Microorganisms persist at record depths in the subseafloor of the canterbury basin

ABSTRACT The subsurface realm is colonized by microbial communities to depths of >1000 meters below the seafloor (m.b.sf.), but little is known about overall diversity and microbial

distribution patterns at the most profound depths. Here we show that not only _Bacteria_ and _Archaea_ but also _Eukarya_ occur at record depths in the subseafloor of the Canterbury Basin.

Shifts in microbial community composition along a core of nearly 2 km reflect vertical taxa zonation influenced by sediment depth. Representatives of some microbial taxa were also cultivated

using methods mimicking _in situ_ conditions. These results suggest that diverse microorganisms persist down to 1922 m.b.sf. in the seafloor of the Canterbury Basin and extend the

previously known depth limits of microbial evidence (i) from 159 to 1740 m.b.sf. for _Eukarya_ and (ii) from 518 to 1922 m.b.sf. for _Bacteria_. SIMILAR CONTENT BEING VIEWED BY OTHERS

METAGENOMIC PROFILES OF ARCHAEA AND BACTERIA WITHIN THERMAL AND GEOCHEMICAL GRADIENTS OF THE GUAYMAS BASIN DEEP SUBSURFACE Article Open access 27 November 2023 MICROBIAL COMMUNITY STRUCTURE

IN HADAL SEDIMENTS: HIGH SIMILARITY ALONG TRENCH AXES AND STRONG CHANGES ALONG REDOX GRADIENTS Article Open access 08 June 2021 DEEP SUBSEAFLOOR SEDIMENTS IN GUAYMAS BASIN HARBOR

COSMOPOLITAN MICROBIOTA AND TRACES OF HYDROTHERMAL POPULATIONS Article Open access 13 September 2024 INTRODUCTION In addition to terrestrial and marine near-surface habitats, the deep

biosphere is considered to be a third realm for microbial life. Subseafloor sediments provide a habitat for large numbers of microbial cells, as revealed by cell counts (Parkes et al., 2000)

or CARD-FISH (Schippers et al., 2005). Although recent data have shown that the global biomass in subseafloor sediments is smaller than given by earlier estimates, the deep subseafloor

biosphere still constitutes a large fraction (2.9 × 1029 cells) of Earth’s living biomass (Kallmeyer et al., 2012). The subsurface microbiota is diverse and complex, hosting metabolically

active communities down to depths of >1000 meters below the seafloor (m.b.sf.), as revealed by molecular, metagenomic and metatranscriptomic studies (Lipp et al., 2008; Roussel et al.,

2008; Biddle et al., 2011; Pawlowski et al., 2011; Orsi et al., 2013a). It harbors representatives from the three domains of life, for example, numerous endemic and/or as yet uncultured

_Archaea_ and _Bacteria_ (for example, Inagaki et al., 2006; Orcutt et al., 2011), in addition to bacterial endospores (Lomstein et al., 2012), protists and fungi belonging to _Eukarya_

(Schippers and Neretin, 2006; Edgcomb et al., 2011; Orsi et al., 2013a, 2013b). Occurrence of capsid-encoding organisms has also been confirmed (Engelhardt et al., 2011). Although in

subsurface sediment shallower than 1000 m.b.sf. background molecular data on bacterial and archaeal lineages exists (for example, Inagaki et al., 2006; Orcutt et al., 2011), most

deep-subsurface microorganisms detected so far were refractory to cultivation (Sass and Parkes, 2011). The diversity of deeply buried microorganisms remains untapped, as subseafloor

prokaryotic culturability in most studies is <0.1% of all microscopically detected cells (D’Hondt et al., 2004). Remarkably, when wide enrichment collections targeting different

physiological groups such as fermenters, sulfate-reducers and methanogens were performed using different subseafloor sediments, these often led to the isolation of the same few ‘generalist’

bacteria (for example, Batzke et al., 2007). In most cases, the retrieved bacterial genera were adapted to a broader spectrum of environmental conditions (for example, broad temperature

range for growth) compared with their surface counterparts (Sass and Parkes, 2011). So far, within subseafloor sediments, active _Bacteria_ have been identified down to 518 m.b.sf. (Bale et

al., 1997), active _Archaea_ down to 1626 m.b.sf. (Roussel et al., 2008) and active microeukaryotes down to 159 m.b.sf. (Orsi et al., 2013a), but we are still eager to know the depth limit

