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Geochemistry and microbiology

The Costa Rica Seismogenesis Project (CRISP) was designed to understand the processes that control fault zone behavior during earthquake nucleation and rupture propagation at erosional subduction zones. The first phase of this project focuses on sampling sediments, fluids, and crustal rocks to fully characterize the eroding material before subduction. Fluids and associated diagenetic reactions are a key component of this study, as they affect hydrological parameters (e.g., permeability and pore pressure) and may regulate the mechanical state of the plate interface at depth. The concentration of dissolved species and their isotopic composition provide critical data for the identification of fluid sources, fluid-rock reactions, pathways of fluid migration, and plumbing of the system. In addition, geochemical data can help characterize the subsurface biosphere and aid in constraining mass balance inventories operating in this subduction zone. To this aim, the geochemical and microbiological sampling and analyses were coordinated so as to generate an integrated plan. Whereas the sampling frequency varies among sites (as described in each site chapter), the sampling package consisted of 5–40 cm long whole rounds for interstitial water and a 2 cm long whole round for a cluster of sediment analyses (Fig. F13). The material from the cluster sample was divided for shipboard and shore-based geochemical and physical property studies. Two routine sediment plugs were collected adjacent to each interstitial water sample for headspace analyses; one was used for standard hydrocarbon concentration monitoring on board and the other was for stable isotope measurements at onshore laboratories. Whole-round samples for microbiological studies were collected only in selected cores; these 5 cm whole rounds were cut adjacent to the interstitial water samples.

Fluid inorganic geochemistry

Interstitial water collection

For interstitial water analyses, whole-round cores were cut on the catwalk, capped, and taken to the laboratory for processing. Samples collected from 0 to 64 mbsf at Sites U1378 and from 0 to 39 mbsf at Site U1379 were processed inside a nitrogen bag to avoid oxidation of redox-sensitive elements. Whole-round samples designated for He isotopic analysis were cut on the catwalk and transferred into a plastic sealable bag initially flushed with ultrahigh purity (UHP) N2. The sample was immediately transferred to a small processing and squeezing station set up on the upper ‘tween deck of the JOIDES Resolution. This deck of the ship is a He-free environment, whereas the shipboard Chemistry Laboratory uses He as a carrier gas for the gas chromatographs. The sample was cleaned, squeezed, and transferred into the copper tubing in a UHP N2 glove bag. All other cores were processed under normal atmospheric conditions. During high-resolution sampling, when there were too many interstitial water cores to process immediately, capped whole-round core sections were stored under a nitrogen atmosphere at 4°C until they were squeezed, which occurred no later than 24 h after core retrieval.

After extrusion from the core liner, the surface of each whole-round interstitial water core sample was carefully scraped with a spatula to remove potential contamination from seawater and sediment smearing in the borehole. In APC cores, ~0.5 cm from the outer diameter, top, and bottom faces were removed, whereas in the XCB and RCB cores, where borehole contamination is higher, as much as two-thirds of the sediment was removed from each whole round. The remaining sediment (~50–300 cm3) was placed into a titanium squeezer, modified after the stainless-steel squeezer of Manheim and Sayles (1974). Gauge pressures up to 30 MPa were applied using a laboratory hydraulic press to extract interstitial water. Interstitial water was passed through a prewashed Whatman No. 1 filter fitted above a titanium screen, filtered through a 0.2 µm Gelman polysulfone disposable filter, and subsequently extruded into a precleaned (10% HCl), plastic syringe attached to the bottom of the squeezer assembly. In most cases, 25–55 mL of interstitial water was collected from each sample after 20–40 min of squeezing. In the deeper sections of the sites, fluid recovery was as low as 0.5 mL after squeezing the sediment for as long as 3 h.

