1 m is required.
Microbiological sampling at Site 1230 covered the shallow sulfate-reducing zone near the sediment/water interface (0-8 mbsf), the transition between the sulfate-rich surface layers and the sulfate-depleted methanogenic deeper sediment layers (8-12 mbsf), and the deep sediment layers down to 268 mbsf. The upper 20 m of the sediment column was characterized by steep biogeochemical gradients. This interval was sampled in at least two sections per core for DNA and FISH-SIMS analysis, measurements of sulfate reduction rates, hydrogen concentration and turnover, methanogenesis rates, acetate turnover, thymidine incorporation, bacterial lipid biomarkers, adenosine triphosphate, FISH, and iron/manganese/sulfur solid phases (Fig. F7) (Cores 201-1230A-1H through 3H). The deeper sediment layers were sampled on average every 15 m (Cores 201-1230A-4H through 38X).
Sampling in Hole 1230B focused on Core 201-1230B-2H. This core included the interface where the steep, decreasing sulfate gradient leads into a zone where sulfate concentrations remained <1 mM but do not approach zero for at least 2 m (~8-10 mbsf). In order to sample this low-sulfate zone in fine spatial resolution, Sections 201-1230B-2H-2, 2H-3, 2H-4, and 2H-5 were divided into three subsections. Each of these subsections was sampled for interstitial water, DNA, and biomarker analyses (Fig. F8). In this way, microbial population gradients obtained from DNA and biomarker analyses can be connected directly to the sulfate profile and other interstitial water data. In addition, the sediment surrounding a gas hydrate nodule in Section 201-1230B-12H-2 was sampled with 5-cm3 syringes (see "Lithostratigraphy" and "Biogeochemistry").
A similar strategy was followed in Hole 1230C. Here, the sampling scheme for Sections 201-1230C-2H-2, 2H-3, 2H-4, and 2H-5 included activity determinations (sulfate reduction measurements and acetate turnover measurements), cultivations, biomarker and DNA analyses, and an interstitial water sample at the top of each section (Fig. F9). The interstitial water sample was analyzed immediately for sulfate to allow precise correlation of the sampling scheme to the sulfate profile in the low-sulfate zone before the actual microbiology sampling started. In particular, the area of the most detailed sampling was centered on the 7.5- to 9.7-mbsf depth interval, where the steep sulfate gradient of the upper sediment column turned into the low-sulfate zone (8.5- to 9-mbsf interval in Section 201-1230C-2H-3). In order to cluster as many samples as possible around this interface, DNA samples and enrichment samples were taken in 5-cm3 syringes (combined enrichments) (Fig. F9) from a single whole-round core (WRC) section rather than as separate WRC samples (Fig. F9). These integrated sampling strategies are necessary at key biogeochemical horizons, when a resolution of 1 m is required.
Samples from the overlying "slop" (liquid mixture of surface sediment and water) were taken from Core 201-1230C-0 for experiments to evaluate the effects of pressure on microbial activity measurements (sulfate reduction, thyridine incorporation, acetate turnover, and bicarbonate reduction) and for total cell counts, FISH, and thermophile enrichments.
Samples of 1-cm3 plugs for total prokaryotic cell enumeration were taken during core processing in the 10°C refrigerator from Holes 1230A, 1230B, and 1230C. From Hole 1230A, 24 samples were taken between 1.11 and 268.65 mbsf. The majority of these samples were processed immediately. From Hole 1230B, 29 samples were taken between the surface of the sediment column and 81.83 mbsf. These samples were concentrated between Section 201-1230B-2H-2 and 2H-5 and were designed to provide high-intensity sampling through the sulfate-methane transition. Most of these will be processed as part of shore-based activities. Two samples were also taken in Hole 1230C at 7.75 and 8.04 mbsf (Samples 201-1230C-2H-3, 25-30 cm, and 2H-3, 54-55 cm). These last samples were specifically targeted at the sulfate-methane transition and will also be processed on shore. Prokaryotes were present in all samples studied, to the depth of 257.70 mbsf (Fig. F10). The highest number of prokaryotes was found near the surface (Sample 201-1230B-1H-1, 0-1 cm), in a sample that contained 4.21 x 108 cells/cm3. The lowest number was at the base of the hole, with 1.28 x 106 cells/cm3 (Sample 201-1230A-37X-1, 10-14 cm). This is a 330-fold difference between the cell concentration of the sediment/water interface and that of this deepest sample.