of the deep subsurface biosphere. Limits to microbial habitability in subseafloor sediments are set by a variety of physical and chemical parameters like temperature, pH, pressure, salinity,

porosity, availability of energy, nutrients and water and maybe also by age as there was water exchange within the sediment pores. The present study site is not characterized by

particularly extreme conditions but stands out from sites previously examined by its depth and low porosity. The depth limit of the deep biosphere remains an important issue to place bounds

on the volume of the subseafloor biosphere and to guide the search for deep life capabilities/adaptation and the role of microorganisms in global nutrient cycles. We hypothesized that life

could exist in even deeper sediments if pore space was sufficient. In this study, we investigated the subsurface microbial communities from a core of 1927 m length collected in the

Canterbury Basin (344 m water depth), off the coast of New Zealand at site U1352, which was drilled during the Integrated Ocean Drilling Program (IODP) Expedition 317 with DS _Joides

Resolution._ Our purpose was to investigate vertical distribution of microbial communities, abundance and evenness of taxa above and below 1000 m.b.sf. depth. We developed a highly stringent

massive parallel tagged-amplicon sequencing of 16S–18S hypervariable regions of small-subunit (SSU) rRNA gene (Supplementary Figures S1 and S2; Supplementary Tables S1–S3), coupled with

cell counts, real-time PCR (phylogenetic and functional genes) and cultivation approaches. This rigorous method was applied to sediment/carbonate rocks spanning epochs from the Holocene to

late Eocene. MATERIALS AND METHODS SITE DESCRIPTION AND SAMPLING Three holes (A, B and C) were drilled at Site U1352 (44°56′26.62′′S; 172°1′36.30′′E), reaching a total depth of 1927.5 m

CSF-A, and thus spanning the Holocene to late Eocene epochs. Fluorescent microspheres were used as tracers for contamination during drilling. Sampling was processed under strict

contamination controls onboard and offshore, and only samples with no detectable contamination were used for this study (Fulthorpe et al., 2011). Onboard, only the center parts of

unconsolidated sediments and intact pieces of rocks that had been exposed to ultraviolet radiation after washing were kept for microbiological analyses, as reported elsewhere (Expedition 317

Scientists, 2011). Subsamples were immediately frozen at −80 °C for onshore molecular analyses, stored at 4 °C under an anaerobic gas phase for later cultivation and stored at 4 °C in a 3%

NaCl/3% formalin solution for cell counting. Detailed information on sampling/subsampling of sediment, on contamination controls and on depth scale terminology are provided in Supplementary

Text. LITHOLOGICAL, PHYSICAL AND GEOCHEMICAL DATA Environmental data were acquired onboard during IODP Expedition 317, as reported elsewhere (Fulthorpe et al., 2011). DNA EXTRACTION, PCR

AMPLIFICATION AND CONTAMINATION CONTROLS DNA extractions were made from 16 samples collected all along the core. In order to avoid contamination, all handling was carried out in a PCR

cabinet exclusively dedicated to low biomass sediment samples (PCR cabinet; CaptairBio, Erlab, Val de Reuil, France), using Biopur 1.5 ml Safe-Lock micro test tubes (Eppendorf, Le Pecq,

France), ultrapure PCR water (Ozyme, St Quentin en Yvelines, France) and ultraviolet-treated (>40 min) plasticware and micropipettes. Negative controls (reaction mixture without DNA) were

included in each set of PCR reactions. In addition, a negative control (for example, negative DNA extraction) was prepared for each work stage, to ensure that no contamination with

exogenous amplifiable DNA occurred during the different stages of sample treatment. The FastDNA Spin Kit for Soil (No. 6560-200, MP Biomedicals, Strasbourg, France) was used to perform DNA

extractions, with few modifications. Detailed information on DNA extractions and PCR amplifications are provided in Supplementary Information. Primer sequences used in this study are

detailed in Supplementary Table S2, and primer sets for direct and nested PCR amplifications are detailed in Supplementary Figure S2. 454-PYROSEQUENCING For each DNA extract, four

independent 25-μl PCR amplifications were run with fusion primer pairs specific for _Bacteria_, _Archaea_ and _Eukarya_, as detailed in Supplementary Table S3. PCR products were pooled two

