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Expedition 337 was the first riser drilling IODP expedition to incorporate extensive shipboard microbiological and molecular biological analyses. Microbiological samples obtained provided a unique and unprecedented opportunity to not only expand the known depth limits of microbial life in the subseafloor, but also—using chemical and molecular tracers—to assess the degree to which contamination introduced by riser drilling operations affects sample quality. In the designated microbiology laboratory on the Chikyu, cell separation and enumeration, DNA extraction, PCR assays on key phylogenetic and metabolic marker genes, molecular fingerprinting analyses, and microscopic observations were all accomplished during the expedition. A wide range of incubations with stable isotope-labeled substrates was initiated to study stable isotope incorporation into nucleic acids and whole cells (for incorporation into lipids and metabolic compounds, see “Organic geochemistry”). Radioactive tracer experiments, which involved 35S-labeled sulfate for determination of potential sulfate reduction rates (pSRRs) and a broad set of 14C-labeled substrates to study potential rates of microbial C-cycling reactions (see “Organic geochemistry” for 14C-experiments), were carried out in a designated radioisotope van. A large number of samples were, moreover, inoculated with media selecting for a wide range of microbial metabolisms, including iron reduction, methanogenesis, and acetogenesis.

The sampling scheme for WRCs and experimental procedures for shipboard and shore-based analyses are illustrated in Figure F55. In total, nearly 300 WRCs out of 32 drill cores were sampled for microbiological investigations (for details, see MBIO_SAMPLE_LIST.XLSX in MBIO in “Supplementary material”). Contaminated outer core layers containing potential drilling mud contamination were removed from all WRCs by sterile spatulas or ceramic knives prior to sampling.

Contamination tests

Chemical tracer

Measured perfluorocarbon (PFC) tracer concentrations fluctuated greatly over time and remained under the target concentration of 1 mg/L for most of the expedition (Fig. F56). During the initial days of the expedition, PFC concentrations in mud tank samples remained remarkably low, despite daily additions of 100 mL of PFC tracer (Table T37), which were, in principle, sufficient to produce PFC tracer saturation (~2 mg/L) (Colwell et al., 1992). The low initial concentration turned out to be, in part, a problem of the extraction method used. Without further PFC tracer addition, measured PFC values increased drastically when a 1 h preincubation step at 80°C with active sample mixing was included prior to the 30 min preincubation without mixing in the autosampler (also see “Microbiology” in the “Methods” chapter [Expedition 337 Scientists, 2013b]). Before coring began, this new preincubation period was increased to 2 h (starting 17 September) to ensure maximum PFC tracer recovery and thus as accurate as possible quantification of drilling mud contamination.

Despite daily PFC additions to active tanks and methodological improvements, PFC concentrations in drilling mud were found to fluctuate considerably on a day-to-day basis. The good agreement between measurements in the physically separated active Tanks 4 and 5, from which we obtained drilling mud samples at the same time of day, indicates that this was not a matter of insufficient homogenization. Instead, these fluctuations are probably to a large extent explained by day-to-day variations in drilling mud production and hence PFC dilution. Because of these fluctuations and the limited sampling frequency of one time per day, the exact PFC concentration in drilling mud at the time of coring was unknown. Fortunately, PFC concentration in active tanks, core liner fluid, and upon recovery in the mud ditch typically exceeded 100 µg/L, allowing calculations of drilling mud intrusion using an assumed constant PFC concentration of 100 µg/L. Because this concentration is lower than most of the measured PFC concentrations, contamination estimates based on this value should be considered conservative.

On average, PFC concentrations in mud fluid from the active tanks, core liner, and upon recovery from the mud ditch show good agreement (typically within a factor of 2), indicating that PFC losses (e.g., because of volatilization during core processing and mud recovery) are minor. The on average slightly lower measured values in the core liner and mud ditch compared to the active tanks might at least in part be due to dilution with core samples during drilling operations.

