The foraging ecology of three fulmarine petrels including Cape petrels, Southern fulmars and Antarctic petrels were investigated at Hop Island during the 2015/16 austral summer. Two datasets were generated: 1) tracking data from Fulmarine petrels, and 2) stable isotope analysis of blood, feathers and egg shells. Tracking data were collected using Ecotone GPS trackers attached to the birds back feathers with tape. Location data has been interpolated using great circle distance to a time step of 15 minutes and include a record of whether the bird dived during that time period or not. Each location point was assigned a breeding stage (incubation or chick rearing) based on individual nest activities. Stable isotope ratios of carbon (13C/12C) and nitrogen (15N/14N) were determined by analysing 1 mg aliquots through continuous flow - elemental analysis - isotope ratio mass spectrometry (CF-EA-IRMS). Isotopic values of blood reflect approximately the last 52 days before sampling and thus the incubation period of all three species. Egg membranes and feathers remain metabolically inert after formation, and hence reflect the trophic niche during the pre-laying and moult period, respectively. We collected moult feathers during the chick-rearing period and therefore assumed that these were formed one year prior to the collection date and thus represent the trophic niche of the chick-rearing period one year earlier (austral summer 2014-15).

Two toxicity tests were conducted in the Davis station laboratories in December 2010. Tests used locally collected amphipods of the species Orchomenella pinguides. The tests were conducted by Bianca Sfiligoj, as part of her PhD research (Sfiligoj 2013), with results published in (Sfiligoj et al. 2015). Field and laboratory work was conducted under project AAS 2933, with analysis and write-up completed under AAS 4100 (both projects CI: King). Details are fully described in the published manuscript provided with this data record; file name: Sfiligoj et al 2015_Ecotoxicology.pdf. A subset of the data is also used in Candy et al. 2015 (Filename: Candy et al 2015_Ecotoxicology.pdf). Data files: Test data are provided in the .xlsx file: 'Orchomenella-Tests-Dec 2010.xlsx'. Each worksheet includes a "This worksheet provides…" description in cell A1. Laboratory notebook records are provided in the scanned file: Sfiligoj-LabBookScan-Davis10-11.pdf. In this notebook, tests are labelled LT1 and LT2 (referred to as: amphipod lentil test 1 and 2); with results recorded on pages: 1-19 and 26-28. Data associated with this record has also been presented at: - Candy SG, Sfiligoj BJ, King CK, Mondon JA (2013) Modelling interval-censored survival times in toxicological studies using generalized additive models, The International Biometric Society Australasian Region Conference 2013, Mandurah, Australia, 1-5 December 2013. - Sfiligoj BJ, King CK, Candy SG, Mondon JA (2012) Development of appropriate bioassay and statistical methods for determining survival sensitivities of Antarctic marine biota to metal exposure, 2nd Society for Environmental Toxicology and Chemistry (SETAC) Australasia Conference, Brisbane, Australia, 4-6 July 2012. - Sfiligoj BJ, King CK, Candy SG, Mondon JA (2012) Development of appropriate bioassay and statistical methods for determining survival sensitivities of Antarctic marine biota to metal exposure, Society for Environmental Toxicology and Chemistry (SETAC) World Congress, Berlin, Germany, 20-24 May 2012.

The dataset comprises scanned copies of the boundaries of Adelie penguin breeding colonies and sections of island coastlines made from aerial photographs taken between 9-15 December 1981. The original tracings by Michael Whitehead were scanned by Colin Southwell.

