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Relatively few geochemical measurements were made during the offshore component of Expedition 302. By contrast, numerous geochemical and mineralogical analyses were made during (and after) the onshore component. However, most of the ensuing geochemical data sets remained incomplete for weeks to months after the onshore component was terminated. Consequently, only the shipboard results could be submitted and discussed in the original draft of this volume.

We raise this issue for three reasons. First, it explains the flow of this discussion, where we have retained the “original” shipboard results and discussion at the beginning and subsequently add shore-based results and discussion. Second, it explains the fairly brief discussion of the results. Third, future MSP expeditions might consider alternative strategies for the onshore geochemical program. The primary problem encountered during Expedition 302 was the significant time lag between collecting sediment samples down cores, processing and analyzing these samples in limited batches, and then collating and checking results. We chose to maximize our productivity, which meant collecting or processing samples while awaiting analytical results and which resulted in a large number of analyses. As alternative strategies, the total number of samples could have been decreased (but with significant idle time), the samples could have been collected and processed before the onshore component commenced (which was partly done using available core catcher and squeeze-cake samples), or the geochemical program could have been staggered after other onshore efforts (particularly the opening of cores, description of the lithology, and collecting of samples).

Background and overall program

For nearly 20 y, well-established procedures were used to collect and analyze gases and pore waters from sediments aboard the JOIDES Resolution (Kvenvolden and McDonald, 1986; Gieskes et al., 1991). Although the “shipboard geochemistry” routine evolved with additional equipment (e.g., an inductively coupled plasma–atomic emission spectrometer, Murray et al., 2000) and new priorities (e.g., high-resolution sampling, D’Hondt, Jørgensen, Miller, et al., 2003), core flow and analytical methods remained essentially the same. There was also a growing tendency to measure an increasing number of chemical species (in both gas and water) in shipboard laboratories.

Sediment core, laboratory space, and scientific personnel were at a premium during Expedition 302. Recovery of a continuous sediment record was the highest scientific priority. This meant that the traditional cutting of whole-round samples from unexamined cores was not permitted. Laboratory space for chemistry on the drillship Vidar Viking was limited to less than half of a 5.5 m × 2.2 m container. This meant that only one person per shift could collect, prepare, and analyze gases and pore waters on the drilling vessel. The Expedition 302 shipboard geochemistry program documented below, therefore, has significant modifications from those of previous international scientific drilling expeditions (i.e., DSDP and ODP legs and IODP expeditions). Key differences include (1) water samples were taken after a core was logged, (2) most analyses were postponed to shore-based activities, (3) samples collected were transferred to the ice-breaker Oden for curation, and (4) shipboard analyses were conducted using different procedures. The overarching theme was to collect and appropriately store a sufficient number of samples to meet IODP and shore-based needs while enabling construction of a continuous sediment section.

Gas analyses

Two samples (HS-1 and HS-2) were collected for headspace gas analyses at selected depth intervals. A “pencil-sized” syringe (soft sediment) or cork borer (hard sediment) was inserted into the working-half end of a freshly exposed core section. After withdrawing the syringe or cork borer, a small amount of sediment was pushed out. This excess was shaved off with a flat spatula so that a sediment volume of 5 mL was obtained.

The two 5 mL sediment borings were extruded into 20 mL glass vials filled with 10 mL of seawater. Both headspace samples were immediately sealed with a septum and metal crimp cap, shaken for 2 min, and turned upside down to prevent gas loss. The samples were stored at room temperature on Vidar Viking and then transferred to Oden.

After 23.5 h, the first headspace sample was heated to 60°C for 20 min. A 1 mL volume of headspace gas was extracted from the vial using an Agilent glass syringe, which was not ideal. The gas volume was injected directly into a portable Agilent 6850 series gas chromatograph (GC) equipped with a 30 m × 0.536 mm stainless steel column packed with GS-Q stationary phase (Agilent 1153432) and a flame ionization detector (FID). Unfortunately, an appropriate calibration standard was not shipped, and the hydrogen carrier gas was exhausted after a few samples. Thus, no shipboard gas analyses are reported.

The second gas sample was frozen for shore-based gas analyses. At BCR, a 1 mL volume of headspace gas was extracted from each vial using a Hamilton gas-tight syringe. This volume was then injected directly into a Thermo-Finnegan trace GC equipped with a 30 m × 0.32 mm AT-Q capillary column and an FID. The carrier gas was helium, and the GC oven was programmed as follows: initial temperature of 90°C, held for 3 min, increased to 110°C over 1 min, held for 6 min, increased to 190°C over 5.3 min, held for 9 min, increased to 220°C over 2 min, and held for 4 min. The retention time for methane was 1.94 min. Data were collected using ChromQuest software. The instrument was calibrated by injecting known volumes of pure methane and pure ethane.

