MICROBIOLOGY

Bacteria play a dominant role in the degradation of organic matter within sediments, and as a consequence, drive chemical changes and early diagenesis. The existence of a deep bacterial biosphere in marine sediments has only recently been established (Parkes et al., 1994), but already the activity of bacteria in depths to 750 mbsf and their direct involvement in geochemical changes have been demonstrated.

Little is understood about what types of bacteria are found deep below the seafloor, what they use for energy and carbon sources and the rate of their activity, how healthy the population is, and how much they affect the environment around them. For example, recent research (Wellsbury et al., 1997) has shown that temperature increases during burial can result in organic matter becoming easier to degrade by bacteria and that bacterial populations and activity can increase in deeper layers below 100 mbsf. Increasing organic matter bioavailability was reflected by increases of up to a thousandfold in substrates for bacterial activity (volatile fatty acids, particularly acetate) in deep sediments (Leg 164; Paull, Matsumoto, Wallace, et al., 1996). Thus, bacterial populations may exist at much greater depths and may even increase with depth, rather than decrease as energy sources "improve" with increasing depth and temperature.

The work undertaken during Leg 190 will identify the bacterial mechanisms important in the environment and determine the impact these have on sediment geochemistry. This investigation has been designed to address the following questions: (1) can we improve methods of culturing deep-sediment bacteria (e.g., sulfate reducers, iron reducers, and oxidizers) and obtain more useful viable counts, and can we isolate new organisms; (2) how many bacteria are present in very deep subsurface sediments, and can we determine when a bacterial cell has died; (3) are there increasing concentrations of labile organic carbon (acetate) with depth, and what affect would this increase have on rates of methanogenesis measured at in situ temperatures and pressures; and (4) using molecular genetics, can we identify and characterize the bacterial populations deep in the sediments, and what biomarkers are present and currently being produced? Leg 190 also provided the deepest samples yet collected for bacterial analyses.

Microbiology samples consisted of plug minicores of a few cubic centimeters from whole-round cores (WRCs), IW splits, and split core samples from the sample table.

Sediment in 1-cm³ plugs was taken for shipboard direct microscopic determination of bacterial numbers. These plugs were taken from the end of selected 1.5-m core sections immediately after the sections were cut on the catwalk and before they were sealed with an end cap. Potentially contaminated sediment was removed with a sterile scalpel, and a sterile 5-cm³ plastic syringe with the luer end removed was used to take a 1-cm³ plug. The syringe was sealed with a sterile Suba-Seal stopper. In a clean area of the laboratory, the 1-cm³ plug was extruded into a sterile serum vial containing 9 mL of filter-sterilized (0.2 µm) 4% formaldehyde in 3.5% NaCl. The vial was crimped and shaken vigorously to disperse the sediment particles.

WRCs were cut from 3.0-m (double) core sections on which the end caps had not been sealed with acetone. The double core sections were removed from the catwalk and brought into the core reception area, where they were cleaned, wiped with ethanol, and placed into a sterile (autoclaved) cutting rig flushed constantly with sterile oxygen-free nitrogen (OFN) to maintain anoxia. The initial cut was made at 1.5 m, and the entire lower section was returned the curator. WRCs of varying lengths were then removed from the base of the retained core section upward. In general, the first WRC was for IW; subsequent WRCs were for microbiology. Each WRC was capped with standard core end caps or sterile (-irradiated) core end caps, where appropriate, under OFN. Core end caps were sealed with insulation tape and the whole WRC was either (1) stored at -80°C, (2) stored in gas-tight aluminum/plastic bags in an anoxic atmosphere (Anaerocult-A, Merck) at -80°C, (3) stored in gas-tight aluminum/plastic bags in an anoxic atmosphere at 4°C, or (4) moved to an anaerobic cabinet for initial microbiological handling. The last 5-cm WRC was cut and left uncapped. It was wrapped in sterile aluminum foil and immediately transferred to the anaerobic cabinet for processing.

IW splits of as much as 3 mL where possible were taken for shore-based acetate analysis (Wellsbury et al., 1997). When squeezed IW volume was unavailable, 10-cm³ of a squeeze cake was taken instead. Samples were stored frozen (-20°C).

After alkalinity was measured in the chemistry lab, ~2.0-mL aliquots of the acidified solution were transferred to glass headspace vials and stored at -20°C for dissolved organic carbon analysis.

The acidified IW samples were sparged with CO₂-free air and oxidized on a platinum catalyst at 680°C in a total carbon analyzer (Shimadzu TOC 5000A). The CO₂ produced upon oxidation was quantified using a nondispersive infrared detector.

Sediment samples were removed from the split-core sampling tables for shore-based lipid analysis. One quarter of a WRC, 30 cm in length, was removed from the working half using latex gloves. Samples were individually wrapped in aluminum foil and stored in nonheat-sealed polyethylene bags at -80°C.