The overall depth profile of cell numbers per cubic centimeter follows a trend observed at other ODP sites (Parkes et al., 1994); many of the data points lie very close to or on the mean regression line for all previously enumerated sites (Fig. F11). There are two increases of note in the Site 1230 record. First, between ~7 and 21 mbsf (Samples 201-1230A-2H-2, 110-115 cm, and 3H-5, 68-78 cm) there is a small increase coincident with the sulfate-methane transition and the upper part of the methanogenic zone. Second, there are nearly sustained constant cell numbers between ~66 and 190 mbsf (Samples 201-1230A-9H-5, 132-138 cm, and 201-1230A-24H-2, 71-77 cm). The reasons for this plateau in prokaryotic numbers are unclear, although it is coincident with a bulge in many of the interstitial water geochemical profiles (see Fig. F3).
Numbers of dividing cells (suggested as an index of growth activity) are typically <10% of the total count. As expected, dividing cells as a percentage of the total count are high near the surface (Fig. F10). They then decrease to ~9% between 20 and 50 mbsf. After a single low datum they then increase to 10%-11% over the depth range of the plateau in total cell numbers. This may indicate that between 60 and 190 mbsf continuing growth contributes to this plateau in prokaryotic cell numbers. Below 190 mbsf, total cell numbers decrease to <1 x 107 cells/cm3 and percent dividing cells becomes unreliable. Nevertheless, there appears to be a substantial decrease in percent dividing cells, and on two occasions zero was recorded.
At 216 mbsf, there is a discontinuity in the sediment column where, during accretion, the upper layers of sediment were removed and replaced by younger sediment, leaving a hiatus of ~4 Ma. When this site was previously cored (Shipboard Scientific Party, 1988) at Site 685, the data were not sufficiently constrained to accurately identify this boundary. If there are missing sediment layers, then prokaryotic cell counts in the lower sediments would appear to be smaller than expected for their current depth in the sediment column. Therefore a step-down in numbers would be visible in the total prokaryote profile. Such a step is observed between ~170 and 230 mbsf (Fig. F12). Taking the tangents of a smoothed curve over the lower and the upper part of the cell number profile, the intersection of the tangents provides an estimate for the boundary of 214 mbsf.
The nature of the sediments at this site made direct counting more problematic than usual. Where subsamples of 15-40 µL can generally be processed, in this case subsample size was restricted to a maximum of 8-12 µL. Consequently, the detection limit increased to 2.5 x 105 cells/cm3 and counting sensitivity decreased. The percentage of dividing cells should be viewed with caution where the direct count is <1 x 107 cells/cm3.
While drilling cores for microbiology, the potential for contamination with bacteria from the surface is highly critical. Contamination tests were continuously conducted using solutes (PFT) or bacterial-sized particles (fluorescent microspheres) to check for the potential intrusion of drill water from the periphery toward the center of cores and thus to confirm the suitability of the core material for microbiological research. We used the chemical and particle tracer techniques described in ODP Technical Note 28 (Smith et al., 2000). Furthermore, the freshly collected cores were visually examined for possible cracks and other signs of disturbance by observation through the transparent core liner. Core sections observed to be disturbed before or after subsampling were not analyzed further. Such disturbance phenomena are critical to the integrity of the core material and therefore also to its usefulness for microbiological studies.
The PFT was injected continuously into the drilling fluid during drilling of Holes 1230A, 1230B, and 1230C (see "Perfluorocarbon Tracer Contamination Tests" in "Microbiological Procedures and Protocols" in "Microbiology" in the "Explanatory Notes" chapter). PFT sampling focused on microbiology cores and especially on sections that were used for slurry preparation and cultivations. Two 5-cm3 subcore samples were taken at the core center and pooled for PFT measurement. In parallel, a 5-cm3 subcore sample was taken at the core periphery, adjacent to the core liner, to compare the PFT concentrations in the center of a core to the PFT concentrations at the periphery of the same core, and to assess the difference in contamination potential. Whenever possible, the samples were taken directly on the catwalk and capped away from nearby cores because the PFT content of catwalk air was usually not detectable.
