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Expedition 337 was the first riser drilling IODP expedition to incorporate extensive shipboard microbiological and molecular biological analyses. These analyses included chemical and microbial contamination tests; molecular ecological studies based on DNA extracted from fresh, unfrozen subseafloor samples; cell detection and enumeration; metabolic activity measurements using various radioactive and stable isotopic tracers (see also “Organic geochemistry”); and a wide range of incubations that included cultivation, nucleic acid–, and whole cell–stable isotope probing (SIP) experiments (for lipid- and metabolic compound-SIP, see “Organic geochemistry”).

The shipboard molecular biological program included DNA extraction, quantitative polymerase chain reaction (qPCR) on bacterial and archaeal 16S rRNA genes, conventional polymerase chain reaction (PCR) assays of functional marker genes and eukaryotic 18S rRNA genes, and most-probable-number PCR (MPN-PCR) of contamination indicator organisms. Molecular fingerprinting assays were used to document changes in community composition with core depth and across changing lithologies and to evaluate the potential contribution of microbial contaminants. Cell enumerations were performed by manual microscopic observation, image-based discriminative cell enumeration (Morono et al., 2009), high-performance flow cytometry (Morono et al., in press), and fluorescence in situ hybridization (FISH). For measurements of potential activity, stable isotope-labeled carbon, nitrogen, oxygen, and hydrogen were added to fresh samples and incubations were initiated on board the ship under anaerobic conditions near in situ temperature. Radioactive tracers, such as 35S-labeled sulfate and 14C-labeled bicarbonate, were handled in the new radioisotope container laboratory on the Chikyu.

Complementary to core samples, a large number of samples were obtained from drilling mud and cuttings to evaluate the risk of background contamination during riser drilling. Analyses performed on these control samples included chemical contamination tests, cell counts, MPN-PCR targeting well-known contaminants, and shore-based incubation experiments. The sampling approaches and methods used are described in the following sections.

Sampling strategy


Cores were sampled for shipboard analyses and shore-based experiments (Table T5). All core sections underwent a nondestructive X-ray CT scan as soon as possible after arrival on deck and before further processing. A core section for a series of high-priority, mainly shipboard microbiological, geochemical, and geophysical investigations (referred to as community WRC hereafter) and a corresponding core section for interstitial water analysis (see “Inorganic geochemistry”) were processed immediately after the CT scan. Processing of community WRCs took place in the QA/QC laboratory on the core processing deck. All remaining core sections were packed into ESCAL bags, vacuum-sealed, and flushed with nitrogen. These core sections then underwent MSCL-W analyses at room temperature (see “Introduction”).

Community WRCs were typically 15 cm long and taken from 1 to 4 depths within each core. All handling of these community WRCs was carried out in a laminar flow hood to minimize sample contamination with cells from laboratory air. Because samples for PFC tracer contamination tests were taken from these community WRCs, rapid processing was necessary. This was to minimize PFC loss to volatilization and PFC diffusion from contaminated outer parts to cleaner inner parts of cores, both of which would lead to inaccurate quantification of core contamination (for more info on PFC method, see “Contamination tests”). In addition to PFC tracer analyses, samples for shipboard microbiological, geochemical, geophysical, and lithologic analyses, as well as shore-based fungal cultivation and solid-phase Fe and S analyses were taken from these community WRCs (Fig. F23; Table T6 for overview).

All other WRC samples were cut by shipboard microbiologists in the QA/QC laboratory after MSCL-W scanning. Redox-sensitive samples used for radiotracer-based activity measurements, shore-based cultivation, and nucleic acid analyses were again packed into ESCAL bags, flushed with N2, vacuum-sealed, and stored at 4°C until further processing; this anaerobic sample handling procedure was performed to prevent or minimize the potentially rapid growth of aerobic and mesophilic microbial contaminants from the riser drilling mud.

Drilling mud and cuttings

In addition to WRCs, samples of drilling mud from the active tanks, core liners, and mud ditch, as well as samples of core cuttings were obtained as follows: active tanks were sampled on a daily basis, mud fluids from the core liner were taken from every core recovered, mud fluids from the mud ditch were taken around the time of core recovery, and cuttings samples were obtained at 50 m depth intervals throughout riser drilling operations. These samples were taken to monitor concentrations of chemical tracer, as well as changes in microbial cell abundance and community structure within drilling mud and cuttings over the course of the expedition. In addition, cuttings from six designated depths (796, 896, 996, 1096, 1996, and 2496 m MSF) were taken for shore-based cultivation experiments.

