IODP Proceedings    Volume contents     Search
iodp logo


Geochemistry and microbiology

The objectives of shipboard geochemical and microbiological sampling during Expedition 307 were to determine the stratigraphic distribution of zones of active microbial reaction and to identify stratigraphic intervals characterized by relatively high rates of hydrocarbon fluid flow. These objectives were achieved through the development of high-resolution vertical profiles of interstitial water and headspace gas geochemistry and the acquisition of sediment samples for a wide suite of microbiological analyses. At Site U1316, depth profiles of dissolved sulfate and methane gas in Hole U1316A were used to establish a scheme for detailed microbiological and biogeochemical sampling and postcruise research in Hole U1316B. Because of drilling contingencies, detailed microbiological and biogeochemical sampling was carried out simultaneously with collection and shipboard analysis of interstitial water at Sites U1317 and U1318.

Interstitial water samples

Shipboard interstitial water samples were obtained from 5–20 cm long whole-round intervals that were cut on the catwalk, capped, and taken to the laboratory for immediate processing. In cases where the time required to obtain pore fluids exceeded the rate at which new cores arrived, interstitial water samples were stored in a refrigerator until processed. After extrusion from the core liner, the surface of each whole-round interval was carefully scraped with a spatula to remove potential contamination. Sediments were then placed in a titanium squeezer, which was modified after the standard stainless steel squeezer of Manheim and Sayles (1974). Interstitial water was passed through a prewashed Whatman number 1 filter fitted above a titanium screen, filtered through a 0.45 µm Gelman polysulfone disposable filter, and subsequently extruded into a precleaned (10% HCl) 50 mL plastic syringe attached to the bottom of the squeezer assembly. After interstitial water collection, the syringe was removed to dispense aliquots for shipboard and shore-based analyses.

Interstitial water analyses

Most interstitial water samples were analyzed for routine shipboard measurements according to standard procedures (Gieskes et al., 1991). Salinity was measured as total dissolved solids using a digital refractometer. The pH was determined by ion-selective electrode. Alkalinity was determined by Gran titration with a Metrohm autotitrator. The acidified alkalinity sample was saved for shore-based phosphate analyses.

Dissolved inorganic carbon was measured using a Coulometrics 5011 CO2 coulometer. An aliquot of 1.0 mL of interstitial water was pipetted into the reaction tube followed by addition of 3.0 mL of 2N HCl after attaching the reaction tube to the coulometer apparatus. The liberated CO2 was titrated, and the end point was determined by a photodetector. Measured concentrations were corrected for the value of the acid blank. Analytical uncertainty, based on repeated measurements of the International Association of the Physical Sciences of the Ocean (IAPSO) standard and reagent-grade calcium carbonate, was ±1%.

Concentrations of chloride and sulfate were determined by manual dilution and injection into a Dionex DX-120 ion chromatograph. Chloride concentrations were also determined by titration with AgNO3. Quantification was based on comparison with IAPSO standard seawater.

Dissolved silica and ammonium concentrations were determined by spectrophotometric methods using a Shimadzu UV Mini 1240 spectrophotometer. Concentrations of Fe, Mn, Ca, Mg, Sr, B, Li, and Ba were determined using the Jobin-Yvon Ultrace inductively coupled plasma–atomic emission spectrometer (Murray et al., 2000). Analytical standards for all elements were then prepared by analyzing mixtures of this master standard and seawater.

Gas analyses

Concentrations of methane through propane hydrocarbon gases were monitored at intervals of 1–2 samples per core. The standard gas analysis program for safety and pollution prevention purposes (Kvenvolden and McDonald, 1986) was complemented by additional headspace analyses following a slightly different approach (Iversen and Jørgensen, 1985; Hoehler et al., 2000; Shipboard Scientific Party, 2003) with the intent to better constrain the concentrations of dissolved gases. Compared to the rapid safety-oriented protocol, which measures methane dissolved in interstitial water, the latter, more time consuming alternative, measuring the additional dissolution of methane adsorbed onto sediment, provides an estimate of total methane.

