Microb Ecol DOI 10.1007/s00248-014-0412-7
SOIL MICROBIOLOGY
Characterization of Chasmoendolithic Community in Miers Valley, McMurdo Dry Valleys, Antarctica Charmaine C. M. Yung & Yuki Chan & Donnabella C. Lacap & Sergio Pérez-Ortega & Asuncion de los Rios-Murillo & Charles K. Lee & S. Craig Cary & Stephen B. Pointing
Received: 28 November 2013 / Accepted: 11 March 2014 # Springer Science+Business Media New York 2014
Abstract The Antarctic Dry Valleys are unable to support higher plant and animal life and so microbial communities dominate biotic ecosystem processes. Soil communities are well characterized, but rocky surfaces have also emerged as a significant microbial habitat. Here, we identify extensive colonization of weathered granite on a landscape scale by chasmoendolithic microbial communities. A transect across north-facing and south-facing slopes plus valley floor moraines revealed 30–100 % of available substrate was colonized up to an altitude of 800 m. Communities were assessed at a multidomain level and were clearly distinct from those in surrounding soils and other rock-inhabiting cryptoendolithic and hypolithic communities. All colonized rocks were dominated by the cyanobacterial genus Leptolyngbya (Oscillatoriales), with heterotrophic bacteria, archaea, algae, and fungi also identified. Striking patterns in community distribution were evident with regard to microclimate as Electronic supplementary material The online version of this article (doi:10.1007/s00248-014-0412-7) contains supplementary material, which is available to authorized users. C. C. M. Yung Nicholas School of the Environment, Duke University, Durham, NC 27708, USA Y. Chan : D. C. Lacap : S. B. Pointing Institute for Applied Ecology New Zealand, School of Applied Sciences, Auckland University of Technology, Private Bag 92006, Auckland 1142, New Zealand S. Pérez-Ortega : A. de los Rios-Murillo Department of Environmental Biology, Museo Nacional de Ciencias Naturales (CSIC), C/ Serrano 115 Duplicado, 28006 Madrid, Spain C. K. Lee : S. C. Cary : S. B. Pointing (*) The International Centre for Terrestrial Antarctic Research, Department of Biological Sciences, University of Waikato, Private Bag 3105, Hamilton 3240, New Zealand e-mail: [email protected]
determined by aspect. Notably, a shift in cyanobacterial assemblages from Chroococcidiopsis-like phylotypes (Pleurocapsales) on colder–drier slopes, to Synechococcuslike phylotypes (Chroococcales) on warmer–wetter slopes. Greater relative abundance of known desiccation-tolerant bacterial taxa occurred on colder–drier slopes. Archaeal phylotypes indicated halotolerant taxa and also taxa possibly derived from nearby volcanic sources. Among the eukaryotes, the lichen photobiont Trebouxia (Chlorophyta) was ubiquitous, but known lichen-forming fungi were not recovered. Instead, fungal assemblages were dominated by ascomycetous yeasts. We conclude that chasmoendoliths likely constitute a significant geobiological phenomenon at lower elevations in granite-dominated Antarctic Dry Valley systems.
Introduction Exposed rocky terrain is a major feature of arid environments worldwide [1]. It is increasingly realized that such rocky substrate supports a near-contiguous microbial colonization that has major importance in weathering, hydrological cycles and nutrient transformation [2]. Although the most visibly obvious colonization is epilithic lichens and mosses on exposed surfaces, subsurface colonization is emerging as a major contributor to lithic biomass in both hot and cold deserts [2, 3]. Such colonization is usually classified according to how microorganisms occupy the rock matrix. Cryptoendolithic communities exploit the pore spaces between crystals in porous and weathered substrates such as sandstone and limestone [4], whereas chasmoendolithic communities develop in the cracks and fissures of porous and nonporous rocky substrates that are exposed to the ambient surroundings [4]. These cryptic modes of colonization are viewed as a stress avoidance strategy where the habitat beneath the rock surface provides buffering from extremes of thermal, moisture, and UV stress as well as