Synchrotron based FTIR macromolecule profiles of 5 diatom species from the AAS_4026 ocean acidification project. Data represent the peak areas for wavenumbers related to key macromolecules. For details on methods see Duncan et al. (2021) New Phytologist. Experimental design and mesocosm set up Mesocosm set up and conditions were as described previously (Deppeler et al., 2018; Hancock et al., 2018). Briefly, a near-shore, natural Antarctic microbial community was collected from an ice-free area among broken fast ice approximately 1km offshore from Davis Station, Antarctica (68° 35ʹ S, 77° 58ʹ E) on 19 November 2014. This community was incubated in 6 x 650L polyurethane tanks (mesocosms) across a gradient of fCO2 levels (343, 506, 634, 953, 1140 and 1641 μatm; denoted M1 – M6). These fCO2 levels corresponded to pH values ranging from 8.17 to 7.57. Temperature was maintained at 0.0 °C ± 0.5 °C and the mesocosms were stirred continuously by a central auger (15 r.p.m.) for gentle mixing and covered with an air-tight lid. Irradiance was initially kept low (0.8 ± 0.2 μmol photons m-2s-1), while cell physiology was left to acclimate to increasing fCO2 levels (over 5 days). When target fCO2 levels were reached in all six mesocosms, light was gradually increased (days 5-8) to 89 ± 16 μmol photons m-2s-1 on a 19 h:5 h light:dark cycle, to mimic current natural conditions. To generate the gradient in carbonate chemistry, filtered seawater saturated with CO2 was added to five of the mesocosms. Daily measurements were taken to monitor pH and dissolved inorganic carbon (DIC). For details of fCO2 manipulations, analytical procedures and calculations see Deppeler et al., (2018). Samples for physiological and macromolecular measurements in this study were taken on day 18, at the end of the incubation period (Deppeler et al., 2018). Cell volume Cell volume was determined for selected taxa from M1 and M6 via light microscopy. Cells were imaged on a calibrated microscope (Nikon Eclipse Ci-L, Japan) and length, width and height (24-77 cells per taxa) determined using ImageJ software (Schneider et al., 2012). Biovolume was then calculated according to the cell morphology and corresponding equations described by Hillebrand et al (1999). Macromolecular content by FTIR The macromolecular composition of the selected diatom taxa sampled from all six mesocosms on day 18 was determined using Synchrotron based FTIR microspectroscopy on formalin-fixed (2% v/v final concentration) cells. Measurements were made on hydrated cells and processed according to previous studies (Sackett et al. 2103; 2014; Sheehan et al. 2020). Briefly, fixed cells were loaded directly onto a micro-compression cell with a 0.3 mm thick CaF2 window. Spectral data of individual cells (between 15-49 cells per taxon per mesocosm) were collected in transmission mode, using the Infrared Microspectroscopy Beamline at the Australian Synchrotron, Melbourne, in November 2015. Spectra were acquired over the measurement range 4000− 800 cm−1 with a Vertex 80v FTIR spectrometer (Bruker Optics) in conjunction with an IR microscope (Hyperion 2000, Bruker) fitted with a mercury cadmium telluride detector cooled with liquid nitrogen. Co-added interferograms (n = 64) were collected at a wavenumber resolution of 6 cm−1s. To allow for measurements of individual cells, all measurements were made in transmission mode, using a measuring area aperture size of 5 × 5 µm. Spectral acquisition and instrument control were achieved using Opus 6.5 software (Bruker). Normalised spectra of biologically relevant regions revealed absorbance bands representative of key macromolecules were selected. Specifically, the amide II (~1540 cm-1), Free Amino Acid (~1452 cm-1), Carboxylates (~1375 cm-1), Ester carbonyl from lipids (~1745 cm-1) and Saturated Fatty Acids (~2920 cm-1) bands were selected. Infra-red spectral data were analysed using custom made scripts in R (R Development Core Team 2018). The regions of 3050-2800, 1770-1100 cm-1, which contain the major biological were selected for analysis. Spectral data were smoothed (4 pts either side) and second derivative (3rd order polynomial) transformed using the Savitzky-Golay algorithm from the prospectr package in R (Stevens and Ramirez-Lopez, 2014) and then normalised using the method of Single Normal Variate (SNV). Macromolecular content for individual taxon was estimated based on integrating the area under each assigned peak, providing metabolite content according to the Beer-Lambert Law, which assumes a direct relationship between absorbance and relative analyte concentration (Wagner et al., 2010). Integrated peak areas provide relative changes in macromolecular content between samples. Because of the differences in absorption properties of macromolecules, peak areas can only be used as relative measure within compounds.

