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doi:10.2204/iodp.proc.317.102.2011

Geochemistry and microbiology

Organic geochemistry

Shipboard organic geochemistry for Expedition 317 included routine sets of analyses for (1) hydrocarbon gas in sediment cores, (2) inorganic carbon content of sediment, (3) total carbon (TC), total nitrogen (TN), and total sulfur (TS) content of sediment, and (4) pyrolysis characterization of sedimentary organic matter using the source rock analyzer (SRA). Most of the procedures and instruments used during Expedition 317 were described by Pimmel and Claypool (2001) and generally are similar to those used during recent IODP expeditions. Comments on routine sampling and deviations from standard practice are noted below and in the individual site chapters.

Hydrocarbon gases

Sediment gas composition was typically determined at each interstitial water sampling point or at least once every core. Two sampling methods were employed: headspace (HS) sampling and gas void (VAC) sampling using a syringe. HS gas is given off after a known quantity of sediment is heated in a vial. VAC sampling is a direct extraction of gas from visible expansion voids within the core liner of the recovered core. Sediment plugs for HS analysis were taken directly after the core was brought onto the catwalk and were prepared for analysis using two methods (Fig. F10):

1. The first sediment plug (code HS), consisting of 5 cm³ of sediment, was usually sampled using a cork borer, put into an HS vial, and crimp-sealed for standard IODP hydrocarbon safety monitoring. When the sediment became too lithified to extract using a cork borer, fragments of core were chiseled out of the core and placed in the vial. HS samples for onboard analyses were heated at 70°C for 30 min before being injected into the gas chromatograph (GC).

2. The second sediment plug (code LIPP1), consisting of 3–5 cm³ of sediment, was sampled as above and put into a vial containing 5 mL of a 1M sodium hydroxide solution, capped with a precleaned butyl stopper (heated with 1M KOH and rinsed three times with nanopure water), and frozen upside down for shore-based analyses of stable carbon isotopes.

Gases obtained by either HS or VAC sampling were analyzed by one of two GC systems: an Agilent/HP 6890 Series II (GC3) or an Agilent/HP 6890A natural gas analyzer (NGA). Gases were introduced by injection from a 5 mL syringe directly connected to the GC system via a 1 cm³ sample loop; helium was used as the carrier gas.

The GC3 system determines concentrations of methane (C₁), ethane (C₂), ethene (C₂=), propane (C₃), and propene (C₃=) with a flame ionization detector (FID) using a 2.4 m × 3.2 mm internal diameter (ID) stainless steel column packed with 100/120 mesh HayeSep R. Helium was used as the carrier gas, and the GC oven temperature was programmed to hold for 0.5 min at 90°C, ramp at 30°C/min to 100°C, ramp at 15°C/min to 110°C, remain at 110°C for 4.5  min, and then ramp at 50°C/min to 150°C with a final holding time of 1.8 min. The FID temperature was 250°C.

The NGA system measures concentrations of C₁–C₇ hydrocarbons with an FID as well as concentrations of N₂, O₂, and CO₂ with a thermal conductivity detector (TCD). TCD separation used three columns: a 6 ft × 2.0 mm ID stainless steel column (Poropak T [50/80 mesh]), a 3 ft × 2.0 mm ID stainless steel molecular sieve column (13X; 60/80 mesh), and a 2.4 m × 3.2 mm ID stainless steel column packed with 80/100 mesh HayeSep R (Restek). FID separation was performed on a DB1 capillary column (60 m × 0.32 mm) with 1.5 µm phase thickness. FID separation used helium as the carrier gas, and the GC oven temperature was programmed to hold for 2 min at 50°C, ramp at 8°C/min to 70°C, and then ramp at 25°C/min to 200°C with a final holding time of 5 min. The FID temperature was 250°C.

