Total and labile particulate element concentrations via ICPMS

Sampling Methodology:

Super GeoFISH samples were collected from surface waters (~2m) using a towed ‘fish’ deployed by Ken Bruland’s lab, and typically used to collect a surface sample upon arrival at the station location, for use as the shallowest sample of the resulting vertical concentration profile for that station. The ‘fish’ water was cleanly pumped directly into the bottom stopcock of a Go-Flo mounted vertically in the clean “bubble” (temporary clean room). Sampling then proceeded identically to procedures used for GO-Flo samples collected at depth, except that the GO-Flo was not inverted for mixing as potential particle settling time was negligible. All surface samples were analyzed by the Twining group at Bigelow Laboratory.

During sampling in the clean van, unfiltered salinity and nutrient samples were first taken from the GO-Flo bottles to create headspace, and then the bottles were inverted slowly three times to re-suspend any large particles that might have settled before sampling. Then the GO-Flo bottles were pressurized to ~0.5 atm with HEPA-filtered air, and filtration commenced using methods similar to those published in Planquette and Sherrell (2012). GO-Flo stopcocks were fitted with an acid-cleaned piece of Bev-a-Line tubing feeding into a polycarbonate elbow that that attached by Luer lock into a 25 mm, polypropylene Swinnex filter holder (http://www.millipore.com/catalogue/module/C160). These filter holders had been loaded before each cast with an acid-cleaned 25mm Supor (Pall Gelman) 0.45um polyethersulfone filter. For three shelf stations, 47mm Supor 0.45 um filters were used instead, as particle concentrations were markedly higher than at other stations. These were supported in 47 mm polypropylene filter holders (MFS; Planquette and Sherrell, 2012). Immediately prior to sampling, the headspace of the filter holder was flushed with seawater to evacuate any air bubbles to prevent air lock or occlusion of the filter surface area by air bubbles during sampling (Planquette and Sherrell, 2012). Filtration commenced with filtrate collected in a plastic bucket, and the filter holders held approximately horizontal such that any residual headspace air bubbles would rise to the top of the filter holder, avoiding occlusion of the filter face. After filtration was complete (filter clogged to <1 drop of filtrate per second) or two hours had passed since the start of filtration, filtration was ceased by closing the stopcock on the GO-Flo bottle. Seawater volume passed through the filter was measured and recorded. The Swinnex filter holders were taken into HEPA-filtered clean space, and excess headspace seawater was removed by gentle vacuum suction through the filter. The filter holders were then transferred in a sealed plastic bag to the HEPA-filtered “bubble” clean room in the ship’s main lab. Working directly under a vertically flowing HEPA hood, the filter holders were opened, and the filter was removed using Tefzel forceps (held by only the edge of the filter). The non-sampled side of the filter was “blotted” by placing onto an acid-cleaned 47mm Supor filter to remove remaining seawater by capillary action. Finally, the visibly dry filter was placed into an acid-cleaned polystyrene PetriSlide (EMD Millipore), with the filter positioned such that the inner rim of the lid was positioned over one edge of the filter, thus acting as a stabilizing device to limit subsequent movement of the filter within the PetriSlide, and stored at -20 degC for transport and storage until analysis at Bigelow Laboratory or Rutgers University.

Analytical Methodology:

Bigelow Laboratory

All digestion steps were performed in a Class-100 clean room using standard clean techniques. Filters were cut in half using a ceramic blade, using a cutting template and a light table to aid precision. One half was digested following the protocol of Berger et al. (2008) to obtain labile particulate concentrations; the other was digested using a 4M HCl, 4M HNO₃, and 4M HF mixture as described in Ohnemus et al. (2014) to obtain total particulate element concentrations. Labile particulate filter halves were leached in a solution of 25pcnt Optima-grade acetic acid and 0.02 M hydroxylamine hydrochloride following the protocol of Berger et al. (2008). One milliliter of this solution was added to the filter stored in a 1.7 mL polypropylene vial. Following the recommendation of Berger et al. (2008), the solution was heated to 95 degC in a water bath for 10 minutes and then allowed to cool to room temperature. The filter was in contact with the acetic acid leach for a total of two hours, after which the filter was removed from the polypropylene vial and the acetic acid/hydroxylamine leachate was centrifuged at 14,000 rpm for 10 minutes to sediment all particles. Without disturbing particles on the bottom of the tube, approximately 0.8 mL of leachate was transferred into an acid-cleaned 7 mL PFA digestion vial. Optima-grade HNO3was added (100 uL) to the digestion vial, which was subsequently heated uncapped at 110 degC to near dryness. Vial contents were redissolved with 2pcnt HNO3 (Optima grade).