C. C. M. Yung et al.
substrate stability [5]. While the biodiversity of cryptoendoliths in both cold and hot deserts is relatively well studied, little is known about chasmoendoliths. Community assembly in cryptoendoliths has been observed to develop in a layered state that has been explained by the ‘microbial cabana’ hypothesis, where upper (nearsurface) colonists that are relatively UV-tolerant protect underlying bands of less tolerant colonists [2]. The Antarctic deserts of the McMurdo Dry Valleys have long been acknowledged to support biologically unique cryptoendolithic communities, and these have been well studied from sandstone substrates that dominate the inland Dry Valleys. Friedmann [6] identified two compositional classes of Antarctic cryptoendoliths: lichenized and cyanobacterial, based largely on studies of sandstone. Endolithic lichenized communities were typically described as comprising the chlorophyte Trebouxia with unidentified mycobionts, while cyanobacterial cryptoendoliths in sandstone were attributed to the cyanobacterial genus Chroococcidiopsis [6]. The advent of molecular tools to resolve community molecular diversity in culture-independent studies has allowed resolution of far greater diversity than was previously appreciated by morphological and cultivation studies [7]. Studies of cryptoendoltihs in Antarctic Beacon sandstone using ‘universal’ 16S/18S ribosomal RNA (rRNA) gene primers revealed complex communities of algae, fungi, and bacteria [8, 9]. A lichendominated community comprised around 70 % algal (Trebouxia) and fungal phylotypes, while the dominant phylotypes in a cyanobacterial endolithic community were identified as cyanobacteria, alpha-proteobacteria, and deinococci. Archaea were not detected in these studies despite the use of primers specifically targeting this domain. Broader comparison with nonpolar locations suggests that cyanobacteria are the keystone and dominant phylum in most endolithic substrates [10–12], and other exposed rocky microbial habitats such as quartz hypoliths [13–17]. This likely reflects the cyanobacterial role as primary producers and their secretion of extracellular polymers prolifically within rocks that are strongly implicated in tolerance to environmental stress in lithic habitats [18]. Previous studies in nonpolar locations have elucidated some aspects of chasmoendolithic biodiversity, notably cyanobacteria including the observation of Chroococcidiopsis and chlorophyte algae [19, 20]. Since the first description of Antarctic cryptoendolithic and chasmoendolithic colonization of sandstone [6]. Only one study has reported multidomain biodiversity in an Antarctic sandstone chasmoendolithic system; this indicated that cryptoendoliths and chasmoendoliths in Beacon sandstone displayed few differences in community assembly [9]. This was explained in terms of the homogeneity between substrate and microenvironment in the two niches. Granite-dominated valleys present a fundamentally different ecological niche both physically and geochemically from
sandstone, and importantly, cryptoendolithic colonization is not a feature unless the rock is highly weathered due to the nonporous nature of the unweathered granite matrix. Ultrastructural and genetic studies suggest granite chasmoendoliths may support a distinctive lichen community [21, 22]. Granitic chasmoendolithic communities are therefore likely a major source of standing biomass in such dry valley systems, alongside open soil communities [9, 23]. In this study, we characterized the distribution and biodiversity of chasmoendoliths in weathered granite of the Miers Valley. The Miers Valley is a long-term study site for the New Zealand Terrestrial Antarctic Biodiversity Survey (NZTABS) in the McMurdo Dry Valleys of Antarctica. Weathered granite is the predominant rocky substrate in this valley system (and in other nearby Dry Valleys) and is characterized by a highly eroded and porous surface layer that is prone to microbial colonization. Characterizing its biology has key importance to understanding regional ecology in granite-dominated Dry Valleys. We describe the occurrence of these communities in relation to valley topography and microclimate, and the multidomain community composition and phylogenetic affiliations as determined by environmental rRNA gene sequences.