We undertook key biogeochemical analyses on the ship and collected appropriate interstitial water subsamples for critical categories of postcruise studies (Table T4); sample allocation was determined based on the interstitial water volume recovered and analytical priorities based on the objectives of the expedition. Because of time constraints imposed by a short expedition with high recovery and the importance of fully constraining the fluid regime at this site, we focused our efforts on collecting samples for postcruise studies and only a limited number of analyses were carried out onboard. Interstitial water subsamples were collected in glass vials for shore-based analyses of halogens, isotopic characterization of the interstitial water (oxygen and hydrogen), and dissolved metabolites (e.g., bicarbonate, sulfate, and sulfide). In addition, interstitial water subsamples were collected for analyses of dissolved volatile fatty acids, dissolved organic carbon (in glass vials and frozen), and noble gases (in copper tubing); sulfur isotope studies (fixed with 5% ZnAc); and minor and trace metal constituents and their isotopes (acidified with ultrapure nitric acid and stored in plastic vials). Shipboard analytical protocols are summarized below.

Shipboard interstitial water analyses

Salinity, alkalinity, and pH were measured immediately after squeezing, following the procedures in Gieskes et al. (1991). Salinity was measured using a Reichert temperature-compensated handheld refractometer. pH was measured with a combination glass electrode, and alkalinity was determined by Gran titration with an autotitrator (Metrohm 794 basic Titrino) using 0.1 N HCl at 20°C. Certified reference material 104 obtained from the laboratory of Andrew Dickson, Marine Physical Laboratory, Scripps Institution of Oceanography (USA), was used for calibration of the acid. International Association for the Physical Sciences of the Oceans (IAPSO) standard seawater was used for calibration and was analyzed at the beginning and end of a set of samples for each site and after every 10 samples.

Subsamples for shore-based sulfate analysis were treated with 5 µL of 10% ZnAc per milliliter of analyte immediately after collection to precipitate ZnS. Sulfate concentrations were determined shipboard on the inductively coupled plasma–atomic emission spectrometer (ICP-AES), as detailed below, as well as with a Dionex ICS-3000 ion chromatograph, which is a reagent-free system combining automated eluent generation and self-regenerating suppression with a conductivity detector. The samples were run using a 1:100 dilution of interstitial water sample with Nanopure-grade water (18.2 MΩ). A standard curve prepared using IAPSO dilutions was run daily, and three IAPSO standards were run after every 10 samples to quantify long-term reproducibility and as quality assurance for each run.

Chloride concentrations were determined using silver nitrate (AgNO3), as detailed by Gieskes et al. (1991).

Ammonium concentrations were determined by the indo phenol blue method using a Milton Roy Spectronic 301 spectrophotometer equipped with a Milton Roy “Mr.Sipper” sample introduction system (Gieskes et al., 1991).

Major (Ca, Mg, K, and Na) cations and sulfate concentrations were analyzed by ICP-AES with a Teledyne Prodigy high-dispersion ICP spectrometer. The general method for shipboard ICP-AES analysis of samples is described in ODP Technical Note 29 (Murray et al., 2000) and the user manuals for new shipboard instrumentation. Samples and standards were diluted 1:200 using 2% HNO3. Each batch of samples run on the ICP spectrometer contains blanks and solutions of known concentrations. Each item aspirated into the ICP spectrometer was counted four times from the same dilute solution within a given sample run.

Following each run of the instrument, the measured raw-intensity values were transferred to a data file and corrected for instrument drift and procedural blank. If necessary, a drift correction was applied to each element by linear interpolation between the drift-monitoring solutions. For many of the runs there was no unidirectional instrumental drift >1% (total, through the entire run of multiple hours), and in such cases no drift correction was employed. After drift correction (where appropriate) and blank subtraction, a calibration line for each element was calculated using the results from the analyses of known solutions. The calibration lines were strongly linear and concentrations deviated only slightly from seawater concentrations. Therefore, for these elements the final concentrations were calculated on the basis of the ratio to the analysis of IAPSO seawater standard. IAPSO seawater standard was measured at least four times through a run (and as many as eight times) each time as an unknown; therefore, it was straightforward to use some IAPSO samples for precise determination of the ratio and other IAPSO samples to provide an independent check on the resultant accuracy and precision.

