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Starting in the 1990s, microbiologists participating in ODP and IODP expeditions have documented the presence of microbial life in deeply buried sediment and the basaltic basement (Fisk et al., 1998; Parkes et al., 1994). Active microbial life has been detected as deep as 1626 mbsf (Roussel et al., 2008), and the introduction of molecular biology into marine ecology has led to great advances in our understanding of microbial life below the seafloor (Biddle et al., 2008; Cowen et al., 2003; Inagaki et al., 2006; Mason et al., 2010). Much of the microbiology performed during ODP and IODP expeditions has concentrated on sediment (e.g., ODP Leg 201 to the Peru margin), with the notable exception of expeditions to the Juan de Fuca Ridge (Cowen et al., 2003; Lever et al., 2006) and Atlantis Massif (Mason et al., 2010). Sampling for microbiological studies during Expedition 330 focused primarily on the microbiology of basement rocks. Differences in microbial population between the overlying sediment and volcaniclastic layers and the basaltic basement were also targeted. Drilling during Expedition 330 provided a unique sample set because at several sites there is a thin layer of sediment overlying volcaniclastic breccia, followed by igneous basement. As much as 505 m of basement was drilled at Site U1374, providing one of the most complete sets of igneous basement samples to date for microbiological investigation

Sediment sampling

Whole-round core samples are required for microbiological studies to avoid contamination introduced by sample handling before and during the core splitting process. For sediment samples, whole rounds were sampled on the catwalk. After the core liner was cut, whole-round sediment samples were cut off with a sterile spatula, transferred onto a precombusted (450°C for 2 h) aluminum foil sheet, and immediately transported to the cold room in the microbiology laboratory, where they were processed.

Sediment samples were collected for molecular biology and cell counts. One of the goals of this sampling was to trace the change, if any, in microbial community composition across the sediment/basement transition. The sampling was carried out on unconsolidated sediment recovered from the uppermost ~10–20 m at each drill site where such sediment was recovered.

One 5 cm whole round of sediment was collected from each core in the 5 cm interval immediately adjacent to the core catcher. Once the samples were transported to the microbiology cold room, a sterile syringe with the top cut off (creating a cookie cutter type of coring device) was used to collect 2 cm3 for cell counts, 2 cm3 for carbon-hydrogen-nitrogen-sulfur and total organic carbon analyses (see “Geochemistry”), and ~20 cm3 for molecular biological work. For cell counts, 2 cm3 of sediment was added to 8 mL of 2% formalin in 3% NaCl (sterile filtered) in a 15 mL Falcon tube.

Molecular biology samples were collected in sterile 5 mL centrifuge tubes and frozen at –80°C for shore-based analysis. After microbiology sampling, an extruded polystyrene foam plug was placed in the void left in the center of the whole round, preventing the collapse of its outer rim (which was likely contaminated by drilling fluids and therefore not used for microbiological analysis) and allowing it to be returned to the curator for further sampling by other laboratory groups.

Igneous rock sampling

For volcaniclastic breccia and basement samples, whole rounds were selected in the core splitting room and collected from the core liner onto precombusted (450°C for 2 h) aluminum foil before the core was cleaned with water. Samples were chosen with a petrologist present to ensure that no samples critical to the main objectives of the cruise were lost because of microbiological sampling, which destroys nearly the entire sample. Before samples were transported to the microbiology laboratory, photographs of each side of the whole-round sample were taken while the sample was on the combusted foil. During sampling and photography, samples were handled with gloved hands only.

Prior work has shown that the interiors of rock cores are generally free from contamination (Lever et al., 2006). Therefore, efforts were taken to sample only the core interiors. Sections that showed some sign of alteration or fluid flow conduits were specifically chosen because these are likely locations for microbial life. Microbiology samples on average were 5–10 cm long. All samples were divided into subsamples for cell counts, molecular biological analyses (DNA and RNA extraction, to be carried out on shore), and in situ stable isotope analysis (δ34S and δ13C). For some samples, stable isotope addition bioassays and cultivation experiments were also performed.

The undisturbed whole round was washed three times with artificial seawater in a fresh, resealable storage bag before subsampling to avoid contamination from drilling fluids. Next, the rock was split with a sterile chisel and sampled from the interior. For those samples on which culturing or stable isotope bioassays were to be performed on board, a portion of the sample was sectioned off immediately, and placed in a N2 environment in a glove bag or the anaerobic chamber to minimize exposure of anaerobic cells to O2 while the rock was being processed. These precautions were taken because basement rocks at Sites 801, 1272, and 1274 have been shown to be anaerobic based on detection of sulfate reduction, a strictly anaerobic process (Alt et al., 2007; Rouxel et al., 2008). Therefore, we assumed that the basement rocks here were also anaerobic. Whenever pieces of the whole round remained, including the outer portions of the rock, they were returned to the curator to be placed back into the working and archive sections of the core.

