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Organic geochemistry

Expedition 337 was the first scientific initiative to drill and study a subseafloor hydrocarbon system with riser drilling technology. In this context, organic geochemists aimed to elucidate the cycling of carbon, including the conversion and transport of hydrocarbons, the flux of both thermogenically and biologically produced organic compounds, their utilization as carbon and energy sources by the deep biosphere, and the impact of deep hydrocarbon sources on the carbon budget of the shallower subsurface.

Methodologically, the shipboard organic geochemistry program of Expedition 337 was challenging in several ways. First of all, this expedition was only the second riser drilling operation in the history of scientific ocean drilling. Riser-specific scientific methods are still under development and require evaluation in terms of QA and QC. This is particularly true for the new types of samples that become available through riser technology (i.e., mud gas and cuttings that arrive on board during the recycling of drilling mud). Second, riser drilling differs from riserless drilling with respect to the potential contamination of samples. Mud gas and cuttings are mixed with the drilling mud during their transport from the bottom of the hole to the rig, and the drilling mud used for riser drilling is more alkaline (pH = 9.3–11.9), more viscous, and has a higher density than seawater used for riserless drilling (see “Drilling mud” in the “Site C0020” chapter [Expedition 337 Scientists, 2013]). The potential contamination of sediment cores by drilling mud needs to be considered in the context of organic and inorganic geochemistry but is of the utmost importance for microbiological investigations and was comprehensively tested in the latter context (see “Microbiology”). Finally, DFA was performed for the first time in the framework of IODP (see “Downhole logging”). For the investigation of dissolved organic matter (DOM), dissolved hydrogen (H2), and hydrocarbon gases, DFA provides large-volume fluid samples that are drawn directly from the formation and recovered at in situ pressure.

Organic geochemists investigated gas, solid-phase, and fluid samples. Where possible, established IODP protocols were used (e.g., Pimmel and Claypool, 2001; Underwood et al., 2009). The gas program included continuous online monitoring of mud gas that was extracted from the drilling mud in a separator unit, sampling of mud gas for shore-based analysis, shipboard analysis of gases in cuttings and sediment core samples (gas phase analysis of samples enclosed in gas-tight headspace vials), and sampling and preservation of solid-phase samples from cuttings and sediment cores for shore-based analysis of sorbed gases. Together with hydrocarbon gases and their stable carbon isotopic composition, H2 and carbon monoxide (CO) were targets of high relevance to the expedition objectives. In addition, O2, Ar, N2, and noble gases were monitored in mud gas.

For solid-phase analyses, samples were collected from cuttings and sediment cores. The solid-phase program comprised the shipboard elemental analysis of sedimentary organic matter (C, N, and S), analysis of inorganic carbon content, and characterization of the kerogen type through Rock-Eval pyrolysis (Pimmel and Claypool; 2001). In addition, lipids, including fossil hydrocarbons, phospholipid fatty acids (PLFAs), and intact polar lipids (IPLs), were extracted from cuttings and sediment cores by accelerated solvent extraction (ASE) for both shipboard and shore-based analysis. These different types of biomarkers will help to characterize the deep coalbed biosphere and to constrain the thermal history of the borehole.

For fluid analyses, the interstitial water of sediment cores and fluids obtained by DFA were sampled, and shore-based analysis of DOM will yield information on concentrations and stable carbon isotopic compositions of organic metabolites and on the molecular composition of DOM.

Fresh sediment samples were taken from sediment cores for shipboard and shore-based incubation experiments to study microbial life. Shipboard 14C-radiotracer experiments were employed to determine rates of microbial metabolic activities, such as methanogenesis, acetogenesis, acetate oxidation, and carboxidotrophic pathways. In addition, 13C, 15N, and deuterium (D) labeled compounds were used for stable isotope probing (SIP) experiments that will serve to track substrate utilization and uptake into cellular biomass (e.g., Morono et al., 2011; Kellermann et al., 2012; Wegener et al., 2012). SIP experiments were conducted in close collaboration with shipboard microbiologists, who on their part performed SIP on nucleic acids (see also see “Microbiology”). The analysis of gas, solid-phase, and fluid samples together with radiotracer and SIP experiments will allow us to track the carbon flow in the deep subseafloor biosphere within and above the Shimokita coalbed and will help us test the hypothesis that biogenic methane is formed in situ within the deeply buried subseafloor coalbeds.

In order to achieve our goals, additional equipment was brought on board the Chikyu and new methods were implemented. In particular, a mud-gas monitoring laboratory was set up in a container next to the rig floor, a radiotracer laboratory was installed in a van next to the core cutting area (see “Microbiology”), new protocols for shipboard lipid biomarker analysis were implemented, and additional third-party tools were employed (i.e., a reduced gas analyzer for the analysis of H2 and CO was provided by the Japan Agency for Marine-Earth Science and Technology [JAMSTEC] Kochi Institute for Core Sample Research, and a radon analyzer came from the JAMSTEC Institute for Research on Earth Evolution [IFREE]).


Sampling and analysis of mud gas

Mud-gas monitoring laboratory

Continuous mud-gas monitoring during riser drilling is a standard procedure in oil and gas operations, where it is used to examine reservoir rocks for hydrocarbons and to fulfill safety regulations. In contrast, in scientific ocean drilling, real-time mud-gas monitoring of geochemical parameters is a new technique to investigate sediments that are drilled without coring. In the framework of IODP, continuous mud-gas monitoring was applied for the first time in 2009, when riser drilling was conducted on the Chikyu during Expedition 319 (Saffer, McNeill, Byrne, Araki, Toczko, Eguchi, Takahashi, and the Expedition 319 Scientists, 2010). Our experimental setup builds on previous experience with scientific real-time mud-gas monitoring and sampling in the context of continental drilling (e.g., Erzinger et al., 2006; Wiersberg and Erzinger, 2007, 2011).

Gas extraction system

Mud gas was extracted from the circulating drilling mud by a degasser. The degasser is composed of an explosion-proof electric motor on the top and a cylinder below (Fig. F21A). Inside the cylinder, a smaller degassing chamber, which connects to both the motor and the cylinder, is deployed in the place where gas is separated from the fluid. The gas is pumped online to the mud-gas monitoring laboratory, where the pumping rate that regulates the vacuum applied to the degassing chamber is set. A stirring impeller, which is operated by a motor, ensures the fluid circulation in the degassing chamber. An inlet and outlet on the cylinder allows drilling mud to flow through the whole system.

The degasser was installed right after the flow splitter, where the upcoming drilling mud is first exposed to the atmosphere (Fig. F21B). Some of the drilling mud flowed directly into a bypass line for the degasser. A safety valve outside the degassing chamber prevented overflow of drilling mud into the system (Fig. F21C). The safety valve is a 1 m long cylinder with a central tube that opens at both ends. Water is filled up to 40 cm in the cylinder and the tube. If the gas pressure from the degasser is not sufficient to sustain the preset pumping rate, air can be sucked in from outside and pushes water to flow from the central tube to the cylinder, where it compensates for the pressure difference. The reverse happens when gas pressure from the degasser exceeds the required level.

