The embryonic development of Antarctic krill (Euphausia superba) is sensitive to elevated seawater CO2 levels. This data set provides the experimental data and WinBUGS code used to estimate hatch rates under experimental CO2 manipulation, as described by Kawaguchi et al. (2013). Kawaguchi S, Ishida A, King R, Raymond B, Waller N, Constable A, Nicol S, Wakita M, Ishimatsu A (2013) Risk maps for Antarctic krill under projected Southern Ocean acidification. Nature Climate Change (in press) Circumpolar pCO2 projection. To estimate oceanic pCO2 under the future CO2 elevated condition, we computed oceanic pCO2 using a three-dimensional ocean carbon cycle model developed for the Ocean Carbon-Cycle Model Intercomparison Project (2,3) and the projected atmospheric CO2 concentrations. The model used, referred to as the Institute for Global Change Research model in the Ocean Carbon-Cycle Model Intercomparison Project, was developed on the basis of that used in ref. 4 for the study of vertical fluxes of particulate organic matter and calcite. It is an offline carbon cycle model using physical variables such as advection and diffusion that are given by the general circulation model. The model was forced by the following four atmospheric CO2 emission scenarios and their extensions to year 2300. RCP8.5: high emission without any specific climate mitigation target; RCP6.0: medium-high emission; RCP 4.5: medium-low emission; and RCP 3.0-PD: low emission (1). Simulated perturbations in dissolved inorganic carbon relative to 1994 (the Global Ocean Data Analysis Project (GLODAP) reference year) were added to the modern dissolved inorganic carbon data in the GLODAP dataset (5). To estimate oceanic pCO2, temperature and salinity from the World Ocean Atlas data set (6) and alkalinity from the GLODAP data set were assumed to be constant. Marine ecosystems of the Southern Ocean are particularly vulnerable to ocean acidification. Antarctic krill (Euphausia superba; hereafter krill) is the key pelagic species of the region and its largest fishery resource. There is therefore concern about the combined effects of climate change, ocean acidification and an expanding fishery on krill and ultimately, their dependent predators—whales, seals and penguins. However, little is known about the sensitivity of krill to ocean acidification. Juvenile and adult krill are already exposed to variable seawater carbonate chemistry because they occupy a range of habitats and migrate both vertically and horizontally on a daily and seasonal basis. Moreover, krill eggs sink from the surface to hatch at 700–1,000m, where the carbon dioxide partial pressure (pCO2 ) in sea water is already greater than it is in the atmosphere. Krill eggs sink passively and so cannot avoid these conditions. Here we describe the sensitivity of krill egg hatch rates to increased CO2, and present a circumpolar risk map of krill hatching success under projected pCO2 levels. We find that important krill habitats of the Weddell Sea and the Haakon VII Sea to the east are likely to become high-risk areas for krill recruitment within a century. Furthermore, unless CO2 emissions are mitigated, the Southern Ocean krill population could collapse by 2300 with dire consequences for the entire ecosystem. The risk_maps folder contains the modelled risk maps for each of the climate change scenarios (i.e. Figure 4 in the main paper, and Figure S2 in the supplementary information). These are in ESRI gridded ASCII format, on a longitude-latitude grid with 1-degree resolution. Refs: 1. Meinshausen, M. et al. The RCP greenhouse gas concentrations and their extensions from 1765 to 2300. Climatic Change 109, 213-241 (2011). Orr, J. C. et al. Anthropogenic ocean acidification over the twenty-first century and its impact on calcifying organisms. Nature 437, 681-686 (2005). Cao, L. et al. The role of ocean transport in the uptake of anthropogenic CO2. Biogeosciences 6, 375-390 (2009). Yamanaka, Y. and Tajika, E. The role

