequences obtained from a sample with PCR methods may not directly correspond to the relative abundance of organisms in the original sample. However, the fact that sequences from Cyanobacteria were the most represented in our study, in both D and R libraries obtained with different primer combinations, together with the direct observation of these prokaryotes in the sample, could indicate that Cyanobacteria were the most abundant organisms present in the dry stromatolite. In addition, the data are in good agreement with that of Souza-Egipsy et al. (2005) who observed filamentous cyanobacteria in the different layers of the stromatolites. In fact, these authors showed diatom decay was much faster than that of cyanobacteria, which were well preserved, and this is also in agreement with our results, since we could not observe diatoms in the dry stromatolite, and plastid sequences were very scare in the libraries (only 2 out of 242). Furthermore, the presence of putatively active Cyanobacteria in the stromatolite could support the theories about the important role of bacteria and their components, such as EPS, in the mineralization processes that have been suggested for Ruidera National Park stromatolites (Souza-Egipsy et al. 2005).
219
Sequences related to endospore forming bacteria also accounted for a large proportion of the DNA clones, while they were much less abundant in the R library. This phenomenon was more noticeable for the Bacillus- and Sporosarcina-related sequences, present in the D library but totally absent from the R library. Although the combination of primers for PCR and RT-PCR and the number of analyzed clones in each library were different, this discrepancy between R and D libraries could also indicate that spore-forming microorganisms were present in the stromatolite but these bacteria were not an active part of the microbiota at the time of sampling. In addition, a likely source of overestimation of this group of clones in the D library is the high number of rRNA gene copies present in sporeforming microorganisms, such as Bacillus or Clostridium spp., which can harbour up to 15 rRNA operons, in contrast with the one or two copies found in some cyanobacteria (Klappenbach et al. 2000). The Cyanobacteria most frequently recovered in this study were Oscillatoriales related to Leptolyngbya spp. Such Leptolyngbya representatives have been isolated from desert soil from different sites in the western United States (Payne et al. 2001), from microbiotic crusts of Natural Bridges National Monument, Utah, USA (Nagy and Johansen 2001), and from different aquatic environments (Miller and Castenholz 2001). Pseudoanabaenaceae family members were also an important part of cyanobacterium-dominated cryptoendolithic communities from the McMurdo Dry Valleys in the Antarctica (de la Torre et al. 2003). Moreover, clones corresponding to Pseudoanabaenaceae family members have been found when analyzing 16S rRNA gene libraries from hot springs (Papke et al. 2003), permanent ice covers (Priscu et al. 1998) and microbial mat samples (Taton et al. 2003) from Antarctic lakes in McMurdo Dry Valleys and east Antarctic lakes (Taton et al. 2006a,b) in which Oscillatoriales constituted the major group of cyanobacterial sequences retrieved. In previously analyzed stromatolites, most of the sequences were also closely related to Cyanobacteria (Neilan et al. 2002), although they varied depending on the sample. For instance, Schizothrix and Solentia have been found to be the dominant cyanobacteria in modern, marine stromatolites at Highborne Cay, Bahamas (Visscher et al. 1998). The genera Euhalothece and Prochloron were found in stromatolites from the hypersaline marine environment of Shark Bay, Australia (Burns et al. 2004). Sequences related to Microcoleus, although present in a small fraction of the analyzed clones, were associated with stromatolites from Hamelin Pool, also in Shark Bay (Papineau et al. 2005). A recent study based on metagenomic and stable isotopic analyses of modern freshwater stromatolites (Breitbart et al. 2009) in Cuatro Ciénagas, Mexico, also indicated that one of the microbialite metagenomes studied, that of Río Mesquites, was dominated by cyanobacterial sequences. Many of the cyanobacterial sequences retrieved from the Ruidera stromatolites were similar (or even identical) to each other and clustered in the tree shown in Fig. 3. Indeed, a low number of cyanobacterial-related genetic groups were recovered, while the diversity displayed for the non-cyanobacterial sequences obtained in this study was considerably higher. Molecular studies based on 16S rDNA amplification in giant microbialites, ikaite tufa columns or hypersaline stromatolites of Shark Bay (Burns et al. 2004; LópezGarcía et al. 2005; Stougaard et al. 2002) have revealed the presence of bacterial lineages similar to those obtained in this work. Highly diverse non-cyanobacterial communities have also been detected in freshwater microbialites from Pozas Azules, in Cuatro Ciénagas, Mexico (Breitbart et al. 2009), in which the metagenome was dominated by “heterotrophic” bacteria. In addition, Laval et al. (2000) studied modern freshwater microbialites from Pavilion Lake, British Columbia, Canada, and also found that Cyanobacteria dominated microbial mats (identified by phenotypic traits), although “numerous heterotrophs” and diatoms were also present. This phenomenon has also been observed in microbial mats (Paerl
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et al. 2001) or even in marine picoplankton (Vaulot et al. 2002), where a low diverse primary autotrophic community supports a productive, functionally and phylogenetically more diverse heterotrophic bacterial community. Finally, it is remarkable that only bacterial sequences could be retrieved from the Ruidera stromatolite. Archaeal sequences were retrieved from stromatolites in the hypersaline marine environment of Shark Bay, Australia (Burns et al. 2004), as well as in metagenomic analysis from Mexican microbialites (Breitbart et al. 2009) in which sequences that could be assigned to Archaea accounted for, respectively, 13% and 5% of the total metagenomes analyzed from Pozas Azules and Río Mesquites microbialites. However, studies of giant microbialites from the highly alkaline Lake Van, Turkey (López-García et al. 2005) or ikaite tufa columns in Greenland (Stougaard et al. 2002) also failed to amplify archaeal 16S rRNA genes, suggesting that, if present, Archaea were not major components of the autochthonous community at the time of sampling.