Gas extracted from the drilling mud traveled through a pipeline (3 mm inside diameter and ~50 m long) to the mud-gas monitoring laboratory, which was set up in a container next to the rig floor. It took ~3 min for gas to arrive at the mud-gas monitoring laboratory (Fig. F21D) (lag time was determined based on the time difference between the start of mud flow and the arrival of mud gas in the monitoring laboratory when drilling resumed after periods in which mud flow had been stopped). For such a short gas traveltime, diffusion loss during transportation is negligible (Wiersberg and Erzinger, 2007). In the mud-gas monitoring laboratory, particles and water vapor were removed from the incoming gas by a mist and moisture remover. Two sampling ports allowed sampling of mud gas either before or after passing the mist and moisture remover. All samples for shore-based analysis were taken from the mud gas before passing the mist and moisture remover. After passing the mist and moisture remover, the dry and clean gas was distributed online to different instruments.

Online analysis of hydrocarbon gases, CO2, CO, Ar, and O2 by gas chromatography

One part of the incoming mud gas was directed to a gas chromatograph (GC)-natural gas analyzer (NGA) (Agilent Wasson ECE 6890N). This instrument is designed to analyze hydrocarbon gases (C1–C5), Ar, O2, N2, CO, and carbon dioxide (CO2). The main component of the GC-NGA system is a GC equipped with a gas sampling port with a multiposition valve. The carrier gas is helium (He). For analysis of hydrocarbon gases, the gas flow is first introduced into a 50 cm capillary column that retains hexane and heavier hydrocarbon components. C1–C5 hydrocarbon gases are then separated by another 49 m capillary column that connects to a flame ionization detector (FID). For analysis of nonhydrocarbon gases, Ar, O2, and CO are separated from the rest of the components by an 8 inch micropack column (Wasson ECE Instrumentation, column Code 2378). CO2 is separated by a 1.27 m capillary column (Wasson ECE Instrumentation, column Code S036). These two columns are connected to a thermal conductivity detector (TCD). The detection limit was 200 parts per million (ppm) for all nonhydrocarbon gases besides CO, which had a detection limit of 400 ppm. The detection limit was <1 ppm for all hydrocarbons. The GC-NGA provides a sensitive method for gas analysis, but the run time of each analysis is rather long (20–30 min), and consequently the temporal resolution and corresponding depth resolution of the mud-gas analysis are low. In addition, separation of Ar and O2 can only be achieved with cryofocusing using liquid N2. This procedure increases the analysis run time to 30–40 min and thus further decreases the depth resolution. Therefore, it was only applied sporadically.

The GC-NGA was calibrated on a daily basis in order to detect any sensitivity changes. Two standards were used. The standard mixture for the calibration of nonhydrocarbon gases contained 1% Ar, CO, Xe, O2, H2, CO2, and He in a balance of N2. The hydrocarbon standard mixture contained 1% C1–C5 in a balance of N2.

GC-FID analysis informs us about the presence of C1–C5 hydrocarbon gases in the formation. However, the interpretation of concentration data is difficult because the gas yield from the drilling mud depends not only on hydrocarbon concentrations in the drilled horizon but also on drilling conditions such as ROP and mud flow. Monitoring of drilling operations is crucial for the comparison and interpretation of quantitative data (see “Recording of online gas analysis and monitoring of drilling operations, time, and depth”). In contrast, the ratio of hydrocarbon gases varies only slightly when drilling parameters change and likely reflects in situ conditions. In particular, the C1/C2+ ratio is a valuable parameter to distinguish between hydrocarbon gases from biogenic and thermogenic sources (Pimmel and Claypool, 2001; Ocean Drilling Program, 1992).

Ar and O2 concentrations allow tracking the introduction of air into the mud-gas monitoring system during drilling operations. Air can be introduced into the borehole when the pipe is broken to recover core, when mud flow is stopped while new pipe connections are made (every 40 m), when pressure drops in the gas separator, and when mud gas is flowing from the bypass line into the flow splitter. Note that CO2 can be analyzed by GC-NGA, but the resulting concentrations are not meaningful because the drilling mud is highly alkaline.

Online analysis of the stable carbon isotopic composition of methane

Another fraction of the incoming mud gas was transferred online to a methane carbon isotope analyzer (MCIA) (Los Gatos Research, Model 909-0008-0000), which detects the concentration and stable carbon isotopic composition of methane on the basis of cavity ring-down spectroscopy technology (cf. van Geldern et al., 2013). This instrument is composed of three parts: the main body of the MCIA, a gas dilution system (DCS-200), and an external pump. The stable carbon isotopic composition of methane is reported in the δ13C notation relative to the Vienna Peedee belemnite (VPDB) standard and expressed in parts per thousand (per mil, ‰), with

δ13C = (Rsample – RVPDB)/RVPDB,


R = 13C/12C, (27)


RVPDB = 0.0112372 ± 2.9 × 10–6. (28)

The precision and analytical error of stable carbon isotope analysis by the MCIA was examined by serial dilution of gas standards (prepared manually). The error was <1‰ for CH4 concentrations of >400 ppm but increased to 3‰ for concentrations ranging from 200 to 400 ppm. The gas dilution system can dilute sample gas 100× with zero-air (hydrocarbon-free compressed air). With such dilution ability, samples with methane concentrations from 200 ppm to 100% can be measured. However, the dilution system did not function because of technical problems and the concentration data from the MCIA were not utilized when they exceeded 1%. Although the quantification of methane by the MCIA is less sensitive than the GC-FID method and higher hydrocarbon gases cannot be analyzed simultaneously, MCIA analyses have the advantage of a short run time (a frequency of 1 measurement/s was chosen here), enabling continuous monitoring of CH4 concentration. Typically, 100–200 measurements were conducted per meter of drilled sediments, depending on the ROP and mud flow rate. The sensitivity of this instrument was checked daily with a gas standard containing 2500 ppm CH413C = –54.5‰ vs. VPDB) and balance air.

Together with C1/C2+ ratios obtained by the GC-FID, the δ13C of methane is an important parameter to distinguish biogenic and thermogenic sources of hydrocarbon gases during mud-gas monitoring (for details see Whiticar, 1999). The classical interpretation is based on the assumption that biological methanogenesis uses CO2 with δ13C values less than –10‰ vs. VPDB. If, however, the pool of dissolved inorganic carbon (DIC) in the sediment is enriched in 13C, isotopic fractionation during methanogenesis will result in methane with similar δ13C values as expected for thermogenic methane (Pohlman et al., 2009). The full interpretation of the stable carbon isotopic composition of methane will require shore-based analysis of the carbon isotopic composition of DIC in interstitial water samples retrieved from sediment cores and DFA.