The data reports the pigment concentrations and results of CHEMTAX analysis for 2 summer seasons in Antarctic. In 2008/09 three experiments in which 6 x 650 l minicosms (polythene tanks) were used to incubate natural microbial communities (less than 200 um diameter) at a range of CO2 concentrations while maintained at constant light, temperature and mixing. The communities were pumped from ice-free water ~60 m offshore on 30/12/08, 20/01/09 and 09/02/09. These experiments received no acclimation to CO2 treatment. A further experiment was performed in 2014/15 using water helicoptered from ~ 1 km offshore amongst decomposing fast ice on 19/11/14. This experiment included a 5 day period during which the community was exposed top low light and the CO2 was gradually raised to the target value for each tank, followed by a two day period when the light was raised to an irradiance that was saturating but not inhibitory for photosynthesis. A range of coincident measurements were performed to quantify the structure and function of the microbial community (see Davidson et al. 2016 Mar Ecol Prog Ser 552: 93–113, doi: 10.3354/meps11742 and Thomson et al 2016 Mar Ecol Prog Ser 554: 51–69, 2016, doi: 10.3354/meps11803). The data provides a matrix of samples against component pigment concentration and the output from CHEMTAX that best explained the phytoplankton composition of the community based on the ratios of the component pigments. For the 2008/09 experiments, samples were obtained every 2 days for 10, 12 and 10 days in experiments 1, 2 and 3 respectively. In 2014/15 samples were obtained from each incubation tank on days 1,3, 5, and 8 during th acclimation period and every 2 days until day 18 thereafter. For each sample a measured volume was filtered through 13 mm Whatman GF/F filters for 20 mins. Filters were folded in half, blotted dry, and immediately frozen in liquid nitrogen for analysis in Australia. Pigments were extracted, analysed by HPLC, and quantified following the methods of Wright et al. (2010). Pigments (including Chl a) were extracted from filters with 300 micro l dimethylformamide plus 50 micro l methanol, containing 140 ng apo-8'-carotenal (Fluka) internal standard, followed by bead beating and centrifugation to separate the extract from particulate matter. Extracts (125 micro l) were diluted to 80% with water and analysed on a Waters HPLC using a Waters Symmetry C8 column and a Waters 996 photodiode array detector. Pigments were identified by comparing retention times and spectra to a mixed standard sample from known cultures (Jeffrey and Wright, 1997), run daily before samples. Peak integrations were performed using Waters Empower software, checked manually for corrections, and quantified using the internal standard method (Mantoura and Repeta, 1997).

The data reports the pigment concentrations and results of CHEMTAX analysis for 2 summer seasons in Antarctic. In 2008/09 three experiments in which 6 x 650 l minicosms (polythene tanks) were used to incubate natural microbial communities (less than 200 um diameter) at a range of CO2 concentrations while maintained at constant light, temperature and mixing. The communities were pumped from ice-free water ~60 m offshore on 30/12/08, 20/01/09 and 09/02/09. These experiments received no acclimation to CO2 treatment. A further experiment was performed in 2014/15 using water helicoptered from ~ 1 km offshore amongst decomposing fast ice on 19/11/14. This experiment included a 5 day period during which the community was exposed top low light and the CO2 was gradually raised to the target value for each tank, followed by a two day period when the light was raised to an irradiance that was saturating but not inhibitory for photosynthesis. A range of coincident measurements were performed to quantify the structure and function of the microbial community (see Davidson et al. 2016 Mar Ecol Prog Ser 552: 93–113, doi: 10.3354/meps11742 and Thomson et al 2016 Mar Ecol Prog Ser 554: 51–69, 2016, doi: 10.3354/meps11803). The data provides a matrix of samples against component pigment concentration and the output from CHEMTAX that best explained the phytoplankton composition of the community based on the ratios of the component pigments. For the 2008/09 experiments, samples were obtained every 2 days for 10, 12 and 10 days in experiments 1, 2 and 3 respectively. In 2014/15 samples were obtained from each incubation tank on days 1,3, 5, and 8 during th acclimation period and every 2 days until day 18 thereafter. For each sample a measured volume was filtered through 13 mm Whatman GF/F filters for 20 mins. Filters were folded in half, blotted dry, and immediately frozen in liquid nitrogen for analysis in Australia. Pigments were extracted, analysed by HPLC, and quantified following the methods of Wright et al. (2010). Pigments (including Chl a) were extracted from filters with 300 micro l dimethylformamide plus 50 micro l methanol, containing 140 ng apo-8'-carotenal (Fluka) internal standard, followed by bead beating and centrifugation to separate the extract from particulate matter. Extracts (125 micro l) were diluted to 80% with water and analysed on a Waters HPLC using a Waters Symmetry C8 column and a Waters 996 photodiode array detector. Pigments were identified by comparing retention times and spectra to a mixed standard sample from known cultures (Jeffrey and Wright, 1997), run daily before samples. Peak integrations were performed using Waters Empower software, checked manually for corrections, and quantified using the internal standard method (Mantoura and Repeta, 1997).

