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doi:10.2204/iodp.proc.310.103.2007

Geochemistry

Offshore interstitial water sampling

Effect of drilling procedure and core flow on pore water quality

Drilling was generally performed with the RCB. In order to minimize damage to the reef and improve the quality of the cores, cuttings were flushed from the pipe with seawater instead of drilling mud. Seawater from the ship’s firepump system was constantly supplied into the Seacore mud-pump container pool. The tank level was controlled by an overflow pipe that drained back into the sea. From this pool, seawater was pumped into the drill pipes with the pump system normally used for drilling mud. During the drilling process, seawater flushes the barrel from the outside and leaves the pipe through a small gap between the barrel shoe and the inner side of the drill bit. Thus, porous or coarse sediments are expected to contain pore waters that consist mainly of exchanged seawater from the drilling process. Loose fine-grained material in cavities or large pores is expected to be flushed away or moved downward the core sequence.

Although plastic-lined core barrels were used, most of the water stayed in the liner until it was capped and sealed in the curation container. After several problems with jammed plastic liners, a test with split-spoon stainless steel liners revealed that the quality of the cores could be significantly improved with their use. Subsequently, an increasing number of cores (see remarks field in OffshoreDIS individual core runs) were drilled using this method, and the cores were transferred on the deck from the split-spoon stainless steel liners into a plastic liner that was cut on one side along its length. The plastic liner was eased open at the cut, and the core and the lower spoon half were inserted. After turning everything upside down so that the cut in the liner pointed upward, the stainless steel spoon was pulled out toward the top. This way, even fragile cores were transferred from split-spoon liners to plastic liners with minimum disturbance. From a geochemical point of view, however, this included the possibility of contamination because the stainless steel liners were reused and the plastic liners were opened. Care was taken to clean the split-spoons from the outside as they were driven out of the barrel with pressurized drilling water. Even so, small amounts of rust, grease, or other contaminants from the outside of the split-spoon may have been transferred to the inner wall of the plastic liner.

After microbiology and pore water sampling, the core was curated, sealed with tape, and refilled with seawater for physical property measurement. Only cores with obvious contents of loose sediments were not refilled. After physical property logging, refilled cores were drained and finally sealed. One of the tap-water outlets in the curation container was connected to the mud container pool so that the same water used for drilling was used to refill cores for physical property measurement. The seawater tap was left running 24 h a day to avoid alteration on the supply tube. During the first days of operation, seawater sampled with a clean bucket from over the ship’s side was used to refill the cores because the drill-water supply was not immediately operational. Because of the open Pacific location, the seawater around Tahiti is extremely clean and low in microbial activity.

One obvious alteration of the seawater used as drilling fluid is that some iron content was acquired in the mud pool, evidenced by brownish stains that developed in the sink after a few days of constantly running seawater. Both seawater and drill water from the mud container pool were sampled several times for reference.

Although this procedure has many potential problems and would perhaps be inappropriate for handling normal marine sediments, it preserved in situ drilling conditions for these very coarse and porous reef materials.

Pore water sampling

During Expedition 310, the priority was to sample good-quality, undisturbed coral reef sequences. In the Expedition 310 Scientific Prospectus, extraction of pore water was planned only for soft-sediment sequences beneath or within the reef sequences (Camoin et al., 2005). Generally, very few of these sediments were sampled. Most of the recovered material was very porous carbonates or coarse volcaniclastic material with cavities, which were flushed with seawater during the drilling process as described above. In some cases, short sequences of fine nonencrusted sediments were sampled that allowed extraction of pore waters.

Preparation

The pH electrode was calibrated, and rhizons were placed in a beaker with pure water. Sample vials were prepared with preservatives as described below.