In the MCIA, individual gas components are not separated prior to analysis and interferences could potentially impact the analysis. Because mud-gas monitoring with the MCIA was used for the first time in the history of scientific ocean drilling, the accuracy of the analysis needs to be confirmed. In order to further verify the analytical results obtained by the MCIA, we preserved samples of mud gas for shore-based stable carbon isotope analysis of methane by isotope ratio monitoring gas chromatography–mass spectrometry (Heuer et al., 2010; Ertefai et al., 2010). Shore-based investigations will also provide information on the stable hydrogen isotopic composition (δD) of methane and potentially on δ13C and δD values of higher hydrocarbon gases, which will help to identify their sources.

Online analysis of gases by process gas mass spectrometer

The third fraction of the incoming mud gas was directed to a process gas mass spectrometer (PGMS) (Ametek ProLine process mass spectrometer) for continuous monitoring of H2, He, O2, Ar, Xe, N2, CO, CO2, methane, ethane, and propane contents. No carrier gas was added, as a vacuum was applied. The PGMS detects gases with a quadrupole mass filter based on the individual molecular masses of the target compounds. A Faraday cup detector provides an optimal scanning range of mass-to-charge ratio (m/z) 1–100 and an optional scanning range of m/z 1–200 with the mass resolution of 0.5 amu at 10% peak height. Input gas flow rate is set to 50 mL/min. For quantification of individual gas species, the PGMS was calibrated on a daily basis using the same standards as for GC-NGA. One standard contained 1% Ar, CO, Xe, O2, H2, CO2, and He in a balance of N2, and the other contained 1% C1–C5 in a balance of N2. Pure N2 and Ar were used for daily background checks. However, the regression line based on the standard could not be used because the concentration of major gas species (N2, O2, Ar, and CO2) in the sample gas was significantly higher than the standard. Therefore, these gas species were calibrated by using the GC-NGA data of sample gas measurements at the same time point. The sensitivity of this instrument is 1 ppm. Although the PGMS is less sensitive than the GC-NGA, it allowed one measurement every 20 s for the selected scan (m/z 1–150) and resulted in better depth resolution.

O2, Ar, and N2 allow us to monitor the input of air during drilling operations (see “Online analysis of hydrocarbon gases, CO2, CO, Ar, and O2 by gas chromatography”), and O2/Ar ratios provide information on corrosive processes at the drill bit and pipe, which might interfere with H2 analysis.

Online analysis of radon

Radon analysis was carried out using a third-party laboratory instrument that was provided by JAMSTEC IFREE. The gas phase concentration of Rn that exsolved from the circulation mud was measured by a stand-alone radon monitor (AlphaGUARD PQ2000 PRO). The apparatus was attached to the auxiliary port of the scientific gas monitoring line parallel to other instruments. It has an ion-counting chamber 650 mL in volume (effective volume is ~500 mL), in which Rn decay is counted directly with a sensitivity of 100 Bq/m3 in the concentration range of 2 to 2 × 106 Bq/m3. Internal temperature, pressure, and relative humidity are recorded automatically. The time-sequential data can be exported to CSV format.

Sampling for shipboard and shore-based analysis

During mud-gas monitoring, discrete samples were taken by IsoTube samplers (Isotech Laboratories, Inc.) for shore-based analysis of δ13C and δD values of hydrocarbon gases, δD values of H2, and quantification of noble gases. In addition, 5 mL of gas was sampled occasionally from the sampling port of the gas flow line and transferred to 20 mL headspace vials for shipboard monitoring of the PFC tracer that was employed by the microbiologists to quantify core contamination with drilling mud (see “Microbiology”).

Recording of online gas analysis and monitoring of drilling operations, time, and depth

During riser drilling, the recovery of gases from drilling mud is affected by drilling operations. Drilling parameters were monitored and recorded in the SSX database. Lag depth was provided by the mud-gas logging contractor (Geoservices). The online data were available in the mud-gas monitoring laboratory and were used as a guide for sampling and data interpretation.

Ship time (UTC + 8) was the primary parameter against which all mud-gas monitoring operations, analyses, and sampling activities were recorded, except for Rn analysis, which used an internal clock set to the time zone UTC + 1. For the interpretation of raw data it is important to note that the internal MCIA clock could not be synchronized and was 90 s ahead of ship time. Because it takes time for mud to be recovered from the bottom of the hole, the depth of the arriving mud gas lags several meters behind the actual depth of the hole bottom at a given time. We used lag Depth A (as recorded in real time in the SSX database) to align data and samples from mud-gas monitoring to sediment depth. Lag Depth A was calculated by the mud-gas logging contractor (Geoservices, Schlumberger) based on mud flow, hole depth, and geometry and was recorded in drilling depth below rig floor (rotary table) (DRF). It can be converted to sediment depth in meters below seafloor (mbsf) by subtracting water depth (1180 m) and distance between water level and rotary table (28.5 m). The converted depth is treated as mud depth below seafloor (MSF).

Aside from drilling parameters, the SSX database also recorded online data that were generated during mud-gas monitoring by the GC-FID (see “Online analysis of hydrocarbon gases, CO2, CO, Ar, and O2 by gas chromatography”), MCIA (see “Online analysis of the stable carbon isotopic composition of methane”), and PGMS (see “Online analysis of gases by process gas mass spectrometer”). However, we noticed that the uploading of data from the instruments to the database suffered from technical problems, and it was not possible to correct for the faulty data. Therefore, we only used raw data as recorded by the individual instruments for further processing and interpretation.

Data were recorded nonstop during operations in Hole C0020A, including periods during which drilling did not advance into the formation and times without mud flow and/or mud-gas recovery. In the course of data processing, the time periods that did not yield meaningful information about the gas content of the geological formation were omitted from the raw data set. Subsequently, the high-resolution online records of the MCIA and PGMS, which yielded as many as 200 data points per drilled meter of sediment, were compiled into 1 m averages.

Background control, quality checks, and comparison of different sampling techniques

Several tests were conducted to account for various potential problems that might arise during geochemical mud-gas monitoring:

  • Background control: the gases that are separated from the drilling mud in the degasser might result from the mixing of drilling mud with sediments, fluids, and gases in the formation, as well as from gases that are originally present in the drilling mud before being sent to the borehole. In order to account for background gases, drilling mud was regularly sampled from the tank and the hydrocarbon gas component was measured on board the ship. Samples were processed in the same way as samples from cuttings and sediment cores for headspace gas analysis with the GC-FID. The available instrumentation did not allow discrete sample analysis to determine background concentrations of other gases that were monitored online in the mud-gas monitoring laboratory.

