IODP Proceedings    Volume contents     Search
iodp logo



Microorganisms are the kinetic controls on both methane production and consumption. Their distribution, identities, and activity must, therefore, be estimated to model the dynamics of the gas hydrate system. Target zones of interest include

  • The top of the sediment column, where microbial sulfate reduction and microbial methane oxidation are coupled;
  • The base of the gas hydrate stability zone (GHSZ), which was a region of increased microbial numbers at Blake Ridge (Wellsbury et al., 2000); and
  • Sediments hosting massive and disseminated gas hydrate, which may also host distinct microbial communities.

In addition to investigating the methanogens, the deep cores provide an opportunity to explore extremophiles, particularly piezophiles. These organisms live optimally at high hydrostatic pressure (e.g., Yayanos, 1995). We are interested in cultivating novel piezophilic microorganisms originating from the top 10 cm and bottom of cored holes at high pressure.

Because good microbiology samples can only be taken when the cores are fresh, the shipboard microbiologists focused on obtaining the best samples possible for future investigations. Cores were sampled for microbiology studies at all sites.

Core handling and sampling

Core collection and retrieval

Because of the need to monitor contamination and control core temperature, cores for microbiological sampling were specified prior to core collection. Two types of tracers monitored drilling fluid (seawater) infiltration into the cores, soluble perfluorocarbons and fluorescent microparticles (microspheres). The perfluoro(methylcyclohexane) tracer was pumped into the drilling fluid (surface seawater) by a variable-speed, high-pressure liquid chromatography pump connected to the mud pump system. The tracer pump operated at a fixed rate relative to the mud pump speed so that the concentration of tracer was always 1 ppmv. The tracer pump was run continuously in the continuously cored hole at each site. The fluorescent microspheres were delivered in a plastic bag designed to rupture on contact with the sediment and attached to the core catcher (see "Contamination tests").

Core temperature is an important consideration when taking microbiological samples. Ideally, cores should be maintained as close to in situ temperatures as possible; in practice, cooling cores below in situ temperatures is acceptable, whereas warming cores above in situ temperatures is not. Whereas our goal was to maintain core temperature at or below in situ temperatures at all times, as a practical matter it is difficult to avoid some warming above in situ temperatures. Shallow cores at the northern Cascadia accretionary prism have in situ temperatures below the temperature of surface seawater in this region (12°–14°C), which was used as drilling fluid and through which the cores traveled to reach the rig floor. To keep warming to a minimum, microbiology cores were retrieved and sent to the catwalk using the expedited core-retrieval protocol implemented during Leg 201 (see "Microbiology" in Shipboard Scientific Party, 2003a), except at Site U1329 (see "Microbiological sampling" in the "Site U1329" chapter). When a core barrel was retrieved, the core was immediately removed from the barrel and sent to the catwalk. Once the core was in the hands of the core technicians, the drilling crew sent the next core barrel down the wireline. This core-handling protocol increased the coring time but was necessary to minimize core warming. Temperatures were monitored using the IR camera on freshly cut core ends of each section identified for microbiology (see "Physical properties"). These images provide a temperature profile of the section immediately prior to moving it to the microbiology sampling area in the hold reefer and are available for postcruise assessment of microbiology sample quality. Examples of core-end temperature profiles at different times are provided in Figures F36 and F37 in the "Site U1327" chapter. For these particular examples, the temperature of the center of the core was ~9 and ~10C, respectively, at times when microbiology samples were typically taken to the hold deck reefer.

Core section subsampling

Core sections were cut on the catwalk following ODP/IODP standards as described in ODP Technical Note 36 (ODP Science Services, 2006), but acetone was not used to seal the end caps of sections dedicated for microbiological sampling. Most sections that were sampled for microbiology had IW whole-round (WR) samples removed from the bottom end. The hold deck reefer on the JOIDES Resolution was maintained at 4°C and served as microbiological laboratory space. All sections used for microbiological study were transported directly from the catwalk to the reefer, where they were subsampled and packaged.

Core sections sampled for microbiology were generally intact and undisturbed. WR samples were removed from these sections in the refrigerated reefer by first cutting the liner with the rotary knife cutter and then fracturing, rather than cutting, the sediment. If the sediment was slurrylike in texture and would not fracture easily, an ethanol-rinsed wide spatula was used to cut through the sediment and contain it in the liner. The WR samples were dropped into sterile plastic bags (Nasco or Fisher) and either frozen, refrigerated, or further subsampled. Although this method leaves most samples in contact with contaminated material near the core liner, a decision was made that, in view of the very slow nature of the diffusion process and the low storage temperatures involved, such treatment was preferable to additional processing under adverse shipboard conditions.