Using the same preincubation method as for drilling mud, PFC concentrations were determined in cuttings and cores. Volumes of drilling mud intrusion and numbers of contaminant cells introduced were subsequently calculated using an assumed PFC concentration of 100 µg/L (0.1 µg PFC/mL) drilling mud and cell concentration of 2.66 × 108 cells/mL drilling mud (mean value of microscopic direct counts) (Table T38, also see “Cell counts,” as well as Smith et al., 2000, for details on calculation method). Cuttings were highly contaminated with drilling mud, with all values except one (Sample 337-C0020A-267-SMW) exceeding 1% (v/v) contamination of pore fluid, and contamination in many samples exceeding 100 µL/g sample (Table T39). Although the outside and exterior of cores typically showed high levels of contamination, the inner portions in most cases showed much lower contamination (Table T40). In many samples obtained from the innermost portions of cores, PFC tracers were even below detection. To estimate the maximum drilling mud contamination and number of contaminant cells that could still be present in samples where PFC was below detection, we inferred the PFC detection limit based on the data distribution of PFC values measured on cores (Fig. F57). Drilling mud contamination of samples, in which PFC was below detection, was conservatively calculated substituting the detection limit value of 8 pg. We then calculated the maximal amount of drilling mud, and hence contaminant cells, within core samples where PFC was below detection (bracketed values in Table T40) based on

  • A detection limit of 8 pg,

  • An assumed concentration of 100 µg PFC/mL drilling mud,

  • The volume of headspace injected into the gas chromatograph with electron capture detector (GC-ECD; 0.5 cm3),

  • The total sample amount (g),

  • The total sample volume (g sample × sample bulk density),

  • The total headspace volume (20 cm3 – total volume of sample), and

  • An average cell concentration of 2.66 × 108 cells/mL drilling mud.

To examine the relationship between drilling mud contamination and environmental variables, we plotted drilling mud contamination versus depth (Fig. F58). Apart from the expected trend of higher contamination in exterior compared to interior parts of cores, the critical importance of vertically resolved contamination measurements was underscored; several cores, from depths spanning almost the entire interval cored, harbored very high drilling mud contamination all the way to the core center. We also examined the relationship between contamination within core interiors and lithology (Fig. F59). Perhaps surprisingly, no clear contamination trends could be attributed to lithology. PFC results for core, drilling mud, and cuttings are summarized in PFC-CORE.XLSX, PFC-CUTTINGS.XLSX, and PFC-DRILLING MUD.XLSX in MBIO in “Supplementary material.”

Cell counts

Cell counts in drilling mud and cuttings

Cell concentrations in drilling mud were enumerated daily. Cell counts ranged from 1.10 × 108 to ~8.37 × 108 microbial cells/mL drilling mud before drilling started and did not change markedly throughout drilling operations. We also observed 1.6 × 107 to 9.8 × 107 cells/mL in cuttings samples from 646 to 1046 m MSF (Table T39). These cell counts are close to or higher than the cell counts observed in the shallower (100~346 mbsf) part of this site obtained during the Chikyu shakedown cruise (Expedition CK06-06). Simple extrapolation from the cell abundance curve from shallower depths predicts that the number of the cells in the formation is <106 cells/cm3. Combined with the fact that high PFC concentrations were detected in cuttings, we attribute these high cell numbers predominantly to contamination with drilling mud. Calculations based on cell numbers in drilling mud samples indicate that roughly 6%–38% of the volume of cuttings consists of drilling mud.

Cell counts in core samples

From the microbiology community WRCs, 2 cm3 was aseptically taken from the innermost parts, immediately fixed, and processed using the cell separation method described in “Microbiology” in the “Methods” chapter (Expedition 337 Scientists, 2013b). In total, 94 samples were taken for cell enumeration (Fig. F55; also see MBIO_SAMPLE_LIST.XLSX in MBIO in “Supplementary material”). Cell separation was done under a clean air-flow safety cabinet with daily intensive cleaning prior to separation. To minimize the contamination risk from stock solutions, small aliquots of these solutions were stored after filtration and underwent cell separation. After cell separation, these solutions were again passed through a 0.22 µm pore cartridge filter and poured directly into sample tubes. The initial attempts to determine cell abundance in core samples during Expedition 337 were performed using an image-based automatic cell enumeration system (Morono et al., 2009; Morono and Inagaki, 2010). However, because of very low cell abundances, entire filter membranes had to be counted, resulting in long processing periods. Therefore, we decided to count microbial cells from core samples using visual direct counts and flow cytometry (FCM). For direct counts, we selectively counted green fluorescent cells under the microscope. Particles that emitted other colors were not counted.

