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Hoffmann, Ralf; Pasotti, Francesca; Vázquez, Susana; Lefaible, Nene; Wenzhöfer, Frank; Braeckman, Ulrike (2018): Sediment properties, benthic biogenic compounds, benthic fauna density and biomasses, and benthic diffusive and total fluxes from three stations (Faro, Creek, Isla D) in Potter Cove, Antarctic. PANGAEA, https://doi.org/10.1594/PANGAEA.886232

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Abstract:
For the determination of sediment properties and biogenic sediment compounds, sediment was sampled with 3.6 cm diameter cores in five replicates by SCUBA divers. Sediment subsamples were taken with cut-off syringes (cross-sectional area = 1.65 cm²) and sliced in 1 cm intervals down to 5 cm sediment depth. Each interval was analyzed for various parameters including median grain size, porosity, photosynthetic pigments, total carbon, total organic carbon and total nitrogen. Sediment samples for photosynthetic pigments were stored at -80°C. Sediment samples of other parameters were stored at -20°C. The median grain size was determined with a Malvern Mastersizer 2000G, hydro version 5.40. Sediment porosity was determined after drying sediment samples over minimum two days at 105°C. The sediment porosity was calculated following Burdrige (2006, Geochemistry of marine sediments, Princeton University Press). Chlorophyll a (Chl a), phaeophytin (Phaeo) and fucoxanthin (Fuco) pigment concentrations were determined by HPLC The total carbon (TC) and total nitrogen (TN) was measured by combustion using an ELTRA CS2000The total organic carbon (TOC) was measured using the same method, but after acidifying the sample (3 ml of 10 M HCl).
For prokaryotic density determination, five replicate sediment sub-samples were taken with cut-off syringes (cross-sectional area = 1.65 cm²), sliced at 1 cm intervals down to 5 cm sediment depth, fixed in a 2% formaldehyde/seawater filtered solution (9 ml) and stored at 4°C. The acridine-orange-direct-count method after Hobbie (1977, https://www.ncbi.nlm.nih.gov/pmc/articles/PMC170856/) was used to stain prokaryotes in the sub-samples and subsequently counted with a microscope (Axioskop 50, Zeiss) under UV-light (CQ-HXP-120, LEj, Germany). For each sample, single cells were counted on two replicate filters and for 30 random grids per filter (dilution factor 4000). Prokaryotic biomass was estimated by the determination of the mean prokaryotic cell volume in the first two centimetres with a "New Portion" grid (Graticules Ltd, Tonbridge, UK), converted into biomass using a conversion factor of 3.0 x 10-13 g C pm-3 after Børsheim et al. (1990, https://www.ncbi.nlm.nih.gov/pmc/articles/PMC183343/) and multiplied with the prokaryotic density.
For the determination of meiofauna density, biomass and identification of meiofauna taxa, five sediment samples were taken with small sediment cores (Ø 3.6 cm). Sediment samples of the first five centimeters were stored in filtered seawater buffered 4% formaldehyde solution at 4°C till the start of the analyses. The samples were sieved over a 1 mm and 32 µm mesh. The sample was then centrifuged three times in a colloidal silica solution (Ludox TM-50) with a density of 1.18 g cm-3 and stained with Rose Bengal after Heip et al. (1985, The Ecology of marine nematodes, Oceanography and Marine Biology - An Annual Review; vol. 23). Afterwards, the taxa were identified and counted. Calcifying organisms (except nematodes and polychaetes) were acidified prior the total organic carbon content of single taxa were analyzed with a FLASH 2000 NC Elemental Analyzer (0.01% detection limit)
The benthic macrofauna was sampled by using a Van Veen grab (530 cm² surface area). At each station, four recovered sediment samples were sieved over a 1 mm mesh, stored in seawater buffered 4% formaldehyde and stained with Rose Bengal after Heip et al. (1985). In the laboratory, the taxa were identified to the lowest possible taxonomic level (at least family level), counted, weighted, and the Shannon-Wiener diversity index (H') calculated in Primer v6.0. Ash-free dry weight (AFDW) was determined by subtracting the ash weight (after combustion at 500°C) from the dry weight (dried for 48 h at 60°C). AFDW was converted into carbon by assuming that 50% of the AFDW is carbon after Wijsman et al. (1999, http://www.jstor.org/stable/24849594). The Van Veen grab sampling results in a strong underestimation of the density of the important Antarctic bivalve Laternula elliptica. Therefore, two rows of eight grids (45 cm x 45 cm) were randomly placed on the seafloor by scuba divers and photos were taken (Nikon D750 with a rectilinear Nikon 16-35 mm lens in a Nauticam underwater housing and two Inon Z-240 strobes). The photos were used to count siphons of L. elliptica to determine the density and to measure the siphon width (maximum distance between outer edges of the two siphons of one individual) at the three sites. Assuming a linear relationship between siphon width and AFDW, a conversion from the siphon width to estimated biomass of L. elliptica was performed. The calculation of the conversion relationship of the siphon width to AFDW was performed on data from the same L. elliptica population. The community bioturbation potential was calculated following formula Queiros et al. (2013, doi:10.1002/ece3.769).