For the required safety analysis, a 3 cm3 bulk sediment sample from a freshly exposed end of a core section was collected upon core removal using a brass boring tool or plastic syringe and then extruded into a 20 mL headspace vial and immediately capped with a silicone/polytetrafluoroethylene (PTFE) septum, which was sealed with an aluminum crimp cap. The vial was then heated to 60°C for ~20 min prior to analysis.

For samples designated for refined headspace analysis, a 5 cm3 sediment sample was collected from a freshly exposed end of a core section using an open-ended plastic syringe. The sample was collected by penetrating the sediment surface while the plunger was maintained at the sediment surface to prevent contamination from atmospheric gases or trapped air bubbles. After sampling, the syringe was extruded until 3 cm3 of sample remained and the excess was shaved off with a flat spatula flush with the end of the syringe barrel to provide an accurate estimate of the sediment volume within the syringe. The remaining 3 cm3 sediment sample in the syringe was extruded into a 20 mL vial containing 5 mL of 1M NaOH. The vial was immediately capped with a silicone/PTFE septum and an aluminum crimp cap. After vigorous manual shaking for 2 min, the vials were shaken automatically for an additional hour and subsequently left to stand for at least 23 h at room temperature prior to analysis by gas chromatograph (GC).

GC analyses of headspace samples for both safety and refined protocols were performed in an identical manner. A 5 mL volume of headspace gas was extracted from the sealed sample vial using a standard gas syringe and directly injected into the GC. The headspace gas samples were analyzed using the GC3 chromatograph, a Hewlett Packard 5890 II Plus GC, equipped with an 8 ft × ⅛ inch stainless steel column packed with HayeSep S (100–120 mesh) and a flame ionization detector (FID). Some samples were analyzed for higher molecular weight hydrocarbons by injection into the natural gas analyzer, a modified Hewlett Packard 5890 II Plus GC with an FID and a thermal conductivity detector. Concentrations of methane, ethane, ethene, propane, and propene were obtained. The carrier gas was helium, and the GC oven was programmed from 100°C (5.5 min hold) to 140°C (4 min hold) at a rate of 50°C/min. Data were processed using a Hewlett-Packard 3365 program.

The safety methane concentrations (ppm) were used to derive the concentration of dissolved methane (µM) as described by equation 1 below. Adsorbed methane (µmol/g) was calculated by subtracting the dissolved methane concentration from the total methane concentration (µM) as determined by the refined (NaOH) protocol and normalizing to mass using density and porosity relationships.

The concentration of dissolved methane, both in the safety and refined protocols, was derived from the headspace concentration by the following equation:

CH4 = [(χM – χbkg) × Patm × VH]/(R × T × ϕ × VS), (1)


  • VH = volume of the sample vial headspace,
  • VS = volume of the whole sediment sample,
  • χM = molar fraction of methane in the headspace gas (obtained from GC analysis),
  • χbkg = molar fraction of methane in headspace gas because of background,
  • Patm = pressure in the vial headspace (assumed to be the measured atmospheric pressure when the vials were sealed),
  • R = the universal gas constant,
  • T = temperature of the vial headspace in degrees Kelvin, and
  • ϕ = sediment porosity (determined either from moisture and density measurements on adjacent samples or from porosity estimates derived from gamma ray attenuation [GRA] data representative of the sampled interval).

Sediment analyses

Only limited analyses were carried out on sediment samples because of the short duration and emphasis on interstitial water and gas geochemistry of Expedition 307. To enable determination of carbonate mass accumulation rates, samples taken immediately adjacent to those used for determining dry bulk density were measured for inorganic carbon content using a Coulometrics 5011 CO2 coulometer. A total of ~10–15 mg of freeze-dried, ground sediment was weighed and reacted with 2N HCl. The liberated CO2 was titrated, and the end point was determined by a photodetector. Calcium carbonate concentration, expressed as weight percent, was calculated from the inorganic carbon content, assuming that all evolved CO2 was derived from dissolution of CaCO3, by the following equation:

CaCO3 (wt%) = 8.33 × inorganic carbon (wt%). (2)

No correction was made for the presence of other carbonate minerals such as dolomite. Analytical uncertainty, based on repeated measurements of reagent-grade calcium carbonate, was ±1%.


