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The response of the benthic microbial community to a controlled sub-seabed CO2 leak was assessed using quantitative PCR measurements of benthic bacterial, archaeal and cyanobacteria/chloroplast 16S rRNA genes. Similarly, the impact of CO2 release on the abundance of benthic bacterial and archaeal ammonia amoA genes and transcripts, and also to the abundance of nitrite oxidizer (nirS) and anammox hydrazine oxidoreductase (hzo) genes and transcripts. Samples were taken from four zones (epicentre (zone 1); 25m distant (zone 2), 75m distant (zone 3) and 450m distant (zone 4)) during 6 time points (7 days before CO2 exposure, after 14 and 36 days of CO2 release, and 6, 20 and 90 days after the CO2 release had ended). Changes to the active community of microphytobenthos and bacteria were also assessed before, during and after CO2 release using Denaturing Gradient Gel Electrophoresis of cyanobacteria/chloroplast 16S rRNA. Changes to the composition of the active bacterial community was assessed first using Terminal Restriction Fragment Length Polymorphism (T-RFLP) of bacterial 16S rRNA. In depth comparisons of possible changes to the active bacterial community at zone 1 and 4 before, during and immediately after the CO2 release was performed using 16S rRNA 454 pyrosequencing. This dataset was created by Plymouth Marine Laboratory (PML) under the program QICS (Quantifying and monitoring environmental impacts of geological carbon storage) which was funded by the Natural Environment Research Council (NERC), with support from the Scottish Government. The results are contained in three text files. QICS project website: www.bgs.ac.uk/qics/home.html. Tait et al. (2015) Rapid response of the active microbial community to CO2 exposure from a controlled sub-seabed CO2 leak in Ardmucknish Bay (Oban, Scotland). IJGGC DOI: 10.1016/ijggc.2014.11.021. Watanabe et al. (2015) Ammonia oxidation activity of microorganisms in surface sediment to a controlled sub-seabed release of CO2. IJGGC DOI: 10.1016/j.ijggc.2014.11.013.
To identify and quantify soil N species over a full growth season, small volumes of soil were removed from each sampling site 5 times during the field season and extracted in the laboratory. Bare soil at higher elevations, namely Observation Bluff, Factory Bluffs, Jane Col and lower parts of Spindrift Col; Soils from below mosses on the Backslope and on Moss Braes. Soils from below higher plant species at Bernsten Point, Factory Bluffs, Moss Braes and North Point. Orthinogenic soils from around penguin colonies at Gourlay Peninsula, Spindrift Rocks and North Point and disturbed soil from around Signy Base were collected. At the same time, soil pore water was extracted using Rhizon soil water samplers. DON (Dissolved organic Nitrogen) and Microbial biomass measurements were made by standard CHCl3 fumigation-extraction techniques. Turnover of DON in the soil was determined by the addition of 14C-labelled plant protein (purified from 14C-labelled algal cells) or 14C-labelled glucose to the soil at a range of concentrations, and their turnover (soil label depletion in combination with NH4 +, NO3-and 14CO2 production) was determined. Gross rates of N mineralization and nitrification were determined using 15N isotope dilution methodology. Laboratory analysis of N speciation and quantification, 14C uptake and respiration, 13C PLFA signatures and 15N analysis was done. Amino acid turnover times have been determined using 14C labelled amino acids. For the final stage of the project a mathematical model to describe plant-soil-microbial N fluxes in Antarctic soils was constructed.
This dataset contains in-stream measurements of sediment porewater nutrients, nitrification rates (and the fraction which is either fully oxidised to nitrate or reduced to N2 gas), and the abundance of microbial 16S rRNA and specific N-cycling genes and transcripts. Sediments were sampled in winter (February 2018) and summer (July 2018), from 12 UK rivers with permeable beds (sand or chalk geology) and a gradient of P concentrations, in the Hampshire Avon catchment, Kent, and Essex. Methods included measurements of porewater nutrients using Skalar SAN++AutoAnalyser, nitrification rates from in-situ ‘push-pull’ injections of 15N-labelled ammonia and -nitrite, and sediment microbial gene and transcript abundance by DNA extraction and qPCR. The work was supported by the Natural Environment Research Council (Grants NE/P01142X/1, NE/P011624/1; A new dynamic for Phosphorus in RIverbed Nitrogen Cycling - PRINCe Full details about this dataset can be found at https://doi.org/10.5285/d432d96c-7aff-45a5-9d4b-37e4065afdd7