  • Mist and moisture remover: the mist and moisture remover is an essential unit in the online mud-gas monitoring system. However, it might cause fractionation both with respect to gas contents and their isotopic composition. In order to test the potential effect of the mist and moisture remover on gas contents and the carbon isotopic composition of methane, gas standards were measured with and without passing through the gas dryer. For methane, the effect of the mist and moisture remover was within the analytical uncertainty: compared to methane in the wet gas, the methane content of the dried gas was 3% lower and δ13C values were 0.4‰ more positive (n = 13). For the PGMS, mist and moisture affected the analysis. When air was measured with and without passing through the gas dryer, H2 and O2 concentrations in the wet gas were both 2% higher than in the dried gas, whereas He and CO2 concentrations in the dried gas were 67% and 84% higher than in the wet gas.

  • Verification of results obtained from the MCIA: the MCIA for online mud-gas monitoring was used for the first time in scientific ocean drilling. In order to confirm its accuracy, samples of gas were taken for shore-based analysis by isotope ratio monitoring gas chromatography–mass spectrometry (Heuer et al., 2010; Ertefai et al., 2010).

Sampling and analysis of sediment cores

Hydrocarbon gases

Concentrations and distributions of light hydrocarbon gases, mainly methane (C1), ethane (C2), and propane (C3), were monitored for each core following standard headspace sampling (Kvenvolden and McDonald, 1986). A 5 cm3 sediment sample was collected with a cork borer from the freshly exposed end of the first section that was cut open in each core. In general, this was the section adjacent to the WRC cut for interstitial water sampling. The sample was extruded into a 24 mL glass vial and immediately sealed with a Teflon-coated septum and metal crimp cap. The exact bulk mass of the wet sample was determined after gas analysis was finished. For C1–C4 hydrocarbon gas analysis, the vial was placed in a headspace sampler (Agilent Technologies G1888 network headspace sampler), where it was heated at 70°C for 30 min before an aliquot of the headspace gas was automatically injected into an Agilent 6890N GC equipped with a packed column (GL HayeSep R) and FID. The carrier gas was He. In the GC temperature program, the initial temperature of 100°C was held for 5.5 min before the temperature was ramped up at a rate of 50°C/min to 140°C and maintained for 4 min. Chromatographic response of the GC was calibrated against five different authentic standards with variable quantities of low-molecular weight hydrocarbons and checked on a daily basis.

Methane concentration in interstitial water was derived from the headspace concentration using the following mass balance approach (Underwood et al., 2009):

CH4 = [χM × Patm × VH]/[R × T × Vpw], (29)


  • VH = volume of headspace in the sample vial,

  • Vpw = volume of pore water in the sediment sample,

  • χM = molar fraction of methane in the headspace gas (obtained from GC analysis),

  • Patm = pressure in the vial headspace (assumed to be the measured atmospheric pressure when the vials were sealed),

  • R = universal gas constant, and

  • T = temperature of the vial headspace in degrees Kelvin.

The volume of interstitial water in the sediment sample was determined based on the bulk mass of the wet sample (Mb), the sediment’s porosity (ϕ, which was extrapolated from shipboard MAD measurements in adjacent samples), grain density (ρs), and the density of pore water (ρpw) as

Vpw = Mpwpw = [ϕ × ρpw]/[(1 – ϕ)ρs] × Mbpw, (30)


  • Mpw = pore water mass,

  • ρpw = 1.000–1.024 g/cm3 (adjusted to salinity based on shipboard data), and

  • ρs = 2.8 g/cm3.

Hydrogen and CO

H2 and CO analyses supplemented the routine shipboard analytical program and utilized a third-party laboratory instrument that was provided by the JAMSTEC Kochi Institute for Core Sample Research. The methodology described below focuses mainly on H2. CO data were also generated during the H2 analysis of the same sample set, but note should be taken that the underlying assumptions in the H2 methodology do not necessarily apply to CO. A similar approach had previously been used on board the Chikyu during Expedition 322 (Underwood et al., 2009).

Dissolved H2 concentrations were monitored using two different headspace equilibration techniques. For the first method, hereafter called the incubation method, ~5 cm3 of sediment was collected from the freshly cut section ends. Samples were transferred into 20 mL headspace vials that were closed with thick blue butyl rubber stoppers (Chemglass Life Sciences, Vineland, New Jersey, USA), crimp capped, and thoroughly flushed with He in order to establish an O2-free gas phase inside the vials. After analysis of the initial H2 concentration, samples were incubated at estimated in situ temperatures and H2 concentrations in the gas phase were monitored as a time series. At each time point, 1 mL of gas was sampled using a gas-tight syringe. In order to maintain a constant pressure inside the vials, the withdrawn amount of gas was substituted by injecting an equal volume of He after each analysis. In principle, the time series is supposed to continue until H2 concentrations reach a constant level that represents a steady state between production and consumption. The incubation method allows the determination of dissolved H2 concentration based on two fundamental assumptions: (1) H2 in the headspace is in equilibrium with dissolved H2 in the pore water, and (2) the incubation of samples in the laboratory allows the establishment of a biologically controlled steady state that is representative of in situ equilibrium (Hoehler et al., 1998).

The incubation method was initially developed for studies of freshwater sediments and microbial cultures (Lovely and Goodwin, 1988; Hoehler et al., 1998). In contrast to these metabolically active systems, deep-marine subsurface sediments host microorganisms that metabolize at very low rates (D’Hondt et al., 2002; Parkes et al., 2005). Therefore, it is unclear whether the required steady state can be reached within an acceptable time frame in the laboratory and whether such a steady state would be representative of in situ conditions. Alternative methods are needed for the determination of dissolved H2 concentration in deeply buried sediments. However, the establishment of suitable methods is not a trivial task because of potentially low in situ H2 concentrations and possible sampling artifacts. An alternative approach is the complete extraction of dissolved H2 into a defined, H2-free gas volume as previously used by Novelli et al. (1987) and D’Hondt et al. (2009), hereafter called the extraction method. In principle, this method is based on the assumption that the initially present H2 exsolves from the liquid phase and can be captured in the defined headspace volume of a closed vial. Dissolved H2 concentration can then be calculated using a mass balance (see below). The suitability of the extraction method for deep biosphere studies has recently been evaluated by Lin et al. (2012).

For the extraction method, ~3 cm3 of sediment was sampled immediately after core recovery from the freshly exposed end of a core section. Typically, this was the same section that was used for hydrocarbon gas analysis. The sample was extruded into a 10 mL headspace vial, which was immediately completely filled with NaCl solution (3.5%), sealed with a butyl stopper (GL Science), and crimp capped. Excess water was allowed to escape through a hypodermic needle. Analysis was conducted as soon as possible after core recovery and sampling. A headspace was created by displacing 5–10 mL of the aqueous phase with an equal volume of H2-free He. The gas-in needle was removed first, and the liquid-out needle connected to a syringe was allowed to equilibrate the pressure in the vial headspace to atmospheric pressure. The volume offset in the liquid-out syringe was recorded. The vial was vortexed, and dissolved H2 was allowed to diffuse out of the interstitial water and equilibrate with the headspace for 20 min before H2 concentrations were analyzed in the headspace gas. Background controls are essential for accurate H2 analysis by the extraction method. The reagent blank (analysis of vials filled with only NaCl solution) was 3.2 nM H2. Samples of drilling mud were processed in the same way.