Core handling and sampling


Microbiological sampling depends on careful and appropriate sample handling techniques. Precise operational definitions for special microbiology handling terminology, such as “clean” or “sterile” are given in the “Explanatory Notes” chapter of the Leg 201 Initial Reports volume (Shipboard Scientific Party, 2003). Because the samples were retrieved from stable sedimentary environments, the prokaryotes are expected to be sensitive to chemical and physical change, in particular to changes in oxygen, temperature, and pressure. Consequently, all samples for microbiology and process studies were transferred from the drilling platform to the hold refrigerator (set to <10°C) as quickly as possible and were kept as whole-core sections until processed. In order to avoid intermittent warming of retrieved cores, IODP’s usual core handling procedure was modified. Efforts were also made to obtain APC cores, even when this led to an increase in core recovery times, as APC cores are generally much less disturbed than XCB cores. RCB cores were used for deeper sections of the sites with partially-lithified to lithified sediments.

In order to ensure that we were indeed analyzing the indigenous prokaryotes and their activities, tests for contamination were conducted during the entire coring process for microbiological samples. Contamination tests were conducted in holes where microbiological sampling occurred using solutes (perfluorocarbon tracer [PFT]) or bacterial-sized particles (fluorescent microspheres) to check for potential intrusion of drill water from the periphery toward the center of the 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., 2000a). 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 sampled further.

Sampling on the catwalk

A limited number of microbiological and related biogeochemical samples were collected on the catwalk as soon as the core was retrieved. After the core was cleaned, the core was visually inspected for signs of disturbance, such as gas voids, cracks, and drilling disturbance. One pair of sections was cut as a 2 m section and a 1 m section and the former was used for subsequent microbiological sampling. Two further sections were selected for a specific sampling request taken three times per core. The top ends of the selected sections were cut and capped (without acetone). The bottom 10–20 cm of the 2 m section was used as an interstitial water biogeochemistry sample. Samples for total prokaryotic cell counts and, where performed, PFT/​fluorescent microsphere contamination checks were immediately collected using 5 cm sterile syringes from the lowermost, freshly cut end. Samples for headspace methane, δ13C of methane, and analysis of higher hydrocarbon gases were taken from the same core section or from the adjacent core top of the next section. The lowermost core end was then sealed with an end cap (without acetone). The microbiological section was quickly transported to the cold room to limit temperature increase. At Site U1318, sections destined for microbiological sampling were first run through the Fast Track MSCL. This 5 to 10 min delay allowed data for correlation purposes to be obtained before the material was sampled.

Whole-round core sampling in the cold room

A considerable proportion of the microbiological work was to be shore based because of time constraints and the necessity to use dedicated, specialized laboratory facilities. Consequently, a number of samples were taken as whole-round cores (WRCs) and stored until after the cruise. Keeping samples cool, processing times short, and minimizing contamination were key criteria for determining how the core sections were taken. To minimize changes in the microbial population, all handling took place in a cold room. The lower refrigerated core room on the hold deck of the ship served as a cold room at <10°C and was equipped with a work bench and working space for two to four persons.