WR samples were subsampled with sterile, truncated 5 mL syringes or alcohol-rinsed spatulas. Subsamples were placed in preservative solutions for direct microscopic counts, fluorescence in situ hybridization (FISH), or microsphere enumeration. WR samples that were stored unpreserved at 4°C were sealed in nitrogen-flushed aluminized polyester heat-seal bags (Kapak Corp.) to maintain an anoxic environment. Samples to be frozen were stored at –80°C.

A glove bag (Coy) containing a nitrogen atmosphere with 5% CO2 and 5% H2 was used for anaerobic handling of core samples to inoculate growth media selective for sulfate-reducing microorganisms to be cultured under high pressure. Subsamples were brought into the glove bag in nitrogen-filled bags, kept cold in an ice box with lid, and quickly processed to minimize warming. All media and sterile tools were also precooled and kept on ice during processing. Hydrogen was present to combine with residual oxygen in a reaction to make water, which was catalyzed by palladium pellets maintained within the bag. The water was absorbed by a desiccant, and, thus, the glove bag's atmosphere was scrubbed of any oxygen. The bag was maintained regularly, and several hours before each use it was flushed with a gas mixture and provided with freshly baked (140°C) catalyst. As an additional precaution to minimize oxygen contamination, tools and glassware to be used for manipulation and storage of samples for strict anaerobic work were stored within the glove bag.

Although cores were processed as quickly and carefully as possible, shipboard handling should not be simply accepted as aseptic. We recommend that investigators receiving samples treat them as potentially contaminated and subsample accordingly whenever possible. Microsphere enumerations, summarized in the relevant site chapters, and additional microsphere evaluations performed postcruise at individual laboratories should be used to evaluate the quality of individual samples.

Contamination and sampling of extended core barrel cores

The XCB coring system is typically used when the APC coring system reaches resistance in the sediment below the seafloor. Although this coring method captures deeper sediment, the quality of the cored sediment is compromised by increased mixing of drill fluid and extraneous sediment (drill slurry). Thus, the WR samples may contain sediment, drilling slurry, or biscuits, sometimes regularly spaced, embedded in lower density drilling slurry. Drilling slurry is soft, moldable, and wet and cracks with a "fluffy" texture, whereas the biscuit is firm and drier and cracks with distinct edges. Figure F10A is an example of an XCB core section where it is difficult to distinguish between sediment and drilling slurry. Figure F10B represents an XCB core section that appears to be mostly sediment in the upper part and mostly slurry surrounding small biscuits in the lower part. Figure F10C shows a typical alternating pattern of biscuits and drill slurry.

Biscuits are coherent pieces of sediment and they are typically less contaminated than the surrounding drilling slurry. Five biscuit/slurry microsphere comparisons (Tables T8 in the "Site U1325" chapter, T9 in the "Site U1326" chapter, and T12 in the "Site U1328" chapter) showed the biscuits remained uncontaminated even if the slurry contained microspheres. Unfortunately, biscuits and slurry were not distinguished during subsampling of XCB cores during the early part of Expedition 311. Samples contaminated with drilling slurry may still be suitable for some purposes (e.g., enrichment cultures) but are inappropriate for others (e.g., biomarker analysis for microbial community structure).

Sampling biscuits within XCB cores is not trivial. In the past, microbiologists have split the core liner while keeping the sediment intact and cleaned off biscuits by hand, in a manner similar to that used by the geochemists when preparing IW samples. This method is not completely satisfactory, as it is time-consuming, has the potential for contamination and warming, and allows oxygen to contact the core unless performed inside the glove bag. Nondestructive testing of XCB sections to reveal biscuits would allow quicker, cleaner processing of core material for microbiology. Gamma density, electrical resistivity, and X-ray density all show potential for distinguishing biscuits from slurry without removing the core from the liner; however, some of these measurements may adversely affect sediment microorganisms.