Formation fluid sampling

Formation fluid samples of ~250 mL volume were obtained from six different depths using the Schlumberger Quicksilver probe. All samples were processed for activity measurements (35S and 14C), shore-based cultivation, cell counts, FISH, and PFC tracer measurements. After separation of gas and liquid components from Quicksilver sampling bottles (see “Sampling and gas analysis in fluids retrieved by DFA”), fluids were immediately transferred into the glove box, where 100 mL was sampled for microbiological examinations. This 100 mL sample was used as follows:

  1. 1 mL was transferred to a 20 mL headspace vial for PFC tracer measurement;

  2. Four 15 mL samples were transferred to sterilized screw-cap glass bottles, flushed with nitrogen gas, and stored at 4°C for shore-based cultivation experiments;

  3. A 5 mL aliquot was fixed with paraformaldehyde (PFA) for cell enumeration and FISH analysis and stored at –20°C;

  4. A 10 mL aliquot of formation fluid was frozen using the alternating magnetic field freezer (Cell Alive System CAS-LAB1-M, ABI, Co., Ltd, Japan) for single-cell analysis;

  5. Four 10 mL headspace vials containing 1 mL of fluid were used for measurement of sulfate reduction rates under N2 and N2/CH4 (1:1), with two replicates for each headspace treatment; and

  6. Ten 5 mL headspace vials were prepared for incubation experiments to determine rates of acetogenesis and methanogenesis in the formation fluid.

Sterile sampling and its tools

Samples for shipboard microbiological analyses were collected using devices that differed depending on sample lithology. Drilling mud was sampled by pipetting, where pipet tips had been cut off to enlarge the opening and facilitate drawing up this viscous fluid; cuttings and unconsolidated sand were sampled by spatulas or cut-off syringes; ceramic knives and cork borers were used for shales, coal, and consolidated sand; very hard samples, such as certain mud-, silt-, and sandstone, were sampled by core drill or handheld power drill; extremely hard samples, such as certain mud- and siltstones, could, however, only be sampled using a diamond-paste band saw system placed in a clean booth within the QA/QC laboratory (Masui et al., 2009). Sampling tools were sterilized by wiping with 70% ethanol followed by flaming and/or autoclaving. Additional cleaning was done periodically by spraying with RNAse AWAY surface decontaminants (Life Technologies).

Contamination tests

Contamination tests were carried out by chemical tracer quantification and by molecular monitoring of microbial communities.

Chemical tracer

Perfluoromethylcyclohexane (PMCH; C7F14), a PFC compound used as a chemical tracer during previous scientific ocean drilling expeditions on the riserless drill ship R/V JOIDES Resolution (Smith et al., 2000a, 2000b; House et al., 2003; Lever et al., 2006), was supplied directly to drilling mud in actively mixed mud tanks, and concentrations were monitored on a daily basis (Table T7). Additions were paused during logging operations and resumed 2 days prior to drilling. PFC tracer measurements followed the protocol outlined in Lever et al. (2006), except for the following modifications:

  1. Headspace vials (20 mL) with silicone septa were used instead of Vacutainer tubes;

  2. Prior to measurement, all samples were preincubated in an HB-80 hybridization incubator (TAITEC, Japan); and

  3. Samples were analyzed by a gas chromatograph (GC) with an electron capture detector (ECD) (Network GC System 6890N, Agilent Technologies) connected to an autosampler (Network Headspace Sampler G1888, Agilent Technologies).

For preincubation, headspace vials were placed in a carousel within a hybridization oven and rotated in horizontal orientation at 20 revolutions per minute (rpm) for 2 h at 80°C. This change to the incubation protocol was performed after a 1 h preincubation step with gentle motion was found to drastically increase (≥36×) the release of PFC tracer from drilling mud into headspace compared to mere incubation without rotation for 5–10 min at 80°C in an incubation oven (as on previous riserless drilling expeditions) or for 30 min at 80°C in the GC-ECD autosampler (as in the Chikyu shipboard protocol; Fig. F24). Even after drilling mud had been incubated for 5 h without movement within the GC-ECD autosampler, headspace PFC tracer concentrations were drastically (≥15×) lower compared to vials that had been rotated for a 1 h period prior to insertion into the autosampler. Because of the anticipated even slower diffusion and volatilization of PFC tracer out of core samples compared to drilling mud, a preincubation time of 2 h with rotation was used in the final protocol, which was applied to all core samples. After this preincubation, headspace vials were rapidly transferred to the autosampler, where they were incubated for an additional 30 min at 80°C prior to measurement.

PFC tracer concentrations, which were calculated according to Smith et al. (2000b), were monitored in the exterior, halfway, and interior of cores (Fig. F23B; Table T6); in drilling mud from active tanks prior to and during drilling; in cuttings; and in mud fluid coming up with the cuttings (Table T7).

