Leg 200 was the first ODP cruise to carry out onboard microbial examinations of water, sediment, and rock samples. Classical cultivation approaches as well as fluorescent in situ hybridization (FISH), different fluorescent staining techniques, and shore-based investigations like deoxyribonucleic acid (DNA)-related analyses, biolipid marker analyses, and various microscopic methods such as electron microscopy (transmission electron microscopy [TEM], environmental scanning electron microscopy [ESEM], and field emission scanning electron microscopy [FE-SEM]) were used.
The microbiology objectives for Leg 200 included
The general goal of the microbiological investigations conducted during Leg 200 was to obtain insights into the mutual interdependent relations between the microbial communities and the geochemical processes within the sedimentary, igneous, and volcanic environments.
Samples were taken for both cultivation and cultivation-independent approaches. Magnetostratigraphic measurements of the first core indicated that during recovery a significant amount of presumably loose sediment was displaced as the APC penetrated into the sediment. The sample from 0 mbsf might therefore be more accurately defined as near surface. From a water depth of 4235.1 m, two near-surface samples were taken from the uncontaminated center part of the core. After removing a thin layer (0.5 cm) of sediment from the section's end, samples were taken from the very upper part and from 5 cm deep in the core under sterile conditions using autoclaved spatulas. The sediment was classified as yellowish brown clay with volcaniclastic turbidites (Fig. F65). The next samples came from distinct layers of black sands at depths of 0.95 and 7.90 mbsf, respectively (Fig. F66). Additional sediment samples, classified as volcaniclastic turbidites, were taken at depths of 5.95 and 6 mbsf (Fig. F67). In order to determine distinctive microbial communities at various depths and within different volcaniclastic layers, a sample of volcaniclastic silt claystone was taken from a depth of 23.94 mbsf (Fig. F68). Further samples were taken in order to compare the microbial communities of the two different tuff layers embedded in volcaniclastic silty claystone from a depth of 32.24 mbsf (classified as altered vitric tuff) and 37.33 mbsf (described as palagonitized crystal vitric tuff) (Figs. F69, F70).
For a comparative determination of total cell counts within the sediments from Hole 1223A, the green fluorescent nucleic acid stains SYTO 9 and SYBR Green I and the red fluorescent stain SYTO 62 were successfully combined with the commonly used DNA stain, 4´,6-diamidino-2´-phenylindole-dihydrochloride (DAPI). The detection limit was calculated to be 1 x 104 cells per gram of wet sediment. Bacteria were present in all sediment samples from Hole 1223A down to 5.95 mbsf. The most abundant bacterial community was present in near-surface sediments (Sample 200-1223A-1H-1, 0-5 cm) with at least 2.3 x 1010 cells per gram of wet sediment. Figure F71 shows an overview of cells and microcolonies stained with the green fluorescent DNA-staining dye SYBR Green I. The inset shows a close up of a bacterial microcolony embedded in extracellular polymeric substances. Microcolonies and microbial aggregates were counted as one cell.
In the sediment layer at 0.95 mbsf (Sample 200-1223A-1H-1, 95-96 cm), the number of total cell counts significantly declined to 1.5 x 108 cells per gram of wet sediment. Population numbers decrease rapidly with increasing depth, confirming the general model for bacterial distributions in marine sediments as previously described by Parkes et al. (1994).
In the sediment layer located at 5.95 mbsf (Sample 200-1223A-1H-4, 145-150 cm), classified as volcaniclastic turbidites, the amount of total cell counts increased again to 3.2 x 109 cells per gram of wet sediment.
To determine the MPN of sulfate-reducing and fermentative bacteria, MPN series were prepared from both sediment samples and vitric tuff. From the altered vitric tuff, additionally 25 distinct microbial colonies could be isolated and further characterized as facultatively anaerobic organisms forming stable cell aggregates under appropriate conditions (Fig. F72). Applying DNA-staining fluorescent dyes and FISH with group-specific oligonucleotide probes, it turned out that both the cell morphology and phylogenetic affiliation of all isolates were very similar. All hybridizations were performed at 48°C under stringent conditions using 35% formamide solution in the hybridization buffer and the corresponding sodium chloride concentration in the washing buffer (Manz, 1999). Strong fluorescent signals could be detected using probes EUB338, SRB385Db, and, to a minor extent, with probe LGCa-c. In contrast to this, no hybridization signals could be monitored after hybridizations with probes ARCH915, EUB338-II, EUB338-III, ALF1b, GAM42a, or CF319a.