by two, so as to have two independent replicates for pyrosequencing. Potential contaminants from lab reagents were excluded through the sequencing of negative-control samples and the removal

of operational taxonomic units (OTUs) containing sequences retrieved in negative controls. Detailed information on 454-pyrosequencing, quality filtration, trimming, clustering and taxonomic

affiliation are provided in Supplementary Text. CELL COUNTS Total prokaryotic cells were enumerated in triplicate from 13 uncontaminated sediment samples collected all along the core, using

the cell extraction protocol (protocol FCM-A) described by (Morono et al., 2011) until step 9. Then, all supernatants containing extracted cells were filtered onto 0.2-μm filters (Anodisc,

Whatman, Versailles, France) and stained with SYBRGreen I (Invitrogen, Cergy Pontoise, France), as described elsewhere (Noble and Fuhrman, 1998). Filters were counted in epifluorescence

mode, with an Olympus BX60 microscope (Rungis, France) (objective × 100, pH3, WIB filter; details in Supplementary Text). REAL-TIME PCR MEASUREMENTS Quantifications of different lineages and

diverse functional genes were performed all along the core by quantitative, real-time PCR. Quantifications of _Bacteria_, _Archaea_, _Eukarya_, JS1-_Chloroflexi_ and _Geobacteriaceae_ were

performed using previously described quantitative PCR assays based on the detection of 16S or 18S rRNA (Schippers et al., 2012). These assays were carried out using the _Taq_Man or SYBRGreen

chemistries. DNA copy numbers were also determined for the following functional genes: _mcrA_ for alpha subunit of the methyl coenzyme M reductase, _dsrA_ for the alpha subunit of the

sulfite (bi)reductase, _aprA_ for the alpha subunit of the adenosine-5′-phosphosulfate reductase and _cbbL_ for the large subunit of the enzyme ribulose-1.5-bisphosphate

carboxylase/oxygenase (RubisCO, form I ‘red-like’), as described elsewhere (Schippers et al., 2012). CULTURES AND APPROACHES USED FOR THEIR ANALYSIS A sediment slurry membrane system was

used for cultivation (Ferrari et al., 2008) (Supplementary Figure S8; details in Supplementary Information). Different anaerobic metabolisms found in the subsurface biosphere were targeted

in culture: fermentation, sulfate reduction and methanogenesis/acetogenesis. Media, culture conditions, viability and identification procedures of cells are described in Supplementary Text.

STATISTICAL ANALYSES Principal component analysis was used to help in visualization of high-dimensional data. An order abundance matrix was combined with environmental parameters, using

XLSTAT, to assess relationships between microbial taxa and ecological variables (Addinsoft USA, New York, NY, USA). A second complementary approach was based on regularized canonical

correlation analyses, which were performed to highlight correlations between the order abundance matrices (X) and the environmental parameters (Y) using the R software CCA package. RESULTS

AND DISCUSSION CORE DESCRIPTION The core lithology was characterized by horizontal gradual layers, from unconsolidated sediments (clay, marl) to carbonate rocks (Figure 1). The core was

composed of three lithological units (UI, UII and UIII). Unit I (0–711 m CSF-A, meters of core depth below seafloor computed by conventional method A, corresponding to m.b.sf. (see ‘IODP

depth scale terminology’ at www.iodp.org/program-policies/) was predominantly characterized by a transition from mud-rich sediment to marl. Unit II (711–1853 m CSF-A) consisted of

hemipelagic/pelagic sediment from calcareous sandy mud to sandy sandstone. Unit III (1853–1924 m CSF-A) was characterized by a sharp change (Marshall unconformity: ∼12 Ma are missing) that

occurred at 1853 m CSF-A and was formed of hemipelagic to pelagic foraminifer-bearing nannofossil limestone of early Oligocene to late Eocene age (Figure 1). The temperature at the bottom of

the hole was estimated to be in the range of 60 °C–100 °C on the basis of thermal conductivity measurements and geochemical results (Fulthorpe et al., 2011). Below 1000 m CSF-A, sediments

were replaced by consolidated sedimentary calcium carbonate rocks with porous horizons of glauconite. Porosity decreased with depth and mean pore size was around 2–4 μm at the hole bottom.