Interstitial water samples

Shipboard interstitial water (IW) samples were collected by two methods: (1) high-pressure squeezing of whole-round intervals (Fig. F8) and (2) Rhizone sampling of intact cores (Fig. F9). Notes regarding both methods are contained in the “Bremen Core Curation Van Handbook.” High-pressure squeezing has been commonplace during most sediment coring expeditions of DSDP, ODP, and IODP. Unfortunately, this procedure destroys the integrity of the sediment core. Rhizone sampling offers an alternative, nondestructive approach for collecting pore waters. However, it was not clear whether this method (Seeberg-Elverfeldt et al., 2005) could extract sufficient quantities of water, especially from indurated sediment. The goals of Expedition 302 cruise provided a good rationale for developing and assessing this new method.

Whole-round intervals for squeezing were usually taken every third or fourth core. Approximately 5–10 cm long intervals were cut from sections in the laboratory after they passed through the MSCL. This procedure diverged significantly from standard ODP protocol, where samples are cut from cores on the catwalk before sectioning and MSCL analyses. However, it ensured that IW samples were taken only after collection of their sedimentary record. A potential disadvantage is that the time between core retrieval and squeezing was 30–40 min longer during Expedition 302 than during typical ODP legs. However, we doubt that this time lag significantly modified pore water chemistry (at least relative to IW collection on previous expeditions) because the low temperatures of the shallow-water column and ship deck would help to preserve water chemistry.

Each whole-round interval was extruded onto a stainless steel tray, where its surface was carefully scraped with a spatula to remove potential contamination. The remaining “sediment plug” was then placed into a titanium squeezer, which is detailed in the “Bremen Core Curation Van Handbook.” Pressures as high as 10 × 104 N (gauge) were applied using the accompanying hydraulic press. Interstitial water was passed through a Whatman no. 1 filter fitted above a titanium screen, a hole in the base of the squeezer, and a 0.45 µm disposable filter and into a 20 or 50 mL plastic syringe.

After IW collection, the syringe was removed from the squeezing apparatus and used to dispense IW aliquots. These aliquots included the following:

  • 1.0 mL untreated for Viking-based salinity;

  • 3.0 mL untreated for Viking-based pH and alkalinity;

  • 2.0 mL untreated in plastic scintillation vials for Oden-based NH4+;

  • 8.0 mL in glass vials to be split on Oden for shore-based measurements of anions (e.g., SO42–), sulfide and sulfur isotopes (2 mL saturated zinc acetate solution added to 2 mL of pore water), and 13C (1 drop of saturated HgCl2 solution added to 2 mL of pore water);

  • 5 mL untreated for shore-based Cl and stable isotope analyses;

  • 5 mL untreated as an archive sample; and

  • 5 mL untreated for shore-based dissolved organic carbon (DOC) analyses.

Ideally, ~30 mL of pore water was taken to meet all requirements. In the case of excess pore water, additional archive samples were made; usually less water was collected, and the priority for making aliquots generally followed the order listed above.

For Rhizone sampling, cores were logged, placed in a rack, and turned so the cutting line was vertical. From one to four holes were drilled through the liner on the top of the working half. Rhizones were then fully inserted into the sediment and connected to evacuated 10 mL syringes. Even with four adjacent Rhizones over 10 cm of core, usually <20 mL of total water was collected. Waters from Rhizone sampling were usually reserved for Viking-based pH and alkalinity measurements, Oden-based NH4+ measurements, and shore-based metal and stable isotope analyses.

Offshore interstitial water analyses

Four types of measurements on water samples were made during the offshore component of Expedition 302: salinity, pH, alkalinity aboard the Vidar Viking, and NH4+ aboard the Oden. All other chemistry was analyzed onshore using preserved water samples and sediment samples.

With regard to the measurements made on the Vidar Viking, it is worth noting two complications. First, on most days, there was a 10°C temperature gradient from the lower bow end of the laboratory to the opposite upper stern end. The average temperature within the laboratory also varied from 11° to 19°C. Second, the laboratory, which was placed on the stern, experienced extreme vibrations, especially when ice was caught in the propellers. At times, an apt analogy for working conditions in terms of sound and motion was sitting on a freight train barreling through level crossings at 120 km/h.

Salinity (S) to the nearest part per thousand was measured using a Krüss Optronic DR 301-95 digital refractometer. Approximately 1 mL of sample was placed onto the prism, the lid was closed, and the salinity (after calibration) was recorded. Further details are presented in the “Bremen Core Curation Van Handbook.” Multiple measurements of Arctic surface seawater consistently gave S = 34 after temperature correction.