Fluid organic geochemistry

Routine analysis of hydrocarbon gas in sediment cores is a part of the standard IODP shipboard monitoring of the cores to ensure that the sediments being drilled do not contain greater than the expected amount of hydrocarbons. The most common method of hydrocarbon monitoring used during IODP expeditions is the analysis of gas samples obtained from either core samples (headspace analysis) or from gas expansion pockets visible through clear plastic core liners (void gas analysis), following the procedures described by Kvenvolden and McDonald (1986).

When gas pockets were detected, the free gas was drawn from the sediment void using a syringe attached to a hollow stainless-steel tool used to puncture the core liner and the void gas was analyzed on the natural gas analyzer (NGA). For headspace analyses, a 3 cm3 bulk sediment sample was collected from the freshly exposed end of a top core section and next to the interstitial water sample, immediately after core retrieval, using a brass boring tool or plastic syringe. The sediment plug was capped with a gray butyl rubber septum and sealed with an aluminum crimp cap. The vial was then heated to 70°C for ~30 min to evolve hydrocarbon gases from the sediment plug. When consolidated or lithified samples were encountered, chips of material were placed in the vial and sealed. For gas chromatographic analysis, a 5 cm3 volume of headspace gas was extracted from the sealed sample vial using a standard gas syringe and analyzed by gas chromatography.

The standard gas analysis program for safety was complemented by collecting an additional headspace sample (same resolution as described above but labeled as NZ) to measure the stable carbon and hydrogen isotope composition at onshore laboratories. The sampling method is the same as that used for the safety analysis, except that the sediment plug is extruded into a 20 cm3 headspace glass vial filled with 10 cm3 of 10% KCl solution containing borosilicate glass beads, immediately capped with a gray butyl rubber septum, and sealed with an aluminum crimp cap. The vial was then vigorously shaken to help dissociate the sediment. Potassium chloride is toxic and was thus used to stop all microbial activity in the sediment. The glass beads (3 mm diameter) were used to help break up the sediment plug during shaking and liberate gas trapped in sediment pore space or adsorbed on particles. The vials were flushed with helium and capped within 1 h prior to sampling in order to remove air from the headspace and ensure the sample is preserved anaerobically. The objective is to preserve CO2 from the interstitial water for stable carbon isotope analysis.

Headspace, NZ, and void gas samples were directly injected into the gas chromatograph–flame ionization detector (GC-FID) or into the NGA. The headspace samples were analyzed using an Agilent/HP 6890 Series II gas chromatograph (GC3) equipped with a 2.4 m × 3.2 mm stainless steel column packed with 100/120 mesh HayeSep R and a FID set at 250°C. The GC3 oven temperature was programmed to hold for 0.5 min at 80°C, ramp at 30°C/min to 100°C, ramp at 15°C/min to 110°C, and remain at 110°C for 4.5 min before ramping at 50°C/min to 150°C, with a final holding time of 1.8 min. Helium was used as the carrier gas. The GC3 system determines concentrations of methane (C1), ethane (C2), ethene (C2=), propane (C3), and propylene (C3=). Alternatively, concentrations of C1–C6 hydrocarbons as well as nonhydrocarbons N2, O2, and CO2 were measured using the NGA system. For hydrocarbon analysis, the NGA consists of an Agilent/HP 6890 Series II NGA equipped with an Agilent DB-1 dimethylpolysiloxane capillary column (60 m × 0.25 mm diameter × 0.25 µm film thickness) fitted with a FID and using helium as carrier gas (constant flow of 2 mL/min). The gas chromatograph oven temperature was programmed to hold for 2 min at 50°C, ramp at 8°C/min to 70°C, and then ramp at 25°C/min to 200°C with a final holding time of 5 min. The FID temperature was 250°C. For nonhydrocarbon gases, thermal conductivity detector (TCD) separation used three columns: a 6 ft × 2.0 mm internal diameter stainless steel column (Poropak T; 50/80 mesh), a 3 ft × 2.0 mm internal diameter stainless steel molecular sieve column (13X; 60/80 mesh), and a 2.4 m × 3.2 mm internal diameter stainless steel column packed with 80/100 mesh HayeSep R (Restek).