DNA-based contamination tracers

In community WRCs (Fig. F23; Table T6), as well as in several samples of drilling mud and cuttings, PFC tracer quantifications were complemented by DNA-based contamination tests. The latter were designed to quantify microbial contamination in cores and involved MPN-PCR assays with group-specific PCR primers (Table T8). Target groups were potential microbial indicators of

  1. Drilling mud viscosifiers (Xanthomonas and Halomonas),

  2. Anthropogenic wastewater (Bifidobacterium, Blautia, and Methanobrevibacter), and

  3. Surface seawater (SAR11 and Marine Group I Archaea).

These were identified as target groups based on past evidence indicating viscosifiers, wastewater, and seawater as the main sources of microbial contamination in cores retrieved by scientific ocean drilling (Masui et al., 2008; Santelli et al., 2010). With the exception of Methanobrevibacter, 16S rRNA genes were targeted for PCR-based gene detection and quantification. In the case of Methanobrevibacter, the gene encoding the alpha subunit of methyl coenzyme M reductase (mcrA) was targeted. For details on DNA extraction method and PCR amplification, see “DNA extraction and purification” and “PCR.”

Cell separation and enumeration

A 2 cm3 core sample was transferred into a sterile 15 mL centrifuge tube containing 8 mL of 3× PBS (pH = 7.5; Invitrogen 70013), with 2% (v/v) neutralized PFA as a fixative, and then thoroughly mixed by vortexing to form a homogeneous suspension. After fixation for 6–12 h at 4°C, samples were washed twice with 10 mL of 3× PBS, resuspended in ethanol:3× PBS (1:1), and stored at –20°C. These suspensions were subjected to cell detachment and separation steps as follows:

  1. Aliquots of 1000 µL of 1:5-diluted PFA-fixed core slurry were pipetted into 15 mL centrifuge tubes. Subsequently, 3000 µL of 2.5% NaCl, 500 µL of detergent mix (100 mM ethylenediaminetetraacetic acid [EDTA], 100 mM sodium pyrophosphate, and 1% [v/v] Tween-80), and 500 µL of pure methanol were added.

  2. Samples were shaken using a Shake Master (Bio Medical Science, Japan) at 500 rpm for 60 min.

  3. Slurry samples were sonicated at 160 W for 10 cycles, each 30 s long, and cooled in an ice-water bath for 30 s between cycles.

  4. An aliquot of 500 µL of 10% hydrofluoric acid (HF) was then added, and samples were incubated at room temperature.

  5. After 20 min, HF treatments were stopped by adding 500 µL of 1.5 M tris(hydroxymethyl)aminomethane.

  6. Samples were transferred to 15 mL tubes, where they were placed on top of four density layers. These density layers consisted of a 30% Nycodenz (1.15 g/cm3), a 50% Nycodenz (1.25 g/cm3), an 80% Nycodenz (1.42 g/cm3), and a 67% sodium polytungstate (2.08 g/cm3) layer. These density layers had been prepared by sequentially overlaying 1 mL of each density solution, starting with the highest density at the bottom.

  7. Cells and sediment particles were then separated by centrifugation with swinging rotors at 6000× g for 1 h at 20°C.

  8. Starting from the top and using a 27G needle, density layers were collected down to where the first particles became visible. The latter often started to be visible in the 80% Nycodenz layer, of which only the particle-free upper part was then collected.

  9. The density layers that contained particles (i.e., typically the polytungstate layer and in some cases part of the 80% Nycodenz layer) and pellets were then washed by resuspension with 5000 µL of 2.5% NaCl and followed by centrifugation at 5000× g for 15 min at 25°C. The recovered supernatant was combined with the one from Step 8.

  10. Pellets were resuspended in 5000 µL of 2.5% NaCl, 500 µL detergent mix, and 500 µL of methanol and shaken at 500 rpm for 60 min at 25°C.

  11. The resuspended sediment was again placed on top of density layers with 30% Nycodenz (1.15 g/cm3), 50% Nycodenz (1.25 g/cm3), 80% Nycodenz (1.42 g/cm3), and 67% sodium polytungstate, and the cell extraction (Steps 6–9) was repeated. Particle-free layers and supernatants were combined with previously obtained ones.

  12. A total of 50% of the resulting total cell extract was passed through a 0.22 µm polycarbonate membrane filter. Cells on the membrane filter were stained with SYBR Green I–staining solution ( of SYBR Green I in Tris-EDTA [TE] buffer). The number of SYBR Green I–stained cells was enumerated by a fluorescent image-based cell counting system (Fig. F25) as described in Morono et al. (2009) and Morono and Inagaki (2010) with a careful check by visual observations.