Materials and Methods Field Location, Microclimate, and Biological Sampling Miers Valley is a granite-dominated valley occupying a maritime location within the McMurdo Dry Valleys Antarctic Special Managed Area. It is a long-term ecological study site for the New Zealand Terrestrial Antarctic Biodiversity Survey (NZTABS, http://nztabs.ictar.aq). The valley lies between the latitudes 78°060 S and 78°070 S and longitudes 163°440 E and 164°120 E and comprises a wide valley floor characterized by moraine deposition, and steep scree and boulder slopes. A field survey was conducted in Miers Valley in December 2009. A single linear transect across the valley floor and up to 800 m on both the north-facing and south-facing slopes was used for sample recovery (S78″04. 608, E163″48.797–S78′06.771, E163″48.284, approximately 9 km). Field estimates of colonization were made along the transect at 200 m altitudinal increments. At each location, visual searching for ten person minutes was undertaken within a 10 m radius of the transect line. Granitic rock was examined for laminar flaking and colonization by chasmoendoliths. A maximum of 100 rocks were examined within each 10-min period. At each location, colonized rock fragments (approx. 50 g each) were aseptically collected from three separate rocks for subsequent laboratory analysis (n=45). All rock samples were stored frozen in the field in darkness, and subsequently, at −80 °C in the laboratory until processed.
Ecology of Antarctic Chasmoendoliths
Surface microenvironment along the transect was characterized using iButton dataloggers (DS1923 Maxim Integrated, Sunnyvale, CA, USA). These were fixed at the ground surface using plastic pegs, and we recorded temperature and relative humidity continuously during the austral summer immediately preceding sampling (November–January 2009) [24]. The granitic nature of all substrates was assessed in the field according to standard USGS mineralogical criteria. Elemental mapping of substrate surfaces to confirm granitic nature was achieved by energy-dispersive X-ray spectroscopy (EDX; Oxford Instruments, INCAx-sight EDS Detectors, INCA Energy Software) under an extra high tension (EHT) of 20 kV using scanning electron microscopy of gold-coated samples (Hitachi S-4800 FEG SEM). Three randomly selected squares of 50×50 mm at a magnification of 25× were scanned for each rock sample. Five iterations were performed automatically for each read. This demonstrated a silicon:oxygen ratio of 1:2 and was indicative of granite (SiO2 >60 %), with trace levels of some metals including aluminum, calcium, potassium, and sodium. No significant variation in mineral composition was detected among samples. Scanning Electron Microscopy Visual characterization of microscale colonization was achieved using scanning electron microscopy [25]. In brief, the pieces of rock were first fixed in glutaraldehyde and then in osmium tetroxide, dehydrated in a series of ethanol solutions, and embedded in LR-White resin. Blocks of resinembedded rock samples were finely polished, carbon-coated, and observed using a Zeiss DMS 960 SEM microscope. DNA Recovery, PCR, and tRFLP Environmental DNA recovery was achieved separately for each sample by lysis in cetyl trimethylammonium bromide (CTAB) with lysozyme and RNAse, followed by phenol:chloroform extraction at 60 °C. Genomic DNA was checked for quality by electrophoresis in 1 % agarose gels and quantified by spectrophotometry (Bio-Rad Smartspec Plus). PCR reactions for terminal restriction fragment length polymorphism (t-RFLP) analysis were carried out using a FAMlabeled forward primer (Table S1). Gel-purified amplicons were digested using three restriction enzymes (HaeIII, Hinf1, and MspI) and the most informative selected for further analysis (MspI for 16S/18S rRNA, HaeIII for ITS). Fragment analysis was achieved by capillary electrophoresis (Applied Biosystems 3730 Genetic Analyzer), using a GeneScan ROXlabeled GS500 internal size standard. The t-RFLP patterns and quality were analyzed using the freeware PeakScanner™ (version 1.0; Applied Biosystems; https://products. appliedbiosystems.com), and a data matrix comprising fragment size and abundance was generated. The software