Three transparent and three black chambers (inner diameter 19 cm, height 33 cm) were carefully pushed into the sediment at each station by SCUBA divers, who took special care to not disturb the sediment surface during the procedure. About 15 cm of sediment and 18 cm of overlying water was enclosed. During the incubation, which lasted 20-22 h and encompassed light and dark hours, cross-shaped stirrers powered by a 12 V lead-acid battery kept the overlying water homogenous. HOBO Pendant loggers (Onset, Bourne, USA) were placed both in situ and on land to record the amount of radiation (150-1200 nm) during the incubation with a temporal resolution of 5 minutes. The enclosed overlying water in the chambers was sampled through valves attached to the chamber lids at the start and end of the chamber incubation, using gas-tight glass syringes. The water samples were kept at in situ temperature and in darkness until further processing, which took place within 1.5 h after the samples were taken.
Subsamples were taken to either determine the oxygen concentration, the concentration of dissolved inorganic carbon (DIC) and the concentration of phosphate, ammonium, nitrite, nitrate, and sulfate. Winkler titration was used to immediately determine the oxygen concentration in the water sample in technical duplicates. For DIC analyses technical triplicates were poisoned with HgCl2 and stored at 4°C until measurement after 6 months. DIC samples were analyzed using an autosampler (Techlab, Spark Basic Marathon) with a digital conductivity measuring cell (VWR, digital conductivity meter, Germany). For nutrient analyses technical triplicates were filtered through a GF/F filter and stored at -20°C until analysis. The samples were analyzed with an autosampler (CFA SAN-plus, Skalar Analytical B.V., Netherlands) for ammonium, phosphate, nitrite and [nitrate + nitrite] concentrations. The nitrate concentration was determined by subtracting the nitrite concentration from the [nitrate + nitrite] concentration. The resulting total fluxes were calculated following Glud et al. (2008, doi:10.1080/17451000801888726).
High-resolution in situ oxygen profiles were measured using a microprofiler. The microsensors were driven from the water phase into the sediment with a spatial resolution of 100 µm and a temporal resolution of 30 seconds. On the profiler electronic unit, three custom made electrochemical O2 microsensors after Revsbech (1989, doi:10.4319/lo.1989.34.2.0474) were mounted and calibrated before deployment in oxygen saturated and oxygen depleted water. The microprofiler was programmed, so microsensors penetrated the SWI around noon at the same or the following day after the deployment. Running average smoothed profiles (https://doi.pangaea.de/10.1594/PANGAEA.885472) were used to calculate the diffusive oxygen uptake (DOU) over the SWI using Fick's first law.