In carbonate rocks, numerous fractures and stylolites were observed (Supplementary Figure S3). Organic carbon content was low (<0.6 wt %), with only a few samples having >1 wt % total

organic carbon (Figure 1). The organic matter quality changed from relatively labile volatile material in the shallower sediments to more stable protokerogen with increasing depth. Methane

and ethane both occurred below 11.7 and 18.2 m CSF-A, and the relative ethane content increased with increasing burial depth and temperature (Figure 1). Low but increasing concentrations of

C3-C5 and occasionally C6 alkanes were also measured with depth. pH values were close to 7.5 and stable from the surface to 1164 m CSF-A. Sulfate concentration decreased gradually in the

first meters of the core and reached the detection limit at ∼16 m CSF-A (the SMTZ (sulfate–methane transition zone), was placed between 15.2 and 16.6 m CSF-A), then it remained close to the

detection limit (∼0.85 mM) down to 1433 m CSF-A (Figure 1). VERTICAL DISTRIBUTION OF CELLS We analyzed and compared cell abundances and cell concentrations reported for different geographic

sites using a standardized procedure based on cell extraction and dissolution of silicates (Noble and Fuhrman, 1998; Kallmeyer et al., 2008; Morono et al., 2011) (Figure 2). Mean cell

numbers decreased with depth from about 1.5 × 106±4.7 × 104 cells cm−3 (_n_=8) at the surface (3.76 and 15.1 m CSF-A) to 2.5 × 104±4.9 × 103 cells cm−3 (_n_=7) within the deepest samples

(1911 and 1922 m CSF-A). The detection limit, calculated in our conditions (Kallmeyer et al., 2008), was 2.94 × 103 cells cm−3. The depth profile (down to 600 m CSF-A) was consistent with

the general depth distribution of prokaryotic cells from other subsurface sediments (Kallmeyer et al., 2012). VERTICAL DISTRIBUTION OF MICROBIAL TAXA It is not clear what controls abundance

of _Bacteria_ and _Archaea_ within deep marine sediments (Schippers et al., 2005; Lipp et al., 2008; Schippers et al., 2012). Here, a real-time PCR approach was applied to quantify

representatives of the three life domains. Calculated detection limits for _Bacteria_, _Archaea_ and _Eukarya_ were respectively 1.6 × 104, 1.1 × 103 and 2.9 × 103 SSU rRNA gene copies per

gram of sediment (wet weight)_. Archaea_ were the most abundant within the first meters, while _Bacteria_ dominated the rest of the core (Figure 2). Archaeal SSU rRNA gene copy numbers

strongly decreased with depth (from 1.8 × 106 to 1 × 103 gene copies g−1, corresponding roughly to 1 × 106 to 6 × 102 cells g−1) and were no longer detectable below 650 m CSF-A. A similar

depth distribution was observed for eukaryotic SSU rRNA gene copy numbers, but abundances were relatively constant with depth (∼104 copies g−1). Bacterial SSU rRNA gene copy numbers were low

(∼106 copies g−1≈2.5 × 105 cells g−1) at the surface and decreased with depth up to 1600 m CSF-A (8 × 104 copies g−1≈2 × 104 cells g−1). Along with these measures, deep sequencing allowed

the detection limits to be lowered and masked lineages to be revealed. We pyrosequenced bacterial (V4-V5), archaeal (V1-V3) and eukaryotic (V1-V3) SSU rRNA gene amplicons from 16 depth

horizons and one negative control, pooled together in one single data set with two PCR replicates per sample to overcome PCR and sequencing errors (Supplementary Figure S1). Sequences were

grouped into OTUs with a 97% identity threshold. Sequence composition of the OTUs was then analyzed, and OTUs entirely composed of sequences that had appeared in a single PCR only were

excluded from the diversity analyses. All the sequences kept appeared at least twice independently. Potential contaminants from laboratory reagents were excluded through the sequencing of

negative-control samples and the removal of OTUs containing sequences retrieved in negative controls. The remaining OTUs were used to calculate non-parametric diversity indices (Figure 3,