  13. Cells in the remaining 50% of the supernatant were trapped onto an Anopore inorganic membrane (Anodisc, Whatman) and stained with SYBR Green I. Membranes with stained cells were then put into 15 mL tubes with TE buffer and sonicated to detach the cells. Cell numbers in these suspensions were quantified using a Gallios flow cytometer (Beckman Coulter, CA) (Fig. F26) placed on an antivibration table.

All cell separation and filtration procedures were performed on clean benches, with great care to avoid contamination. Minimum quantification limit (MQL) of the cell count was estimated by blanks. Samples were processed in batches of ten plus two additional blank samples, in which sterile-filtered 2.5% NaCl solution replaced the sediment slurry. The blank was calculated as the average of all blanks processed during the expedition. The MQL was set as the blank value plus three times the standard deviation of the blank.

DNA extraction and purification

DNA extraction protocols are known to vary in total DNA yields and in efficiency of lysing different groups of organisms. Thus, we used three different DNA extraction methods: a hot alkaline lysis protocol, a chemical lysis protocol, and a modification of the chemical lysis protocol that included the separation of extracellular and intracellular DNA pools. All three protocols had been successfully tested across a wide range of subseafloor samples prior to the expedition.

Hot alkaline lysis protocol

This protocol was used for core samples only. To 2 g of core sample, 6 mL of 6.25 mM EDTA (pH = 8.0) was added and warmed to 70°C for 10 min. These suspensions were then amended and mixed with 800 µL of each 10% sodium dodecyl sulfate and 10 M NaOH and incubated for 20 min at 70°C. Suspensions were then centrifuged at 10,000× g for 1 min at 25°C and supernatants transferred to clean tubes. Pellets were washed with 4 mL of warmed (70°C) double-distilled water, centrifuged at 10,000× g for 1 min at 25°C, and supernatants combined with previously obtained ones. The supernatants were neutralized with 6 mL of a solution containing 1 M HCl and 0.3 M Tris-HCl (pH = 8.0), and 1 volume of phenol/chloroform/isoamylalcohol (24:24:1) was added. The resulting mixture was shaken manually and then centrifuged at 10,000× g for 10 min at 25°C. Aqueous supernatants were transferred to clean tubes, mixed by manual shaking with one volume of chloroform/isoamylalcohol (24:1), and centrifuged at 10,000× g for an additional 10 min at 25°C. Aqueous phases were collected and DNA was precipitated by adding volume of 3 M sodium acetate, 3 µL of ethachinmate (Nippon Gene) as co-precipitant, and 2.5× volumes of 99.5% ethanol. The recovered DNA pellet was dissolved in 5 mL of a × TE buffer containing 10 µM Tris-HCl and 1 µM EDTA (pH = 8.0) and purified using an Aurora DNA purification system (Boreal Genomics).

Chemical lysis protocol

This protocol was used to extract DNA from core samples, cuttings, and drilling mud. Amounts of 1 g, 0.2 g, and 100 µL were used per extraction from cores, cuttings, and drilling mud, respectively. Extractions from cuttings and drilling mud followed the same protocol, except in the initial steps, whereas extractions from cores followed the same protocol as for cuttings, except that all reagents were increased proportionally to the larger amount of sample used (i.e., by a factor of 5; more information below).

Cuttings samples were placed into 2 mL screw-cap tubes filled to ~20% with 0.1 mm diameter zirconia/silicate beads (Biospec Products, USA) and briefly mixed by tapping or vortexing with 100 µL of 100 mM deoxyribonucleotide triphosphate (dNTP) solution. A volume of 500 µL of lysis solution (30 mM Tris-HCl, 30 mM EDTA, 800 mM guanidium hydrochloride, pH raised to 10.0 with NaOH, and 2% Triton X-100) was then added and homogenized with tube contents by manual shaking or brief vortexing. Screw-cap tubes were then taped horizontally onto a Vortex-Genie Pulse (Scientific Industries, Inc., USA) and shaken at the maximum speed setting (3000) for 10 min. Treatments of drilling mud samples differed as follows: only 10 µL of 100 mM dNTP solution was added; the lysis solution contained 0.5% Triton X-100; and the resulting mixture of drilling mud, dNTP solution, and lysis solution was vortexed for only 10 s at the maximum speed.