Perl and R were then used to identify true peaks and bin fragments of similar size [26]. The relative abundance of a true terminal restriction fragment within a given t-RFLP pattern was generated as a ratio of the respective peak area. A virtual digest using HaeIII and MspI was carried out on sequences retrieved from the bacterial, archaeal, and eukaryal clone libraries. This allowed the assignment of phylogenetic identity to individual t-RFLP peaks. After discarding ambiguous t-RFLP profiles and negative results, final n = 44 (bacteria), n=21 (archaea), and n=29 (eukarya). Sequence Acquisition Samples that were most parsimonious to all others within a given NMDS cluster (see below) were selected for clone library construction. Each amplicon was gel-purified and used as template for construction of clone libraries (Qiagen PCR Cloning plus kit, CA, USA), using domain specific primers (Table S1). Plasmids were extracted from positive transformants (Mini-M™ Plasmid DNA extraction system, Viogene, Taiwan) and screened by restriction fragment length polymorphism (RFLP) using the restriction endonucleases MspI, HaeIII, and CfoI. We set a sampling cut-off of 250 clones (bacteria) and 100 clones (archaea and bacteria), based upon richness estimates obtained from studies of similar locations [9]. At least three samples from each distinct RFLP pattern were sequenced where possible, using the BigDye Terminator Cycle Sequencing kit (Applied Biosystems, CA, USA; Applied Biosystems 3730 Genetic Analyzer). Phylotypes were delineated on the basis of 97 % sequence similarity using the freeware DOTUR [27]. All sequences generated by this study have been deposited in the NCBI GenBank database under accession numbers KC476241–KC476287 and KF294470–KF294500. Screening for possible chimeric sequences was made using Chimera_Check available on the Ribosome Database Project website (http://rdp.cme.msu.edu. html). Approximate phylogentic affiliations were then determined by BLAST searches of the NCBI GenBank database (http://www.ncbi.nlm.nic.gov). Estimates of clone library sampling effort were made using the freeware EstimateS [28]. Sampling effort was assessed by calculation of Coverage and Rarefaction curves, estimates of library richness were made using the nonparametric estimators ACE and Chao 1. All OTU delineation was made on the basis of sequenced phylotypes. Phylogenetic and Statistical Analyses Multiple alignments were created with reference to selected GenBank sequences using BioEdit v7.0.9.0 [29]. The alignments were tested against prescript models of evolution using the softwares PAUP* 4.0b10 [30] and Modeltest v3.0 [31]. The criteria described by the most appropriate evolutionary
C. C. M. Yung et al.
model were input for maximum likelihood analysis using Genetic Algorithm for Rapid Likelihood Inference (GARLI) Version 0.96 Beta [32]. Robustness of furcated branches was supported by both bootstrap values (1,000 replicates) determined using PAUP* 4.0b10 and Bayesian posterior probabilities [33] calculated using Mr. Bayes v3.0b4 [34]. Values (in percent) were shown on all branch nodes supported by more than 50 % of the trees. Statistical analysis was conducted as follows: alpha diversity indices (Shannon’s index, Simpsons diversity index, Pielou’s evenness) were calculated using untransformed data. Differences stated as significant were tested using one-way and two-way analysis of variance (ANOVA), or analysis of similarity (ANOSIM). The ANOSIM produces a statistic R which is based on the difference of mean ranks between groups and within groups [35]. A value approaching to 1 indicates the assemblage composition is totally different, where a value of 0 indicates no difference. Multivariate analysis of diversity data was performed on square-roottransformed diversity data and on nontransformed normalized data for environmental variables. Nonmetric multidimensional scaling (NMDS) ordinations were used to visualize Bray Curtis similarities (diversity data) and Euclidean distances (environmental data). All analyses were performed using Primer v6.1.6 [35]. All results stated as significant have a confidence level of P