Pore water sampling using rhizon samplers

For closed liners, a standard 3.8 mm diameter drill bit was used to drill a hole in the plastic liner. A spacer on the drill bit prevented it from going into the core material. The half-split liners were sampled through the split before sealing. Core catcher samples were sampled in a split liner after they were delivered from the drill floor. If necessary, a 2.5 mm wide stainless steel stick was used to prepare a hole in the sediment. A rhizon sampler was carefully pushed into the sediment and connected to a 50 or 20 mL disposable syringe. Because rhizons with female Luer adapters were used, no Luer-Luer adaptors were necessary. Vacuum was established by pulling the syringe plunger and keeping it open with a wooden spacer. In most cases, the syringe was filled after 30 min. If there was still pore water flow, the syringe was emptied into a 20 mL scintillation vial (Greiner, polypropylene) and reattached. If the pore water flow through the rhizon was slow, the syringe was taped to the liner in order to allow core-curation procedures to continue. Filtering was generally not necessary because the maximum pore width of the rhizons is 0.2 µm. The samples were refiltered with 0.45 µm disposable syringe filters (Nalgene 25 mm, nylon) if the rhizons turned out to be broken. Broken rhizons can easily be detected, as the vacuum can not be maintained when the porous tube is damaged. If detected early enough, these rhizons were replaced. When pore water collection was very slow, two rhizons were used in the same sampling interval to speed up the sampling procedure. Rhizons retain 130 µL of liquid when wet and are readied for use by being left in a beaker with pure water. Usually the first drops are discarded to avoid collecting this water and rinsing the system. However, at very low expected sample volumes it was decided to leave this water in the sample and allow for the dilution.

Sampling of exchange water

Because of the lack of proper pore water samples to test the equipment, drainage water from the core after physical property measurement was collected in a beaker, sampled with a disposable syringe, and filtered with 0.45 µm Nalgene nylon filters. Because the refill seawater had some time (generally >2 h) for diffusive exchange with smaller pores even from very porous cores, the measured data were recorded and entered into OffshoreDIS with the remark “exchange water after flushing and draining cores with drill water” and a sampling depth that covered the whole-core section length. Some indication of the in situ pore water conditions may be extracted from this procedure if the exchange waters significantly differ from the refill water.

Sampling with the Bremen squeezer

The Bremen pore water squeezer was set up and operational, but no suitable samples of stiff but compressible material were recovered that would have given better pore water extraction than with the rhizons.

Labeling

Both the syringes and sample vials were first hand-labeled with hole, core, section, and sampling depth information, and then the sample was entered into OffshoreDIS. The primary bar code sample label was used for the 20 mL scintillation vial. Samples were split and labeled using the following abbreviations:

• DW = drilling water (seawater) from the mud pool tank/seawater tap;
• SW = seawater taken from over the ship’s side; and
• TW = tap water from the ship’s freshwater supply.

Samples were measured for pH, alkalinity, ammonia, and chlorinity in the curation container. The results were entered into a worksheet to calculate alkalinities and both calibrated ammonia and chloride concentrations. The results, the total sample and split-sample volumes, and the type and amount of added preservatives were entered into OffshoreDIS. Split-sample labels from OffshoreDIS were glued to the appropriate vials and lined with transparent tape (Tesafilm). If label fields failed to print correctly, they were hand-corrected with a permanent marker. Samples other than pore water were only hand-labeled.

Sample splitting

If possible, sample splits were filled straight from the syringe. Where exact amounts were needed (alkalinity and sulfide splits), a 1000 µL Eppendorf adjustable pipette was used to transfer samples from the primary sample vial. Table T4 shows the sample-split priority.

Samples for isotope analysis were put into vials and sealed without headspace. The sample split for oxygen isotope analysis was taken only if a sufficient amount of sample was collected. Because the exact amount of acid added, and thus the sample dilution, is known, alkalinity splits were labeled and stored as well, especially when the total sample volume collected was low. Wheaton ampules were sealed by welding the glass with a small torch.

The minimum sample amount from which all offshore parameters were measured was 400 µL. In this case, pH and alkalinity were measured from a 185 µL sample split, and ammonia and chloride were measured from a 1 in 5 dilution (100 µL sample + 400 µL pure water). Because rhizons contain 130 µL of pure water when wet, this had to be considered as dilution of the original sample.

Pure water

Pure water was generated in the curation container from the ship’s tap water with a Seradest USF 3000 purifying cartridge. The conductivity of the water was controlled to be <0.1 µS (>10 MΩ) by a LFM C1 conductivity detector. For microbiological use, an Aquafine SL-10 A ultraviolet sterilization unit and a Seralplus 0.2 µm filter cartridge served as extra purifying steps. Pure water for laboratory use was produced in batches of 10 L and stored in a carboy. On 3 November 2005, the ship received new freshwater supplies, which turned out to have a much higher chloride content (>1000 ppm compared to 300 ppm Cl before). This was possibly due to seawater contamination in the tank of the supply ship. Whatever the cause, no change in the purified water quality was detected. On 11 November, the ship’s freshwater supply was switched to another tank, which had even lower chloride contents (25 ppm).