After these initial extraction steps, protocols for cuttings, cores, and drilling mud followed the exact same protocol. Samples were frozen at –80°C. After >20 min, the fully frozen samples were transferred to a shaker incubator (Iwashiya Bio Science, Japan) and shaken at 120 rpm for 1 h at 50°C. Samples underwent one more freeze cycle followed by incubation at 50°C, and were then centrifuged at 9500× g for 10 min at 4°C. Supernatants were transferred to clean vials and washed twice with 1 volume of chloroform/isoamylalcohol by vortexing for 10 s followed by centrifugation at 9500× g for 10 min at 4°C. The final supernatant was mixed with linear polyacrylamide (LPA) (final concentration 20 µg/mL) and 0.2 volumes of 5 M sodium chloride. After gentle mixing by tapping, 2.5 volumes of 99.5% ethanol were added, tubes were inverted five times for homogenization, and DNA was precipitated by incubation in the dark at room temperature for 2 h. After these 2 h, tubes were centrifuged at 14,000× g for 30 min, supernatants were poured and pipetted off, and pellets were redissolved in 100 µL molecular grade water after drying for 5 min in a laminar flow hood. The redissolved pellets were purified using a Norgen CleanAll DNA/RNA cleanup and concentration kit (Norgen Biotek Corp, Canada). All purified DNA extracts were stored at –80°C until PCR-based analysis.

Extracellular DNA extraction

Several studies over the past decades have reported a large fraction of DNA in marine sediment to be extracellular rather than intracellular, from living organisms (e.g., Ogram et al., 1987; Dell’Anno and Danovaro, 2005; Corinaldesi et al., 2011), and suggested that this extracellular pool might represent a genetic archive of past environmental change (e.g., Willerslev et al., 2003). To extract extracellular DNA, 5 cm3 of sediment from designated 10 cm WRCs were homogenized with 5 cm3 of carbonate dissolution/phosphate binding solution (0.47 M sodium acetate, 0.47 M glacial acetic acid, 10 mM EDTA, 100 mM sodium metaphosphate, and 3% NaCl) and rotated in a carousel at 1 rpm at room temperature. After 1 h, 40 mL of 10× TE buffer (pH = 10.0) (300 mM tris-HCl, 10 mM EDTA, and 3% NaCl, with pH raised with NaOH) were added, mixtures inverted five times or, if necessary, vortexed to ensure homogenization, and rotated for one additional hour. Samples were then centrifuged at 9500× g for 30 min at room temperature and supernatants containing extracellular DNA transferred to separate vials. Both vials (i.e., supernatants containing extracellular DNA and sediment pellets containing intracellular DNA) were then frozen at –80°C for comparative phylogenetic studies of extracellular and intracellular DNA pools at the on shore laboratory.


PCR-based quantifications

We quantified bacterial and archaeal 16S rRNA gene copies by qPCR using a StepOnePlus real-time PCR system (Life Technologies Japan, Tokyo, Japan) provided by the JAMSTEC Kochi Institute for Core Sample Research (Fig. F27) and SYBR Green I chemistry (Morrison et al., 1998). For amplification, the SYBR Premix DimerEraser kit (Takara Bio, Shiga, Japan) was used (5 µL SYBR Premix Dimer Eraser, 0.2 µL of ROX reference dye, 0.3 µL of 10 µM forward and reverse primer, 2 µL template, and 2.2 µL nuclease-free water). The standard consisted of plasmids containing full 16S rRNA gene inserts of uncultured bacterial clone N_194 (not deposited) and uncultured archaeal clone 1H3M_ARC08 (JN229535) for Bacteria and Archaea, respectively. The qPCR protocol consisted of

  1. Initial denaturation for 30 s at 95°C;

  2. 50 cycles of 5 s denaturation at 95°C, 30 s annealing (see Table T8 for Tm), and 40 s elongation; and

  3. A final elongation step of 3 min at 72°C. qPCR primers used are shown in Table T8.

MPN-PCR was performed to quantify drilling mud–derived microbial contaminants in cuttings and cores. All PCR amplifications were performed on Veriti Thermal Cyclers provided by the JAMSTEC Kochi Institute for Core Sample Research using Takara Ex Taq polymerase kits (Takara Bio, Shiga, Japan) following the manufacturer’s suggestion, except that a bovine serum albumin (BSA) concentration of 1 mg/mL was included. All seven groups of contamination indicator organisms (see “Contamination tests” for more information) were amplified with group-specific PCR primers (Table T8). PCR protocols consisted of

  1. Initial denaturation for 2 min at 98°C;

  2. 50 cycles of 15 s denaturation at 95°C, 30 s annealing (see Table T8 for Tm), and 30 s elongation at 72°C; and

  3. A final elongation step of 5 min at 72°C.

A 10× dilution series of extracts was prepared, with the lowest dilution yielding correctly sized PCR amplicons used to estimate minimum target gene concentrations in samples.