We experimented with a modification of the geochemical paring method with an XCB WR sample (311-U1328C-27X-3, 55–70 cm; 297.0 mbsf) that had already been sampled for shipboard high-pressure culturing experiments. The anaerobic chamber was turned into a temporary cold box by filling it with blue ice packets to keep the sediment cool. The WR sample was extruded onto a sheet of ethanol-sterilized aluminum foil in the glove bag and inspected for the biscuit. Once the biscuit was identified and isolated from the surrounding slurry, biscuit fragments were immediately placed into sterile, anoxic, 50 mL screw-cap tubes and temporarily placed in ice buckets until they could be frozen at –80°C. When the frozen biscuit fragments were ready for sampling deoxyribonucleic acid (DNA) for analysis, the fragment was cracked open and subsampled from the center to obtain an undisturbed sample for microbial DNA. This method is the most reliable way to find the appropriate sediment for phylogenetic studies. Reexamination of XCB gas hydrate WR samples from previous sites should ideally have been done immediately but will be done on shore because of the uncertain condition of the ship's anaerobic glove bag and the difficulty in chilling all the exposed sediment in this glove bag while searching for biscuits.

In XCB cores, it is best to obtain at least 10–15 cm of a WR sample, instead of 5 cm, to increase the chances of obtaining intact biscuits. These WR samples are better suited for "spiraling," a slightly less invasive, quick, and simple technique that can be performed immediately as core sections arrive in the cold room. The 10–15 cm XCB WR sample was secured to the liner clamp in the hold deck reefer and the cutting end was uncapped. Using a small ethanol-sterilized spatula, the sediment at the end was inspected by observation and tactilely (cracking, texture, scrapings, etc.) as it remained in the liner. If it appeared to be drill slurry, then a ~1–2 cm ring of the core liner (containing the slurry) was removed with the core liner cutter. Usually when cutting a relatively thin section of core liner, 1–2 cm will be nicked but cannot be completely removed as a ring. This is because the physical interference between the liner clamp and core liner cutter restricts blade contact. However, slowly unwinding the nicked liner in a spiral motion until the liner breaks off works well. The sediment at the exposed end was inspected again and if the sediment appeared relatively dry, firm, and dense, the WR sample was immediately subsampled for DNA analysis, frozen at –80°C, capped, sealed in a nitrogen-flushed bag, heat-sealed, and stored at 4°C. An XCB WR sample that did not contain any dense, firm, dry sediment at either end was either rejected or saved at the discretion of the microbiologist. Spiraling may help minimize the error of subsampling slurry for DNA analysis and help eliminate WR subsamples of pure slurry. It can result in the loss of culturable sediment unless the removed slurry is captured in a sterile bag flushed with nitrogen. The chance of capturing quality "live" sediment samples can increase under the strict conditions of a chilled, sterilized, anoxic chamber. However, during this expedition, the available hold deck reefer was equipped for spiraling and an intact chilled anaerobic glove bag was equipped for completely dissecting the WR samples.

Shipboard microbiological procedures and protocols

Contamination tests

The greatest challenge for subsurface microbiological investigations is verification that observed populations and activities are in situ and not the result of the drilling and sampling process itself, including introduced microbial contaminants. Chemical (perfluorocarbon) and particulate (latex microsphere) tracers were used during coring to test for the potential intrusion of drilling fluid and assess the suitability of the core material for microbiological research. The presence or absence of these two tracers also acts as a quality assurance check on core-handling methods. These tracer techniques were used during Leg 201 and are described in ODP Technical Note 28 (Smith et al., 2000b).

Perfluorocarbon tracer

Perfluorocarbon tracer (PFT) was continuously fed into the drilling fluid at a concentration close to the limit of solubility (1 g/g) and well above the detection limit for GC analysis of that material (1 pg/g). Samples for PFT analysis were taken from all cores intended for microbiological studies. Syringe subcores were taken from the interior (to monitor intrusion) and exterior (to verify delivery) of a freshly broken core or biscuit surface, extruded into HS vials, and sealed with polytetrafluoroethylene (PTFE) septa. Air samples were occasionally taken to monitor the background level of PFT in the hold deck reefer or on the catwalk. Samples were kept at –80°C until ready for GC analysis. The method described in Smith et al. (2000b) was used for PFT analysis, with minor modifications. The same GC setup was used, but the instrument was modified by installation of a 1 cm3 sample loop and injector valve to standardize injection volumes.