Data were collected and evaluated with an Agilent ChemStation data-handling program. For both systems, chromatographic response was calibrated to nine different gas standards with variable quantities of low molecular weight hydrocarbons, N₂, O₂, CO₂, Ar, and He and checked on a daily basis. Gas components are reported as parts per million by volume (ppmv) of the injected sample. Methane in the uppermost HS samples is also expressed as millimoles per liter of pore volume (mM), assuming a porosity of 0.45, a sample volume of 5 cm³, and a vial volume of 21.5 cm³:

Sampling for carbonate and organic matter analyses

Sediment samples collected for shipboard analysis (nominally 10 cm³ wet volume; typically ~3 g dry mass) were selected from the working half of the core on the sampling table in collaboration with sedimentologists. Samples were selected based on (1) major lithology, so that questions regarding carbonate content could be answered, and (2) any darker, more organic-rich lithologies, based on visual differences. Samples were freeze-dried for at least 12 h. Dried samples were crushed to a fine powder and carefully homogenized with a mortar and pestle in preparation for carbonate and organic matter analyses.

Inorganic carbon and carbonate

The inorganic carbon (IC) content of sediment samples was determined by coulometry using a UIC 5011 CO₂ coulometer in which samples of ~10 mg of freeze-dried, ground sediment were reacted with 1N HCl. The liberated CO₂ was back-titrated to a colorimetric end-point. The carbonate content of sediment (in weight percent) was calculated from IC content by assuming that all carbonate occurs as calcium carbonate:

Accuracy was ensured during analysis batches by running a carbonate standard (100 wt% CaCO₃) every 10 samples and continuing analysis only if values were between 99 and 101 wt%. Typical precision was assessed using 11 replicate analyses of a carbonate sample (Table T8), which gave a standard deviation of 1.2 for a sample with a mean carbonate content of 66.4 wt% and a coefficient of variation of 0.018.

Elemental analyses

TC, TN, and TS contents of sediment samples were determined with a ThermoElectron FlashEA elemental analyzer 1112 equipped with a ThermoElectron packed column (CHNS/NCS) and a thermal conductivity detector (TCD). An aliquot of 8–12 mg of freeze-dried, ground sediment was weighed into tin cups, one small spatula of vanadium pentoxide catalyst was added, and the sample was combusted in a stream of oxygen at 900°C. The reaction gases were passed through a reduction chamber to reduce nitrogen oxides to nitrogen; they were then separated by the GC before detection by the TCD.

All measurements were calibrated to a sulfanilamide standard (N = 16.27 wt%, C = 41.84 wt%, and S = 18.62 wt%) that was run every 10 samples. Analyses were only continued if standard data varied by <1% from these values for N and C. Based on the drift of N, C, and S in the sulfanilamide standard in typical batches, S content varies more than that of N and C. Analyses were only continued if standard data for S varied by <10%.

Typical precision was assessed using 10 replicate analyses of a rock standard from Weatherford Laboratories (99986; PWDR5) (Table T9) having a nominal total organic carbon (TOC) content of 3.11 wt% and an IC content of 0.43 ± 0.02 wt% based on coulometry (TC = 3.54 wt%). These replicate analyses show standard deviations of 0.032, 0.046, and 0.032 wt% for N, C, and S, respectively. The coefficients of variation are 0.055, 0.013, and 0.020 wt% for N, C, and S, respectively.

TOC_DIFF content was calculated as the difference between TC and IC from coulometry:

Organic matter characterization

The type and quantity of organic matter in sediments were evaluated by pyrolysis assay using the SRA (Weatherford Laboratories). Between 60 and 150 mg of freeze-dried, ground sediment was weighed into SRA crucibles. Volatile hydrocarbon (HC) content was released when the sample was heated at 340°C for 3 min as the S₁ peak (mg HC/g rock). Hydrocarbons were released during the pyrolysis of kerogen as the temperature was increased from 340° to 640°C at 25°C/min as the S₂ peak (mg HC/g rock). The nominal temperature of the maximum rate of hydrocarbon yield during S₂ analysis is T_max. CO₂ (as mg C/g rock) released during pyrolysis between 340° and 390°C is the S₃ peak. CO₂ (as mg C/g rock) produced by oxidizing the pyrolysis residue at 580°C is the S₄ peak, but this is not directly reported. TOC_SRA was calculated from S₁, S₂, and S₄, assuming that S₁ and S₂ are 83% carbon:

The carbon-normalized hydrogen index (HI) (mg HC/g C) and the oxygen index (OI) (mg CO₂/g C) were calculated from pyrolysis values:

and

All measurements were preceded by a blank and then calibrated to a rock standard from Weatherford Laboratories (99986; PWDR5); the same standard was used for quality control (QC) every 10 samples. Analysis was only continued if QC data fell within the permitted range for this standard, as defined by Weatherford Laboratories (Table T10). Typical precision was assessed using eight replicate analyses of the rock standard. Coefficients of variation for this data set fell between 0.005 (T_max) and 0.1 (OI) (Table T10). In practice, S₃ and OI were occasionally more variable than permitted for the 99986 standard when used as a QC standard in batches of samples; the range of error was as high as ±50%.

Total organic carbon measurement

TOC was measured using two completely independent methods during Expedition 317. In the first method, TOC_SRA was derived directly from the SRA as the sum of pyrolysis carbon (0.83 × [S₁ + S₂]/10) and residual carbon (S₄/10). The S₄ parameter is the oxidizable (at 580°C) residual carbon remaining after pyrolysis. This technique has the advantage of being derived directly during an analytical run on one instrument.

In the second method, TOC_DIFF was derived from the difference between TC (measured on the elemental analyzer) and IC (measured by coulometry). This technique has the advantage of being derived partly from the traditional elemental analysis approach but the disadvantage of relying on two separate instruments, because carbon associated with carbonate content must be subtracted from TC.

To provide constraints on the difference between these methods and advise IODP of the best approach for shipboard TOC measurements, a small batch of 10 samples from Hole U1351A was selected for offline decarbonation. This was achieved by reacting the rock powder with gently warmed 2M HCl in a beaker to remove the carbonates, after which the samples were washed, filtered, freeze-dried, and recrushed. The decarbonated fractions were then analyzed by both the elemental analyzer and the SRA (Table T11).

The 10 selected samples have variable carbonate contents (2.4–31.4 wt%) and thus variable original IC contents (0.29–3.76 wt%; Fig. F11). Like the main data set, the subset has considerably higher TOC_SRA than TOC_DIFF (Fig. F11A). TOC measured on the decarbonated samples (TOC_{decarbonated TC}) does not equal TOC_DIFF but is variable, expressed as a difference of –1.0 to 0.1 (Table T11). This difference does not correlate with the original IC contents (Fig. F11B), meaning that the variability is not related to carbonate content. Furthermore, no correlation exists between carbonate content and the original difference between TOC_DIFF and TOC_SRA. A good correlation is present between TOC_SRA and TOC_{decarbonated TC} (Fig. F11C), having almost the same offset as TOC_DIFF versus TOC_SRA. TOC_{decarbonated TC} is more closely correlated with the measurement of TOC by the SRA on decarbonated samples (TOC_{SRA [decarbonated]}), but an offset remains whereby the SRA gives higher TOC values than the elemental analyzer (Fig. F11D).

TOC_{SRA (decarbonated)} is considerably reduced (20%–70%) compared to original TOC_SRA (Table T11). Removing carbonate should lead to an increase in TOC in the residue, not a decrease, so bulk removal of carbonate using the technique described above clearly alters and removes a portion of organic matter. This means that the technique commonly used to measure TOC (bulk offline removal of carbonates by acid digestion followed by TC measurement) may be invalid. It may be better to remove carbonates using acid treatment in the tin cups to avoid the partial removal of organic matter.

In conclusion, the different TOC measurements reflect inherent differences in the way the two instruments (CHNS elemental analyzer and SRA) measure organic carbon content. The temperature of combustion is higher for the elemental analyzer (900°C) than for the SRA (580°C), and the two instruments are calibrated in quite different ways. The difference does not relate to carbonate content. We recommend that bulk carbonate removal not take place prior to TOC measurement. In order to determine the correct TOC profile for Site U1351 (Fig. F11C–F11D), low TOC% standards must be measured. At the time of measurement of Expedition 317 samples, the only TOC standard available in the laboratory was the rock standard from Weatherford Laboratories (99986; PWDR5), which has a TOC of 3.11%.