Alkalinity and pH were measured together using an electrode calibrated to known standards of 4.00 and 7.00 pH, and the “fast titration” method of Grasshoff (1983) was followed. A sample of 2.00 to 3.00 mL was pipetted into a 5.0 mL cup with a magnetic stir bar. The electrode was then inserted into the solution, the solution was stirred, and the pH was recorded. A burette was filled with 0.01M HCl. A small polytetrafluoroethylene (PTFE) microtube was attached to the burette at one end and the electrode to the other end so that acid could be slowly dispensed into the sample. While the solution was still being stirred, a titration was performed to a pH of ~3.8. The amount of acid added and the final pH value were recorded. Alkalinity was determined through equations outlined in the “Bremen Core Curation Van Handbook.” Multiple measurements of Arctic surface seawater gave a pH of 7.8 ± 0.1 and an alkalinity of 2.1 ± 0.1 mM.

Ammonium (NH4+) concentrations were determined by the “Teflon tape gas separator method” (Hall and Aller, 1992). In general, this technique works as follows. An alkaline buffer comprising Na citrate dihydrate in a 10 mM NaOH solution is separated from a 1 mM HCl solution by a Teflon membrane. When samples are added to the buffer, NH4+ ions in the sample are transformed to NH3. Ammonia (NH3) then passes through the membrane, where it dissolves, causing a conductivity change. The change in conductivity can be calibrated to the amount of NH4+ ion. During Expedition 302 cruise, 0.5 mL of sample was injected into the buffer solution. Each sample was analyzed twice, and the values were averaged. The estimated error in these measurements is ±5 µM.

Onshore interstitial water analyses

Using analytical equipment housed in the Department of Geosciences, University of Bremen, some aliquots of IW samples were analyzed for a suite of dissolved species. These included major and trace elements by inductively coupled plasma–optical emission spectrometer (ICP-OES), chloride by titration, sulfate by ion chromatography, and phosphate by photometry. Deionized water used during these analyses was Milli-Q 18 M water.

Dissolved cations were analyzed using a PerkinElmer Optima 3300 R simultaneous ICP-OES (​koelling/​icpoes.html). Most samples used for these analyses were those spiked aboard Vidar Viking with concentrated HNO3. However, alkalinity splits were examined in some cases. In a first run, aliquots of samples were diluted 1:10 (or 1:5 for alkalinity splits) with deionized water and analyzed three times for B (249.772 nm), Ca (317.933 nm), Fe (259.939 nm), K (766.490 nm), Mg (279.077 nm), Mn (259.372 nm), S (181.975 nm), Si (251.611 nm), and Sr (421.552 nm) using a cross-flow nebulizer. In a second run, aliquots of samples were diluted 1:20 (or 1:10 for alkalinity splits) with deionized water and analyzed three times for Al, Ba, Co, Cu, Ni, Pb, V, Y, Zn, and Zr using an ultrasonic nebulizer CETAC USN 5000AT. Concentrations of elements were determined by comparison to a curve defined by multiple dilutions of a multielement standard prepared from single-element standards with a 0.5M NaCl solution as a matrix background. Concentrations of all elements analyzed in the second run were at or near the detection limit, so they are not reported.

Chloride and bromide were collectively measured by titration with AgNO3. Precisely 100 L of sample was added to 5 mL deionized water and then spiked with 100 L potassium chromate indicator solution. This was titrated with a 0.1M AgNO3 solution until the color changed from bright yellow to light orange. The titration was calibrated by 20 replicate analyses of International Association of Physical Sciences Organizations (IAPSO) standard, which gave an average sum for Cl and Br of 0.563 mM. All samples were analyzed three times; the precision of these replicates was consistently within 2 mM.

Sulfate was measured using an ion chromatography system comprising a Knauer autosampler 7500, a Thermo Separation Products pump, a 250 mm × 4 mm GA-1 anion column packed with 100 m PRP-x100 beads, and a Knauer ultraviolet-visible light detector operating at 288 nm (​koelling/​so4.html). The eluant used in this system was a 3 mM potassium hydrogen phthalate (KHP) solution adjusted to pH = 6.8 with KOH, with 5 mL/L of methanol added for sterilization. Samples were diluted 1:20 with deionized water. Concentrations were determined by comparison to IAPSO seawater, and dilutions of this standard were prepared using sulfate-free artificial seawater. Estimated errors in sulfate measurements were ~1 mM.

Dissolved phosphate was analyzed by the molybdenum blue method using a Merck portable photometer SQ118 (see​koelling/​po4_fotom.html). To precisely a 1 mL sample in a microcuvette, 50 L of ammonium molybdate solution and 50 L of ascorbic acid solution were added. Samples were shaken and placed in the dark for 10 min. They were then analyzed at a wavelength of 820 nm. Concentrations were determined by comparison to a curve defined by seven dilutions of a phosphate standard.