Molecular biology

Samples were placed in 5 mL autoclaved centrifuge tubes and immediately stored at –80°C for DNA analysis or were placed in 15 or 50 mL centrifuge tubes with LifeGuard Soil Preservation Solution (MoBio Laboratories, Inc., Carlsbad, CA) and held at 4°C overnight before freezing at –80°C for RNA analysis. LifeGuard protects the viability of microbial cells while keeping them in stasis, allowing for efficient DNA and RNA extractions from samples collected in the field. These samples will be analyzed during postexpedition research.

Stable isotope analysis

Analysis of in situ δ34S fractionation in the rocks can indicate the presence of microbially mediated sulfate reduction (Alt et al., 2007; Rouxel et al., 2008), even in basalts older than 50 Ma. Analysis of in situ δ13C fractionation provides insight into the production mechanism of carbon present in the rocks (Mason et al., 2010). Samples for stable isotope analysis were subsamples of the whole round sampled for molecular biology. At least 20 cm3 of rock chunks were placed in Whirl-Pak bags and set in the anaerobic chamber in the cold room for a few hours, after which the bags were sealed in the chamber (so that the atmosphere of the closed bag remained anaerobic) and stored at 4°C. Isotopic analysis will be carried out as part of postexpedition research. Alteration rims were specifically targeted for this work because they are more likely to harbor microbes (Fisk et al., 1998).

Cultivation experiments

Based on prior work with both subsurface and surface-exposed basalts, the functional groups of microbes likely to be found in the subsurface basalts along the Louisville Seamount Trail include sulfur oxidizers, sulfate reducers, methanogens, iron reducers, and iron oxidizers (Cowen et al., 2003; Mason et al., 2009, 2010; Santelli et al., 2009). Media targeting enrichment of these groups were prepared prior to the expedition and used to culture subsurface microbes. Details of the media recipes can be found in Table T14. Samples used for culturing were maintained in an anaerobic environment following collection and splitting of whole rounds into rock fragments for culturing and molecular biology sampling. Rock fragments were placed in 5 mL serum vials filled with different media and sealed. The vials were kept at 4°C (the assumed temperature of the volcanically inactive subsurface seamount environment) until the end of the expedition and then were shipped in refrigerated containers for onshore assessment of growth. If any enrichments prove successful, further steps will be taken to isolate and characterize pure cultures of microorganisms during postexpedition research.

Stable isotope bioassays

13C-labeled bicarbonate was used to measure incorporation rates of inorganic carbon into subsurface microbes as an indicator of autotrophic production in this environment. In addition, 13C-labeled glucose and acetate were used to measure incorporation rates of organic carbon by heterotrophic microbes, 15N-labeled ammonia and nitrate were used to estimate rates of N uptake by subsurface microbes, and 34S elemental sulfur was used to measure rates of sulfur cycling.

In each experiment, 100 mL of artificial seawater (Sigma-Aldrich; St. Louis, Missouri [USA]) modified with 50 µM sodium nitrate, 3 µM potassium phosphate, and 0.5 µM ammonium chloride was added to a precombusted serum vial. We herein refer to this as the basic seawater medium. Sigma sterile seawater is collected from surface water in the Gulf Stream, which is low in N and P. However, deep seawater has N and P concentrations closer to those of the modified Sigma sterile seawater. Note that the composition of pore water in the subsurface rocks collected during Expedition 330 is unknown, so we chose to simulate typical deep seawater. Enough rock chips were added to cover the bottom of the vial. This amount depended on the amount obtained during sample processing.

A more persuasive “enhanced extraction” method for obtaining large amounts of uncontaminated rock samples for these bioassays was developed during sample processing of Sample 330-U1374A-15R-2, 79–89 cm. For all other (nonstable isotope addition bioassay) microbiology sampling during Expedition 330, subsamples were retrieved only from the centermost portion of the whole-round sample to avoid potential contamination from drilling fluids on the outside of the whole-round sample. This method provided enough uncontaminated sample suitable for cell counts, culturing experiments, and molecular biology analysis. However, analysis of stable isotope addition bioassays requires more rock than can be collected solely from the center of the whole-round sample. Therefore, the following enhanced extraction method was carried out on all samples for stable isotope analyses from Holes U1374A–U1377A:

  1. The whole-round sample was rinsed three times with sterile filtered seawater in the same manner as all other microbiology samples.