Acknowledgments This work was supported by grants CGL2006-12714-C02-01 from the Spanish Ministry of Science and Education. We are grateful to Dr. Ignacio Luque for his advice with the cyanobacterial cultures and DNA extraction, to Jesús Fernández Lentisco for his help with Linux, to Cristina Almansa for her help with confocal microscopy, and to Asunción de los Ríos for her critical reading of the manuscript and very helpful comments.
References Alonso, C., 2005. Identity and activity of marine microbial populations as revealed by single cell techniques. Ph.D. Thesis, Max Planck Institute for Marine Microbiology, Bremen, Germany. S.F. Altschul, T.L. Madden, A. Schäffer, J. Zhang, W. Miller, D.J. Lipman, Gapped BLAST and PSI-BLAST: a new generation of protein database search programs, Nucleic Acids Res. 25 (1997) 3389–3402. R.I. Amann, W. Ludwig, K.H. Schleifer, Phylogenetic identification and in situ detection of individual microbial cells without cultivation, Microbiol. Rev. 59 (1995) 143–169. K.E. Ashelford, N.A. Chuzhanova, J.C. Fry, A.J. Jones, A.J. Weightman, At least 1 in 20 16S rRNA sequence records currently held in public repositories is estimated to contain substantial anomalies, Appl. Environ. Microbiol. 71 (2005) 7724–7736. K.E. Ashelford, N.A. Chuzhanova, J.C. Fry, A.J. Jones, A.J. Weightman, New screening software shows that most recent large 16S rRNA gene clone libraries contain chimeras, Appl. Environ. Microbiol. 72 (2006) 5734–5741. M. Breitbart, A. Hoare, A. Nitti, J. Siefert, M. Haynes, E. Dinsdale, R. Edwards, V. Souza, F. Rohwer, D. Hollander, Metagenomic and stable isotopic analyses of modern freshwater microbialites in Cuatro Ciénagas, Mexico, Environ. Microbiol. 11 (2009) 16–34. B.P. Burns, F. Goh, M. Allen, B.A. Neilan, Microbial diversity of extant stromatolites in the hypersaline marine environment of Shark Bay, Australia, Environ. Microbiol. 6 (2004) 1096–1101. A. Cifuentes, J. Antón, S. Benlloch, A. Donnelly, R.A. Herbert, F. Rodríguez-Valera, Prokaryotic diversity in Zostera noltii-colonized marine sediments, Appl. Environ. Microbiol. 66 (2000) 1715–1719. J.R. de la Torre, B.M. Goebel, E.I. Friedmann, N.R. Pace, Microbial diversity of cryptoendolithic communities from the McMurdo Dry Valleys, Antarctica, Appl. Environ. Microbiol. 69 (2003) 3858–3867. E.F. DeLong, Archaea in coastal marine environments, Proc. Natl. Acad. Sci. U.S.A. 89 (1992) 5685–5689. B. Díez, C. Pedrós-Alió, T.L. Marsh, R. Massana, Application of denaturing gradient gel electrophoresis (DGGE) to study the diversity of marine picoeukaryotic assemblages and comparison of DGGE with other molecular techniques, Appl. Environ. Microbiol. 67 (2001) 2942–2951. ˜ M.A. García del Cura, J.A. González, A. Fernández, S. Ordónez, An ultramodern (post1947) stromatolitic system developed in a small pool, upper Guadiana river (Central Spain), in: IAS-ASF-IGCP 380 International Workshop on Microbial Mediation in Carbonate Diagenesis, vol. 26, Publication ASF, Paris, France, 1997, pp. 27–29. F. García-Pichel, A. López-Cortés, U. Nübel, Phylogenetic and morphological diversity of cyanobacteria in soil desert crusts from the Colorado Plateau, Appl. Environ. Microbiol. 67 (2001) 1902–1910. Ø. Hammer, D.A.T. Harper, P.D. Ryan, PAST: paleontological statistics software package for education and data analysis, Paleontol. Electrón. 4 (2001) 9.