In addition to the above measurements, aliquots that received zinc acetate were examined for precipitates. Samples with a white precipitate should indicate the original presence of dissolved hydrogen sulfide; samples with a reddish orange precipitate should indicate the original presence of dissolved iron.

Onshore sediment analyses

A suite of samples was taken for analyses of major element contents, minor element contents, carbonate content, organic carbon content, and mineralogical composition. The samples generally included one 10 cm3 aliquot of sediment from available cores, all squeezecakes, and a few selected horizons of interest (e.g., rocks and nodules). A second suite of samples was also taken for analyses of carbonate content, organic carbon content, organic matter properties, and mineralogy. These samples generally came from available core catchers. Sediment samples were frozen, freeze-dried to remove water, and ground by hand with an agate mortar and pestle. Rock samples were air-dried, photographed, and cut to obtain small fragments. Many fragments were examined by scanning electron microscope (SEM) (see “Supplementary Material”); some fragments were also ground for compositional analyses. All preparation and analyses for the first suite of samples were performed at the Department of Geosciences, University of Bremen. The second set of samples was prepared and analyzed at Alfred Wegner Institute (AWI) (Germany), except for the mineralogy, which was conducted at University of Bremen.

Samples were analyzed for major and minor elements using a Spectro XEPOS portable energy dispersive polarization X-ray fluorescence analyzer, a system specifically designed for rapid analysis of marine sediment chemistry (e.g., Wien et al., in press). Approximately 5 g of dried and powdered sample was packed into a cuvette with a mylar foil bottom. Ten or eleven cuvettes of samples were then placed into an autosampler along with one or two cuvettes of three different standards to assess accuracy and precision. Results of 16 total replicates of U.S. Geological Survey MAG-1 standard, 6 total replicates of Bremen internal standard MAX, and 3 total replicates of Bremen internal standard CAMAX, are presented in Table T7. We note that the system has problems analyzing samples with high sulfur contents.

Sediment samples were analyzed for contents of organic carbon, carbonate, and sulfur using a LECO CS-200 IH carbon-sulfur analyzer at University of Bremen or a LECO CC-125 at AWI (see “Organic geochemistry”). Approximately 50 mg of dried, ground sample was weighed in a ceramic cup and heated in a furnace. The evolved CO2 and SO2 were then measured with a nondispersive infrared detector. A second aliquot of ~90 mg was weighed in a ceramic cup, reacted with 12.5% HCl twice, washed with deionized water twice, and reanalyzed as above. The CO2 measured in the second run was assumed to come from organic carbon. The analytical precision is about ±0.02% absolute.

Samples were analyzed for their mineralogical composition as described in “X-ray diffraction” in “Lithostratigraphy.”

Several pieces of rocks, concretions, or nodules were examined using a Camscan CS 44 SEM equipped with a Phillips energy dispersive system (EDX-PV 98) to obtain semiquantitative elemental analyses. Samples were mounted on carbon-coated stubs and lightly coated with gold.

Organic geochemistry

Total organic carbon (TOC) content was determined on bulk ground samples using a LECO CS-125 analyzer. The analytical precision is about ±0.02% absolute.

Rock-Eval pyrolysis was conducted on bulk sediment samples (with TOC content >0.3 wt%) to determine the amount of hydrocarbons already present in the sample (S1 peak in milligrams hydrocarbons per gram sediment), the amount of hydrocarbons generated by pyrolytic degradation of the kerogen during heating to 550°C (S2 peak in milligrams hydrocarbon per gram sediment), the amount of carbon dioxide generated during heating to 390°C (S3 peak in milligram carbon dioxide per gram sediment), and the temperature of maximum pyrolysis yield (Tmax value in degrees Celsius) (Espitalié et al., 1977; Peters, 1986). As further indicators for the composition of the organic matter, the Rock-Eval parameters hydrogen index (HI) and oxygen index (OI) values were used (cf. Tissot and Welte, 1984; Stein, 1991, and further references therein). HI corresponds to the quantity of pyrolyzable hydrocarbons per gram TOC (mg HC/g C), and OI corresponds to the quantity of carbon dioxide per gram TOC (mg CO2/g C). HI values <100 indicate a dominantly terrigenous (higher plant) source of organic matter, whereas HI values of 200–400 indicate the presence of major amount of aquatic algae (marine or freshwater) and/or microbial biomass. Tmax values <435°C are an indication of fresh immature organic matter, whereas Tmax values >435°C point to the presence of more mature and/or refractive organic matter.