Although cell abundance was generally low (Fig. F60), cell numbers were above the minimum quantification limit (MQL) even in deeper layers. The cell counts as well as the MQL obtained by FCM are generally higher than the microscopic counts. This is mainly due to the existence of noncellular particles that fluoresce in green color. There was no problem avoiding these particles during counting under a microscope. However, these unspecific signals were difficult to eliminate from the FCM data. In addition, we saw these noncellular particles in blank samples, suggesting that these particles might be generated through the sample preparation protocol. Although we could not solve this problem on board the ship, we will try to eliminate it in shore-based work. Seven and five blanks were counted by microscope and FCM, respectively, and resulted in MQLs of 115 and 6350 cells/cm3, respectively.

In core samples from intervals 337-C0020A-25R-3, 140–141 cm (1999 m CSF-B); 32R-1, 144–145 cm (2457 m CSF-B); and 32R-2, 43–65 cm (2458 m CSF-B), we found microbial cells attached to surfaces of mineral particles (Fig. F61). By comparison, we did not see any particles with attached cells in blank samples. Considering the size of these particles (i.e., 10~20 µm in diameter), it seems unlikely that they were introduced via drilling mud intrusion. Instead, because these samples were all from sandy silt or sandstone, they could be evidence of indigenous microbes residing on mineral surfaces.

DNA extraction

Hot alkaline lysis protocol

Thirty-eight out of 47 microbiology community WRCs (MBIO, Fig. F55) were used for DNA extraction using the hot alkaline lysis protocol. For each extraction, 2–5 cm3 of sample was used. Apart from the extract from the coal layer at Section 337-C0020A-24R-5, which had a black color, crude extracts generally had a brown color. To prevent PCR inhibition, we purified all extracted DNA with an Aurora system (Boreal genomics, Vancouver, Canada). For this purpose, the Aurora low molecular weight soil protocol (without the wash block) was used according to the manufacturer’s instructions. For samples with dense black color, the Aurora 0.7-53kb DNA from soil with enhanced contaminant rejection protocol was used. The final volume of purified DNA solution was ~60 µL and was used for subsequent quantitative polymerase chain reaction (qPCR) and terminal restriction fragment length polymorphism (T-RFLP) analyses.

Chemical lysis protocol

DNA extractions using the chemical lysis protocol were performed on 13 drilling mud samples from active Tank 4, 12 unwashed cuttings samples, 4 washed cuttings samples, and 11 of the 47 microbiology community WRCs (MBIO, Fig. F55). Drilling mud samples were chosen at time points spanning the entire expedition to monitor changes in microbial communities and DNA signatures within drilling mud as background controls for DNA detected in cores. DNA was also extracted from unwashed cuttings obtained throughout the expedition, both as additional background controls for DNA contamination in cores and to examine the potential for cuttings to provide information on indigenous deep subseafloor microorganisms. It was anticipated that DNA of strictly anaerobic microorganisms, which might be below detection in drilling mud, could perhaps be detected in cuttings. Under this scenario, useful phylogenetic and/or metabolic information on in situ communities could be obtained from cuttings originating from drilling intervals without WRC sampling, such as the depth interval from 646 to 1046 m MSF (i.e., Unit I). For comparison to unwashed cuttings, DNA was also extracted from four samples of washed cuttings obtained during the early stage of the expedition.