Sample temperature

In situ temperatures of the sample could not be measured. From logging data we learned that they differ little from the water temperature, which is between 26° and 28°C in the working area. During core retrieval from the barrel, there may have been significant warming on the deck for daytime samples (0900–1800 h) because of outside temperatures of up to 40°C in the shade. In the curation container, samples were thermally equilibrated to temperatures between 23° and 26°C. Apart from some air conditioning failures, the curation container temperatures were constant at 25° ± 1°C.

pH value

The pH value was measured with a Mettler Toledo InLab 423 microcombination glass electrode with a 3 mm tip. In 2 mL Eppendorf cups, the pH value and the alkalinity were determined from 1 mL of sample. When the sample volume was low, the amount needed could be reduced to 185 µL using a conical 1.5 mL Eppendorf cup. A constant reading was achieved by turning the vial around the electrode with a special magnetically driven vial holder rather than stirring the sample with a perflurotetrafluoroethylene (PTFE)-coated stir bar. The pH meter was calibrated twice a day using Merck Certipur pH buffer solutions (pH = 4.01, 7.01, and 10.03). Temperatures were measured prior to pH measurement and manually entered into the pH meter. The instrument shows the pH with a resolution of 0.001 pH units. The measurement has an accuracy of better than ±0.02 pH units.

Alkalinity

Alkalinity was determined by titration with 0.01M HCl. The equivalence point was detected by titrating a 1 mL sample with 0.01M HCl while controlling the pH value. Titration was stopped at pH < 3.9. An Accustep digital burette with a 5 mL tip was used for titration. With this setup, a minimum of 10 µL increments may be titrated at an accuracy of ±5 µL when using a small-diameter tube that is placed in the solution to be titrated. The algorithm used to calculate alkalinity accounts for the activity of seawater and dilution by the titration solution so that the results are stable for different endpoint pH values. The measurement has an accuracy of better than 0.2 mmol/L. For the titration, a 0.3 mm internal diameter PTFE tube from the digital burette was placed in the liquid before the titration started. Both the PTFE tube and the pH electrode were rinsed with pure water and carefully dried with lab tissues before the measurement. The magnetically driven rotating vial holder is described in more detail in the ESO curation container cookbook, and the algorithm is derived from Grasshoff et al. (1983).

Ammonia

Ammonia was detected using the PTFE tape gas separator technique as described in the curation container cookbook. With this technique, ammonia is stripped from a 100 µL sample by an alkaline carrier solution (0.2M Na citrate in 10 mM NaOH), passes a 200 mm × 5 mm PTFE membrane area as NH₃, and is redissolved as NH₄⁺ in an acidic solution (1 mM HCl). The NH₄⁺ causes a conductivity signal in the acidic carrier that is detected with an Amber Science 1056 conductivity meter with a model 529 temperature-compensated micro flow-through cell. The conductivity signal was recorded with a Knauer strip-chart recorder, and peak height was analyzed manually. This method is very precise and stable, practically insensitive to matrix changes, and shows a linear conductivity response for ammonia concentrations between 10 and 1000 µM. The detection limit is 5 µM, and accuracy is better than 2%. Generally, measurements were made with the original sample. Only if the sample volume was extremely low was the sample diluted to gain a sufficient amount of sample for analysis. Ammonia calibration standards were freshly prepared from a 5.56 mM (100 ppm) standard solution. In order to avoid unnecessary time for preparation of calibration standards and calibration prior to measurements, only one standard was measured to check instrument function and a proper calibration was performed only if samples showed a signal above the detection limit.

A 300–400 µL sample split was taken from the primary sample vial with a Hamilton 1000 µL precision glass syringe and injected onto a 100 µL loop with a Rheodyne high-pressure liquid chromatography (HPLC) valve. The valve was then opened to the carrier solution stream to start analysis. The Hamilton syringe was rinsed with pure water before and after analysis.