Acridine Orange Direct Counts

Total bacterial abundance and numbers of dividing/divided cells were determined using acridine orange as a fluorochrome dye with epifluorescence microscopy (Fry, 1988). Fixed samples were mixed thoroughly, and a 5- to 10-µL subsample was added to 10 mL of 2% (by volume) filter-sterilized (0.1 µm) formaldehyde in 3.5% NaCl. Acridine orange (50 µL of a 1-g/L filter-sterilized [0.1 µm] stock solution) was added, and the sample was incubated for 3 min. Stained cells and sediment were trapped on a 0.2-µm black polycarbonate membrane (Osmonics, USA). Excess dye was removed from the membrane by rinsing with a 10-mL aliquot of 2% (v/v) filter-sterilized formaldehyde in 3.5% NaCl. The membrane was mounted for microscopic analysis in a minimum of paraffin oil under a coverslip.

Mounted membranes were viewed under incident illumination with a Zeiss Axiophot microscope fitted with a 50-W mercury vapor lamp, a wide-band interference filter set for blue excitation, a 100× (numerical aperture = 1.3) Plan Neofluar objective lens, and 10× eyepieces. Bacterially shaped fluorescing objects were enumerated, with the numbers of cells on particles doubled in the final calculations to account for masking. Dividing cells (those with a clear invagination) and divided cells (pairs of cells of identical morphology) were also counted.

Cell Viability

Subsamples (5-20 µL) of the slurries were filtered through a 0.22-µm membrane filter (Millipore Corp.). Filters were stained with 100 µL of Live/Dead BacLight Kit for 20 min then dried and mounted with nonfluorescent immersion oil on glass microscope slides and examined with epifluorescence microscopy (420-nm excitation filter; 590-nm barrier filter) at 1000× using an epifluorescent microscope.

Two approaches were used to assess bacterial metabolic activity, fluorescene diacetate (FDA) and 5-cyano-2,3-ditolyl tetrazolium chloride (CTC). Subsamples (5-20 µL) were filtered through a 0.22-µm membrane filter (Millipore Corp.). These were stained with FDA-dimethyl sulfoxide (12-µM final concentration) for 20 min. Filters were dried and mounted with nonfluorescencing immersion oil on glass microscope slides and examined under 1000× magnification. FDA-stained microorganisms were viewed using a 520-nm barrier filter.

Diluted slurries were incubated in triplicate with 5% CTC in sterile test tubes for 4 hr at room temperature. After incubation, selected samples were counterstained with 50 µg/L of the DNA-binding fluorochrome 4´,6-diamidino-2-phenylindole (DAPI) to determine the total cell abundance in the same preparation. Subsamples were filtered (0.22 µm, Millipore Corp) and dried and mounted with nonfluorescent immersion oil on glass microscope slides and examined under 1000× magnification. CTC-treated preparations were viewed using a blue (420 nm) excitation filter used in combination with a 590-nm barrier filter. CTC- and DAPI-stained microorganisms in the same preparation could be viewed simultaneously with a 365-nm excitation filter and a 400-nm cutoff filter.

Enrichment Cultures

A 5-cm WRC was placed in the anaerobic chamber, and 1-cm³ plugs were taken with sterile syringes to inoculate enrichment cultures. Anaerobic growth media for sulfate-reducing, iron-oxidizing, iron-reducing, and denitrifying bacteria (Widdel and Bak, 1992) were utilized. An additional 2-cm³ plug was removed for shore-based DNA/RNA analysis. The residue of the WRC was removed from the core liner using a sterile steel spatula, leaving the outer, potentially contaminated sediment adjacent to the core liner. The sediment was stored in plastic bags with an oxygen scrubber at -80°C for shore-based fatty acid analysis.

The uncapped WRC was subsampled in the anaerobic cabinet for viable counts of sulfate-reducing and iron-reducing bacteria. Viable counts were also initiated for Methanococcus sp. Additional plugs (3 cm³) were collected from the WRC using sterile syringes and then placed in plastic tubes and frozen at -80°C for shore-based DNA extraction and analysis. The residue was stored in a plastic ZipLok bag at -80°C for shore-based analysis of nucleic acids.

To confirm the suitability of the core material for microbiological research, contamination assays were conducted to quantify the intrusion of drill water using the chemical and particulate tracer techniques described in ODP Technical Note 28 (Smith et al., 2000), except that the extraction of the particulate tracer, fluorescent microspheres, has been modified to increase the sensitivity of the assay. The modification consists of suspending the sediment sample in a solution of saturated NaCl. The solution is centrifuged for 5 min (Marathon 10K, 2800 × g), and the supernatant containing the microspheres is filtered through polycarbonate filters (0.2-µm pore size). The microspheres are counted, and data are reported as number of microspheres per gram of sediment.