DNA extractions were performed in parallel on the contaminated exterior and the typically cleaner inner core portions, resulting in a total of 22 DNA extractions from cores. Samples were chosen based on depth, lithology, and contamination of the core interior, with the aim of extracting DNA from samples spanning the entire depth interval sampled, from all main lithologies, and from samples with minimal drilling mud intrusion. Two cores were dominated by fine to medium sandstone (Sections 337-C0020A-10R-2 and 16R-2), two by fine sandstone (Sections 28R-6 and 30R-4), two by siltstone (Sections 3R-3 and, 20R-3), two by coal (Sections 15R-2 and 24R-3), and three by shale (Sections 7R-1, 15R-5, and 26R-7). Drilling mud contamination was below detection in the inner portion of seven of these samples (Sections 7R-1, 10R-2, 15R-5, 20R-3, 26R-7, 28R-6, and 30R-4) but was always detectable in the exterior portion (Tables T40, T41), with estimated contaminant cells in the exterior exceeding 105 cells in all except Section 28R-6, with 5.6 × 103 cells/cm3.

Extracellular DNA extraction

Samples for shipboard extracellular DNA extraction (MLWR, Fig. F55) were obtained from 22 WRCs. Of these, all but one highly contaminated sample from Section 32R-2 were processed, typically within 24 h of sampling. Several core samples consisting of unconsolidated sand, fine sandstone, and shale were observed to contain intact snail and clam shells, infilled fossilized worm burrows, and terrestrial plant debris, making them promising samples for the planned exploration of extracellular DNA pools as genetic archives of ancient biological communities.


Sources of DNA in drilling mud

Most probable number PCR assays were performed on DNA extracts from drilling mud, cuttings, and cores using PCR primer pairs targeting phylogenetic clusters ubiquitously detected in surface seawater (SAR11 and Marine Group I [MGI] Archaea), viscosifiers used during riser drilling (Xanthomonas and Halomonas), and anthropogenic wastewater (Bifidobacterium, Blautia, and Methanobrevibacter) (Table T41). All three indicator organism groups yielded successful PCR amplification; however, the frequency of detection and abundance varied tremendously. Xanthomonas and Halomonas were detected in the vast majority of drilling mud and cuttings samples, two of 11 core interiors, and six of 11 core exteriors. SAR11 was found in all the initial samples but not later on during the expedition. The MGI Archaea, Bifidobacterium, Blautia, and Methanobrevibacter were only detected a few isolated times and at low abundances. These results clearly suggest that the biggest risk of core DNA contamination obtained by riser drilling during Expedition 337 was from microbes that have been linked to mud viscosifiers used to prepare drilling mud (Masui et al., 2008). Microbes from surface seawater represented a significant source of contamination in the initial phase but less so later on, whereas the risk of contamination with human sewage–derived DNA was low.

Open questions remain regarding the variables explaining the observed DNA contamination trends. Considering the chemically harsh environment of drilling mud, with highly alkaline pH (9–11.5) and hypersaline (578–847 mM K+ to 1150 mM Na+) conditions, one might expect seawater- and human sewage–derived microorganisms to die and their DNA to disappear (e.g., due to hydrolysis by DNAses or uptake by surviving organisms) within a short time after drilling mud preparation. The observed disappearance of DNA indicative of these organisms within the initial drilling period is consistent with this interpretation. Yet, fresh drilling mud was also prepared later during the expedition and hence one would expect a reemergence of DNA indicative of seawater microbes during this later stage of the expedition. Since the mud in the initial drilling period of operations was prepared prior to Expedition 337, the seawater in the tank was taken from a different geographic location, possibly influencing the variability of contaminant cell abundance in the mud over the course of drilling operations.

Whether the observed high abundances of Xanthomonas and Halomonas DNA can be attributed to living organisms or high inputs of DNA included in viscosifiers is also not clear. Xanthomonads are used industrially in the synthesis of xanthan gum, a component of the drilling mud gelling agent bentonite, and could potentially grow within drilling mud using organic compounds supplied as gelling agents as an energy source. However, past efforts to grow Xanthomonads from drilling mud using standard Xanthomonas enrichment assays have failed (Masui et al., 2008), suggesting that alive xanthomonads are absent and that Xanthomonas DNA detected may be left over from the production process of bentonite. The presence of Halomonas is perhaps easier to explain: members of this versatile group are widely distributed in seawater, can tolerate high salinities and pH, grow aerobically and anaerobically, and utilize a wide range of organic carbon compounds as energy sources. Thus, though it remains to be shown, it is possible that the broadly adapted and opportunistic Halomonas can grow within drilling mud and account for a significant fraction of the high cell densities observed there.