These data are linked to what appears to be an unfinished report/paper by Pat Quilty. An extract of the unfinished report is available below, and the full document is included in the data download. These data are also linked to a collection in the biodiversity database, and are also related to another record (both listed at the provided URLs). Foraminiferids are recorded from samples collected on Mac. Robertson Shelf and Prydz Bay, East Antarctica in 1982, 1995 and 1997. Most are identifiable from previous literature but a new enrolled biserial agglutinated genus is noted but not defined. Distribution is related to oceanographic factors. The Mac. Robertson Shelf-Prydz Bay region off the East Antarctic coast is that segment of the southern Indian Ocean between latitudes 66 degrees and almost 70 degrees S, and longitudes 60 degrees and 80 degrees E. It includes Mac. Robertson Shelf, the continental shelf, bounded seaward by the 500 m isobath, and Prydz Bay, the deepest re-entrant into the east Antarctic shield and the outlet for the Lambert Glacier at its southern end. The Lambert Glacier is the world’s largest glacier and drains some 1 000 000 km2 of East Antarctica. The marine region studied here covers some 140 000 km2. Several research cruises to the region have collected sediment samples that yielded modern and recycled foraminiferid faunas. The modern component of the faunas has not been recorded in detail previously. This paper records the details of the taxonomy and distribution of species collected during marine geology/geophysics cruises that provided the foraminiferids discussed in Quilty (1985, 2001), O’Brien (1992), O’Brien et al. (1993, 1995) and Harris et al. (1997). The geophysical results and interpretations of the 1982 voyage of MV Nella Dan are described by Stagg (1985) and this provides also the general setting and nomenclature of Prydz Bay. Two cruises (1995 and 1997) of RSV Aurora Australis collected samples and these provided the basis for Quilty’s records of foraminiferids and other components on a sample-by-sample basis in O’Brien et al. (1995) from 51 samples, and from a further 27 samples reported in Harris et al. (1997). The 1995 cruise also yielded the recycled foraminifera recorded by Quilty (2001) and the Mesozoic material documented by Truswell et al. (1999). Neither of these cruise records provided details of the faunas to the level covered here. Further studies for the region are given in the results of ODP Legs 119 and 188. The impetus for conducting this review comes from two sources. Firstly, few foraminiferids have been documented from this region, and even fewer have been figured. Secondly, 2007-2008 was designated the [fourth] International Polar Year (IPY) and one of the major programs is the Census of Antarctic Marine Life (CAML), a component of the global Census of Marine Life (CML). This paper is a contribution to that project. Included in the review are faunas from the modern environment and some which may be ‘Late Cenozoic’ in which the faunas are of the same species as the modern and in which data from the modern can be, and have been, used to infer past environments (Fillon 1974, Kellogg et al. 1979, Ward and Webb 1986). The aims of this paper are: - to document the species of foraminifera recovered from geology/geophysics cruises to the Mac. Robertson Shelf and Prydz Bay region, offshore East Antarctica (Fig. 1); - to make the nomenclature of species recorded consistent with latest taxonomic practice; - to characterise the faunas by diversity and dominance factors; and - to discuss the controls on the distribution of faunas recorded.

This dataset contains scanned copies of the RMT and bottom trawl logs from Voyage 6 1990-91 (AAMBER2) of the Aurora Australis. This was primarily a marine science voyage. Surveys of krill, other zooplankton and pelagic fish were taken in Prydz Bay, Antarctica between January and February 1991. 177 midwater trawls were successfully completed at 59 stations. Midwater fish were sampled using an International Young Gadoid Pelagic Trawl (IYGPT). At each station, hauls were taken at depths of 20-30m, approximately halfway down the water column, and 20-30m above the bottom. At six stations, the lowest sample was duplicated using a light fitted to the net. Where samples were made off the shelf, standard depths of 20-30m, 400m, and 800m were fished. All hauls were of 30 minutes fishing time. Bottom trawls were made using a 35m headline length otter trawl fitted with 40cm diameter bobbin gear. A 2" mesh cod end liner was used to retain small fish. On both nets, a Simrad trawl surveillance sonar was used.