A common feature shared by all bacteria within the sediment samples was the small average cell size of 1 µm at best, which is in good agreement with the results reported by Torella and Morita (1981). For a comprehensive review on sediment bacteria, see Nealson (1997). In contrast to the widely used acridine orange staining technique, the application of new fluorescent dyes with higher quantum yields (e.g., SYBR Green I and SYTO 9) in combination with objectives of numerical apertures >1.2 enabled the clear visualization of bacterial cells with average cell size of 1 µm or less. Hence, the staining techniques performed in this study led to total cell counts in the sediments that range about one order of magnitude higher than the previously reported bacterial cell counts in sediments from deep marine environments (Parkes et al., 1994; Cragg et al., 1996). One has also to keep in mind that marine bacterial communities might be associated with high numbers of bacteriophages, with average particle sizes ranging up to 0.5 µm. These bacteriophages may significantly contribute to the total cell counts as determined by high-performance nucleic acid stains.
In this study, all colonies obtained from the altered vitric tuff proved to be mixed cultures consisting of at least three different bacterial specimens as revealed by FISH. This is further evidence that the classical approach to obtaining a pure culture (that is transferring a single colony three times in order to obtain a pure culture) is insufficient to get pure cultures from highly complex, heterogeneous environmental samples. Using FISH, strong fluorescent signals could be detected with the 16S ribosomal ribonucleic acid (rRNA)-targeted oligonucleotide universal probe EUB338 (Amann et al., 1990) specific for Bacteria (Fig. F73). No hybridization signals could be monitored after hybridization with the universal probe ARCH915, specific for the domain Archaea (Burggraf et al., 1994), or the probes EUB338-II and EUB338-III, targeting Planctomycetales and Verrucomicrobia (Daims et al., 1999). As the main phyla of bacteria common in seawater are affiliated within the alpha- and gamma-Proteobacteria (Gauthier et al., 1995; Gonzales and Moran, 1997), the subclass-specific probes ALF42a and GAM1b (Manz et al., 1992) were used to specify the isolated bacteria in terms of potential allochthonous contaminants. Because no fluorescent signals could be detected after application of these probes, it seems very unlikely that the newly isolated bacteria belong to bacterial genera known to be fast-growing heterotrophic contaminants from ambient seawater (e.g., Pseudoalteromonas and Roseobacter group). Positive hybridization signals were also not observed in additional hybridizations performed with probe CF319a/b (Manz et al., 1996), which is specific for the Cytophaga-Flavobacteria group, known to be common autochthonous inhabitants within marine microenvironments. The most abundant part of the investigated microcolony, however, showed strong fluorescent signals after hybridization with the probe SRB385dD (Rabus et al., 1996), which was assigned to sulfate-reducing bacteria (Fig. F74). Only a few, but strong, fluorescent signals were achieved by the application of three probes LGCa-c (Meier et al., 1999), initially designed for the specific detection of Gram-positive bacteria with low G+C content of DNA (Fig. F75). Interestingly, the sum of the hybridized cells did not cover all cells visualized by DNA staining. This indicates the presence of further bacterial cells within the microcolony that have not been characterized yet.
Besides the onboard investigations performed during Leg 200, additional microbiology data will be generated in shore-based studies. The major effort will be to determine the bacterial composition of the sediments as well as the altered and palagonitized crystal vitric tuff by DNA extraction and subsequent phylogenetic analyses. The characterization of bacteria, isolated from the anaerobic enrichment cultures and from the aerobic colonies grown on solid agar plates containing different levels of substrates, will be continued. Proxies of past microbial activities in the samples will be investigated by extended lipid biomarker analysis for the presence of both bacterial and archaeal traits.
Based on the rRNA sequences of the isolates and clones, additional isolate-specific probes will be developed to recover the cultivated isolates and sequenced clones by FISH. Using these probes, the spatial distribution of the targeted organisms will be studied within the fixed material and within thin sections of both the sediments and the volcaniclastic tuff. Sample material will be embedded in different mounting media, and the spatial bacterial distribution within the thin sections will be investigated by the combination of FISH and wide-field deconvolution epifluorescence microscopy (Manz et al., 2000). To elucidate the nature of the sulfate-reducing bacteria as shown by FISH in the microcolony obtained from altered vitric tuff, further shore-based investigations will be carried out using recently developed genus- to species-specific probes (Manz et al., 1998).