Quantifications of bacterial and archaeal 16S rRNA genes were performed on DNA extracts obtained from cores using the alkaline lysis and chemical lysis methods (Fig. F62). Negative controls, in which molecular-grade water had been substituted for sample material during extractions, were also examined.

Because of background bacterial contamination from extraction and PCR reagents, amplification of bacterial DNA was also observed in negative controls for the DNA extraction. Therefore, we set the detection limit for bacterial gene detection in DNA extracted from cores to 2× the copy number obtained in negative controls (1.5 × 104 copies/cm3). Archaeal qPCR did not show any amplification from negative controls and could thus detect as few as 1 copy of rRNA genes per reaction.

For both extraction methods, we observed that bacterial DNA from cores amplified at a lower PCR amplification rate than bacterial DNA from extraction negative controls or PCR negative controls, indicating PCR inhibition by co-extracted substances for both the hot alkaline lysis and chemical lysis methods—despite purification by the Aurora system and Norgen Cleanup kit, respectively. Thus, estimated copy numbers most likely underestimate actual copy numbers in cores. More accurate quantifications will be attempted on shore by further improvements to the DNA extraction and purification methods, as well as a newly developed digital PCR technique that is less affected by PCR inhibitors, such as humic acids (Hoshino and Inagaki, 2012).

Overall, our qPCR data confirm the very low in situ cell abundances of microorganisms determined by cell counts (Fig. F62). Highly elevated bacterial gene copy numbers of >106 cm–3 were only found in Sections 337-C0020A-1R-2, 2R-3, and 5R-3 (Table T41), which were highly contaminated with drilling mud all the way to the innermost parts according to PFC analyses (Table T40). All other cores harbored estimated bacterial gene copy numbers between 103–106 cm–3 of sample. Archaeal 16S rRNA gene copy numbers were consistently lower than bacterial copy numbers by two or more orders of magnitude. Several of the core samples with bacterial 16S rRNA genes detected by general bacterial primers also tested positive for 16S rRNA genes of Xanthomonas and Halomonas, indicating drilling mud contamination. The degree of contamination (i.e., whether contaminant DNA dominates or is only a minor fraction of total DNA in these samples) will be evaluated by next-generation DNA sequencing on shore.

Bacterial and archaeal gene copy numbers were converted to cell numbers using standard conversion factors, in which 4.07 and 1.76 copies of 16S rRNA genes were assumed per bacterial and archaeal cell, respectively (values from Ribosomal Database Project,; Fig. F62). Both DNA extraction methods showed good agreement in estimates of bacterial and archaeal cell numbers, in spite of using different primer pairs. Interestingly, our data suggest that archaeal abundance showed no clear depth-related trend, whereas an overall decrease in bacterial abundance occurred with depth. The fact that Archaea were detectable to greater depths than Bacteria is attributable to the much higher detection limit of Bacteria that resulted from the higher level of bacterial background DNA in both PCR reaction and DNA extraction negative controls. In addition to digital PCR quantification, the use of larger core volumes for DNA extractions and improvements to the DNA purification procedure in shore-based investigations will hopefully enable us to reliably quantify Bacteria to greater depths than during the expedition.

DNA fingerprinting techniques

T-RFLP, which is a molecular fingerprinting technique to monitor differences in microbial community structures across different habitats, was performed on 37 microbiology community samples, 10 drilling mud samples, and 3 extraction negative controls. Full 16S rRNA gene amplification was tested by PCR using bacterial and archaeal domain-specific primers. Full archaeal 16S rRNA genes were not amplifiable from any of the samples, but we were able to amplify bacterial 16S rRNA genes. Yet, checks with gel electrophoresis revealed only faint bands of amplified DNA in DNA extracts from below Core 337-C0020A-20R (1962 m CSF-B).