  2. The outside of the rock was flamed with a propane blowtorch by laying the whole round on its side in the rock box, flaming it for 5 s, and then rotating it. All four quadrants of the rock were flamed to ensure the entire outside of the whole round was sterilized. The top and bottom of the whole round were also flamed for 5 s.

  3. An ethanol- and flame-sterilized chisel was used to remove the outer portion of the whole round, and rock chips from the inside of the rock were selected and preserved for cell counts and molecular biological analysis. Sections of the whole round were also preserved for shore-based analysis of in situ 34S and 13C.

  4. The remainder of the rock was broken into rock chips ~2–3 cm in diameter with the sterilized chisel. These chips were then placed in unused Whirl-Pak bags.

  5. The Whirl-Pak bags were placed between two autoclaved polytetrafluoroethylene Delrin plugs inside an autoclaved section of core liner (~20 cm long) and crushed using a Spex 3624B X-Press hydraulic press, with pressure not exceeding 2 tons.

  6. The rocks were removed from the bags and collected in unused 50 mL centrifuge tubes for transportation to the radiation van, where they were used in the stable isotope addition bioassays.

The advantages of the enhanced extraction method are twofold. First, sterilizing the outside of the entire whole round allows samples from all parts of the rock to be collected with minimal fear of contamination, which allows collection of a greater volume of rock than that which could be obtained solely from the interior of the whole-round sample. Although some of the rock chips collected (those pieces containing portions of the outside of the whole round) will contain no live microbes because of the sterilization, the gain of volume versus the loss of cells makes this a valuable sampling method. Second, using the hydraulic press allows a large volume of small rock chips to be collected, which is important because the mouth of the serum vial used for these incubations is only 17 mm wide. Enough rock material was collected using the enhanced extraction method to add 15–30 cm3 per serum vial for bioassays, starting with Site U1374.

Following sampling using the enhanced extraction method, two different sets of treatments were prepared for the stable isotope bioassays: (1) 13C bicarbonate, 15N ammonia, and 34S sulfate, which targeted metabolic rates specifically among autotrophic microbes, and (2) 13C glucose, 13C acetate, 15N nitrate, and 34S, which targeted metabolic rates among heterotrophic microbes. Varying concentrations of each stable isotope were added during the expedition as experimental strategies were adjusted. For the range of concentrations for each label, see Table T15.

After a stable isotope label was added to each vial, the vials were sealed with a rubber stopper and crimped. Incubations were kept at 4°C until termination. At the end of each incubation, 5 mL of fluid was removed from each vial with a syringe and transferred to a crimp-sealed 20 mL serum vial containing 1 mL of 2 g/30 mL NaOH solution (for dissolved inorganic carbon), and the excess fluid was poured into a 50 mL Falcon tube. The remaining rock chips were collected in a separate Falcon tube.

For each stable isotope bioassay, there were four time points (t0t3). The first (t0) was when the rock was collected during routine microbiology sampling, which provided the background stable isotope fractionation value. No manipulation of this rock was performed after sampling. The remaining time points were taken from the incubation vials as described above. The second time point (t1) was taken after 2 weeks (most were processed on the ship), the third time point (t2) was taken after 2 months, and the fourth time point (t3) was taken after 6 months. Both t2 and t3 were sent to shore, and the incubations will be terminated during shore-based research. The long duration of the experiments allows for detection of potentially slow metabolisms of subsurface microbes.

Cell counts

During Expedition 330, we attempted to enumerate cell abundances for the basaltic rock samples to determine biomass concentrations in the subsurface basalt environment. Crushed and powdered basalt (1 cm3) samples were fixed with 4 mL of 4% paraformaldehyde in 3% NaCl at 4°C overnight. The fixed basalt suspensions were then subjected to cell counting according to the following protocol:

  1. After vigorous stirring of the paraformaldehyde-fixed slurry, 100 µL of the suspension was immediately dispensed in 10 mL of filtered 1× phosphate-buffered saline (PBS) buffer (pH 7.4).

  2. The second suspension was placed in an ultrasonic bath for 30 s.

  3. The ultrasonically treated suspension was filtered through a 25 mm diameter, 0.2 µm pore size black polycarbonate filter underlain by 25 mm diameter, 0.45 µm pore size cellulose.

  4. Cells on the polycarbonate filter were stained with filtered 1× SYBR Green I (Invitrogen; Ann Arbor, Michigan [USA]) DNA stain in filtered 1× tris-EDTA (TE) buffer for 5 min.