The Davis Aerodrome Project (DAP) collected a range of environmental survey data over several field seasons to support a comprehensive environmental assessment of the proposed aerodrome. This data includes flora, fauna, soils, lake ecosystem, nearshore, marine, air quality and meteorological information which has been collected by a number of different methods, and extends across the current Davis Station, proposed aerodrome and supporting infrastructure footprint (Ridge Site), previous sites considered for the aerodrome (Heidemann Valley, Adams Flat), as well as locations across the Vestfold Hills away from any of the proposed developments. This dataset contains long-term underwater acoustic recordings made for the Australian Antarctic Division’s Davis Aerodrome Project 5097 environmental assessments. Calibrated measurements of sound pressure were made at two sites in the vicinity of Davis Research station (approx. 5km west of the station and one in Long Fjord to the north of station). The attached data was downloaded from the instrument deployed west of Davis Station. Data was recorded over 7 months using a custom moored Autonomous Multichannel Acoustic Recorder (AMAR G4) designed and manufactured by JASCO Applied Science following specifications provided by the Australian Antarctic Division. These moored acoustic recorders were designed to operate for year-long, near shore, Antarctic deployments. The moorings were deployed through the ice during the 2021 winter and one retrieved during the 2021/22 summer when the seaice was clear of the surface. The Autonomous Multichannel Acoustic Recorder is a fully autonomous underwater sound and data recorder. The acoustic recorder included a factory calibrated M36-100 hydrophone, data acquisition electronics and solid state digital storage (SDHC) to reduce power consumption and mechanical self-noise (e.g. from hard-drives with motors and rotating disks). Batteries, SDHC cards and electronics were placed in watertight pressure sealed PVC housing rated to a depth of 250 m. The moorings were secured to the seafloor by weights and suspended up into the water column by a string of floats attached to the top of the structure to separate the recorder and hydrophone from sea-bed. The hydrophone was securely mounted to the base of the AMAR housing. All connections between mooring components where taped with protective coverings to reduce mechanical self-noise from movement of the structure. The target noise floor of each recorder was below that expected for a quiet ocean at sea state zero. The data for each recording site comprise folders of 24-bit WAV audio files recorded on a duty cycle with two different sample rates. The duty cycle recorded for 60 s at a sample-rate of 512 kHz, followed immediately by a 580 s at a sample-rate of 32 kHz, and then 280 s off before repeating. The names of each WAV file correspond to instrument serial number followed by the start time (in UTC) of the file as determined by the AMAR’s real-time clock e.g. AMAR897.20210722T061621Z.wav would correspond to a wav from AMAR serial number 897 that starts at 06:16:21 on 22 July 2021 (UTC).

This data set was collected during an ocean acidification mesocosm experiment performed at Davis Station, Antarctica during the 2014/15 summer season. It includes: - description of methods for all data collection and analyses. - diatom cell volume - bulk silicification - species specific silicification via fluorescence microscopy - bulk community Fv/Fm on day 12 - single-cell PAM fluorometry data (maximum quantum yield of PSII: Fv/Fm) A natural community of Antarctic marine microbes from Prydz Bay, East Antarctica were exposed to a range of CO2 concentrations in 650 L minicosms to simulate possible future ocean conditions up to the year ~2200. Diatom silica precipitation rates were examined at CO2 concentrations between 343 to 1641 micro atm, measuring both the total diatom community response and that of individual species, to determine whether ocean acidification may influence future diatom ballast and therefore alter carbon and silica fluxes in the Southern Ocean. Described and analysed in: Antarctic diatom silicification diminishes under ocean acidification (submitted for review) Methods described in: Antarctic diatom silicification diminishes