Table T4 shows the cold room sample processing and cutting. For processing convenience, some of the physical property samples (e.g., oedometry and microtomography) were also sectioned in the cold room. The subsectioning equipment was the standard IODP core cutter coupled with a clean wire or blade. Samples were taken for measurement of bacterial activities (anaerobic oxidation of methane, sulfate reduction, methanogenesis, thymidine incorporation, and hydrogenase activity), capped, packed in a gas-tight aluminum bag in a nitrogen atmosphere with an oxygen scrubber sachet (Merck “Aerocult A”), and stored at 4°C. Samples for fluorescence in situ hybridization and virology were collected and stored in the same way. Samples for gas analysis were cut and sealed in a tin with Milli-Q water with the addition of 20 mL of 1% sodium azide to prevent bacterial activity. The samples for DNA, lipid biomarker, and amino acid analyses were capped, placed in polyethylene bags, and frozen in a –80°C freezer. Samples for oedometry and microtomography were capped and stored at 4°C. Standard IODP core end-cap color codes were maintained with the top of the section blue and the base of the section orange. Cut WRC samples were stored with a clear upper cap and orange lower cap indicating orientation, and the base of the residual core was capped with an orange cap before return to the core laboratory for reintroduction into the standard core handling process.

Total cell counts

The most immediate method to visualize and quantify the deep biosphere is total prokaryotic cell counts using the nucleic acid stain acridine orange. These counts have been made on a wide range of ODP sediment cores, including cores from the Peru margin and the equatorial Pacific (Parkes et al., 1994; D’Hondt et al., 2004). In general, these counts have demonstrated an exponential decrease of prokaryotic cells with depth. The method detects sediment layers of increased cell density that often coincide with particular geochemical conditions that are conducive to prokaryote growth (Parkes et al., 2000). The AODC enumeration method was used at all sites during this leg.

Procedures and protocols

Potentially contaminated sediment was removed with a sterile scalpel. A 1 cm3 minicore was then taken with a sterile 5 mL plastic syringe. The syringe was sealed with a sterile stopper. In a clean area of the laboratory, the 1 cm3 plug was extruded into a sterile serum vial containing 9 mL of 2% (v/v) filter sterilized (0.2 µm) formaldehyde in 3.5% NaCl. The vial was crimped and shaken vigorously to disperse the sediment particles.

Total prokaryotic cell numbers and numbers of dividing or divided cells were determined using acridine orange as a fluorochrome dye with epifluorescence microscopy (Fry, 1988). Fixed samples were mixed thoroughly, and a 5–20 µL subsample was added to 10 mL of 2% (v/v) formaldehyde and 2% (v/v) acetic acid filter-sterilized (0.1 µm) in 3.5% NaCl. Acetic acid dissolves a substantial amount of carbonate, allowing larger samples to be processed, therefore increasing accuracy and giving a lower detection limit. 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 removed on a 0.2 µm black polycarbonate membrane. Excess dye was flushed from the membrane by rinsing with a further 10 mL aliquot of 2% (v/v) filter-sterilized formaldehyde plus 2% (v/v) acetic acid in 3.5% NaCl, and 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 100 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× oculars. Prokaryote-shaped fluorescing objects were counted, with the numbers of cells on particles doubled in the final calculation to account for masking by sediment grains. The detection limit for prokaryotic cells was estimated at 1 × 105 cells/cm3 (Cragg, 1994).

The percentage of cells involved in division has been suggested as an indication of growth, although the assessment of dividing cells has never had a standardized approach in the literature. Dividing cells were defined operationally as those having clear invagination. A divided cell is operationally defined as a visually separated pair of cells of identical morphology. The percentage of cells involved in division is then calculated as follows:

Percentage of cells involved in division = [number of

dividing cells + 2 (number of divided cell pairs)] ×

100/total number of prokaryotic cells.

Perfluorocarbon tracer contamination tests

In each hole chosen for microbiological subsampling, PFT was continuously fed into the seawater drill fluid at a tracer concentration of 1 mg/L seawater drill fluid. Concentrations of PFT were measured in all sections used for microbiological studies. A 5 cm3 subcore from the same section was routinely taken, as described by Smith et al. (2000a). Air samples were occasionally taken to monitor the ambient concentration of PFT on the catwalk. The concentrations of PFT at the outer periphery of the cores and in the drill fluid were measured to verify delivery of the PFT. During APC, the concentration of PFT was measured in a syringe sample taken adjacent to the core center. During XCB coring and RCB drilling of harder sediment, chunks of intact core adjacent to the center of the core were collected for PFT measurement, thus providing data on the minimal size and quality of intact core pieces that could be confidently sampled for microbiological investigations.