Fluorescent microparticle tracer

Latex fluorescent microspheres (Poly Sciences, Inc.) (YG; 0.5 m diameter) were used as a particulate tracer complementary to the volatile PFT. A 2 mL aliquot of microsphere stock (2.69% solids) was diluted with 40 mL of distilled water, sonicated for 2 min, and heat-sealed into a 0.12 L (4 oz) sterile plastic bag (Nasco). The bag was then attached as described in Smith et al. (2000a, 2000b) to the inside of the core catcher and positioned to rupture upon impact of the core tube with bottom sediments, where the microspheres mix with seawater and coat the outside of the core as it is pushed into the liner. During core processing, subsamples of sediments were collected from outer and inner layers for microscopic examination. Weighed samples were mixed thoroughly with saturated sodium chloride solution to extract microspheres. The slurry was then centrifuged to separate the liquid phase (Marathon 21K; 5 min; 1000 relative centrifugal force), the supernatant was filtered onto a black polycarbonate filter (Millipore; 0.2 m pore size), and the filter was mounted on a clean slide for microscopic examination. Microspheres in slide preparations were counted using a Zeiss Axioplan fluorescence microscope equipped with the Zeiss Number 9 filter set (BP 450–490; LP 520), and the number of spheres observed was used to quantify contamination in spheres per gram of sample. Comparison of microsphere numbers between paired samples from inner and outer core layers provides a relative measure of fluid intrusion. A sample with many spheres in outer layers and few or none within may be considered of "higher quality" than one with very few spheres in the outer layers and few or none within.

Enrichment cultures

Liquid culture enrichments were started for anaerobic and aerobic piezophiles. Samples for anaerobic enrichments were taken as quickly as possible from cores that had been maintained at low temperatures and oxygen-free conditions. Subsamples for enrichment of aerobic microorganisms were separately kept cold in 15 mL sterile tubes. For sediment transfers, subcores were brought into the glove bag in nitrogen-filled sealed bags and insulated on ice; work was performed quickly to minimize warming. All media and sterile tools were also precooled during processing. In the glove bag, anaerobic sulfate-reducing bacteria (SRB) presumed to be in 1 g of sediment were vigorously mixed, serially diluted 10-fold three times, and inoculated in cold Widdel's medium (R.J. Parkes, pers. comm., 2005) amended with acetate, lactate, or formate as an energy source. Aerobic heterotrophic microorganisms were enriched in Zobell/10 medium lacking yeast extract (F. Malfatti, pers. comm., 2005) outside the glove bag. Aerobic Actinomycetes (e.g., Salinospora spp.) was also selected and enriched in A/10 medium (Mincer et al., 2002; P. Jensen, pers. comm., 2005). These cultures (SRBs, heterotrophs, and Actinomycetes) were maintained at 55.1 MPa of pressurized water in an attempt to discover novel piezophiles. The pressure vessels were incubated at 4°C until shipment at the end of the voyage to Scripps Institution of Oceanography (La Jolla, California) for enrichment and isolation study.

Enrichment culturing for methanogens began in the anaerobic chamber. Samples for methanogen enrichments were taken from cores that had been maintained at low temperature and placed in oxygen-free conditions as quickly as possible. For sediment transfers, subcores were brought into the glove bag in nitrogen-filled bags and insulated on blue ice. Approximately 1 g from the final inner pared core sample was placed in a serum vial containing 9 mL of MSH medium (Ni and Boone, 1991) without KCl. The vial was stoppered and shaken vigorously for 10 s. H2 was added (2 atm), and cultures were incubated statically at 20°C (Mikucki et al., 2003).

Summary of sampling for shore-based studies

The bulk of the samples taken were for shore-based investigations. WR samples taken for lipid analysis, DNA extraction, and organic matter analysis were stored in sterile plastic bags at –80°C. Subsamples preserved for direct counts were fixed in a final concentration of 1% glutaraldehyde, sonicated, filtered onto 0.2 m black polycarbonate filters (Lunau et al., 2005), or preserved in 1:1 ethanol and phosphate-buffered saline (PBS). Subsamples for FISH were fixed in 2% formalin and stored in a 1:1 solution of ethanol-PBS (Boetius et al., 2000); all of these samples were stored at –20°C. Cores to be used for culture-based analyses and for methods that require living cells as starting material (e.g., rate measurements) were stored at 4°C under nitrogen either in triple-ply aluminized polyester heat-sealed bags or in sterile plastic bags inside canning jars that were flushed with nitrogen before sealing.