The bacterial community structure of drilling mud samples showed consistent peak locations, suggesting that the microbial community composition of drilling mud was stable throughout the entire drilling period. Terminal restriction fragments (T-RFs) of 181, 553, and 572 base pairs (bp) were observed in all drilling mud samples (Fig. F63) and could thus be used to assess drilling mud contamination in cores. We also obtained bacterial PCR products from negative controls of the DNA extraction. These negative controls provided 200, 666, and 780 bp T-RFs in T-RFLP profiles. Moreover, a T-RF of 233 bp was detected within PCR negative controls, though this amplicon was not detected by agarose gel electrophoresis.

T-RFLP results from cores show only a few peaks, indicating low diversity of indigenous bacterial communities and/or very low copy numbers of amplifiable template DNA in the initial PCR (Fig. F64). T-RFLP profiles also indicate that contamination during drilling, DNA extraction, and from PCR reagents was significant because T-RFs characteristic of drilling mud and DNA extraction negative controls were observed in most core samples. Certain samples contained T-RFs that could not be attributed to contamination, however; for example, a T-RF of 566 bp was unique to DNA extracts from Samples 13R-2, 105–120 cm, to 32R-2, 43–65 cm. Additional T-RFs of 99 and 763 bp appeared only from Samples 19R-8, 60–77 cm, 20R-3, 13–23 cm, and 20R-7, 62–76 cm. Our results from T-RFLP thus demonstrate that, despite low yields, DNA of indigenous bacterial cells was obtained by the shipboard molecular biology program.

Functional genes

We performed PCR assays targeting functional marker genes of methanogenesis and anaerobic methanotrophy (methyl coenzyme M reductase [mcrA]), dissimilatory sulfate reduction (dissimilatory sulfate reductase [dsrB]), and acetogenesis (formyl tetrahydrofolate synthetase [fhs]) on DNA extracts from drilling mud, cuttings, and, in the case of mcrA, cores (Table T41). Because of time constraints, we were unable to check for dsrB and fhs presence in cores during the expedition, but we plan to do this soon after returning to our home laboratories.

Although mcrA was consistently below detection in DNA extracts from drilling mud, we were able to amplify PCR product of the right amplicon size for mcrA in two cuttings samples (Samples 337-C0020A-30-SMW and 61-SMW) and the interiors of two cores (Sections 16R-2 and 20R-3; Table T41). dsrB was not detectable in any drilling mud or cutting samples. fhs was detectable in most cuttings but below detection in drilling mud after 50 cycles of PCR amplification. After an additional 50 PCR cycles, however, amplicons of the right fragment size were also found in DNA extracts from drilling mud.

Our shipboard results from PCR assays on metabolic marker genes are further evidence that DNA belonging to indigenous microorganisms was extracted on board the ship. mcrA genes of probably methanogenic microorganisms were solely detected in core samples and cuttings—not in drilling mud. fhs genes were found to be in vastly higher abundance in cuttings compared to drilling mud. These results also suggest that, despite being highly contaminated with drilling mud, cuttings can provide potentially useful genetic information on anaerobic microorganisms inhabiting the deep subseafloor in Hole C0020A.

Potential sulfate reduction rates

A total of 28 WRCs were sampled for pSRR measurements, covering the entire drilling interval at Site C0020 (Table T42). For each sample depth, two sets of duplicate samples (A, B and C, D) were obtained. Additionally, a set of samples (A–D) was obtained from the pooled master sample (1950–2000 msbf). Sample volumes were ~5 cm3 and measured sample weights ranged from 4.4 to 9.8 g depending on density and moisture. A and B samples were incubated with N2 headspace, whereas 15 mL of CH4 (99.9%) were added via syringe to the N2 headspace of C and D samples, resulting in an increased pressure of ~2 bar. An additional set of samples (A, B, C, and D), obtained from mud tanks that were actively circulating drilling mud before sampling, were incubated as contamination controls.