Functional marker genes

Functional marker genes indicative of microbial energy metabolism were PCR-amplified via conventional PCR (Table T8). All PCR amplifications were performed on Veriti Thermal Cyclers provided by the JAMSTEC Kochi Institute for Core Sample Research using Takara Ex Taq polymerase kits (Takara Bio, Shiga, Japan) following the manufacturer’s suggestion, except that a BSA concentration of 1 mg/mL was included. mcrA genes of in situ populations of methanogenic and anaerobic methane-oxidizing archaea were the primary target groups and amplified with two primer pairs. Additional targets were (1) formyl tetrahydrofolate synthetase (fhs) genes found in acetogens and other C1-compound metabolizing microbes and (2) dissimilatory sulfate reductase (dsrB) genes of sulfate-reducing microbes. PCR protocols consisted of

  1. Initial denaturation for 2 min at 98°C;

  2. 50 cycles of 15 s denaturation at 95°C, 30 s annealing (see Table T8 for Tm), and 60 s elongation at 72°C; and

  3. A final elongation step of 5 min at 72°C.

PCR products were checked using 2% (w/v) low melting point agarose in a tris-acetic acid-EDTA (TAE) buffer and visualized with a SYBR Safe DNA gel stain (Invitrogen, USA) provided by the JAMSTEC Kochi Institute for Core Sample Research.

Molecular fingerprinting assays

Terminal restriction fragment length polymorphism (T-RFLP) analyses of PCR-amplified 16S rRNA genes were used to evaluate changes in bacterial and archaeal communities along environmental variables (e.g., depth, geochemistry, and lithology) and to evaluate whether amplified DNA derives from the indigenous microbial community or drilling mud. We followed a published general T-RFLP procedure (Singh et al., 2006; Suzuki et al., 1998). The detailed steps used during Expedition 337, which were modified from the original published protocols, are outlined below.

DNA was PCR-amplified using Veriti Thermal Cyclers with the 16S rRNA gene primer sets 27F-926R and 21F-958R to target the domains Bacteria and Archaea, respectively (Table T8). Both forward primers were 5′-end labeled with 6-carboxyfluoresce (6-FAM). PCR reaction mixtures consisted of 0.3 µM of each primer, 0.2 mM deoxyribonucleoside triphosphate, 1.6 µL of PrimeSTAR GXL polymerase, 1× PrimeSTAR GXL buffer (Takara Bio, Shiga, Japan), and 2 µL of extracted DNA. PCR amplification protocols consisted of 32 cycles for Bacteria and 40 cycles for Archaea, whereby each cycle consisted of 10 s denaturation at 98°C, 15 s annealing at 56°C, and 15 s elongation at 68°C.

PCR products were examined by gel electrophoresis on 2% (w/v) agarose gel followed by staining with SYBR Safe DNA dye. Gel-excised PCR products were purified with a NucleoSpin gel and PCR cleanup kit (Takara Bio, Shiga, Japan) according to the manufacturer’s instructions. Purified PCR products were then digested by the restriction enzyme Hha I (cleavage site [GCG′C], where ′ shows the site of cleavage) in a hybridization oven for 6 h at 37°C. After DNA digestion, restriction enzymes were inactivated by 20 min of incubation at 65°C and digested PCR amplicons precipitated after adding volume 3 M sodium acetate solution and 2 volumes of 99.5% ethanol. For the precipitation, mixtures were incubated for >1 h in a –20°C freezer followed by centrifugation at 12,000× g for 30 min. After decanting the supernatants, DNA precipitates were rinsed with 70% ethanol and air dried.

Air-dried DNA fragments were dissolved in 10 µL of deionized formamide supplemented with 0.3 µL of the internal GeneScan 1200 LIZ size standard (Applied Biosystems, Foster City, California, USA). After denaturing the fragments for 5 min at 94°C followed by immediate chilling on ice, the lengths of terminal restriction fragments (T-RFs) were analyzed by electrophoresis on an ABI 3130 XL Genetic Analyzer (Applied Biosystems) provided by the JAMSTEC Kochi Institute for Core Sample Research (Fig. F28). The injection time was 15 s and the run time was ~2 h. Microbial communities were compared in terms of genetic composition (size of peaks), richness (number of peaks), and evenness (height of peaks) based on the T-RFs.