Supplementary Figures S4 and S6) and compared with the SILVA 111 database for taxonomic affiliation. Pyrosequencing results were congruent with the data discussed above. Archaeal sequences

could not be amplified and sequenced for samples <634 m CSF-A, as observed with real-time PCR analyses. The non-detection of archaeal 16S rRNA genes <650 m CSF-A using two different

amplification methods suggests that _Archaea_ are likely rare or absent at great depths in the Canterbury Basin. Eukaryotic sequences were detected down to 1740 m CSF-A, and bacterial

sequences were found up to the maximal depth of 1922 m CSF-A. The observed species richness (that is, number of OTUs) was extremely low in comparison with other microbial habitats

investigated so far, including extreme environments (for example, Roalkvam et al., 2012). Indeed, only 198, 16 and 40 unique bacterial, archaeal and eukaryotic OTUs, at 3% dissimilarity

level, were detected in the entire cored sequence (Supplementary Figure S4, Supplementary Tables S4 and S5). Chao1 estimator revealed a vertical decrease in microbial richness with

increasing depth (Figure 3). Richness estimates for _Archaea_ and _Eukarya_ dropped off gradually with depth and reached only two and four OTUs, respectively, at the deepest depth for which

a PCR signal was obtained. Beta diversity estimators (that is, diversity among samples) revealed a strong differentiation between communities with depth and a strong vertical structuration

(Supplementary Figure S5). Archaeal diversity showed high abundances of MBG-B (Marine Benthic Group B) and MCG (Miscellaneous Crenarchaeotal Group), two archaeal groups typically found in

subseafloor sediments (Lloyd et al., 2013). Representatives of the as-yet-uncultured lineages MBG-B, MBG-E (Marine Benthic Group E) and MCG were the predominating taxa in surficial layers,

while MCG was the most consistently detected archaeal lineage down to 346 m CSF-A (Figure 2). MBG-B and MCG members are heterotrophic _Archaea_ frequently found in surficial marine sediments

(Biddle et al., 2006; Lloyd et al., 2013). _Thermococcales_ dominated archaeal diversity of the sediment horizon at 634 m CSF-A. Methanogens and anaerobic methanotrophs were not detected,

in agreement with the real-time PCR analysis for _mcrA_. Their absence from the data set might be due to the intervals sampled, which do not correspond to the SMTZ. In _Eukarya_, few protist

OTUs (Stramenopiles and uncultured _Eukaryota_) were detected down to 583 m CSF-A. Sequences affiliated with the bacterivorous protists _Bicosoecida_ were detected at 346 m CSF-A, raising

the question of the existence of a subsurface complex trophic web. In agreement with recently published papers (Edgcomb et al., 2011; Orsi et al., 2013a, 2013b), fungi appeared to be the

most frequently detected eukaryotes in the Canterbury Basin, with 56–100% of the SSU rRNA gene sequences. Different shifts between _Ascomycota_ and _Basidiomycota_ were observed along the

core (Figure 2). _Tremellomycetes_ (order _Tremellales_), _Sordariomycetes_ and _Eurotiomycetes_ dominated shallow depths while _Saccharomycetes_ were detected at depths between 630 and 1365

 m CSF-A. Deeper layers were dominated by _Wallemiomycetes_, _Microbotryomycetes_ and _Tremellomycetes_ (order _Filobasidiales_, not found at shallow depths). These heterotrophic fungi have

been described in deep sediments of other locations (for example, Nagano and Nagahama, 2012; Richards et al., 2012) and demonstrated to be active members of microbial communities (Orsi et

al., 2013b). So, fungi represent an important component of sediment ecosystems through their impact on nutrient cycling and mineral weathering. _Bacteria_ were dominated by _Chloroflexi_ and

_Proteobacteria_, two heterotrophic bacterial groups well represented in subsurface sediments (Figure 2). They comprised 67% of the sequences and 69% of the OTUs in total. However, the

abundances of the two phyla were negatively correlated. _Chloroflexi_ dominated microbial communities at shallow depths (>600 m CSF-A), and their abundances and richness decreased

rapidly. Reciprocally, _Proteobacteria_ were found all along the core, but their relative abundance showed a sharp increase <343 m CSF-A. Among the other lineages observed in this study,

_Planctomycetes_, _Nitrospirae_ and the candidate division OP9 were major contributors of the amplicon pool at shallow depths. Below 600 m CSF-A, _Acidobacteria_, _Firmicutes_ (a phylum

containing spore-formers) and two loosely defined groups of uncultured _Bacteria_ (ML635J-21 and MLE1-12) were the most consistently detected lineages. Real-time PCR quantification of the