To each sample, 3.7 MBq of 35S Na2SO4 (30 µL of aqueous solution) was added, except for Sample 337-C0020A-30R-3, 72–77 cm, where 7.4 MBq was added. For drilling mud samples, 2.1 MBq was added. The incubation time for all samples incubated before 11 September 2012 was 10 days. Sample 30R-2, 72–77 cm, and the drilling mud samples were incubated for 9 and 6 days, respectively. Samples obtained from the pooled master sample (1950–2000 m CSF-B) were incubated for 5 days. Three incubation temperatures were chosen to account for increasing in situ temperature with depth. Temperature I (~25°C, room temperature) was used for samples from Cores 1R through 6R (1276.70–1495.00 m CSF-B), Temperature II (35°C, Incubator I) was used for samples from Cores 8L and 9R through 22R (1607.16–1977.84 m CSF-B), and Temperature III (45°C, Incubator II) was used for samples from Cores 23R through 32R (1984.13–2456.63 m CSF-B) (Table T42) and for the pooled master sample. Drilling mud samples were incubated at room temperature.

Incubation for shore-based cultivation of deep subseafloor microbes

Six unwashed samples of cuttings from 696.5 to 1206.5 m MSF and formation fluid obtained from the Quicksilver probe at 1279.5, 1844.0, and 1978.0 m WMSF were used as inoculum to enrich for anaerobically respiring microbes such as methanogens, homoacetogens, and ferric iron reducers. Formation fluid from 1489.3, 1808.0, and 1901.2 m WMSF and all WRCs obtained during the expedition, were stored under anaerobic conditions at 4°C for shore-based cultivation. After the shipboard incubation of unwashed cuttings, growing cells were confirmed based on culture turbidity and microscopic observations of enriched cells. We observed particularly high turbidity and many cells in media targeting iron reducers (i.e., media of ferric citrate, lepidocrocite, goethite, hematite, and magnetite) and homoacetogens (i.e., medium of H2/CO2 plus bromoethanesulfonic acid [BES]) (Fig. F65). In addition, there seemed to be cell growth with media targeting hydrogenotrophic and methylotrophic methanogens (i.e., media with H2/CO2 and methanol, respectively). Because these first-generation enrichment cultures likely contain carbon and energy sources originally present in unwashed cuttings, it is uncertain whether the targeted microbes grew on the substrates supplied in the media or substrates present in drilling mud. Continuous subculturing will be conducted on shore in order to remove the effects of substrates other than the ones supplied through media.

Using light microscopy, fungal sporelike structures were identified in incubated core samples (Sections 337-C0020A-8L-5, 10R-2, 13R-1, 14R-2, 15R-3, 15R-5, 20R-7, and 28R-3) and seawater (Table T43). Furthermore, based on size, morphology, and motility, several types of microorganisms were discriminated. Motile filaments were observed in high abundance in drilling mud (Sample 337-C0020A-245-LMW) and core Sections 7R-1, 15R-7, and 25R-2 and in low abundance in seawater and core Sections 13R-1, 15R-3, and 20R-7, suggesting origin from drilling mud. We will conduct shore-based PCR assays to target fungal DNA and discriminate between indigenous and contaminant communities.

Additional sampling for shore-based microbiological investigations

Single-cell analyses of carbon and nitrogen assimilation rates of subseafloor autotrophic and heterotrophic microbial communities

Six WRC samples and the pooled master sample were used for detecting incorporation of stable isotope-labeled substrates into whole cells by nanoscale secondary-ion mass spectrometry (NanoSIMS): intervals 337-C0020A-4R-1, 100–141.5 cm; 4R-2, 0–47 cm (1376 m CSF-B); 8L-4, 50–100 cm (1606 m CSF-B); 14R-1, 40–80 cm (1820 m CSF-B); 15R-3, 20–48 cm (1921 m CSF-B); 18R-1, 71–114 cm (1946 m CSF-B); 25R-2, 50–60 cm; 26R-4, 40–140 cm (2114 m CSF-B); and 32R-4, 70–141 cm (2461 m CSF-B). After whole-round cutting, samples were flushed with N2, vacuum-sealed, and stored at 4°C. Utilizing aseptic techniques and within 24–48 h after retrieval, core surfaces were removed three successive times to remove contaminated outer parts—either within an aerobic clean bench or nitrogen glove bag, Sections were freshly packaged, reflushed with N2, vacuum-sealed, and stored at 4°C until inoculation.