Chlorinity

Although chloride content is not an ephemeral parameter and may be measured months later without loss in precision, it is usually measured offshore because freshwater influence from meltwater, decay of gas hydrates, or submarine freshwater supplies may be of major scientific interest. Chloride content of samples was detected from a 100 µL split by titration with AgNO₃ using a KCrO₄/​K₂Cr₂O₇ indicator solution. A total of 5 mL of pure water and 100 µL of indicator solution were placed in a glass beaker on a magnetic stirrer with a white surface. Using an Eppendorf EDOS digital burette system with a 2 mL tip, 0.1M AgNO₃ was titrated into the solution until a color change from milky yellow to orange brown was detected visually. The instrument and operator titration factor was determined by repeated measurement of International Association of Physical Sciences Organizations (IAPSO) seawater standard (556.8 mM). The accuracy of this method is ±2%, which is in the range of the total variation of chloride contents in the samples, so it is recommended that the samples be remeasured onshore with a more precise method (i.e., ion chromatography).

Onshore interstitial water analyses

Using analytical equipment housed in the Department of Geosciences, Bremen University (www.geochemie.uni-bremen.de/​koelling/​index.html), aliquots of IW samples taken during drilling operations were analyzed for a suite of dissolved species to complement shipboard analyses. Detection limits and reproducibility of IAPSO seawater measurements are given for each analyte in Table T5.

Cations measured by inductively coupled plasma–optical emission spectrometry

Dissolved cations were measured using a PerkinElmer Optima 3300 R simultaneous inductively coupled plasma–optical emission spectrometry (ICP-OES). All samples used for these analyses were acidified directly following shipboard sampling with concentrated HNO₃. Samples were diluted 1:10 using 0.65% (0.14M) HNO₃ prepared with subboiling distilled HNO₃ and 18 MΩ water. First, sample aliquots were analyzed for major elements utilizing a cross-flow nebulizer for B (249.772 nm), Ca (317.933 nm), K (766.490 nm), Mg (279.077 nm), S (181.975 nm), Si (251.611 nm), and Sr (421.552 nm), measuring the intensity at each wavelength in triplicate. In a second run, the same sample aliquots were analyzed for trace elements using a CETAC USN 5000AT ultrasonic nebulizer for Al (396.153 nm), Ba (455.403 nm), Cd (228.802 nm), Co (228.616 nm), Cr (267.716 nm), Cu (324.752 nm), Fe (239.562 nm), Li (670.784 nm), Mn (259.372 nm), Mo (202.031 nm), Ni (231.604 nm), U (385.958 nm), V (310.230 nm), and Zr (343.823 nm), measuring the intensity at each wavelength in triplicate.

Calibration standards for the major elements were prepared with high-purity single-element standards (Spectrascan by Teknolab, Norway) using a 0.5M NaCl (Merck certipur) solution as the matrix, as shipboard salinity measurements had determined all samples had seawaterlike salinities. Calibration standards for the trace element analyses were prepared by multiple dilutions of a multielement standard (Merck Certipur multielement standard IV) prepared with a 0.6M NaCl solution as the matrix.

Bromide, chloride, and sulfate by ion chromatography

Bromide, chloride, and sulfate were measured using a Metrohm Advanced Compact IC 861 ion chromatography system with chemical suppression comprising a Metrohm 837 eluant degasser, a 4 mm × 100 mm MetrosepA Supp5 anion column, and a wide-range conductivity detector. The eluant used in this system was a 3.2 mM sodium carbonate/​1 mM sodium hydrogen carbonate solution. IW samples were diluted 1:150 with 18 MΩ water. Concentrations were determined by comparison to IAPSO seawater and dilutions of this standard.

Phosphate by photometry

Dissolved phosphate was analyzed by the molybdenum blue method using a Merck portable photometer SQ118. A 1 mL aliquot of sample was placed in a microcuvette, and 50 µL of ammonium molybdate solution and 50 µL of ascorbic acid solution were added. Samples were shaken and placed in the dark for 10 min and then analyzed at a wavelength of 820 nm. Concentrations were determined by comparison to a curve defined by seven dilutions of a phosphate standard (Merck Certipur).

In addition to the above measurements, aliquots that received zinc acetate were examined for precipitates. None of the samples showed any traces of precipitate, indicating that hydrogen sulfide was below the detection limit in all samples.

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