With three exceptions, low levels of potential seawater contamination (<0.05 µL seawater/g sediment) were found for the center portions of all tested APC cores of Holes 1230A, 1230B, and 1230C (average = 0.08 µL seawater/g sediment) (Table T8). The three exceptions, which may have been contaminated, are samples from Cores 201-1230A-13H and 22H and 201-1230C-1H. The outer portions of all tested cores had significantly high levels of PFT tracer and potential seawater contamination (mostly >0.15 µL seawater/g sediment; average = 0.71 µL seawater/g sediment). XCB cores showed higher PFT concentrations than did APC cores (for center portions, average = 0.24 µL seawater/g sediment; for outer portions, average = 2.74 µL seawater/g sediment). Three of the four XCB cores sampled exhibit PFT concentrations in the center that suggest potential prokaryotic cell contamination from seawater drilling fluid (>0.2 µL seawater/g sediment). In almost all cases for both APC and XCB cores, the PFT content and the estimated potential seawater contamination levels were higher in the periphery of the core than in the center.
Of the 11 slurry samples taken from Site 1230 (Table T9), only the slurries from Cores 201-1230A-1H, 31X, and 38X showed concentrations of PFT that suggest potential microbial contamination from seawater drilling fluid. Assuming 5 x 108 prokaryotic cells/L surface seawater, each 0.1 µL of seawater contamination may represent as many as 50 contaminating cells if the sediment is porous enough to allow cells to travel with the PFT.
In order to assess the relationship of PFT contamination to particulate contamination (such as microorganisms), nine of the sediment samples for which PFT concentrations had been determined were further analyzed for bead contamination (Table T8). No beads were found in APC samples with low PFT values (corresponding to 0.02 µL seawater contamination/g sediment). Six beads were found in 50 fields counted for a sample with as much as 0.22 µL seawater contamination/g sediment. This latter count represents 76 beads/g sediment. Considering the extremely high number of beads deployed (1.7 x 1011 in this case), 76 beads/g sediment most likely represents far fewer actual contaminating microorganisms. The results of these comparisons suggest that samples with PFT contaminations representing less than ~0.02 µL contaminating seawater/g sediment were not contaminated with microorganisms during the drilling and sample processing.
Fluorescent microspheres (beads) were deployed on all seven cores from which slurries were intended to be made at this site (Table T10). However, because of variable sediment quality, on two occasions slurries were made from adjacent cores in which beads had not been deployed. Thus, only five slurries could be examined for bead contamination (Table T10). For each slurry two subsamples were processed: (1) a sample of the slurry to check contamination and (2) a scraping from the outer surface of the core to confirm deployment of beads.
Bead deployment was confirmed from the outer core scrapings in five of six cores that were sampled for slurry preparation (Table T10). Beads were detected in one slurry sample (Sample 201-1230A 38X-1, 65-71 cm) at low levels, but nevertheless at levels that suggested potential contamination. This was the only XCB core used for slurry preparation, and XCB cores, because of the way they are obtained, have a higher potential for contamination compared to APC cores. The slurries made from Sections 201-1230C-2H-2 through 2H-5 were low volume and insufficient for bead analysis. PFT analysis however, suggested that Core 201-1230C-2H was free of contamination.
Depths were selected for cultivation in order to cover the entire drilled sediment column (Hole 1230A) and especially the sulfate-methane transition zone (Hole 1230C). Additionally, samples taken close to hydrate layers were used for enrichments. The media inoculated are selective for fermenters, sulfate reducers, methanogens, and various anaerobic chemolithoautotrophic and heterotrophic prokaryotes that use iron(III) or manganese(IV) as an electron acceptor. Quantitative cultivations (MPN experiments) and enrichment cultures were started at temperatures between 4° and 80°C (Table T11). As it is known that aerobic prokaryotes may also be present in strictly anoxic environments, several MPN experiments for aerobic prokaryotes (media with suffix -ox) were started at this site to compare with the anoxic assays (Table T11).
13C substrate incubations were initiated for postcruise analysis by FISH-SIMS using material from Cores 201-1230A-2H, 9H, 15H, and 38X. In these cases, 10 mL of the master slurry was injected into each bottle. The 13C substrates used were methane, acetate, and glucose. Glucose was used in Cores 201-1230A-9H, 13H, 15H, and 38X. Acetate was used at all depths except Core 201-1230A-38X, with multiple bottles at each depth. Methane was used at all depths, with multiple bottles used for samples from Cores 201-1230A-2H and 38X.