Total particulate metals were determined by digestion of the second filter half. The filter was transferred to a rigorously cleaned 22-mL PFA vial, 2 mL of a solution containing 4M HCl, 4M HNO3, and 4M HF (all Optima grade) was added to completely cover the filter piece, and the vial was tightly capped and heated to 110 degC for 4 hours. This procedure has been determined to be adequate for digestion of all particulate material, while allowing the Supor filter to remain intact (Ohnemus et al. 2014). Following heating, the acid solution in the bomb was poured into a second PFA vial, leaving the filter piece behind. To ensure complete transfer of acid, the bombs were thoroughly rinsed with 3 × 0.5 mL aliquots of ultrapure water which were poured into the secondary vial. The secondary vial was then heated to dryness and the contents redissolved with 2 mL of a 50 pcnt Optima-grade HNO3 + 15 pcnt Optima-grade H2O2 (v/v of concentrated reagents) solution. This solution was again dried down and the contents redissolved with 2 pcnt HNO3.

Filter halves from depths typically greater than 150m outside the oxygen minimum zone (stations 15-36) were digested only in the HCl/HNO3/HF solution to provide a total particulate element fraction; no corresponding labile particulate concentration data have been collected for these samples.

All digests were analyzed using a Finnigan-MAT Element2 HR-ICP-MS at the University of Maine following the protocols outlined in Twining et al. (2011). The instrument is equipped with a cyclonic nebulizer, an autosampler contained under a HEPA filter, and nickel cones. Ba-137, Cd-111, La-139, Th-232, and Y-89 were analyzed in low-resolution mode, and the remaining isotopes were analyzed in medium-resolution mode. Y-89 was analyzed in both low- and medium-resolution modes, and the values reported are derived from the low-resolution mode values. Multiple isotopes were analyzed for some elements, and the isotopes used to derive the reported concentration are as follows for each element: Fe (56 and 57), Cu (63 and 65), Ni (60 and 62), Zn (66, 64 and 68). Mean agreement was 1.7 pcnt for the two Cu isotopes, 3.2 pcnt for the two Fe isotopes, 7.6 pcnt for the two Ni isotopes, and 1.3 to 2.9 pcnt for the Zn isotopes. Between low and medium instrument resolution modes, mean agreement was 0.4pcnt for Y-89. Quantification was performed by external calibration, and In-115 was used as an internal standard to correct for variations in instrumental sensitivity during analyses.

Cs-133, spiked during the initial sample digestions, was used as a process recovery monitor, but no samples were discarded or corrected using the Cs recoveries, as typical Cs recoveries were 95-105pcnt.

Rutgers University

Samples were analyzed in the Sherrell laboratory at Rutgers University. Frozen filters were cut in half using a ceramic rotary blade; a filter-cutting template was illuminated on a light table for guidance during cutting, and filter cutting error performed on blank filters was found to be ≤2% by weight. One filter half was used for sample digestion (reported here), and the other filter half was used for archiving or for acid leaching of “labile” metals (to be reported at a later date). For digestion, filter halves were placed into the bottom of acid-clean 15 mL PFA vials (Savillex), and 0.4 mL of Milli-Q ultrapure water was added to the top. Once it was fully wetted, the filter half was pulled up the side of the vial and adhered to the wall, curved side toward bottom of vial. Then 0.6 mL of a solution containing 16.7 pcnt (v/v) hydrofluoric acid (HF, Optima grade, Fisher) and 83.3 pcnt (v/v) nitric acid (HNO3, Optima grade, Fisher) was added by pipet to each vial, aiming at the adhered filter half. The final digestion acid mixture was thus 1.0 mL of a solution containing 8M HNO₃ and 2.9M HF. The vial was then capped tightly and placed on a Teflon hotplate at least 2 cm from other vials to allow air circulation. These “bombs” were refluxed at 135 degC for 4 hours. After cooling, solution was gathered to the bottom of the vials, lids were removed, and the digest solution was evaporated until ~5-10 µL of solution remained. At that point, 100 uL of concentrated HNO3 was added, and the solution was re-evaporated until ~5-10 µL of solution remained. The additional HNO3 dry-down encourages evaporative loss of HF. Evaporating to dryness was avoided to prevent “baking” sample residue onto the Teflon surface, thus aiding in complete re-dissolution and minimizing carry-over to subsequent sample digestions. Finally, the remaining droplet was brought up in 3.0 mL of 5 pcnt HNO3 (v/v) and transferred to a 15 mL acid-cleaned polypropylene centrifuge tube for archiving prior to analysis.

Sample analysis was completed on a Finnigan-MAT Element-1 inductively coupled plasma mass spectrometer, employing a PFA Microflow nebulizer (ESI), an Apex and ACM sequential desolvation system (ESI) to reduce molecular oxide ion formation, and Ni cones. Sample solutions were diluted five times from the archived digest solutions (to reduce chemical matrix effects) and were quantified using nine-point, multi-element standard curves with acid matrix identical to that of samples and concentrations bracketing the range of the samples. Single-point standard additions were run every 10 samples to check for accuracy, and analytical replicates were made every 10 samples to monitor analytical precision.

To monitor overall process recovery, a high Cs spike was added at the start of sample digestion. Analysis of Cs in the final digest solution showed that recovery was always 100±5%, so no samples were discarded or corrected for process recovery.