  5. The stained filter was washed twice with 5 mL of filtered 1× PBS buffer.

  6. The washed filter was mounted on a slide with one drop of immersion oil and covered with a cover glass.

  7. The SYBR-stained cells were directly observed by using a Zeiss Axiophot epifluorescence microscope with a band-path filter slit (excitation at 470 nm; fluorescence > 515 nm) at 1000× magnification (100× objective; 10× eyepiece).

For each cell count, at least 20 microscopic fields were observed, and cell-shaped forms that produced bright green fluorescence were enumerated as cells.

The minimum detection limit was estimated by counting blank filter samples that were treated with 100 µL of filtered 1× PBS with 4% paraformaldehyde, as described above. Cell numbers in the blank filters were calculated as the average of all blanks processed during the expedition. The minimum detection limit was set to be the blank value plus three times the standard deviation. For completely negative control samples, crushed basalt pieces were combusted at 500°C for 3 h and then counted using the same method.

Unfortunately, shipboard cell counts on rock samples proved impossible because of a combination of fluorescence from the rocks themselves and focusing difficulties caused by ship movement. These two factors made it extremely difficult to distinguish between microbial cells and rock particles. We will endeavor to develop successful counting methods for subsurface rock samples during postexpedition research.

Quantitative polymerase chain reaction (qPCR), a modification of classic PCR, is essentially a fluorogenic assay used to quantify the number of target genes, and hence cells, in a given environmental sample. During postexpedition research, qPCR will be applied to the samples collected. Then the two biomass estimation methods (cell counts and qPCR) will be compared (Einen et al., 2008; Santelli et al., 2008).

Contamination testing of drilling fluids

As part of the drilling process, a huge amount of surface seawater is injected into each borehole and is the major source of contamination with microorganisms in the cores collected. As a check for contamination, the microbial composition of the drilling fluid (surface ocean water) was assessed, and organisms present in both the drilling fluid and rock samples will be considered a sign of contamination. These organisms will be presented as contaminants, not subsurface residents. If there are any samples in which all of the organisms detected were also detected in the drill fluid, these samples will not be included in further analyses. Although it is possible that some species may be present both in surface seawater and the subsurface biosphere, we feel it will be more informative and conclusive to focus on microbes known to be present in subsurface samples only. Water samples were collected directly from the injection pipe in sterile bottles with screw caps and were handled using sterile equipment. Microorganisms present in the drilling fluid were extracted by filtration using a vacuum pump through 0.2 µm pore polycarbonate filters. Filters were frozen in cryotubes at –80°C for shored-based DNA extraction and analysis.

Fluorescent microsphere contamination testing

Bags of yellow-green fluorescent microspheres (Fluoresbrite carboxylate microspheres; Polysciences, Inc., 15700) with a diameter of 0.52 ± 0.01 μm were occasionally used (i.e., for 1–2 cores per site) as a particulate tracer that mimics the movement and dispersal of microbial cells in the drill pipe during coring. The microspheres were then counted in core samples, in drill fluid caught as the drill core arrived on the rig floor, and in each of three sterile seawater rinses of the whole-round samples.

The concentration of microspheres was set at 1010 spheres/mL (Smith et al., 2000), and the microspheres were deployed in Whirl-Pak bags containing 40 mL of microsphere suspension in deionized water (2 × 1011 microspheres in a 40 mL bag). The bag was then heat-sealed, leaving some extra plastic (not filled with beads) at each end. By attaching the loose plastic ends using para-aramid synthetic cord, the bag was wedged into a shim above the core catcher and stretched across the throat of the core barrel. Cores were consequently forced to burst through the bead bag when a core was taken.

The microspheres are highly fluorescent (458 nm excitation; 540 nm emission) and appear bright green when they are observed by epifluorescence microscopy (Smith et al., 2000). Concentrations of fluorescent microspheres in core samples were quantified using a Zeiss Axiophot epifluorescence microscope fitted with a mercury lamp (HBO 100 W), a blue filter set, and a 100 Å Plan-NEOFLUAR oil-immersion objective. Nonfluorescent immersion oil was used for all observations. Aliquots (100 μL) of the crushed rock suspension or sediment slurry were resuspended in 10 mL of filtered 1× PBS solution and filtered onto black, 25 mm diameter polycarbonate filters (0.2 μm pore size) in a filtration tower. The filters were then mounted on microscope slides with a drop of nonfluorescent immersion oil and covered with a coverslip. The microspheres on the filter were then counted using the epifluorescence microscope. Microsphere abundance on the filters was determined by averaging the total number seen in at least 20 randomly selected fields of view.