Inorganic geochemistry: interstitial water analyses

Sampling

Interstitial water was extracted from 10–15 cm long whole-round samples (labeled red/orange for interstitial water in Fig. F10). Whole-round interstitial water samples were collected at a rate of 3–6 per core where recovery permitted in the shallow (~25–85 m deep) hole at each site dedicated to geochemistry, microbiology, and geotechnical whole-round sampling. The depth of each dedicated hole was intended to cover sediments from the seafloor to the sulfate/methane interface and an approximately equal distance below. In the main deep hole at each site, one whole-round interstitial water sample was taken per core until geochemical profiles stabilized. Thereafter, whole-round samples were collected once every other core until total depth was reached or until the cores became too lithified (Site U1352). These whole-round samples were cut on the catwalk, capped, and taken to the laboratory for immediate processing. This high-resolution sampling technique enabled the creation of high-resolution chemical profiles for the interstitial waters at each site.

In the dedicated hole, one 30 cm long whole-round sample was taken per core in order to provide sufficient interstitial water for shore-based analysis (LIPP4; Fig. F10). In addition, a separate 10 cm whole-round sample (code ISHI) was taken from the same sections (Fig. F10) to meet a shore-based sample request for more interstitial water.

When too many samples needed to be processed immediately during high-resolution sampling, capped whole-round core sections were stored in the cold room until they were squeezed (no longer than 12 h after core retrieval). Gloves were used during sample processing. After extrusion from the core liner, the surface of each whole-round sample was carefully scraped with a clean spatula to remove potential contamination from seawater and sediment smearing in the borehole. For APC cores, 1 cm was removed from the outer diameter and from the top and bottom faces of the samples. For XCB cores, where borehole contamination is higher, as much as 30% of the sediment was removed from each whole-round sample.

The remaining sediment (~50–300 cm³) was placed in a titanium squeezer modified after the stainless steel squeezer of Manheim and Sayles (1974). In most cases, gauge pressures as high as 20 MPa were applied using a laboratory hydraulic press to extract interstitial water. Interstitial water was passed through a prewashed Whatman Number 1 filter fitted above a titanium screen, filtered through a 0.45 µM polysulfone disposable filter (Whatman Puradisc PES), and subsequently extruded into a prewashed (10% HCl) 50 mL plastic syringe attached to the bottom of the squeezer assembly. In most cases, 20–40 cm³ of pore water was collected from each sample, which required squeezing the sediment for 20–40 min. Interstitial water subsamples collected from the syringe were immediately analyzed for salinity, pH, alkalinity, and sulfate. The remaining interstitial water was divided into aliquots and stored in several vials in a freezer for other shipboard and shore-based analyses. Any filter cake that remained after whole rounds were squeezed for interstitial water was divided into subsamples for shore-based analyses.

Salinity, pH, and alkalinity analyses

Interstitial water analyses followed the procedures outlined by Gieskes et al. (1991), Murray et al. (2000), and user manuals for new shipboard instrumentation, with modifications as indicated. Interstitial water was routinely analyzed for salinity with a Reichert temperature-compensated manual refractometer, previously calibrated using the International Association for the Physical Sciences of the Oceans (IAPSO) seawater standard. Alkalinity and pH were measured immediately after squeezing by Gran titration with a Metrohm autotitrator. The IAPSO seawater standard was used for the standardization of alkalinity. Variation for alkalinity was 2%–3% based on the IAPSO standard run after every fifth sample.

Ion chromatograph anion and cation analyses

Sulfate, chloride, magnesium, calcium, sodium, and potassium concentrations in interstitial water were determined with a Dionex ICS-3000 ion chromatograph on 1:200 diluted aliquots in 18 MΩ water. The IAPSO seawater standard was used to standardize measurements made on the ion chromatograph by running it after every fifth sample. The coefficient of variation based on seven repeat analyses of an interstitial water sample for anions and cations measured on the ion chromatograph was 0.002% (Table T12). Any batches of samples with >2% drift for the IAPSO standard were rerun. Acceptable 2% limits for the IAPSO seawater standard are given in Table T13.