In both the incubation and extraction methods, dissolved H2 concentration was determined based on the H2 concentration in the headspace gas, which was analyzed by gas chromatography with a Reduced Gas Detector using a SRI 8610C (SRI Instruments). Samples are injected into a flow of carrier gas and separated on a packed column before they react with a heated bed of mercuric oxide and form mercury vapor that is subsequently detected in a photometer cell. For H2, the reaction is H2 + HgO(solid) H2O + Hg(vapor). The instrumental detection limit, evaluated statistically by a serial dilution of the primary standard with He, is ~300 parts per billion (ppb). The instrument was operated using a column temperature of 75°C and He as a carrier gas and was calibrated with a 3 ppm H2 primary standard on a daily basis. Typically, 3 mL of gas sample was injected to thoroughly flush the 1 mL sample loop and the tubing between the injection port and the loop.

The incubation and extraction methods use different approaches to deduce the dissolved H2 concentration from the analyzed headspace concentration, but for both methods, the first step is to convert H2 concentration in the headspace from molar fractions to molar concentration ([H2]g):

[H2]g = χH2 × P × R–1 × T–1, (31)


  • [H2]g = expressed as nmol/L,

  • χH2 = molar fraction of H2 in the headspace gas (in ppb, obtained from GC analysis),

  • P = total gas pressure (in atm) in the headspace (1 atm),

  • R = universal gas constant, and

  • T = temperature of the gas phase in degrees Kelvin.

For the incubation method, the concentration of H2 dissolved in the interstitial water ([H2]incub, expressed in nmol/L) is assumed to be in equilibrium with the gas phase and calculated as

[H2]incub = β × [H2]g, (32)

where β is an experimentally determined solubility constant corrected for temperature and salinity (Crozier and Yamamoto, 1974). The value of β is 0.01555 for seawater (salinity = 33.7 parts per thousand) at 19.3°C.

For the extraction method, the concentration of H2 dissolved in the interstitial water ([H2]extract, expressed in nmol/L) is determined based on mass balance:

[H2]extract = ([H2]g × Vg + [H2]aq × Vaq) × Vs–1 × ϕ–1, (33)


  • [H2]g = calculated using Equation 31;

  • [H2]aq = H2 concentration in the aqueous phase (obtained from Equation 32, substituting [H2]aq with [H2]incub and β = 0.0061 for saturated NaCl solution [salinity = 264‰] based on the extrapolated value of Wiesenburg and Guinasso [1979]);

  • Vg = volume of the headspace;

  • Vaq = volume of the aqueous phase, including the pore water and the solution added;

  • Vs = volume of the sediment sample; and

  • ϕ = sediment porosity.

Lithification of sediments hampered the accurate measurement of sampled sediment volumes. Therefore, Vs was obtained based on the volume of NaCl solution that was added initially to completely fill the 14 mL headspace vial.

Gas analysis during downhole fluid analysis and sampling

In situ fluid analysis

DFA enables in situ fluid analysis as well as retrieval of fluid samples. In this manner, organic geochemistry was included in the downhole logging program. For more information see “Downhole logging.”

Sampling and gas analysis in fluids retrieved by DFA

To determine the contents and isotopic compositions of dissolved volatile components in a fluid sample taken by DFA, we extracted the dissolved gases in the fluid under vacuum conditions (Saegusa et al., 2006). About 250 mL of sampled fluids was recovered in a gas-tight cylinder and transferred to a preevacuated glass extraction bottle (~350 mL), leaving ~100 mL headspace in the extraction bottle. In order to facilitate the extraction of the dissolved gases from the fluid into the headspace, the extraction bottle was ultrasonicated at 25°C for 5 min. To determine the total gas volume in the fluid, the pressure was measured by a pressure gauge. The extracted headspace gas was sampled into preevacuated stainless steel bottles and/or glass bottles and vacuum glass vials. After extraction of dissolved gases, the fluid sample was further processed by inorganic geochemists (see “Inorganic geochemistry”) and microbiologists (see “Microbiology”).

Solid phase

Solid-phase sampling

Solid-phase sampling from cuttings

When drilling without coring, cuttings were sampled for both shipboard and shore-based investigations of the solid phase. For Unit I, in which there was no core retrieval, cuttings were sampled every 50 m for lipid biomarker work. The cuttings surfaces were contaminated by drilling mud or other organic materials used for drilling operations. To minimize signals from contaminants, volumes of 50 cm3 cuttings were rinsed first with seawater obtained from the ship’s seawater tap and then with freshwater. Subsequently, we ultrasonicated the cuttings three times, each time for 3 min, using organic solvents in the order of methanol, 1:1 v/v methanol/dichloromethane, and dichloromethane. The cleaned pellets were dried at 60°C and hand ground into powder for elemental, carbonate, Rock-Eval, and biomarker analyses. For Units II–IV, freshwater-washed cuttings were selected only from a few depths that are close to the coring intervals, vacuum-dried, and homogenized for subsequent analyses.

Solid-phase sampling from cores

In order to characterize the sedimentary organic matter and to analyze biomarkers in the solid phase of sediment cores (RCB and LDC), samples were taken from the WRCs dedicated for microbial community analysis (“community WRC”) (see “Microbiology”). After the other small-volume samples were taken from the community WRCs, the remaining sediment was visually examined, the contaminated materials were removed, and the cleaned sediment was split into two aliquots. One aliquot (10–50 cm3) was stored in a plastic bag and kept at –20°C for shipboard lipid biomarker analysis. The other aliquot (10–70 cm3) was transferred into a high-density polyethylene can and kept at –20°C for shore-based supplementary lipid analyses and aqueous extraction of DOM. The shipboard sediment samples were vacuum-dried and ground into powder using either mortars or a SPEX CertiPrep 6850 freezer mill (SPEX CertiPrep Inc., Metuchen, New Jersey, USA), depending on the lithology of the sample. After homogenization, a small aliquot (3–5 g) of sediment was transferred into a glass vial and stored at room temperature for elemental, carbonate, and Rock-Eval analyses. The rest of the sediment powder was stored at –20°C before solvent extraction. In order to increase depth resolution, additional samples were taken in between community WRCs for elemental and Rock-Eval analysis.

For shipboard and shore-based experiments with live sediment, WRCs were first cut from undisturbed core sections after careful X-ray CT scanning; the sections of core liner containing the sediment were then closed with end caps, placed in air-tight bags, flushed with N2, and finally stored at 4°C until further processing by the individual investigators.