To measure PFT concentrations, we used a HP-PLOT/AL203 “S” deactivated column with film thickness = 50 µm, length = 15 m, phase ratio = 12, and column internal diameter = 0.25 mm. The inlet temperature was 180°C with 10 psi, the detector temperature was 250°C, and the column temperature was 100°C for 8 min and then ramped up 50°C/min to 200°C. The PFT peak was at a retention time of 5.7 min. We used a 1 mL injection. Larger injections resulted in loss of material.

Procedures refined during Leg 201 (Shipboard Scientific Party, 2003) were employed. The sample headspace vial was first baked at 80°C for 10 min. Clean nitrogen gas was injected onto the column to ensure that no PFT peak resulted from residual PFT in the syringe or in the GC. After a clean run was achieved, the sample was injected using a 1 mL plastic syringe. A new 1 mL syringe was used for each sample. For best results, background air samples were taken regularly from the same location used for capping headspace vials, ideally on the catwalk when no core was present.

Fluorescent microparticle tracer

The procedure for assessing particle contamination was adapted from that used during Leg 190 (Smith et al., 2000a, 2000b). A plastic bag with a suspension of micrometer-sized fluorescent spheres was positioned within the core catcher. The bead suspension was 30 mL containing 2.1 × 1011 microspheres, giving a concentration of ~7 × 106 microspheres/µL. The beads were released inside the core barrel as it hit the sediment, maximizing the effectiveness of the beads as tracers of potential bacterial contamination. A 5 cm3 subcore was routinely taken from the cut core-end adjacent to the interstitial water WRC, midway between the core center and the periphery. This sample was mixed with 15 mL of saturated sodium chloride solution and shaken on a wrist-action shaker to disperse the sediment plug. The suspension was centrifuged (Marathon 10K; 5 min; 2800 × g), to separate the microspheres (density = 1) from the sediment particles. The supernatant was filtered onto black polycarbonate filters (0.2 µm pore size). Fluorescent microspheres were counted under ultraviolet light, and data are reported as number of microspheres per cubic centimeters of sediment.

Evaluating contamination tests

When comparing the results of both contamination tests (the presence of beads and PFT concentration inside 5 cm3 subcores), one should consider the following. First, in contrast to the beads, PFT can travel through very small pore spaces and is found in the laboratory air and on the hands of anyone who has handled a core liner. Therefore, although its presence at high concentrations in a sediment sample (>0.1 ng PFT/g sediment) may suggest contamination, it is not necessarily an indication that microorganisms from the drilling fluid have in fact contaminated the sample. The PFT detection limit reported for Leg 201 sites was not set as a lower limit of the ability to detect PFT by the GC, but as a lower limit of ability to confidently assess the presence of PFT in real samples given the uncertainty inherent in subtracting background levels of PFT and the reliability of the integration of small GC peaks. The absence of PFT from a sample indicates that contamination by drill water has not occurred.

Second, whereas the number of microspheres (2.1 × 1011 microspheres/30 mL bag) deployed is equivalent to the number of bacteria in ~400 L of seawater (assuming 5 × 108 bacteria/L), microsphere deployment does not produce a uniform dispersion along the core. Although PFT can usually be found in sediment samples taken from the edge of cores, the same is not true for microspheres. At this point, without knowing the factors that control the final concentration and distribution of microspheres along the core barrel, one should consider the microspheres as a qualitative rather than a quantitative measure of contamination. The presence of microspheres in sediment samples away from the edge of a core is a strong indication that contamination by prokaryote-sized particles from the drilling water has occurred, although the absence of microspheres in the contamination sample is not proof of lack of contamination in adjacent samples. However, microspheres are inert and have a historical presence, thus, any researcher receiving samples can independently check for the presence of microspheres in their specific subsamples.