Spectrophotometry analyses

Phosphate, ammonium, and silica concentrations in interstitial water were determined using an OI analytical discrete analyzer (DA3500) spectrophotometer unit, which is an automated system that controls sample analysis and reagent aspiration, dispensing, heating, and mixing.

In the phosphate method, orthophosphate reacts with Mo(VI) and Sb(III) in an acidic solution to form an antimony-phosphomolybdate complex. Ascorbic acid reduces this complex to form a blue color, measured at 880 nm. Potassium phosphate monobasic (KH₂PO₄) was used to produce a calibration curve and as an internal standard.

In the ammonium method, phenol undergoes diazotization, and the subsequent diazo compound is oxidized by sodium hypochlorite to yield a blue color, measured spectrophotometrically at 640 nm. Ammonium chloride (NH₄Cl) was used to produce a calibration curve and as an internal standard.

In the silica method, silica in solution as silicic acid or silicate is reacted with a molybdate reagent in acid media to form the β-molybdosilicic acid. The complex is reduced by ascorbic acid to form molybdenum blue, measured at 420 nm. Synthetic seawater containing sodium silicofluoride (Na₂SiF₆) was used to produce a calibration curve and as an internal standard.

The reproducibility for phosphate and silica concentrations in interstitial water is ~5% and that for ammonium is ~20%.

Minor element analyses by inductively coupled plasma–atomic emission spectroscopy

Minor elements in interstitial water were determined by inductively coupled plasma–atomic emission spectroscopy (ICP-AES) with a Teledyne Prodigy high-dispersion ICP-AES. Minor elements (Mn, Fe, B, Sr, Ba, Si, and Li) were analyzed as described by Murray et al. (2000) by preparing calibration standards in an acidified (2% HNO₃, by volume) sodium chloride matrix (25 g NaCl/L). Samples and standards were diluted 1:10 using the 2% HNO₃. To control for instrumental drift and improve precision, an internal standard (10 ppm yttrium) was added to the solution before digestion by 2% HNO₃. Drift correction was made when necessary using the factor from a drift monitor solution (middle value standard solution), which was analyzed after every seventh sample. The coefficient of variation based on duplicate samples is typically <5% (Table T14).

Microbiology

The Canterbury Basin is a promising place in which to expand our knowledge of the deep biosphere because it is in a complex setting representing the history of life under a variety of environmental constraints. The Canterbury Basin is heavily influenced by the input of terrestrial organic matter and is therefore an excellent end-member environment to complement the marine settings that have been studied during previous drilling expeditions. The microbiology program during Expedition 317 sought to determine the following:

1. Depth profiles of total prokaryotic and viral cell counts (SYBR Green cell counting) and intact polar lipid (IPL) concentrations to quantify living biomass;

2. The identities and distribution of microbial and unicellular eukaryotic groups present at each site using 16S/18S ribosomal deoxyribonucleic acid (rDNA)/ribosomal ribonucleic acid (rRNA) (small subunit ribosomal RNA/DNA) clone libraries or tags 454 pyrosequencing (or alternatively, using catalyzed reporter deposition–fluorescence in situ hybridization [CARD-FISH] with probes specific to domain, phylum, and family regions), quantitative polymerase chain reaction (Q-PCR), and IPL chemotaxonomy; and

3. The activity of dominant groups, functional gene clone libraries, enrichment-isolations of strains under diverse nutritional (with diverse carbon and energy sources) and physiological (notably under high pressure) conditions, stable isotope probing (SIP), determination of main carbon turnover processes by δ¹³C analysis of IPLs and correlation to major carbon pools, and analysis of changes in dissolved organic matter pore water constituents by Fourier transform–ion cyclotron resonance–mass spectrometry during incubation with various substrates.

Core handling and sampling

To ensure the preservation of microbial communities, a special sampling strategy was required. This sample strategy was well described for ODP Leg 201 (Shipboard Scientific Party, 2003a).