Solid-phase analysis

Total carbon, nitrogen, and sulfur contents

Total carbon (TC), total nitrogen (TN), and total sulfur (TS) concentrations were determined using a Thermo Finnigan Flash EA 1112 carbon-hydrogen-nitrogen-sulfur (CHNS) analyzer. Calibration was based on the synthetic standard sulfanilamide, which contains 41.81 wt% C, 16.27 wt% N, and 18.62 wt% S. About 20–50 mg of sediment powder was placed in a tin container and weighed for TC and TN analyses. For TS analysis, the same amount of sediment powder was put into a tin container, weighed, and mixed with an equivalent mass of V2O5 catalyst. Sediment samples were combusted at 1000°C in a stream of O2. Nitrogen oxides were reduced to N2, and the mixture of CO2, N2, and SO2 was separated by GC and detected by TCD on the CHNS analyzer. Standard deviations of TC, TN, and TS for the samples were less than ±0.1%. Accuracy of the analyses was confirmed using soil NCS reference material (Thermo Scientific, Milan, Italy), sulfanilamide standard (Thermo Scientific), and JMS-1 reference material.

Inorganic carbon, organic carbon, and carbonate content

With the same set of samples used for elemental analysis, we determined inorganic carbon concentrations using a Coulometrics 5012 CO2 coulometer. About 10–20 mg of sediment powder was weighed and reacted with 2 M HCl. The liberated CO2 was titrated, and the change in light transmittance was monitored with a photodetection cell. The weight percentage of calcium carbonate was calculated from the inorganic carbon content, assuming that all the evolved CO2 was derived from dissolution of calcium carbonate, by the following equation:

CaCO3 (wt%) = inorganic carbon (wt%) × 100/12. (34)

No correction was made for the presence of other carbonate minerals. Standard deviations for repeated analyses on individual samples were less than ±0.1 wt%. NIST-SRM 88b and JSD-2 (standard reference materials) were used to confirm accuracy. Total organic carbon (TOC) contents were calculated by subtraction of inorganic carbon from TC contents as determined by elemental analysis.

Characterization of type and maturity of organic matter by Rock-Eval pyrolysis

Rock-Eval pyrolysis was used to characterize the type and maturity of the sedimentary organic matter and to identify its petroleum potential. In principle, Rock-Eval pyrolysis utilizes the sequential heating of a sample in the inert atmosphere (He) of a pyrolysis oven to quantitatively and selectively determine (1) the free hydrocarbons contained in the sample and (2) the hydrocarbon- and oxygen-containing compounds (CO2) that are created during cracking of the kerogen in the sample. In addition, the shipboard instrument, a Rock-Eval6 Standard can also be used to oxidize and quantify the residual organic carbon (i.e., the organic matter remaining after pyrolysis).

Rock-Eval pyrolysis yields the following basic parameters (Espitalié et al., 1977; Peters, 1986):

  • S1 = amount of free hydrocarbons (gas and oil) in the sample volatilized at temperatures <300°C (expressed in milligrams of hydrocarbon per gram of sediment).

  • S2 = amount of hydrocarbons generated through volatilization of very heavy hydrocarbon compounds (>C40) and the pyrolytic cracking of nonvolatile organic matter (in milligrams of hydrocarbon per gram of sediment). S2 is an indication of the quantity of hydrocarbons that the sediment can potentially produce should burial and maturation continue.

  • S3 = amount of CO2 (in milligrams CO2 per gram of sediment) produced during combustion of the sample. S3 is an indication of the amount of oxygen in the kerogen.

  • Tmax = temperature at which the maximum release of hydrocarbons from cracking of kerogen occurs during pyrolysis. Tmax is an indication of the stage of maturation of the organic matter.

  • HI = hydrogen index (HI = [100 × S2]/TOC; in milligrams of hydrocarbon per gram of TOC). HI correlates with the H/C ratio, which is high for lipid- and protein-rich organic matter of marine algae.

  • OI = oxygen index (OI = [100 × S3]/TOC; in milligrams CO2 per gram of TOC). OI is a parameter that correlates with the O/C ratio, which is high for polysaccharide-rich remains of land plants and inert organic material (residual organic matter).

  • PI = production index (PI = S1/[S1 + S2]). PI is used to characterize the evolution level of the organic matter by the proportion of free hydrocarbons present.

  • PC = pyrolyzable carbon (PC = 0.083 × [S1 + S2]). PC corresponds to the carbon content of hydrocarbons volatilized and pyrolyzed during the analysis.

  • RC = residual organic carbon.

Samples of ~60 mg of dry sediment were obtained from the same dried and homogenized bulk sample that had been used for elemental analysis. The temperature program of the pyrolysis oven used the following procedures. For 3 min, the oven was kept isothermally at 300°C and the volatilized free hydrocarbons were measured as the S1 peak (detected by the FID). The temperature was then increased from 300° to 550°C at 25°C/min. The hydrocarbons released from this thermal cracking were measured as the S2 peak (by the FID), and the temperature at which S2 reached its maximum was recorded as Tmax. The CO2 released from kerogen cracking was trapped in the 300°–390°C range. The trap was heated and the released CO2 was detected as the S3 peak (by the TCD). The residual organic matter was oxidized in an oxidation oven kept at 600°C.

Lipid analysis

Analysis of hydrocarbon biomarkers in drilling cuttings is routine and has been demonstrated for commercial drilling (Peters et al., 2005), but this is not the case for PLFAs. Shipboard work focused on preparation of total lipid extracts (TLEs), which were split into two aliquots. One aliquot (20%) was used for shipboard analysis of fossil hydrocarbon molecular markers and free fatty acids. The other aliquot (80%) was stored at –20°C and shipped to the Center for Marine Environmental Sciences (MARUM; University of Bremen, Germany) where it will be used for shore-based analysis of PLFAs and IPLs. Shipboard analysis of fossil hydrocarbon markers aimed to characterize the sources of sedimentary organic matter and to construct a thermal history of Site C0020. Shore-based lipid analysis will be carried out to characterize the downcore distribution of PLFAs and IPLs. PLFAs are fatty acids derived from ester-linked phospholipids; IPLs include phospholipids and other types of membrane lipids with the polar head groups remaining attached to the core lipids. These polar lipids are found to degrade rapidly after cell death or lysis (Harvey et al., 1986; Logemann et al., 2011) and have thus been used as proxies for extant prokaryotic biomass (e.g., Mills et al., 2006; Lipp et al., 2008). However, in deep subseafloor sediments, a fossil contribution of typical archaeal IPLs of >50% may complicate their use as quantitative proxies for live biomass (Xie et al., 2013). Results from the shore-based PLFA and IPL analyses will be compared with microbiological data to provide a complementary estimate of bacterial biomass. The analytical scheme for biomarker analysis is shown in Figure F22.