JS1-_Chloroflexi_ group confirmed these results as ∼103–106 SSU rRNA gene copies g−1 were detected between the sediment surface and 1532 m CSF-A. _Deltaproteobacteria_ were detected above

the SMTZ and at great depths. Genes encoding a functional dissimilatory sulfite (bi)reductase (_dsrA_), a key enzyme of dissimilatory sulfate reduction frequently encountered among

_Deltaproteobacteria_, was quantified above the SMTZ and in layers up to 1000 m deep in the sediment. The gene became undetectable below this depth, either because it may decrease below the

detection limit or because the detected _Deltaproteobacteria_ cannot respire sulfate. DIVERSITY AND ENVIRONMENTAL FACTORS Principal component analyses coupled with regularized canonical

correlation analyses were performed to visualize relationships between environmental factors and microbial taxa. We first evaluated the relationships between all environmental parameters

measured (that is, depth, pH, salinity, porosity, alkalinity and concentrations of calcium, calcium carbonate, ammonium, magnesium, sulfate, inorganic carbon, organic carbon, methane and

ethane) to design a network of correlations. Only the six most explanatory variables were kept (Supplementary Figure S6). This complementary analysis reinforced the conclusion about

microbial distribution pattern and vertical community composition, depth being defined as a main factor explaining diversity changes (Supplementary Figure S7). HANDLING DEEPLY BURIED

MICROORGANISMS Cultivation approaches allowed prokaryotic and eukaryotic strains to be grown, corresponding to a fraction of the microbial communities detected all along the core,

underlining that these microorganisms were viable. Fungal strains were obtained at 21–765 m CSF-A, using elevated hydrostatic pressure to mimic _in situ_ conditions (Figures 4a–c,

Supplementary Table S6). Sequencing of the ITS1 rRNA regions allowed identification of a _Cadophora_ representative that had already been found in extreme environments, that is, Antarctic

environments (Tosi et al., 2002) and deep-sea hydrothermal vents (Burgaud et al., 2009) (Supplementary Table S6). Fifty-seven anaerobic fungi, currently under description, have also been

isolated from these sediments (Rédou and Burgaud, unpublished data). In addition to the important finding that living fungi could be cultivated from the sediment samples, microbial colonies

were grown anaerobically at 60–70 °C from calcareous chalk/limestone samples collected at 1827 and 1922 m CSF-A (Figures 4d and e), using a microcultivation method (Supplementary Figure S8).

The microcolonies were successfully transferred to liquid media and subcultured. From the different tests performed, it was impossible to grow true methanogens and true sulfate-reducers.

Only bacterial fermentative strains degrading the organic compounds supplied (that is, low quantity of yeast extract) have grown. Within these subcultures, mean cell densities were low,

around 4 × 105 cells ml−1 and growth rates were slow (in 2.5 years of culture, only 6–9 subcultures at 1/40 or 1/50 have been performed). Cells were able to grow at atmospheric pressure and

at the estimated _in situ_ pressure (22 MPa). They were composed of viable very small rods, coccobacilli and cocci of 300–800 nm in diameter, often forming aggregates (Figures 4f–i). These

small sizes and this cellular organization as consortia raises questions about the living conditions of these cells and their (in)dependence with regard to other cells. The smallest diameter

of a cell that assures its viability was calculated as∼200 nm (Velimorov, 2001). The major lineages identified in DNA and RNA libraries from these subcultures belonged to _Alpha_-, _Beta_-,

_Gamma-proteobacteria_, _Actinobacteria_ and _Armatimonadetes_ (Figures 5). With the exception of _Armatimonadetes_, all these taxa were detected from pyrosequencing in crude samples from

1827 to 1922 m CSF-A. The majority of the sequences had relatives recovered from environments with similar physical–chemical characteristics (Lin et al., 2006; Mason et al., 2010; that is,

hot and reduced habitats) compared with the Canterbury subseafloor. Considering the ‘ubiquity’ of these taxa, one can hypothesize that they are generalist bacteria, which would have been

maintained during progressive burial of sediments or by transportation through circulating fluids. They might have acquired metabolic capabilities enabling them to resist the associated

environmental changes. However, this hypothesis needs to be analyzed in detail. Furthermore, similar SSU rRNA gene sequences do not automatically correspond to identical physiologies,

identical phenotypes or similar functions. IMPACT OF POTENTIAL CONTAMINANTS ON NATIVE MICROBIAL POPULATIONS Contamination is a crucial issue when working with subseafloor sediments. In

general, contamination during drilling is still difficult to predict. During IODP Expedition 317, the level of contamination during drilling was evaluated by using fluorescent microspheres,

and only samples with no detectable contamination were kept for microbiological analyses. Nevertheless, samples without microspheres are not necessarily uncontaminated (Smith et al., 2000).