Generally, when a core arrived on the core deck it was immediately sectioned into 1.5 m sections (Fig. F10). In Sections 1, 3, and 5 of every core from the hole dedicated for whole-round sampling and in Section 3 of every 5–6 cores from the main hole(s), two neighboring whole-round (code WRND) core samples, each 10 cm long, were sliced using sterilized spatulas for microbial analysis: one for IPL analysis (code LIPP5) and one for microbial characterization (code ALA) (Fig. F10). The LIPP5 whole round was capped without further treatment at both ends, sealed in a plastic bag, labeled, and immediately frozen in a –80°C freezer until shipment on dry ice to the shore-based laboratory. The ALA whole round was capped at both ends using sterilized end-caps and transferred to the cold chamber (set to <8°C) for further subsampling. The end of the section adjacent to the ALA whole round (top of Sections 2, 4, and 6 of cores from the dedicated hole; top of Section 4 of cores from the main hole[s]) (Fig. F10) was sampled for perfluorocarbon contamination checks. In the hole dedicated to whole-round sampling, one additional whole-round sample (LIPP6) was taken per core (Fig. F10) under sterile conditions and transferred into an anaerobic chamber in the cold room (<8°C) for subsampling. After the seawater-contaminated outer layer of sediment was removed, 30 cm³ of the LIPP6 sample was transferred to a sterile 30 mL plastic tube (Sarstedt, Germany) and immediately frozen at –80°C for determination of the microbial community composition; the remaining sample was transferred into a sterile 1 L Schott bottle sealed with a butyl stopper for storage at 4°C under a nitrogen atmosphere until shipment at 4°C (incubation sample). The standard ODP/IODP bags previously used for transporting samples under a nitrogen atmosphere at 4°C were not used because they were found to leak (Lipp et al., 2010).

The ALA whole-round samples for microbiological characterization were subsampled under sterile conditions in the anaerobic chamber in the cold room (<8°C) as follows:

Samples for molecular analysis (100–150 cm³; DNA extraction) were taken from the inner part of the ALA whole-round cores with a sterile 30 mL cut-off syringe, transferred into three 50 mL polypropylene tubes (Sarstedt, Germany), and directly frozen at –80°C for onshore analysis. In order to limit RNA degradation, five 2 cm³ samples for RNA extraction were taken as previously described and immediately frozen at –80°C.

Samples for CARD-FISH analysis (1 cm³) were taken with a sterile 2 mL cut-off syringe as previously described by Pernthaler et al. (2001) and stored in a –20°C freezer until shipment. The cell walls of the microorganisms have to be permeabilized to the point that the oligonucleotide probes pass through the cell walls and the cellular ribosomal RNA content is preserved. This was done by fixing cells in buffered formaldehyde solution (1 cm³ of sediment fixed in 9 mL of sterile 3% formalin/3% NaCl solution at room temperature for ~4 h), followed by two washing steps with phosphate buffered saline (PBS) 1× and the addition of 4 mL of a PBS:ethanol (1:1) solution. Samples were then frozen at –20°C until processing in onshore laboratories.

Samples for cultures (6–10 cm³) were taken from the inner part of the core with a 5 mL cut-off sterile syringe and placed into 100 mL Schott vials before storage at 4°C. The samples were flushed with nitrogen for several minutes to eliminate any biogas from the anaerobic chamber that would facilitate the growth of methanogenic or acetogenic microorganisms.

Samples for cell counts (1 cm³) were taken with a sterile 2 mL cut-off syringe and placed in 15 mL polypropylene tubes (Sarstedt, Germany) containing 9 mL of 3% formalin/3% NaCl solution and stored at 4°C until analysis. This slurry represented a 1:10 dilution of the original sample.

Additional samples (1 cm³) for monitoring the infiltration of the 0.5 µm fluorescent microsphere beads were taken on the ALA whole rounds with a sterile 2 mL cut-off syringe from the outer and inner parts of the core and placed in 15 mL polypropylene tubes (Sarstedt, Germany) containing 9 mL of 3% NaCl solution and stored at 4°C until bead counting (Fig. F10).