SOIL MICROBIOLOGY
Characterization of Chasmoendolithic Community in Miers Valley, McMurdo Dry Valleys, Antarctica Charmaine C. M. Yung & Yuki Chan & Donnabella C. Lacap & Sergio Pérez-Ortega & Asuncion de los Rios-Murillo & Charles K. Lee & S. Craig Cary & Stephen B. Pointing
Received: 28 November 2013 / Accepted: 11 March 2014 # Springer Science+Business Media New York 2014
Abstract The Antarctic Dry Valleys are unable to support higher plant and animal life and so microbial communities dominate biotic ecosystem processes. Soil communities are well characterized, but rocky surfaces have also emerged as a significant microbial habitat. Here, we identify extensive colonization of weathered granite on a landscape scale by chasmoendolithic microbial communities. A transect across north-facing and south-facing slopes plus valley floor moraines revealed 30–100 % of available substrate was colonized up to an altitude of 800 m. Communities were assessed at a multidomain level and were clearly distinct from those in surrounding soils and other rock-inhabiting cryptoendolithic and hypolithic communities. All colonized rocks were dominated by the cyanobacterial genus Leptolyngbya (Oscillatoriales), with heterotrophic bacteria, archaea, algae, and fungi also identified. Striking patterns in community distribution were evident with regard to microclimate as Electronic supplementary material The online version of this article (doi:10.1007/s00248-014-0412-7) contains supplementary material, which is available to authorized users. C. C. M. Yung Nicholas School of the Environment, Duke University, Durham, NC 27708, USA Y. Chan : D. C. Lacap : S. B. Pointing Institute for Applied Ecology New Zealand, School of Applied Sciences, Auckland University of Technology, Private Bag 92006, Auckland 1142, New Zealand S. Pérez-Ortega : A. de los Rios-Murillo Department of Environmental Biology, Museo Nacional de Ciencias Naturales (CSIC), C/ Serrano 115 Duplicado, 28006 Madrid, Spain C. K. Lee : S. C. Cary : S. B. Pointing (*) The International Centre for Terrestrial Antarctic Research, Department of Biological Sciences, University of Waikato, Private Bag 3105, Hamilton 3240, New Zealand e-mail: [email protected]
determined by aspect. Notably, a shift in cyanobacterial assemblages from Chroococcidiopsis-like phylotypes (Pleurocapsales) on colder–drier slopes, to Synechococcuslike phylotypes (Chroococcales) on warmer–wetter slopes. Greater relative abundance of known desiccation-tolerant bacterial taxa occurred on colder–drier slopes. Archaeal phylotypes indicated halotolerant taxa and also taxa possibly derived from nearby volcanic sources. Among the eukaryotes, the lichen photobiont Trebouxia (Chlorophyta) was ubiquitous, but known lichen-forming fungi were not recovered. Instead, fungal assemblages were dominated by ascomycetous yeasts. We conclude that chasmoendoliths likely constitute a significant geobiological phenomenon at lower elevations in granite-dominated Antarctic Dry Valley systems.
Introduction Exposed rocky terrain is a major feature of arid environments worldwide [1]. It is increasingly realized that such rocky substrate supports a near-contiguous microbial colonization that has major importance in weathering, hydrological cycles and nutrient transformation [2]. Although the most visibly obvious colonization is epilithic lichens and mosses on exposed surfaces, subsurface colonization is emerging as a major contributor to lithic biomass in both hot and cold deserts [2, 3]. Such colonization is usually classified according to how microorganisms occupy the rock matrix. Cryptoendolithic communities exploit the pore spaces between crystals in porous and weathered substrates such as sandstone and limestone [4], whereas chasmoendolithic communities develop in the cracks and fissures of porous and nonporous rocky substrates that are exposed to the ambient surroundings [4]. These cryptic modes of colonization are viewed as a stress avoidance strategy where the habitat beneath the rock surface provides buffering from extremes of thermal, moisture, and UV stress as well as