Potential sulfate reduction rates

After the X-ray CT scan and MSCL-W measurements, a 5–10 cm long WRC sample was taken from each core to determine potential sulfate reduction rates (pSRR). The WRC was transferred to a glove box, the contaminated outer part (~1 cm) removed, and the cleaner innermost part homogenized or, if necessary, crushed. Approximately 5 mL of innermost sample was placed in a preweighed 20 mL crimp vial and flushed with N2. Four replicate vials were prepared for each sample. After weighing the samples, slurries were prepared by adding 5 mL of sterile, anoxic salt medium (see Table T9 for composition) through the septa. Sulfate-depleted samples, as from the subseafloor at Site C0020, bear the risk that indigenous sulfate-reducing microorganisms immediately consume all radioactive tracer, resulting in unrealistic turnover rates. To prevent this from happening, media were supplied with an additional background concentration of 1 mM Na2SO4. All vials were autoclaved, and solutions were filtered through sterile syringe filters (0.20 µm pore size) prior to use.

In the radioisotope laboratory on the Chikyu, a 15 mL volume of methane (99.9%, standard gas) was injected into the headspace of two vials to produce two duplicate incubation sets (i.e., two vials with N2 headspace and two vials with N2/CH4 headspace [50/50]). The addition of CH4 to N2 in the headspace increased the pressure (to ~2 bar), which increased the dissolution of CH4 into the medium. Subsequently, 30 µL of radiolabeled Na2SO4 (3.7 MBq) was injected and samples vigorously shaken. Samples were incubated for 10 days at temperatures within the in situ range. Samples from between 1200 and 1600 m CSF-B were incubated at room temperature (~25°C), samples from between 1600 and 2000 m CSF-B were incubated at 35°C, and samples from below 2000 m CSF-B were incubated at 45°C. After 10 days of incubation, 3 mL of 20% (w/v) zinc acetate solution was injected into each vial and vials were shaken before they were opened to trap produced H2S gas. Slurries were then transferred to 50 mL centrifuge tubes containing 7 mL of 20% (w/v) zinc acetate solution. Tubes were shaken, frozen immediately at –20°C to stop microbial activity, and shipped to Aarhus University, Denmark, after the expedition for analysis of pSRR using the cold chromium distillation method published by Kallmeyer et al. (2004).

Onboard incubation for shore-based microbiological cultivation experiments

Anaerobic incubations of core, cuttings, and formation fluid samples with media targeting subseafloor microorganisms involved in the metabolism of C1 and C2 compounds (e.g., methanogenesis, homoacetogenesis, ferric iron reduction coupled to acetate oxidation, and syntrophic oxidization of volatile fatty acids; Table T10) were initiated on board the ship. Additional WRCs were stored under anaerobic conditions at 4°C for shore-based cultivations.

Inoculum and incubation media

Unwashed cuttings or formation fluids were mixed with anaerobic freshwater basal media (Table T11) or seawater basal media (Table T12). Various combinations of electron donors and acceptors were added to enrich for hydrogenotrophic and aceticlastic methanogens, homoacetogens, ferric iron reducers, and syntrophic volatile fatty acid oxidizers (Table T8). Reducing reagents (i.e., Na2S [1.25 mM] and cysteine-HCl [1.7 mM]) and resazurin were added to ensure that media were completely anoxic. Enrichments of other hydrogenotrophic or chemolithoautotrophic microbes were performed with media containing 40 mM BES (2-bromoethanesulfonic acid), which is a methanogen-specific inhibitor (Smith, 1983). Anaerobic media (10 mL) were placed in glass test tubes sealed with butyl rubber septa and headspaces were flushed with N2:CO2 (80:20 [v/v]) and/or H2:CO2 (80:20 [v/v]). Samples were incubated at temperatures within the in situ temperature range. Unwashed cuttings samples from 696.5 to 1206.5 m MSF were incubated at 25°C. Formation fluid at 1279.5 m WMSF was incubated at 37°C, and the other fluid samples of 1844.0 and 1978.0 m WMSF were incubated at 50°C.

Incubation and microscopic observation of fungi

For shipboard enrichments of fungi, 1–3 mL of core sample was mixed with 5–13 mL of autoclaved seawater in 50 mL tubes and vortexed for 1–3 min. Each tube was then N2-flushed, sealed with tape, and incubated at 28°C while shaking at 100 rpm for 5–15 days. After this incubation period, samples were vortexed again and allowed to settle for 30 min. Supernatants were then examined by light microscopy by placing one drop on a glass slide.

Sampling for shore-based investigations

Pooled master sample for multiple analyses

Eleven WRC samples from the coalbed and intercalated sandstone and clay layers from 1950 and 2000 m CSF-B were used to prepare a pooled master sample (Table T13). The pooled master sample was prepared after carefully removing the outer 1 cm of the WRC samples and powdering the inner, potentially uncontaminated part of the cores. Core from sand layers was excluded from this mixture because of the higher contamination risk, as determined by PFC tracer measurements. This pooled master sample was used to prepare slurries for shore-based microbial activity measurements, incubation and cultivation experiments, and shipboard radiotracer incubations.