Rock sampling

Sometimes the bottoms of the deepest holes were characterized by hard or cemented intervals that required a different sampling strategy. After core retrieval on the catwalk, LIPP5 and ALA whole-round rock samples were immediately taken into the cold room. The LIPP5 whole round sample was directly frozen at –80°C, and the ALA whole round was subsampled under a laminar flow fume hood. The rock samples were carefully cleaned under ultraviolet (UV) light by scraping off the outside sediment layers using sterile spatulas before being washed with a sterile 3% NaCl solution. Afterward, the cleaned samples were exposed to UV radiation for 15 min on sterile aluminum foil. The scraping of the outside surfaces, combined with washing and exposure to UV light, removed surface contamination. Rocks were then crushed into several pieces by wrapping them in sterile aluminum foil and breaking them into pieces with several forceful hammer strokes. The rock samples were subsequently subdivided for DNA/RNA extractions, cell counts, CARD-FISH, and cultures, as described above. Rock samples for contamination testing using both chemical and particulate tracers were taken from the outer and inner parts of the whole round. This sampling strategy did not allow preservation of the samples in an anaerobic atmosphere. Consequently, the samples used for enrichment cultures were immediately gassed with pure sterile nitrogen for ~30 min. All materials used for sampling (spatulas, tweezers, hammer, etc.) were sterilized for 45 min in an autoclave at 135°C.

Contamination tests

The amount of sample contamination during drilling and handling was evaluated by running two types of tests as previously described for ODP Legs 185 (Smith et al., 2000) and 201 (Shipboard Scientific Party, 2003a), and for IODP Expedition 301 (Expedition 301 Scientists, 2005).

Perfluorocarbon tracers

Perfluorocarbon tracer (PFT; perfluoromethylcyclohexane, C₆F₁₁CF₃; Oakwood Products, 003295) has a molecular weight of 350.06 g/mol and a density of 1.78 g/mL. Its solubility is ~2 mg/L in water and 104 mg/L in methanol. During drilling, the rate of the tracer injection was adjusted to maintain a final concentration of ~1 mg/L in the drilling fluid using shipboard rig instrumentation software to control pumping rates. Using sterile cut-off 3 mL syringes, ~3 cm³ subcores were taken from the center of the sediment cores and from the periphery of the sediment cores at the core liner. In this way, parallel data sets were collected to determine the extent of contamination at the periphery of the sediment core, along the core liner, and in the center of the core (Smith et al., 2000). PFT concentration was analyzed using the GC (HP6890). The column used on the GC was an HP-PLOT/AL₂O₃ (15 m length × 0.25 mm ID × 5 µm film thickness). The inlet temperature was 170°C, and the inlet pressure was 16.23 psi. The detector temperature was 275°C. The column oven temperature was held at 100°C for 3.5 min and then ramped at 50°C/min to 200°C. At this time, 500 µL of headspace was injected, resulting in a PFT peak eluting at ~8.5–9.2 min. For GC calibration, standard solutions were made by diluting the perfluoromethylcyclohexane in methanol to 10^–3, 10^–5, 10^–7, 10^–9, and 10^–11. Attempts were made to construct a standard curve by plotting the peak area versus the amount of perfluoromethylcyclohexane injected.

Particulate tracer: fluorescent microspheres

A plastic bag containing a suspension of submicron-sized fluorescent microspheres (2 × 10¹¹ microspheres/20 mL bags; Carboxylate YG 0.50 µm microspheres; Polyscience, USA) was introduced into the core catchers of all core barrels from which microbiological samples would be taken (Smith et al., 2000). During drilling, the beads were released inside the core barrel as the sediment entered and were dispersed onto the outer surface of the core. When the core was retrieved, 1 cm³ samples for particulate tracer analysis were taken at the periphery of the core, adjacent to the outer PFT samples, and placed in 15 mL polypropylene tubes (Sarstedt, Germany) containing 9 mL of sterile 3% NaCl solution before storage at 4°C. The infiltration of fluorescent microspheres into the inner part of the core was checked on the samples taken near the samples taken for cell counts. The total number of microspheres was determined by epifluorescence microscopy using a Zeiss Axioplan 2 imaging microscope (Germany) equipped with a blue filter (17 FITC, 460/402 nm) at 1000× magnification.