C. C. M. Yung et al.
substrate stability [5]. While the biodiversity of cryptoendoliths in both cold and hot deserts is relatively well studied, little is known about chasmoendoliths. Community assembly in cryptoendoliths has been observed to develop in a layered state that has been explained by the ‘microbial cabana’ hypothesis, where upper (nearsurface) colonists that are relatively UV-tolerant protect underlying bands of less tolerant colonists [2]. The Antarctic deserts of the McMurdo Dry Valleys have long been acknowledged to support biologically unique cryptoendolithic communities, and these have been well studied from sandstone substrates that dominate the inland Dry Valleys. Friedmann [6] identified two compositional classes of Antarctic cryptoendoliths: lichenized and cyanobacterial, based largely on studies of sandstone. Endolithic lichenized communities were typically described as comprising the chlorophyte Trebouxia with unidentified mycobionts, while cyanobacterial cryptoendoliths in sandstone were attributed to the cyanobacterial genus Chroococcidiopsis [6]. The advent of molecular tools to resolve community molecular diversity in culture-independent studies has allowed resolution of far greater diversity than was previously appreciated by morphological and cultivation studies [7]. Studies of cryptoendoltihs in Antarctic Beacon sandstone using ‘universal’ 16S/18S ribosomal RNA (rRNA) gene primers revealed complex communities of algae, fungi, and bacteria [8, 9]. A lichendominated community comprised around 70 % algal (Trebouxia) and fungal phylotypes, while the dominant phylotypes in a cyanobacterial endolithic community were identified as cyanobacteria, alpha-proteobacteria, and deinococci. Archaea were not detected in these studies despite the use of primers specifically targeting this domain. Broader comparison with nonpolar locations suggests that cyanobacteria are the keystone and dominant phylum in most endolithic substrates [10–12], and other exposed rocky microbial habitats such as quartz hypoliths [13–17]. This likely reflects the cyanobacterial role as primary producers and their secretion of extracellular polymers prolifically within rocks that are strongly implicated in tolerance to environmental stress in lithic habitats [18]. Previous studies in nonpolar locations have elucidated some aspects of chasmoendolithic biodiversity, notably cyanobacteria including the observation of Chroococcidiopsis and chlorophyte algae [19, 20]. Since the first description of Antarctic cryptoendolithic and chasmoendolithic colonization of sandstone [6]. Only one study has reported multidomain biodiversity in an Antarctic sandstone chasmoendolithic system; this indicated that cryptoendoliths and chasmoendoliths in Beacon sandstone displayed few differences in community assembly [9]. This was explained in terms of the homogeneity between substrate and microenvironment in the two niches. Granite-dominated valleys present a fundamentally different ecological niche both physically and geochemically from
sandstone, and importantly, cryptoendolithic colonization is not a feature unless the rock is highly weathered due to the nonporous nature of the unweathered granite matrix. Ultrastructural and genetic studies suggest granite chasmoendoliths may support a distinctive lichen community [21, 22]. Granitic chasmoendolithic communities are therefore likely a major source of standing biomass in such dry valley systems, alongside open soil communities [9, 23]. In this study, we characterized the distribution and biodiversity of chasmoendoliths in weathered granite of the Miers Valley. The Miers Valley is a long-term study site for the New Zealand Terrestrial Antarctic Biodiversity Survey (NZTABS) in the McMurdo Dry Valleys of Antarctica. Weathered granite is the predominant rocky substrate in this valley system (and in other nearby Dry Valleys) and is characterized by a highly eroded and porous surface layer that is prone to microbial colonization. Characterizing its biology has key importance to understanding regional ecology in granite-dominated Dry Valleys. We describe the occurrence of these communities in relation to valley topography and microclimate, and the multidomain community composition and phylogenetic affiliations as determined by environmental rRNA gene sequences.