We use RNA sequencing to investigate which genetic/physiological pathways in Antarctic krill are affected by increased CO2 levels. We carried out larval CO2 exposure experiments in March 2012 at the AAD aquarium. Two developmental stages were used (Calyptopis I and Furcilia V) and three CO2 levels (control, 1000 and 2000 ppm). These were short term experiments (2 days) - since initial longer experiments starting with fertilized eggs resulted in differences in developmental stages between treatments and control which could confound the data. RNA was extracted from larvae and high-throughput RNA sequencing (RNA-seq) was carried out on 6 samples (2 stages * 3 treatments). Sequencing was carried out on an Illumina sequencer (Genome Analyzer II). We collected ~ 60 million sequence reads per sample (Data in FASTA format each read gives 100 base pairs of sequence), so a total of ~360 million reads (36 billion bp of data).

Metadata record for data from ASAC Project 2518 See the link below for public details on this project. Global climate change will lead to a reduction in the duration and thickness of sea ice in coastal areas. We will determine whether this will lead to a decrease in primary production and food value to higher predators. Project objectives: Our primary objective is to determine what effect will declining sea ice cover have on Antarctic coastal primary production? Hypotheses to be tested A decrease in sea ice algal production will lead to a net reduction in total primary production. A decrease in sea ice will result in less water column stratification which will reduce the significance of phytoplankton blooms. Less sea ice will lead to a change in phytoplankton bloom composition away from diatoms towards un-nutritious nuisance blooms such as Phaeocystis Benthic microalgal production will increase Seaweed production will increase slightly A decrease in sea ice thickness will increase ice algal production (as they are generally light limited) Ice algae, benthic microalgae, and phytoplankton will acclimate to an elevated light climates by changing their photosynthetic efficiency and capacity Ice algae, benthic microalgae, and phytoplankton will acclimate to an altered light quality. To answer these questions we will also need to determine: What is the total annual primary production at coastal Antarctic sites; this consists of the contributions from the sea ice algal mats, benthic microalgal, seaweed and phytoplankton? What is the effect of major environmental variables, such as UV, salinity, currents oxygen toxicity, cloud cover, nutrient availability and temperature on production. What is the inter-annual variability in primary production? A broader scale issue that our data will contribute to providing answers to is the question What effect will changing primary production have on higher trophic levels? Taken from the 2009-2010 Progress Report: Progress against objectives: The 2009/10 field and laboratory season focused on the second of our primary questions, i.e. 'What is the effect of major environmental variables, such as UV, salinity, currents oxygen toxicity, cloud cover, nutrient availability and temperature on production'. In particular we focused on light and light transmission though the sea ice. The science program AAS2518 was executed at Casey station from 11 Nov to 5 Dec 2009. The project was split into a field and a lab-based component. In situ spectral light transmission data were collected on first year sea ice within the vicinity of Jack's Hut. Ice cores were collected and transported to the laboratory at Casey station for spectral attenuation profiles within sea ice, and for measurements of spectral absorption by particulate and dissolved organic matter. Overall, the program was successful: in situ sea-ice spectral transmission data was collected in combination with vertical profiles of absorption coefficients of particulate (algae and detritus) and dissolved organic matter. Samples for analysis of photosynthetic pigments were collected and shipped to Sydney. Their analysis is underway. Due to logistical issues associated with the collection and transport of sea ice cores, the protocol for vertical profiling of spectral attenuation was modified (see below) and analysis of the data is currently underway. The field component of the program was successful as spectral transmission data was collected for first year sea-ice, and the chosen site contained a thriving sea ice algal community for bio-optical measurements. It was initially planned to sample multiple sites offering a range of varying sea-ice thickness, but this was not possible during this campaign. Many sites in the vicinity of Casey station had already started to melt and break up, so that for logistical and safety reasons the area around Jack's hut was the only workable option. The field period instead spanned ~ 20 days during the melt period at Jack's,