For the calculation of the diffusive flux of sulfate, DIC, and nutrients, sediment was sampled with cores (10cm diameter) with predrilled holes at 1 cm depth intervals that were sealed with diffusion-tight tape. The porewater was extracted using Rhizons (type: core solution sampler, Rhizosphere Research Products, filter pore diameter of 0.1 mm) connected to 10 mL Luer lock syringes. The Rhizons were horizontally inserted into the cores and by creating a permanent vacuum in the syringes, porewater was extracted. The first drops were used to rinse the syringe and then discarded. The extracted pore water was split for sulfate analyses (sample fixed in 5% ZnAc, stored at 4°C), DIC analyses (sample fixed in HgCl2, stored at 4°C) and nutrient analyses (frozen at -20°C). DIC and nutrients were analyzed as described above. Sulfate was analyzed by using non-suppressed ion chromatography with the Methrom 761 Compact IC equipped with a Metrosep A SUPP 5 column (Methrom, Herisau, Switzerland). From the resulting depth profiles, diffusive fluxes were calculated using the same formula as for the DOU calculation, but with Ds of the specific molecule.
Coverage:
Median Latitude: -62.230013 * Median Longitude: -58.652454 * South-bound Latitude: -62.235580 * West-bound Longitude: -58.662060 * North-bound Latitude: -62.225110 * East-bound Longitude: -58.641750
Date/Time Start: 2015-02-09T00:00:00 * Date/Time End: 2015-03-04T00:00:00
Size:
11 datasets

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Datasets listed in this publication series

  1. Hoffmann, R; Pasotti, F; Vázquez, S et al. (2018): Benthic diffusive fluxes from three stations (Faro, Creek, Isla D) in Potter Cove, Antarctic. https://doi.org/10.1594/PANGAEA.886223
  2. Hoffmann, R; Pasotti, F; Vázquez, S et al. (2018): Benthic fauna density and biomasses from three stations (Faro, Creek, Isla D) in Potter Cove, Antarctic. https://doi.org/10.1594/PANGAEA.886224
  3. Hoffmann, R; Pasotti, F; Vázquez, S et al. (2018): Benthic total fluxes from three stations (Faro, Creek, Isla D) in Potter Cove, Antarctic. https://doi.org/10.1594/PANGAEA.886231
  4. Hoffmann, R; Pasotti, F; Vázquez, S et al. (2018): Laternula elliptica photo survey at three stations (Faro, Creek, Isla D) in Potter Cove, Antarctic. https://doi.org/10.1594/PANGAEA.886226
  5. Hoffmann, R; Pasotti, F; Vázquez, S et al. (2018): Pigment concentrations in sediments from three stations (Faro, Creek, Isla D) in Potter Cove, Antarctic. https://doi.org/10.1594/PANGAEA.886227
  6. Hoffmann, R; Pasotti, F; Vázquez, S et al. (2018): Porewater nutrient profiles at three stations (Faro, Creek, Isla D) in Potter Cove, Antarctic. https://doi.org/10.1594/PANGAEA.886228
  7. Hoffmann, R; Pasotti, F; Vázquez, S et al. (2018): Porosity and median grain size from three stations (Faro, Creek, Isla D) in Potter Cove, Antarctic. https://doi.org/10.1594/PANGAEA.892730
  8. Hoffmann, R; Pasotti, F; Vázquez, S et al. (2018): Prokaryote biomass from three stations (Faro, Creek, Isla D) in Potter Cove, Antarctic. https://doi.pangaea.de/10.1594/PANGAEA.892736
  9. Hoffmann, R; Pasotti, F; Vázquez, S et al. (2018): Prokaryote cell volume from three stations (Faro, Creek, Isla D) in Potter Cove, Antarctic. https://doi.org/10.1594/PANGAEA.892731
  10. Hoffmann, R; Pasotti, F; Vázquez, S et al. (2018): Prokaryote density at three stations (Faro, Creek, Isla D) in Potter Cove, Antarctic. https://doi.org/10.1594/PANGAEA.886229
  11. Hoffmann, R; Pasotti, F; Vázquez, S et al. (2018): Sediment carbon and nitrogen content at three stations (Faro, Creek, Isla D) in Potter Cove, Antarctic. https://doi.org/10.1594/PANGAEA.886230