Data were collected using the Hewlett Packard 3365 Chemstation data processing program. Chromatographic response is calibrated to nine different gas standards with variable quantities of low molecular weight hydrocarbons, N2, O2, CO2, Ar, and He and checked on a daily basis. The gas concentrations for the required safety analyses are expressed as component parts per million by volume (ppmv) relative to the analyzed gas. The internal volumes of 15 representative headspace vials were carefully measured and determined to average 21.5 ± 0.18 mL. This volume was taken as a constant in calculations of gas concentrations.

The volumetric units were converted to millimolar concentration units (mM) to facilitate comparisons with dissolved interstitial water constituents as follows:

CH4 = χM × Patm × VH × R–1 × T–1 × ϕ–1 × 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 gas chromatograph analysis),

  • Patm = pressure in the vial headspace (obtained from the bridge),

  • R = the universal gas constant,

  • T = temperature of the vial headspace in degrees Kelvin, and

  • ϕ = sediment porosity (determined either from moisture and density [MAD] measurements on adjacent samples or from porosity estimates derived from gamma ray attenuation [GRA] data representative of the sampled interval).

Sediment geochemistry

To complement the interstitial water analyses, a cluster of subsamples was collected from a single whole round adjacent to the interstitial water whole round for sediment characterization. Shipboard measurements of cluster samples included geochemical analysis following procedures described below, as well as XRD, porosity, and bulk density analysis following procedures described in “Lithostratigraphy and petrology” and “Physical properties.” In addition, sediment subsamples were taken from the cluster for shore-based studies that include cation exchange capacity, biomarkers, major and minor element chemistry, high-pressure and -temperature experiments, and isotopic characterization of iron and molybdenum (Table T5).

For the shipboard sediment geochemistry, 5 cm3 of sediment was freeze-dried for ~24 h, crushed to a fine powder using a pestle and agate mortar, and subsampled to analyze total carbon, total inorganic carbon and total nitrogen.

Elemental analysis

Total carbon and total nitrogen of the sediment samples were determined with a ThermoElectron Corporation FlashEA 1112 CHNS elemental analyzer equipped with a ThermoElectron packed column CHNS/NCS and a TCD. Approximately 10–15 mg of freeze-dried, ground sediment was weighed into a tin cup and the sample was combusted at 900°C in a stream of oxygen. The reaction gases were passed through a reduction chamber to reduce nitrogen oxides to nitrogen and were then separated by the gas chromatograph before detection by TCD. All measurements were calibrated to a standard, sulfanilamide, which was run every five samples. The detection limit was 0.001% for total nitrogen (instrument limit) and 0.002% for total carbon (procedural blank, measured as an empty tin cup).

Total inorganic carbon was determined using a Coulometrics 5011 CO2 coulometer. Approximately 10–15 mg of freeze-dried, ground sediment was weighed into a glass vial and acidified with 2M HCl. The liberated CO2 was titrated, and the corresponding change in light transmittance in the coulometric cell was monitored using a photodetection cell. The weight percent of calcium carbonate was calculated from the inorganic carbon (IC) content using the following equation:

CaCO3 (wt%) = IC (wt%) × 100/12.

NIST-SRM 88b (Standard Reference Material) was used to confirm accuracy. Standard deviation for the samples and standards is less than ±0.1 wt%. Total organic carbon content was calculated by subtraction of inorganic carbon from total carbon.