Cell counts

IODP provides an outstanding opportunity to explore the deep biosphere. Microbiological data collected from 25 ODP and IODP expeditions worldwide mostly comprise total cell counts performed by cell staining with 4′,6-diamidino-2-phenylindole (DAPI), acridine orange (AO; Daley and Hobbie, 1975), and SYBR Green (Noble and Fuhrman, 1998). However, the enumeration of total cell numbers in marine sediment samples still represents a major challenge because the nonspecific binding of fluorescent dye and/or autofluorescence from sediment particles strongly hampers the recognition of cell-derived signals. Lately, new cell extraction procedures have been developed (Lunau et al., 2005; Kallmeyer et al., 2008), but these can be time consuming on board ship.

Recently, Morono et al. (2009) developed a highly efficient and discriminating detection and enumeration technique for microbial cells in sediments using dilute (1%) hydrofluoric acid (HF). This weak acid reduces nonbiological fluorescent signals, such as that from amorphous silica, and enhances the efficiency of cell detachment from particles.

Procedure

Under an aspirant hood, 50 µL of sediment slurry (described above) was mixed with 450 µL of an HF solution (1.0% [wt/v] HF and 3% [wt/v] NaCl) in a plastic test tube and then incubated for 20 min at room temperature. The HF reaction was stopped by adding 2 mL of stop solution (1M tris-hydrochloric acid [HCl; pH 8.0], 0.125M CaCl₂, and 25% methanol). Subsequently, the mixture was sonicated at 20 W for 5 min using an ultrasonic bath (Fisher, FS14H).

A volume of 50–750 µL of sonicate was mixed with 2.5 mL of 3% NaCl and directly filtered through a 0.22 mm pore size black polycarbonate membrane without centrifugation. In order to eliminate potential carbonate particles and/or precipitates, the membrane was treated with 1 mL of 0.1M HCl for 5 min on the filtration device. The membrane was then washed with 5 mL of TE buffer (10 mM tris-HCl and 1.0 mM ethylenediaminetetraacetic acid [EDTA]; pH 8.0) and air dried.

A volume of 80 µL of SYBR Green (25×) was added to the filtered cells, and the sample was incubated at room temperature for 15 min, followed by air drying. Subsequently, the filter was mounted on the slide with 40–50 µL of a mounting solution containing an antifade agent ([50% glycerol, 50% PBS 1×, 0.05M Na₂HPO₄, and 0.85% NaCl; pH 7.5] and 0.1% p-phenylenediamine [in a 1:2 mixture] made daily from a 10% aqueous stock). The cells were counted using epifluorescence microscopy with a blue filter set as described above. Two hundred FOVs were usually counted per sample.

Enrichment cultures

The deep marine subsurface biosphere is a natural habitat characterized by complex conditions that are often difficult to reproduce in the laboratory. In addition, the mean generation times of the populations have been estimated to be very long. This explains why there are only a few reports of successful cultivation and isolation in pure cultures (D'Hondt et al., 2002; Batzke et al., 2007). However, cultivation is the easiest way to access a microorganism's physiology, and we have a lot to learn from microbial isolates originating from the deep biosphere. For this reason, during this expedition we enriched various types of microorganisms likely to grow in this environment. Several enrichment media were prepared on board, based on an artificial salt solution detailed in Table T15. Different substrates were added according to the requirements of the physiological type of the targeted prokaryotes. The enrichment cultures were 1% inoculated from 25% sediment slurries prepared by adding a sterile 3% (w/v) NaCl solution to the samples collected for culturing. The enrichments were prepared under a laminar flow hood located in the cold room and flushed with pure N₂. The growth of cells was observed by microscopy, and successful enrichments were shipped at 4°C.

We targeted different groups of prokaryotes: methanogens, acetogens, sulfate-reducers, and fermenters. We used several substrates and incubated the cultures at near the in situ temperature (Table T16). The turbidity of the culture, together with photonic and fluorescence microscopy, were used to detect positive growth within the cultures. In addition, methanogens were recognized by the autofluorescence of their F420 cofactor when exposed to UV light.

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