Shore-based DNA extraction, functional gene characterization, and organic acid analyses

Remains of the 10 cm WRC for extracellular DNA extraction were transferred to sterile whirlpak bags and frozen at –80°C for further shore-based DNA extractions and functional gene assays, analyses of amino acid enantiomers, and measurement of amino acid, muramic acid, and dipicolinic acid concentrations (Lomstein et al., 2012).

Shore-based 35S incubations

From each WRC sample used for shipboard 35S incubations, a 50 cm3 volume of clean inner core was packed in a gas-tight ESCAL bag, flushed with N2, vacuum-sealed, and stored anoxically at 4°C.

Hydrogenase activity measurements

Samples for hydrogenase activity measurements were handled in an anoxic glove box and taken from the same WRCs used for pSRR measurements. A 50 cm3 volume of clean inner core was packed in a gas-tight ESCAL bag, flushed with N2, vacuum-sealed, and frozen at –80°C.

Cultivation experiments

WRC samples were taken for shore-based cultivation experiments. The heavily contaminated outermost parts were discarded and the clean inner portions were stored by placement either in glass bottles, where they were flushed with N2 and sealed with butyl rubber stoppers, or gas-tight bags, which were flushed with N2 and vacuum-sealed. All samples were stored at 4°C until shore-based cultivation studies targeting microbes (e.g., methanogens, homoacetogens, ferric iron reducers, dehalogenating bacteria, carboxidotorophs, syntrophs, and fungi) (see Table T10). For shore-based fungal cultivation, 20 cm3 core samples obtained from each community WRC were sealed in a bag, flushed with N2, and stored at 4°C.

Single-cell analyses of carbon and nitrogen assimilation rates of subseafloor microbes

We set up incubations for shore-based examinations of microbial physiologies and substrate-uptake ratios by NanoSIMS and other single-cell–based molecular ecological techniques. Contaminated surfaces of WRCs were peeled off with sterile ceramic knives, and less contaminated inner core parts were crushed with a sterile hammer under anaerobic conditions. Crushed samples were distributed to 50 cm3 sterilized glass vials with rubber stoppers for incubation with a wide range of stable isotope-labeled substrates (Table T14). After flushing with argon gas, vials were sealed with screw caps and stored at 4°C. Substrates were added to achieve equimolar 13C (mixture of 15 µM each of 13C and natural abundance isotope ratio C-bearing substrates), 15N (mixture of 1.5 µM of 15N and natural abundance isotope ratio N-bearing substrates), and deuterium (20 vol% in water). For each vial, 19.8 mL anaerobic artificial seawater (1% PBS, 20% D2O, 30 g/L NaCl, 12 g/L MgCl2, and 3 g/L KCl) and 0.2 mL liquid and/or 5 mL gas substrate (50 vol% of 13C or 15N diluted with natural abundance isotope ratio C- or N-bearing substrates) were added. All reagents and gas components had been filtered through 0.2 µm polyvinyllidene fluoride membranes prior to use.

Additionally, a coal sample was prepared for shipboard stable isotope incubation. A WRC was dabbed with Kimwipes soaked in anaerobic Milli-Q water in a glove box and then gently broken into 1–2 cm thick fragments to create artificial fractured surfaces. UV-sterilized 47 mm polycarbonate and cellulose acetate membranes were soaked with substrate and placed between core-breaking points, and the whole thing was “sandwiched” and held together by wrapping the WRC fragments with parafilm. Additional pieces of coal were placed in 500 mL wide-neck glass bottles. Then 300 mL of 3× PBS amended with either 1 mM 13C-bicarbonate and 0.1 mM of 15N-ammonium or 1 mM 13C-acetate and 0.1 mM of 15N-ammonium was added to the bottles and incubated at 51.0°C.

Environmental tag-sequencing and metagenomics

Samples obtained during Expedition 337 provided an unprecedented opportunity to study genetic function and evolution in deeply buried microbial communities with state-of-the-art sequencing techniques. The contaminated outermost 0.5 cm of 20 cm long WRC samples, which had been taken from representative lithologies, were removed, and the less contaminated core interiors were placed into autoclaved perfluoroalcoxy alkane jars and quickly frozen at –100°C. These deep-frozen molecular samples were transferred to the JAMSTEC Kochi Institute for Core Sample Research, where they will be used for molecular ecological studies through 16S rRNA gene-tagged deep sequencing and whole-genome shotgun sequencing.