under ocean acidification (submitted for review) Location: Prydz bay, Davis Station, Antarctica (68 degrees 35'S, 77 degrees 58' E) Date: Summer 2014/2015 Worksheet descriptions: Bulk silicification - raw data Measured total and incorporated biogenic silica using spectrophotometer for all tanks on day 12 after 24 h incubation with PDMPO - raw data Bulk Fv/Fm - dark-adapted maximum quantum efficiency of PSII (Fv/Fm) on whole community - raw data Measured Fv/Fm of individual cells from 3 mesocosm tanks. Single-cell silicificiation, Fluorescence microscopy - raw data Measured autofluorescence and PDMPO fluorescence of individual diatoms from 6 mesocosm tanks Single-cell PAM, dark-adapted maximum quantum efficiency of PSII (Fv/Fm) - raw data Measured Fv/Fm of individual cells from 3 mesocosm tanks. Cell volume Calculated cell volume (um3) of 7 species from minicosm tanks 1 and 6 - raw data Abbreviations: Fv/Fm Maximum quantum yield of PSII PDMPO 2-(4-pyridyl)-5-((4-(2-dimethylaminoethylaminocarbamoyl)methoxy)phenyl)oxazole Tant Thalassiosira antarctica DiscLg Large Discoid centric diatoms Stella Stellarima microtrias Chaeto Chaetoceros spp. Prob Proboscia truncata Pseu Pseudonitzschia turgiduloides FragLg Fragilariopsis cylindrus / curta Centric Large Discoid centric diatoms LargeThalassiosira Large Discoid centric diatoms

Experimental Design A six-level, dose-response ocean acidification experiment was run on a natural microbial community from nearshore Antarctica, between 19th November and 7th December 2014. Seawater was collected from approximately 1 km offshore of Davis Station, Antarctica (68◦ 35’ S, 77◦ 58’ E), pre-filtered (200 μm), and transferred into six 650 L tanks (minicosms) located in a temperature-controlled shipping container. Six CO2 levels were achieved by altering the fugacity of carbon dioxide (ƒCO2) within each minicosms. The ƒCO2 was adjusted stepwise to the target concentrations for each minicosm (343, 506, 634, 953, 1140, 1641 μatm) over a five-day period using 0.2 μm filtered seawater enriched with CO2. This acclimation to CO2 was conducted at low light (0.9 ± 0.2 μmol m−2 s−1) so there was low growth of the phytoplankton. Light levels were then increased over a further two days to 90.52 ± 21.45 μmol m−2 on a 19:5 light/dark non-limiting light cycle. After this acclimation period, the microbial community was allowed to grow for 10 days (days 8-18), during which the ƒCO2 levels within each minicosm was adjusted daily to maintain the target ƒCO2 level for each minicosm, and light levels were kept constant. No nutrients were added during the experiment. For a more detailed description of minicosm set-up, lighting and carbonate chemistry see; Davidson, A. T., McKinlay, J., Westwood, K., Thomson, P. G., van den Enden, R., de Salas, M., Wright, S., Johnson, R., and Berry, K.:Enhanced CO2 concentrations change the structure of Antarctic marine microbial communities, Mar. Ecol. Prog. Ser., 552, 93-113, 2016. Deppeler, S. L., Petrou, K., Westwood, K., Pearce, I., Pascoe, P., Schulz, K. G., and Davidson, A. T. Ocean acidification effects on productivity in a coastal Antarctic marine microbial community, Biogeosciences, 15(1), 2018. Sample Collection Samples of 40-400 L were collected and sequentially size-fractionated filtered onto 293 mm biomass filters with 3.0 and 0.1 μm pore-sized polyethersulfone membrane filters (Pall XE20206 Disc 3.0 μm Versapor 293 mm and 656552 Disc 0.1 μm Supor 293 mm) using the design of the Global Ocean Sampling expedition (Rusch et al., 2007). Samples were collected on days 0 (immediately after seawater collection), 12 (mid-exponential growth) and 18 (end of experiment). On day 0, 400 L of seawater was collected from the reservoir tank (pre-filtered 200 μm), from which all the minicosms were filled, to allow characterisation of the initial community. This sample was collected from the reservoir, and not the minicosms, due to the large volume needed to collect sufficient microbial biomass on the filters. On day 12 and 18, 40 L was collected from each minicosm for filtration. The later samples were of a smaller volume due to the increase in biomass in the minicosms during the experiment, meaning less volume of water was required to gain sufficient material on the filters to perform molecular