Samples were obtained in an anaerobic glove box using a wide spatula and hammer to break cores into centimeter-sized pieces. Pieces were then divided evenly among 120 50 mL glass vials and sealed with butyl rubber stoppers. Within 24 h, samples were flushed with filtered Ar to remove H2 and N2 from headspace and stored at 4°C until substrate addition.

Vials were filled to achieve an estimated sample volume of 5–10 cm3 per incubation (depending on total sample volume). Substrates were added to establish concentrations of 13C (mixture of 15 µM each of 13C and natural-abundance isotope-ratio C-bearing substrates), 15N (mixture of 1.5 µM of 15N and natural-abundance isotope-ratio N-bearing substrates), and 20 vol% deuterium in water. Killed controls were autoclaved before any seawater or substrate was added. Samples were stored at 4°C on board the ship and during shipping to home institutions, where they were then incubated at near in situ temperatures.

Sandwich experiment

The coal WRC from Section 337-C0020A-18R-1 was prepared for shipboard stable isotope incubation. The section was dabbed with KimWipes soaked in anaerobic Milli-Q water in a glove box, and then gently broken into 1–2 cm thick sections to create artificial fracture surfaces. UV-sterilized 47 mm polycarbonate and cellulose acetate membranes were soaked with substrate and placed between core breaking points, and then “sandwiched” back together by wrapping them with Parafilm. Labeled substrates were added with 1 µM carbon and/or 0.1 µM nitrogen according to Table T10 in the “Methods” chapter (Expedition 337 Scientists, 2013b), with the exception of cellulose, of which a suspension containing 176 mg was placed on the polycarbonate membrane. “Sandwiches” were anaerobically incubated in the dark at 42°C without shaking.

Hydrogenase activity measurements

Samples for hydrogenase activity measurements were sampled from WRCs also used for pSRR incubations. The potentially contaminated outer 1 cm of 28 WRCs was carefully removed in an anaerobic glove box using sterilized spatulas and knifes. Approximately 50 cm3 was packed in ESCAL bags, flushed with nitrogen, vacuum-sealed, and stored at –80°C for shore-based tritium incubation experiments.


Data obtained during Expedition 337 demonstrate the suitability of PFC tracers to monitor contamination during riser drilling operations and indicate that the majority of core samples obtained have low levels of contamination at the core center. The successful extraction of cells and DNA demonstrates that the monitoring of microbial populations in cores obtained by riser drilling is possible on board the ship. Community fingerprinting analyses suggest that a substantial fraction of the DNA detected within cores belongs to indigenous communities of microorganisms. Although high cell numbers found in sandy and less consolidated layers were almost certainly the result of sample contamination by drilling mud, cell numbers only slightly above the detection limit in other cores—all the way to Core 337-C0020A-32R at 2460 m CSF-B—are less likely to result from drilling mud contamination. Shore-based molecular analyses will reveal the extent to which cells and DNA detected indeed belong to in situ microbial communities, and, if so, what their metabolic potential is based on functional gene and metagenomic analyses.

A broad range of experiments investigating microbial community zonation and activity in Expedition 337 core samples was initiated on board the ship. In the upcoming months and years, these experiments will produce novel insights to the life histories of subseafloor microbes in the deep core samples and coalbeds at Site C0020. The focus on combined nanoSIMS and single-cell approaches is likely to provide altogether novel insights to the modes of biomass assimilation and energy production in these fascinating, deeply buried microbial communities from the deepest borehole sampled by scientific drilling to this date.