Core handling and sampling

For all microbiological analysis conducted for this expedition, it was imperative to use proper, careful handling techniques. Subseafloor microorganisms are expected to be sensitive to chemical and physical changes caused by the relatively undisturbed conditions to which they are accustomed. Hence, great care was taken to avoid any unnecessary disturbance of the samples prior to analysis. After sectioning of the core, whole-round subsamples were cut with a sterile spatula—or for highly compacted samples, separated from the rest of the core using a sterile chisel and hammer—capped, and taken to the Microbiology Lab for sampling. In the lab, all samples were immediately transferred to a cold storage room (<10°C) until further processing. Samples destined for oxygen-sensitive analyses were transferred into an anaerobic chamber within the cold room as soon as possible until processed. Whole-round samples were further subsampled for different microbiological methods. In all cases, sterile technique was a primary concern. Prior to subsampling, the whole-round sample was cleaned by removing the top ~1 cm of sediment with a flame-sterilized spatula. Immediately after cleaning, sterile, cut-off syringes (minicores) were pushed into the fresh surface of the whole round and samples were removed and stored as required for the different analyses. When sediment became too compacted to push the syringes in, subsamples were removed with a sterile spatula or chisel into Falcon tubes. The outer ~1 cm of the whole round, which was against the nonsterile core liner, was not used for subsampling. Minicore samples destined for shore-based DNA analysis were sealed into plastic bags and stored at –80°C for transportation to shore. Minicore samples for shore-based cultivation analysis were placed into polyester film bags, flushed three times with nitrogen, sealed under a nitrogen atmosphere to limit contamination of oxygen into the sample, and stored at 4°C. Minicores (3 cm3) taken for cell enumeration were extruded directly into a fixative solution containing 2% paraformaldehyde in sterile, filtered seawater. For these, only the middle 2 cm3 of the minicore was extruded into the fixative, to obtain the cleanest sample available.

Cell enumeration

The slurry produced by addition of sediment from the minicores or powdered samples to the fixative solution was shaken vigorously to disperse the sediment. A small aliquot of this slurry (50 µL) was filtered through a black 0.2 µm pore-size polycarbonate filter. Premixing of the aliquot with a few milliliters of sterile filtered seawater added to the filter tower apparatus ensured even distribution of the sample on the filter. The dry filter was then mounted on a glass slide with 20 µL of a staining solution and a glass coverslip. The staining solution consisted of SYBR Green I (1:40 dilution), glycerol, Vectashield mounting media, and phenylenediamine (1%) in a 3:3:3:1 volumetric ratio. Slides were incubated with the staining solution at room temperature in the dark for 1–2 h and stored in the freezer overnight before enumeration to reduce background fluorescence. Blanks were also prepared in the same way but omitting addition of sample. Cells were enumerated by the average number of SYBR Green I–stained particles in a microscopic field using an epifluorescence microscope (ZEISS Axioplan 2 imaging microscope), and images were taken with a ZEISS AxioCamHRc camera. The blanks provided an estimate of the level of background and/or contaminating cells, and final cell counts were adjusted accordingly.

Contamination analyses

During ODP Leg 201 (D’Hondt, Jørgensen, Miller, et al., 2003), contamination concerns for microbiological sampling of cores were addressed through the development of two separate tests for assessing contamination of the cores from the drilling processes: (1) the use of perfluorocarbon tracer added to the drilling fluid and (2) the use of microbe-size fluorescent microsphere beads introduced at the drill bit (Smith et al., 2000a, 2000b). For this expedition, contamination was assessed using the fluorescent microsphere test. Microspheres were deployed during XCB and RCB coring in any core from which microbiological samples were to be taken. Quantifying the number and lateral extent of microspheres introduced into the cores assessed contamination. Samples taken for this analysis were 3 cm3 minicores, when possible. When sediment was too compacted for minicores, samples were removed with a sterile spatula or chisel. All contamination samples were prepped in the same way as those for cell counting described above, except for the replacement of the staining solution with a simple solution of 1:1 phosphate-buffered saline and glycerol. Microspheres were enumerated by taking the average number in a microscopic field using an epifluorescence microscope. Microspheres were not deployed during APC coring because of concerns of physical damage to the core from the microsphere packaging; however, previous perfluorocarbon tracer tests of APC core have consistently shown that the centers of the APC cores are relatively pristine (House et al., 2003).

In addition to the shipboard checks, contamination will be further assessed postcruise by comparison of the microbial communities in the sediment samples with those in the drilling fluid using molecular methods. With these data, the genomic signatures of the contaminating drilling fluid microorganisms can sometimes be subtracted from those of the subseafloor communities.