Contamination generally decreases from the exterior to the interior of both sediment and rocks cores (for example, Lever et al., 2006). In consequence, only the interior of sediment cores

and intact pieces of rocks that had been exposed to ultraviolet light after washing were used for the analyses. In addition, for molecular experiments deeply frozen samples of >1 cm in

diameter were sterilized by flaming. Afterwards, all possible contaminations during the wet-lab steps have been strictly controlled and minimized (see Supplementary Text). The cutting-edge

strategy applied for the pyrosequencing and bioinformatic analyses allowed removing potential spurious sequences and OTUs likely to contain contaminants by sequencing of negative controls, a

duplicate procedure and an associated bioinformatics pipeline. In addition to these precautions, the level of potential contamination of our samples was estimated by calculating the number

of contaminating cells per gram of sediment and per gram of sedimentary rock based on the mean contamination values with drilling fluids and mean cell abundances in surface waters reported

in the literature. The mean potential contamination was estimated as (i) 0.011±0.018 μl of drilling fluid per gram for unconsolidated sediments drilled using advanced piston coring (APC) and

(ii) 0.027±0.029 μl g–1 for rocks collected using rotary core barrel (Lever et al., 2006). Considering these levels of contamination, mean cell counts of 5 × 105 cells ml–1 in surface

waters in the ocean (Whitman et al., 1998) and average densities of 1.85 g cm–3 in sediments and 1.99 g cm–3 in sedimentary rocks at site U1352, potential contamination of the interior of

the core sample should be expected very low with 5–11 cells g–1 of sediment only. A second reported estimate indicates that <50 cells g–1 of sediment contaminated APC core centers drilled

with _Joides Resolution_ and that XCB cores were generally more contaminated with contamination levels 3–10 times higher in XCB cores than in APC core centers (House et al., 2003).

Considering these different estimates of potential contamination, the observed cell counts at site U1352 were 2–5 orders of magnitude higher in the studied samples. If contamination cannot

be excluded, in the worst case, non-indigenous cells represent only up to 1% of total cells in the sample. Therefore, it is most likely that >99% of the counted cells are native to the

sampled sediment/rocks. This implies that the vast majority of the prokaryotic and eukaryotic DNA subjected to pyrosequencing was therefore derived from the sediment native cells. By

extension, assuming that most of the prokaryotic DNA extracted from sediment samples is from native cells, the fact that cultivated bacteria match OTUs abundant in the crude sediment samples

supports the idea that these cultivated strains are isolates of native bacteria. Consequently, the potential impact of contaminants on each category of data (cell counts, molecular data and

cultures) is likely very low. ECOLOGICAL IMPLICATIONS AND FUTURE PROSPECTS We have underlined that the subseafloor of the Canterbury basin hosts microorganisms that comprise _Bacteria_,

_Archaea_ and _Eukarya._ Some of these microorganisms are alive and, at least to a certain extent, revivable. The communities exhibit a quite low phylogenetic diversity, but this does not

necessarily correspond to a low functional diversity. This poor diversity could be explained if natural selection has produced (i) taxa adapted to harsh subsurface conditions (that is,

specialists), which would be expected in the case of a low connectivity among habitats; and/or (ii) taxa with a broad physiological plasticity, allowing them to survive in a diversity of

nutritional and physical–chemical conditions (that is, generalists). In fact, some taxa detected through their 16S/18S rRNA gene sequences are thought to be endemic to subsurface habitats,

while others seem ubiquitous and are consistently encountered in common and extreme environments. The bacterial strains in cultures are related to opportunistic or generalist taxa isolated

from a broad array of redox environments, which raises the question of the existence of microbial metabolic versatility and also questions the species concept, as behind a given name or a

given OTU can lay a variety of microorganisms with different ecological lifestyles. Metabolic versatility has already been demonstrated in well-known taxa. For example, some _Thermococcales_