S.A. Havemann, J.S. Foster, Comparative characterization of the microbial diversity of an artificial microbialite model and a natural stromatolite, Appl. Environ. Microbiol. 74 (2008) 7410–7421. T. Ishida, M.M. Watanabe, J. Sugiyama, A. Yokota, Evidence for polyphyletic origin of the members of the orders of Oscillatoriales and Pleurocapsales as determined by 16S rDNA analysis, FEMS Microbiol. Lett. 201 (2001) 79–82. J.A. Klappenbach, J.M. Dunbar, T.M. Schmit, rRNA operon copy number reflects ecological strategies for bacteria, Appl. Environ. Microbiol. 66 (2000) 1328–1333. B. Laval, S.L. Cady, J.C. Pollack, C.P. McKay, J.S. Bird, J.P. Grotzinger, D.C. Ford, H.R. Bohm, Modern freshwater microbialite analogues for ancient dendritic reef structures, Nature 407 (2000) 626–629. M.K. Liviatis, A molecular test of cyanobacterial phylogeny: inferences from constraint analyses, Hydrobiologia 468 (2002) 135–145. P. López-García, J. Kazmierczak, K. Benzeara, S. Kempe, F. Guyot, D. Moreira, Bacterial diversity and carbonate precipitation in the giant microbialites from the highly alkaline Lan Van, Turkey, Extremophiles 9 (2005) 263–274. W. Ludwig, O. Strunk, R. Westram, L. Richter, H. Meier, Yadhukumar, A. Buchner, T. Lai, S. Steppi, G. Jobb, W. Förster, I. Brettske, S. Gerber, A.W. Ginhart, O. Gross, S. Grumann, S. Hermann, R. Jost, A. König, T. Liss, R. Lüßmann, M. May, B. Nonhoff, B. Reichel, R. Strehlow, A. Stamatakis, N. Stuckmann, A. Vilbig, M. Lenke, T. Ludwig, A. Bode, K.H. Schleifer, ARB: a software environment for sequence data, Nucleic Acids Res. 32 (2004) 1363–1371. S.R. Miller, R.W. Castenholz, Ecological physiology of Synechochoccus sp. strain SH94-5, a naturally occurring cyanobacterium deficient in nitrate assimilation, Appl. Environ. Microbiol. 67 (2001) 3002–3009. I.P. Miskin, P. Farrimond, I.M. Head, Identification of novel bacterial lineages as active members of microbial populations in a freshwater sediment using a rapid RNA extraction procedure and RT-PCR, Microbiology 145 (1999) 1977–1987. G. Muyzer, T. Brinkhoff, U. Nübel, C. Santegoeds, H. Schäfer, C. Wawer, Denaturing gradient gel electrophoresis (DGGE) in microbial ecology, in: A.D.L. Akkermans, J.D. van Elsas, F.J. Bruijn (Eds.), Molecular Microbial Ecology Manual, Kluwer Academic Publishers, Dordrecht, The Netherlands, 1998, p. 3.4.4/1-3.4.4/27. G. Muyzer, E.C. de Waal, A.G. Uitterlinden, Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA, Appl. Environ. Microbiol. 59 (1993) 695–700. M.L. Nagy, J.R. Johansen, Cyanobacterial flora of microbiotic crusts of Natural Bridges National Monument, Utah, J. Phycol. 37 (2001) 38. B.A. Neilan, B.P. Burns, D.A. Relman, D.R. Lowe, Molecular identification of Cyanobacteria associated with stromatolites from distinct geographical locations, Astrobiology 2 (2002) 271–280. B. Nelissen, R. de Baere, A. Wilmotte, R. de Watcher, Phylogenetic relationships of nonaxenic filamentous cyanobacterial strains based on 16S rRNA sequences analysis, J. Mol. Evol. 42 (1996) 194–200. B. Nogales, R.B. Moore, E. Llobet-Brossa, R. Rosselló-Mora, R. Amann, K.N. Timmis, Combined use of 16S ribosomal DNA and 16S rRNA to study the bacterial community of polychlorinated biphenyl-polluted soils, Appl. Environ. Microbiol. 67 (2001) 1874–1884. U. Nübel, F. García-Pichel, G. Muyzer, PCR primers to amplify 16S rRNA genes from cyanobacteria, Appl. Environ. Microbiol. 