Materials and Methods Field Location, Microclimate, and Biological Sampling Miers Valley is a granite-dominated valley occupying a maritime location within the McMurdo Dry Valleys Antarctic Special Managed Area. It is a long-term ecological study site for the New Zealand Terrestrial Antarctic Biodiversity Survey (NZTABS, http://nztabs.ictar.aq). The valley lies between the latitudes 78°060 S and 78°070 S and longitudes 163°440 E and 164°120 E and comprises a wide valley floor characterized by moraine deposition, and steep scree and boulder slopes. A field survey was conducted in Miers Valley in December 2009. A single linear transect across the valley floor and up to 800 m on both the north-facing and south-facing slopes was used for sample recovery (S78″04. 608, E163″48.797–S78′06.771, E163″48.284, approximately 9 km). Field estimates of colonization were made along the transect at 200 m altitudinal increments. At each location, visual searching for ten person minutes was undertaken within a 10 m radius of the transect line. Granitic rock was examined for laminar flaking and colonization by chasmoendoliths. A maximum of 100 rocks were examined within each 10-min period. At each location, colonized rock fragments (approx. 50 g each) were aseptically collected from three separate rocks for subsequent laboratory analysis (n=45). All rock samples were stored frozen in the field in darkness, and subsequently, at −80 °C in the laboratory until processed.
Ecology of Antarctic Chasmoendoliths
Surface microenvironment along the transect was characterized using iButton dataloggers (DS1923 Maxim Integrated, Sunnyvale, CA, USA). These were fixed at the ground surface using plastic pegs, and we recorded temperature and relative humidity continuously during the austral summer immediately preceding sampling (November–January 2009) [24]. The granitic nature of all substrates was assessed in the field according to standard USGS mineralogical criteria. Elemental mapping of substrate surfaces to confirm granitic nature was achieved by energy-dispersive X-ray spectroscopy (EDX; Oxford Instruments, INCAx-sight EDS Detectors, INCA Energy Software) under an extra high tension (EHT) of 20 kV using scanning electron microscopy of gold-coated samples (Hitachi S-4800 FEG SEM). Three randomly selected squares of 50×50 mm at a magnification of 25× were scanned for each rock sample. Five iterations were performed automatically for each read. This demonstrated a silicon:oxygen ratio of 1:2 and was indicative of granite (SiO2 >60 %), with trace levels of some metals including aluminum, calcium, potassium, and sodium. No significant variation in mineral composition was detected among samples. Scanning Electron Microscopy Visual characterization of microscale colonization was achieved using scanning electron microscopy [25]. In brief, the pieces of rock were first fixed in glutaraldehyde and then in osmium tetroxide, dehydrated in a series of ethanol solutions, and embedded in LR-White resin. Blocks of resinembedded rock samples were finely polished, carbon-coated, and observed using a Zeiss DMS 960 SEM microscope. DNA Recovery, PCR, and tRFLP Environmental DNA recovery was achieved separately for each sample by lysis in cetyl trimethylammonium bromide (CTAB) with lysozyme and RNAse, followed by phenol:chloroform extraction at 60 °C. Genomic DNA was checked for quality by electrophoresis in 1 % agarose gels and quantified by spectrophotometry (Bio-Rad Smartspec Plus). PCR reactions for terminal restriction fragment length polymorphism (t-RFLP) analysis were carried out using a FAMlabeled forward primer (Table S1). Gel-purified amplicons were digested using three restriction enzymes (HaeIII, Hinf1, and MspI) and the most informative selected for further analysis (MspI for 16S/18S rRNA, HaeIII for ITS). Fragment analysis was achieved by capillary electrophoresis (Applied Biosystems 3730 Genetic Analyzer), using a GeneScan ROXlabeled GS500 internal size standard. The t-RFLP patterns and quality were analyzed using the freeware PeakScanner™ (version 1.0; Applied Biosystems; https://products. appliedbiosystems.com), and a data matrix comprising fragment size and abundance was generated. The software