strains, which are usually fermenters that reduce sulfur compounds, can grow in oligotrophic conditions or can oxidize carbon monoxide (Sokolova et al., 2004). Heterotrophy is likely to be

the major mode of carbon assimilation within microbial communities of subsurface marine sediments (Batzke et al., 2007). Our culture data support this hypothesis. Genome and metagenome

analyses would allow functions to be predicted on a finer scale to assess and hypothesize the individual ecological functions within the analyzed habitat or ecosystem (Vandenkoornhuyse et

al., 2010). The detection of fungal sequences at great depths and our success in the cultivation of fungal strains leads us to ask what role they play in deep carbon cycling and what

involvement they have in dynamics/regulation of prokaryotic populations, if they are active _in situ_. The broad polyphasic approach developed in this study provides direct evidence that

viable microorganisms can be present in rocks that are hardened but not totally cemented, where stylolites and micro-fluid circulations exist. Our data demonstrate that the combination of

physical, chemical and energetic constraints encountered from 0–1922 m CSF-A in the subseafloor of the Canterbury Basin still allow microorganisms to persist down to at least 650, 1740 and

1922 m CSF-A for _Archaea_, fungi and _Bacteria_, respectively. It extends the subseafloor sedimentary depths at which subseafloor organisms are known to be present to 1740 m for fungi and

to 1922 m for _Bacteria_. Nevertheless, one cannot exclude that some of the detected sequences belong to microorganisms in dormancy. More extensive sequencing efforts will be required, that

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95: 6578–6583. Article  CAS  Google Scholar  Download references ACKNOWLEDGEMENTS Samples, shipboard facilities and expedition support were provided by IODP. We thank the co-chiefs, crew and

shipboard scientific parties of IODP Expedition 317. The Joint Research Unit UMR 6197 (CNRS-Ifremer-UBO), LUBEM, GDR Ecchis, EU program MaCuMBA, DIVVIR project of the FRB and the BGR

supported molecular and cultural post-cruise analyses. The study was supported by grants from the French Ministry of Higher Education and Research to MCC, VR and FG; from the Région Bretagne

to FG; and from the DFG to AS (Grant SCH535/7-2) and to JSL (Grant HI616/11-1). We thank reviewers for their constructive comments. We thank also C Struckmeyer, M Guégan, H Leclerc, C

Argouarch, S Coudouel, A Dheilly and O Quenez for their contribution to this work. AUTHOR INFORMATION Author notes * Gaëtan Burgaud and Alexis Dufresne: These authors contributed equally to

this work. AUTHORS AND AFFILIATIONS * Université de Bretagne Occidentale (UBO, UEB), IUEM—UMR 6197, Laboratoire de Microbiologie des Environnements Extrêmes (LMEE), Plouzané, France

Maria-Cristina Ciobanu, Sarah Ben Maamar, Frédéric Gaboyer, Odile Vandenabeele-Trambouze, Mohamed Jebbar, Anne Godfroy & Karine Alain * CNRS, IUEM—UMR 6197, LMEE, Plouzané, France

Maria-Cristina Ciobanu, Frédéric Gaboyer, Odile Vandenabeele-Trambouze, Mohamed Jebbar, Anne Godfroy & Karine Alain * Ifremer, UMR6197, LMEE, Plouzané, France Maria-Cristina Ciobanu, 

Frédéric Gaboyer, Odile Vandenabeele-Trambouze, Mohamed Jebbar, Anne Godfroy & Karine Alain * Université de Brest, UEB, Laboratoire Universitaire de Biodiversité et d’Ecologie

Microbienne EA 3882, IFR148 SFR ScInBioS, ESIAB, Plouzané, France Gaëtan Burgaud, Vanessa Rédou & Georges Barbier * Université de Rennes I, CNRS, UMR 6553 ECOBIO, Rennes, France Alexis

Dufresne & Philippe Vandenkoornhuyse * Bundesanstalt für Geowissenschaften und Rohstoffe (BGR), Hannover, Germany Anja Breuker & Axel Schippers * Department of Geosciences and MARUM

Center for Marine Environmental Sciences, Organic Geochemistry Group, University of Bremen, Bremen, Germany Julius Sebastian Lipp Authors * Maria-Cristina Ciobanu View author publications

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KEYWORDS * deep biosphere * subsurface life * eukaryote * record depth