analysis. The filter membranes containing the concentrated microbial biomass were stored in 15 mL of storage buffer, flash frozen in liquid nitrogen and stored at - 80◦C. The storage buffer was freshly prepared on each sampling day with a mixture of 2.5 mM EGTA, 2.5 mM EDTA, 0.1 mM Tris-EDTA, RNA Later (0.5x house prepared), 1 mM PMSF and Protease Inhibitor Cocktail VI (Ng et al., 2010). Between samples the filtration apparatus was sequentially washed with 2 x 25 L 0.1 M NaOH, 2 x 25 L 0.07% Ca(OCl)2 and 2 x 25 L fresh water. All samples were stored and transported at -80◦C to the Australian Antarctic Division, Hobart, Australia for DNA extraction. DNA Extraction and Sequencing The DNA was extracted from half of each filter (3.0 and 0.1 μatm per sample) via the method described in Rusch et al. (2007). In short, the filters were cut into small pieces and agitated in a lysozyme and sucrose buffer for 60 minutes and underwent three freeze/thaw cycles in a Proteinase K solution. This was followed by a gentler agitation at 55◦C for 2 hrs to remove all contents from the filter membranes. DNA was then separated using buffer saturated phenol, pelleted and washed in alcohol. The final DNA pellet was dissolved and stored in a 3 M sodium acetate (pH 8.0) and 100% ethanol solution and stored at - 80◦C. The DNA was transported and stored at 4◦C to the University of Queensland, St Lucia, Australia for sequencing within two months of extraction. Eukaryotic 18S rRNA genes (V8-V9 regions) were amplified using polymerase chain reaction (PCR) with the primers V8f (5’ - AT AAC AGG TCT GTG ATG CCC T - ’3) and 1510r (5’ - CCT TCY GCA GGT TCA CCT AC - ’3) (Bradley, 2016). The 16S rRNA genes V8 region were amplified using PCR and primers 926F (5’-AAA CTY AAA KGA ATT GAC GG-3’) and 1392wR (5’-ACG GGC GGT GTG RC-3’) (Engelbrektson et al., 2010). PCR was performed using 1 or 1.5 μL of sample DNA, 2.5 μL 1x PCR buffer minus Mg+2 (Invitrogen), 0.75 μL MgCl2, 0.5 μL deoxynucleoside triphosphate (dNTPs, Invitrogen), 0.125 μL U Taq DNA Polymerase (Invitrogen), 0.625 μL of forward/reverse primer and made up to the final volume of 25 μL using molecular biology grade water. Forward and reverse primers were modified at the 5’-end to contain an Illumina overhang adaptor with P5 and i7 Nextera XT indices, respectively. The PCR thermocycling conditions were as follows: 94◦C for 3 min, 35 cycles of 94◦C for 45 sec, 55◦C for 30 sec, 7◦C for 10 min and a final extension of 72◦C for 10 min. Amplifications were performed using a Vertiti®96-well Thermocycler (Applied Biosystems) and success, amplicon size and quality was determined by gel electrophoresis. The resultant amplicons were purified using Agencourt AMPure magnetic beads (Axygen Biosciences), dual indexed using Nextera XT Index Kit (Illumina). The indexed amplicons were purified using Agencourt AMPure XP beads and quantified using PicoGreen dsDNA Quantification Kit (Invitrogen). Equal concentrations of each sample were pooled and sequenced on an Illumina MiSeq at the University of Queensland’s School for Earth and Environmental Science using 30% PhiX Control v3 (Illumina) and a MiSeq Reagent Kit v3 (600 cycle; Illumina). Bioinformatics Sequencing data and runs were merged to produced single FASTQ file for 16S and 18S rDNA per sample and imported in QIIME2 (v2019.9) (Caporaso et al., 2010). A modified version of the UPARSE analysis pipeline was used to analyse the data. Specifically, the primer sequences were removed from forward reads of the 16S rDNA and reverse complement of the 18S rDNA Illumina read pairs, and chimeras removed using UCHIME2 (Edgar, 2016). These were then trimmed to a length of 200 bp and high-quality sequences identified using USEARCH (v10.0.240) (Edgar, 2010). Duplicate sequences were removed and a set of unique operational taxonomic units (OTUs) were generated using USEARCH employing a 97% OTU similarity radius. Mitochondrial and chloroplast OTUs were classified and removed from the 16S rDNA sequence data using the BIOM tool suite (McDonald et al., 2012). Representative OTU sequences were assigned taxonomy using SILVA132 (Quast et al., 2012) and PR2 (Guillou et al., 2012) for the eukaryotic group Bacillariophyceae (diatoms). Taxonomic assignments were validated against microscopy identifications conducted