Perl and R were then used to identify true peaks and bin fragments of similar size [26]. The relative abundance of a true terminal restriction fragment within a given t-RFLP pattern was generated as a ratio of the respective peak area. A virtual digest using HaeIII and MspI was carried out on sequences retrieved from the bacterial, archaeal, and eukaryal clone libraries. This allowed the assignment of phylogenetic identity to individual t-RFLP peaks. After discarding ambiguous t-RFLP profiles and negative results, final n = 44 (bacteria), n=21 (archaea), and n=29 (eukarya). Sequence Acquisition Samples that were most parsimonious to all others within a given NMDS cluster (see below) were selected for clone library construction. Each amplicon was gel-purified and used as template for construction of clone libraries (Qiagen PCR Cloning plus kit, CA, USA), using domain specific primers (Table S1). Plasmids were extracted from positive transformants (Mini-M™ Plasmid DNA extraction system, Viogene, Taiwan) and screened by restriction fragment length polymorphism (RFLP) using the restriction endonucleases MspI, HaeIII, and CfoI. We set a sampling cut-off of 250 clones (bacteria) and 100 clones (archaea and bacteria), based upon richness estimates obtained from studies of similar locations [9]. At least three samples from each distinct RFLP pattern were sequenced where possible, using the BigDye Terminator Cycle Sequencing kit (Applied Biosystems, CA, USA; Applied Biosystems 3730 Genetic Analyzer). Phylotypes were delineated on the basis of 97 % sequence similarity using the freeware DOTUR [27]. All sequences generated by this study have been deposited in the NCBI GenBank database under accession numbers KC476241–KC476287 and KF294470–KF294500. Screening for possible chimeric sequences was made using Chimera_Check available on the Ribosome Database Project website (http://rdp.cme.msu.edu. html). Approximate phylogentic affiliations were then determined by BLAST searches of the NCBI GenBank database (http://www.ncbi.nlm.nic.gov). Estimates of clone library sampling effort were made using the freeware EstimateS [28]. Sampling effort was assessed by calculation of Coverage and Rarefaction curves, estimates of library richness were made using the nonparametric estimators ACE and Chao 1. All OTU delineation was made on the basis of sequenced phylotypes. Phylogenetic and Statistical Analyses Multiple alignments were created with reference to selected GenBank sequences using BioEdit v7.0.9.0 [29]. The alignments were tested against prescript models of evolution using the softwares PAUP* 4.0b10 [30] and Modeltest v3.0 [31]. The criteria described by the most appropriate evolutionary
C. C. M. Yung et al.
model were input for maximum likelihood analysis using Genetic Algorithm for Rapid Likelihood Inference (GARLI) Version 0.96 Beta [32]. Robustness of furcated branches was supported by both bootstrap values (1,000 replicates) determined using PAUP* 4.0b10 and Bayesian posterior probabilities [33] calculated using Mr. Bayes v3.0b4 [34]. Values (in percent) were shown on all branch nodes supported by more than 50 % of the trees. Statistical analysis was conducted as follows: alpha diversity indices (Shannon’s index, Simpsons diversity index, Pielou’s evenness) were calculated using untransformed data. Differences stated as significant were tested using one-way and two-way analysis of variance (ANOVA), or analysis of similarity (ANOSIM). The ANOSIM produces a statistic R which is based on the difference of mean ranks between groups and within groups [35]. A value approaching to 1 indicates the assemblage composition is totally different, where a value of 0 indicates no difference. Multivariate analysis of diversity data was performed on square-roottransformed diversity data and on nontransformed normalized data for environmental variables. Nonmetric multidimensional scaling (NMDS) ordinations were used to visualize Bray Curtis similarities (diversity data) and Euclidean distances (environmental data). All analyses were performed using Primer v6.1.6 [35]. All results stated as significant have a confidence level of P