63 (1997) 3327–3332. ˜ S. Ordónez, M.A. García del Cura, Recent and tertiary fluvial carbonates in Central Spain, Spe Publs Int Ass Sediment 6 (1983) 485–497. ˜ S. Ordónez, J.A. González Martín, M.A. García del Cura, Tipología y génesis de depósitos tobáceos fluvio-lacustres: el sistema tobáceo de Las Lagunas de Ruidera, Cuadernos de Geología Ibérica 22 (1997) 333–348. ˜ S. Ordónez, J.A. González Martín, M.A. García del Cura, H.M. Pedley, Temperate and semi-arid tufas in Pleistocene to Recent fluvial barrage system in the Mediterranean area: The Ruidera Lakes Natural Park (Central Spain), Geomorphology 69 (2005) 332–350. H.W. Paerl, T.F. Steppe, R.P. Reid, Bacterially mediated precipitation in marine stromatolites, Environ. Microbiol. 3 (2001) 123–130. D. Papineau, J.J. Walker, S.J. Mojzsis, N.R. Pace, Composition and structure of microbial communities from stromatolites of Hamelin Pool in Shark Bay, Western Australia, Appl. Environ. Microbiol. 71 (2005) 4822–4832. R.T. Papke, N.B. Ramsing, M.M. Bateson, D.W. Ward, Geographical isolation in hot spring Cyanobacteria, Environ. Microbiol. 5 (2003) 650–659. M. Payne, J.R. Johansen, S.L. Boyer, Taxonomic resolution of Leptolyngbya (Cyanophyta) utilizing the 16S rRNA gene sequence, J. Phycol. 37 (2001) 40–41. J.C. Priscu, C.H. Fritsen, E.E. Adams, S.J. Giovannoni, H.W. Paerl, C.P. McKay, P.T. Doran, D.A. Gordon, B.D. Lanoil, J.L. Pinckney, Perennial Antarctic lake ice: an oasis for life in a polar desert, Science 280 (1998) 2095–2098. R. Rippka, Recognition and identification of Cyanobacteria, Methods Enzymol. 167 (1988) 28–67. V. Souza-Egipsy, M.A. García del Cura, C. Ascaso, A. de Los Ríos, J. Wierzchos, J.A. González-Martín, Interaction between calcite and phosphorous in biomineralization processes in tufa carbonates, Int. Rev. Hydrobiol. 91 (2005) 222–241. P. Stougaard, F. Jørgensen, M.D. Johnsen, O.C. Hansen, Microbial diversity in ikaite tufa columns: an alkaline, cold ecological niche in Greenland, Environ. Microbiol. 4 (2002) 487–493. A. Taton, S. Grubisic, E. Brambilla, R. de Wit, A. Wilmotte, Cyanobacterial diversity in natural and artificial microbial mats of Lake Fryxell (McMurdo Dry Valleys, Antarctica): a morphological and molecular approach, Appl. Environ. Microbiol. 69 (2003) 5157–5169. A. Taton, S. Grubisic, P. Barthasart, D.A. Hodgson, J. Laybourn-Parry, A. Wilmotte, Biogeographical distribution and ecological ranges of benthic Cyanobacteria in East Antarctic lakes, FEMS Microbiol. Ecol. 57 (2006a) 272–289.
F. Santos et al. / Systematic and Applied Microbiology 33 (2010) 209–221 A. Taton, S. Grubisic, D. Ertz, D.A. Hodgson, R. Piccardi, N. Biondi, M.R. Tredici, M. Mainini, D. Losi, F. Marinelli, A. Wilmotte, Polyphasic study of Antarctic cyanobacterial strains, J. Phycol. 42 (2006b) 1257–1270. J.R. Thompson, L.A. Marcelino, M.F. Polz, Heteroduplexes in mixed-template amplifications: formation consequence and elimination by “reconditioning PCR”, Nucleic Acid Res. 30 (2002) 2083–2088. D. Vaulot, K. Romari, F. Not, Are autotrophs less diverse than heterotrophs in marine picoplankton? Trends Microbiol. 10 (2002) 266–267. P.T. Visscher, R.P. Reid, B.M. Bebout, S.E. Hoeft, I.G. Macintyre, J.A. Thompson Jr., Formation of lithified micritic laminae in modern marine stromatolites (Bahamas): the role of sulfur cycling, Am. Mineral. 83 (1998) 1482–1493.
221
M. Wagner, G. Rath, R. Amann, H.P. Koops, K.H. Schleifer, In situ identification of ammonia-oxidizing bacteria, Syst. Appl. Microbiol. 17 (1995) 251–264. J.J. Walker, N.R. Pace, Phylogenetic composition of Rocky Mountain endolithic microbial ecosystems, Appl. Environ. Microbiol. 73 (2007) 3497–3504. M.R. Walter, Introduction, in: M.R. Walter (Ed.), Stromatolites, Elsevier, Amsterdam, The Netherlands, 1976, pp. 1–3. E. Yergeau, K.K. Newsham, D.A. Pearce, G.A. Kowalchuk, Patterns of bacterial diversity across a range of Antarctic terrestrial habitats, Environ. Microbiol. 9 (2007) 2670–2682.