on the same samples (Chapter 3, Hancock et al. 2018) as well as phylogenetic trees built in iTOL (Letunic and Bork, 2006). Residual eukaryotic chloroplast and mitochondrial sequences were removed from the 16S rDNA data. Other obvious contaminants were removed manually including: Escherichia-Shigella (16S rDNA OTU75) and Saccharomycetales (18S rDNA OTU7, 146 and 160). Escherichia-shigella was removed as this group likely represents external contamination, similarly Saccharomycetales are yeast and are obvious skin-driven contaminants. A total of 9448 OTUs were identified from the 16S rDNA reads and 232 OTUs from the 18S rDNA read data. The number of reads were rarefied to 1300 and 1200 reads per sample for the 18S and 16S rDNA datasets respectively. The following samples were removed due to lack of extracted, amplified and/or sequenced DNA, or due to low quality reads and/or low read numbers: 18S, 3.0 μm, day 18, 634 μatm ƒCO2 treatment 18S, 0.1 μm, day 12, 343 μatm or control ƒCO2 treatment 18S, 0.1 μm, day 18, 343 μatm or control ƒCO2 treatment 16S, 0.1 μm, day 18, 506 μatm ƒCO2 treatment Statistical Analysis The minicosm experiment was based on a repeated measure design, therefore due to being a dose-response experiment with no replication, no formal statistics could be undertaken on the interactions between time and ƒCO2. The richness (number of taxa) and evenness (equivalent to abundances within a sample) of the eukaryotic and prokaryotic microbial communities within each minicosm over time was estimated using three different alpha diversity indexes: observed number of OTUs (Sobs) (DeSantis et al., 2006), the Chao1 estimator of richness (Colwell et al., 2004), and Simpson’s diversity index and Berger-Parker index which account for both richness and evenness (Simpson, 1949; Berger and Parker, 1970) using QIIME2. Clustering and ordinations were performed on Bray-Curtis resemblance matrices of the rarefied, square-root transformed OTU data as per Chapter 3 (Hancock et al., 2018). In brief, hierarchical agglomerative cluster analyses were performed using group-average linkage, and significantly different clusters were determined using similarity profile permutations method (SIMPROF) (Clarke et al., 2008). Both unconstrained (non-metric multidimensional scaling, nMDS) and constrained (canonical analysis of principal coordinates, CAP) ordinations were performed using the Bray-Curtis resemblance matrixes (Kruskal, 1964a,b; Oksanen et al., 2017). The constraining variables in the CAP analysis were ƒCO2, Si, P and NOx. All cluster and ordination analyses were performed using R v.1.1.453 (R Core Team, 2016) and the add-on package Vegan v.2.5-3 (Oksanen et al., 2017). A full description of the statistical methods used for this paper is described in; Hancock, A. M., Davidson, A. T., McKinlay, J., McMinn, A., Schulz, K. G., and van den Enden, R. L. Ocean acidification changes the structure of an Antarctic coastal protistan community, Biogeosciences, 15(1), 2018.

Oceanographic measurements were conducted in the vicinity of the Amery Ice Shelf on two cruises, during the southern summers of 2000/2001 and 2001/2002. A CTD transect parallel to the front of the Amery Ice Shelf was occupied on both cruises, including repeat occupations on each cruise. A total of 100 CTD vertical profile stations were taken near the ice shelf, most to within 20 m of the bottom, and over 1150 Niskin bottle water samples were collected for the measurement of salinity, dissolved oxygen, nutrients, helium, tritium, oxygen 18 and biological parameters, using a 12 bottle rosette sampler mounted on either a 24 or 12 bottle frame. On the first cruise, an additional 39 CTD stations were occupied around an experimental krill survey area in the vicinity of Mawson. Additional CTD stations were taken at the end of each cruise for calibration of CTD instrumentation from borehole sites on the Amery Ice Shelf. Near surface current data were collected on both cruises using a ship mounted ADCP. An array of 9 moorings comprising current meters, thermosalinographs and upward looking sonars were deployed along the ice shelf front in February 2001 during the first cruise, and retrieved on the second cruise in February 2002. A summary of all data and data quality is presented in the data report.