Solvent extraction of the solid phase and splitting of TLE. Both cuttings and samples taken from sediment cores were used for lipid analysis. The cuttings surfaces were contaminated by drilling mud or other organic materials used for drilling operations. To minimize signals from contaminants, we ultrasonicated the cuttings three times, for 3 min each time, using organic solvents in the order of methanol, 1:1 v/v methanol/dichloromethane, and dichloromethane. The cleaned pellets were dried at 60°C and hand ground into powder for lipid extraction.

The pulverized samples were subjected to ASE using an Accelerated Solvent Extractor 200 (Dionex, Osaka, Japan) and subsequently split into two aliquots as shown in Figure F22. Before extraction, known quantities of 1-alkyl-2-acetyl-sn-glycero-3-phosphocholine and 1,2-dihenarachidoyl-sn-glycero-3-phosphocholine were added for shore-based quantification of IPLs. The samples were loaded into 11 mL extraction cells and extracted twice under the following conditions:

  • Solvent = dichloromethane/methanol (9:1 v/v).

  • Temperature = 80°C.

  • Pressure = 1000 psi.

  • Preheat time = 1 min.

  • Heat-up time = 5 min.

  • Static extraction time = 5 min.

  • Cycles = 3.

  • Flush volume = 20%.

  • Purge time = 90 s.

To minimize and monitor the level of contamination during this procedure, all extraction cells were preprocessed using the same program before use, and an empty extraction cell was routinely processed along with the samples to make the procedural blank. The obtained TLE was evaporated under a stream of N2 in a Zymark TurboVap LV (Sotax Corporation, Hopkinton, Massachusetts, USA) and transferred to preweighed glass vials. After determination of dry weight, the extracts were split into two aliquots for shipboard and shore-based analysis, respectively, and stored at –20°C until further processing.

Subsequently, the TLE split for the shipboard analysis of hydrocarbon molecular markers was subjected to silica gel column chromatography to separate saturated, aromatic, and polar fractions; a maltene-asphaltene separation was not performed. The saturated hydrocarbon fraction was eluted with hexane, the aromatic hydrocarbon fraction was subsequently eluted using 3:1 v/v hexane/dichloromethane, and the polar fraction was recovered using 1:1 v/v dichloromethane/methanol (on Fig. F22 these three fractions are denoted as F1, F2, and F3, respectively). The polar fraction was dried under N2 prior to the addition of a nonadecanoic acid standard and derivatization with BSTFA (N,O-bis[trimethylsilyl]trifluoroacetamide) to convert alcohols and carboxylic acids into their trimethylsilyl (TMS) ethers and esters. The standard 5β-cholane was added to fractions for quantification.

Instrumental analysis. Lipids were analyzed using an Agilent 5973 mass spectrometer linked to an Agilent 6890N GC (Agilent Technologies Inc., Santa Clara, California, USA). Separation of compounds was achieved using a HP-5MS column (30 m × 0.25 mm, 0.25 µm film thickness; Agilent Technologies Inc.) with He as the carrier gas. Samples were injected into a split/splitless injector at 300°C in splitless mode. Slightly different temperature programs were used to analyze different compound classes. For aliphatic and aromatic hydrocarbons, the oven temperature was set at 60°C upon sample injection and held for 1 min, increased to 150°C at 10°C/min, increased further to 310°C at 25°C/min, and held isothermal for 25 min. For fatty acids, the initial oven temperature was set to 80°C and the oven was kept at 310°C for 22.5 min. The mass spectrometer was operated in the selected ion monitoring (SIM) mode (ionization energy = 70 eV, operating in SIM mode, dwell time = 0.1 s/ion) for quantification. Compounds were identified by comparison of retention times to well-characterized samples, and selected samples were also analyzed in the scan mode to aid identification. The following standards were used to estimate the response factors of analytes in each compound class: anthracene for polycyclic aromatic hydrocarbons; C20, C28, and C31 n-alkane standards for n-alkanes; and C16, C18, and C20 n-alkanoic acids for alkanoic acids. Concentrations of sterenes, steranes, and hopanoids are reported relative to an internal 5β-cholane standard. The TMS ester of fatty acids was quantified in SIM mode with m/z 117 (O = C+OSi[CH3]3) as the target ion.

Sample processing for shipboard and shore-based incubation experiments

Studies using 14C radioisotope tracers

Shipboard incubations for the radiotracer experiments with 14C labeled substrates were conducted by two methods: under in situ pressure using a pressurized incubation chamber and under atmospheric pressure.

Incubation under atmospheric pressure. WRCs were cut into 10 cm sections. Ten samples (2 mL) were taken from each WRC by open-cut 2.5 mL disposal plastic syringe or cork borer and transferred into 5 mL glass vials and capped with butyl rubber stoppers and aluminum caps in an anaerobic glove box. These samples were taken from the center of the WRC, avoiding sediment near the core liner to minimize the possibility of contamination. The vials were brought out of the glove box, and then 1 mL of anaerobic medium was added to make a slurry using a plastic disposable syringe via the butyl rubber stopper.

Radioisotope tracers were injected into slurries in the radioisotope container laboratory on the Chikyu. A total of 50 µL of dissolved radioisotope tracers (sodium [14C]-bicarbonate, [1-14C]-acetate, and [2-14C]-acetate, 0.1 MBq in deoxygenated, deionized water) were separately injected into each vial using a glass microsyringe to determine hydrogenotrophic methanogenesis, acetoclastic methanogenesis, and syntrophic acetate oxidation. For carbon monoxide and methane turnover, 0.5 mL of gaseous radioisotope tracers ([14C]-methane and [14C]-carbon monoxide, 1 MBq) were injected by a plastic disposable syringe. The samples were incubated during the expedition at various temperatures (10°, 20°, and 40°C) depending on the estimated temperature at the drilled depth and sent to KCC for shore-based measurements.

Incubation in pressurized chamber. WRCs from selected lithologies were cut into 20 cm sections. From each 20 cm section, 10 samples (10 mL) were then taken by cork borer and transferred into a stainless steel chamber in an anaerobic glove box. The chamber consists of a Teflon-coated stainless steel tube (125 mm × 9.4 mm inside diameter) and two needle valves (Swagelok Tube Fitting, USA) attached at both sides of the tube. Pressurization and injection of radioisotope tracers were conducted in the radioisotope container laboratory. One side of the chamber was connected with a high-pressure liquid chromatograph (HPLC) pump (JUSCO Corporation, Japan) with a 1 inch HPLC line. The valve connected with the HPLC pump was opened, and the inner part of the chamber was pressurized by inflow of medium. The pressure was set to the estimated in situ pore water pressure (10–22 MPa). A total of 500 µL of dissolved radioisotope tracers and gaseous radioisotope tracers (sodium [14C]-bicarbonate, [1-14C]-acetate, and [2-14C]-acetate, 0.5 MBq in deoxygenated, deionized water; [14C]-methane and [14C]-carbon monoxide, 0.5 MBq) were separately injected into each chamber via a sample loop. After pressurizing, the valve was closed and the chamber was disconnected and stored at different temperatures (10°, 20°, and 40°C) depending on the estimated in situ sediment temperature. The samples were sent to KCC for shore-based analysis.

Studies using stable isotope probing

Two types of SIP experiments were designed and carried out by organic geochemists during Expedition 337 to investigate biomass production and substrate degradation, respectively. SIP experiments conducted by microbiologists, involving nucleic acid-SIP, are discussed in “Microbiology.” The goal of the “biomass production” experiments was to assess the lipid production potential in the subseafloor sediment, whereas the “substrate degradation” experiments aimed at assessing the potential degradation rates of compounds that are relevant to the coalbed system. The WRC samples for the SIP experiments were chosen from the least disturbed core segments according to the images of X-ray CT scan. The WRCs were transferred into ESCAL bags, which were evacuated and flushed with N2 for a total of three cycles before being heat-sealed and stored at 4°C before further processing. The experiments usually started on board the ship within 2 days after core retrieval and were designed to continue for several months postcruise at MARUM.

Biomass formation experiments. Two types of “biomass formation” experiments were started on board the ship:

  • Type A: intact sediment chunks were incubated using the dual isotope labeling method introduced by Wegener et al. (2012) (i.e., D2O and NaH13CO3 without addition of organic substrates). We assumed that such an incubation condition induces minimal disturbance to microorganisms and their habitats. Therefore, the lipid production rates derived from the labeling results can be considered as reasonable estimates of the in situ rates.

  • Type B: sediment was homogenized and slurried with medium containing D2O, NaH13CO3, and with/without unlabeled organic substrates. Microbial activity is intended to be stimulated because of increased contact with the aqueous phase and/or the presence of labile organic matter. The incubation condition is likely to stimulate microbes that are dormant in situ, and the derived lipid production rates will be taken as indication of microbial vitality were the microbes given more water and/or more degradable carbon sources.

Shipboard salinity measurements did not indicate the presence of freshwater; therefore, we prepared an artificial seawater medium with the following salt ingredients and a salinity close to that of seawater for the experiments: 26.4 g NaCl, 11.3 g MgCl2·6H2O, 1.5 g CaCl2·2H2O, 0.68 g KCl, and 0.099 g KBr per liter. NH4Cl and KH2PO4 were then added to a final concentration of 4.7 and 1.5 mM, respectively. To avoid dilution of the 13C label, no NaHCO3 was added to the medium. Resazurin was added as a redox indicator (final concentration = 1 mg/L). The medium was reduced with HCl-amended Na2S·9H2O to a final concentration of 1 mM and neutralized with 1 M NaOH. The stock solutions of D2O, NaH13CO3, glucose, and amino acid mixture were filter-sterilized (0.2 µm), and O2 was removed by bubbling with N2.

To start the experiments, the core segment was extruded from the core liner on a clean bench, visually inspected and contaminated material removed, and transferred immediately into a glove box before further processing. Two WRC samples containing coal were processed directly in the glove box to avoid extensive oxidation because of the highly fractured nature of the coal. Within the glove box, ~50 cm3 of core sample, used as the time-zero sample for the Type A experiments, was transferred to a high-density polyethylene can and later stored at –20°C. Another 600 cm3 of core sample was split into two 500 mL wide-neck Schott bottles for the Type A experiments, whereas the remaining sediment (~400–500 cm3) was powdered over a sterilized tungsten mortar for the other experiments. Anoxic medium was added to fill ~80% of the headspace of the wide-neck bottles, which were then sealed with rubber stoppers and removed from the glove box. To make the sterilized control, one of the bottles was supplemented with zinc acetate to a final concentration of ~10% and pasteurized at 80°C for 8 h. D2O and NaH13CO3 solution were then added.

About 50 cm3 of the homogenized sediment was saved as the time-zero sample for the Type B experiments. The remaining sample powder was mixed with the anoxic medium to a volume ratio of 1:0.8. Slurry aliquots of 90 mL sediment slurry were distributed to 100 mL Schott bottles, which were sealed with rubber stoppers and removed from the glove box. Two of the bottles were incubated only with D2O and NaH13CO3, whereas the others were amended with glucose or amino acid mixture (both to a final concentration of 1 mM) as separate substrates in addition to the labels.

For both experiments, the estimated labeling strengths were ~20% for D2O and 50% for NaH13CO3, with the latter assuming an alkalinity of 10 mM in interstitial water (the highest alkalinity encountered in 337 was 14.2 mM). The actual values will be determined using samples taken from the aqueous phase of the incubation at the start of the experiments. The samples will be incubated at estimated in situ temperatures for up to 6 months and analyzed for lipid isotopic values at MARUM.

Substrate degradation experiments. The following three 13C-labeled substrates were used in the “substrate degradation” experiments:

  1. 13C-labeled lignin purified from Zea mays,

  2. Methoxy-13C vanillin as the model monomer for plant-derived aromatic compounds, and

  3. 1,2-13C hexadecane as the model compound for aliphatic hydrocarbons.

Progress of biodegradation will be monitored by the δ13C values of methane and DIC in the gaseous and aqueous samples taken from the sediment slurries. Once active biodegradation is confirmed, single-cell nanoscale secondary-ion mass spectrometry (NanoSIMS) analysis will be carried out using the solid phase in order to evaluate the extent of carbon assimilation from the labeled substrates into biomass (see “Microbiology”).

The anoxic medium described above (see “Biomass formation experiments”) was also used for the experiments. In a glove box, an aliquot of the homogenized sediment (~100 cm3) was mixed with medium to a volume ratio of 1:1. Aliquots of 10 mL sediment slurries were transferred into 16 mL glass tubes, which were sealed with rubber stoppers and exported from the glove box. 13C-labeled lignin was added as suspension to a final concentration of 0.06 mg/mL slurry for sediment and 1 mg/mL slurry for coal samples. The dosages of methoxy-13C vanillin, amended as aqueous solution, were 0.06 mg/mL slurry for sediment and 0.7 mg/mL slurry for coal samples. The stock solution of 1,2-13C hexadecane was prepared anoxically using 2,2,4,4,6,8,8-heptamethylnonane, an inert and nontoxic organic carrier, as the solvent. This labeled aliphatic hydrocarbon was added to a final concentration of 85 µg/mL slurry for both sediment and coal samples. After label addition, the time-zero tubes were immediately sampled for the gaseous and aqueous phase, whereas the remaining solid-phase slurry was fixed with 2% paraformaldehyde and preserved in phosphate-buffered saline (PBS)/ethanol (1:1 v/v) at –20°C until analysis. The other sample tubes will be incubated at estimated in situ temperatures for as long as 6 months, with methane and DIC isotopic values monitored regularly at MARUM.