This data may be used to reproduce the analyses (including figures and tables), of 'Two scales of distribution and biomass of Antarctic krill (Euphausia superba) in the eastern sector of the CCAMLR Division 58.4.2 (55°E to 80°E)'. The data describe krill biomass density distribution and krill net samples (krill total length and krill wetmass) collected during the 2021 TEMPO voyage on R/V Investigator. During the TEMPO voyage krill biomass was estimated using observations from two sampling instruments: a calibrated EK80 scientific echosounder operating at 120 kHz and an rectangular midwater trawl (RMT 1+8). The supporting data sets, all in CSV format, are split by instrument type. The EK80 has two datafiles: krill_density.csv – krill areal density from the TEMPO transects, and krill_swarms.csv -krill swarms detected during the TEMPO transects. The RMT1+8 has four datafiles net_locations.csv krill_lengths.csv krill_wet_mass_to_length.csv krill_wet_mass_to_length_model_predictions.csv The fields (columns) in each data file are: krill_density.csv "lat_M" – centre latitude of an echo integration interval [degrees] (dd.ddddd) WGS84 spheroid (GPS latitude) "lon_M" - centre longitude of an echo integration interval [degrees] (dd.ddddd) WGS84 spheroid (GPS longitude), "areal_biomass_density_g_per_m2" – Echo integration interval krill areal biomass density [g wet-mass / m^2] "daynight" – flag for when the sampling took place [day/night] "survey" – Either the main survey for the TEMPO biomass survey or the smaller-scale ‘Mawson box’ survey krill_swarms.csv "transect" – transect number "lat" – latitude [degrees] (dd.ddddd) WGS84 spheroid (GPS latitude) "swarm_depth_m" – mean depth of a krill swarm [m] "daynight" – flag for when the sampling took place [day/night] ”volumetric_density_g_per_m3" – krill swarm internal volumetric biomass density [g wet-mass / m^3] net_locations.csv "station" – Station name for net trawl R for routine haul, T for target trawl "lat" – mean latitude of a net trawl [degrees] (dd.ddddd) WGS84 spheroid (GPS latitude) "lon" – mean longitude of a net trawl [degrees] (dd.ddddd) WGS84 spheroid (GPS longitude) "daynight" – flag for when the sampling took place [day/night] krill_lengths.csv "station" – Station name for net trawl R for routine haul, T for target trawl "total_length_mm” – total length of an individual krill [mm] krill_wet_mass_to_length.csv "total_length_mm" – total length of an individual krill [mm] "wet_mass_g" - wet-mass an individual krill [g] krill_wet_mass_to_length_model_predictions.csv "total_length_mm" - total length of an individual krill [mm] "predicted_wet_mass_g" – predicted mean wet-mass an individual krill of length ("total_length_mm" ) [g] "LB_wet_mass_g" – Lower bound (lower 95% confidence interval) for the predicted_wet_mass_g [g] "UB_wet_mass_g"– Upper bound (upper 95% confidence interval) for the predicted_wet_mass_g [g]

A meta-analysis was undertaken to examine the vulnerability of Antarctic marine biota occupying waters south of 60 degrees S to ocean acidification. Comprehensive database searches were conducted to compile all English language, peer-reviewed journals articles and literature reviews that investigated the effect of altered seawater carbonate chemistry on Southern Ocean and/or Antarctic marine organisms. A document detailing the methods used to collect these data is included in the download file.

Antarctic krill (Euphausia superba) have a keystone role in the Southern Ocean, as the primary prey of Antarctic predators. Any decreases in krill abundance could result in a major ecological regime shift, but there is currently limited information on how climate change may affect krill. Increasing anthropogenic carbon dioxide (CO2) emissions are causing ocean acidification, as absorption of atmospheric CO2 in seawater alters ocean chemistry. Ocean acidification increases mortality and negatively affects physiological functioning in some marine invertebrates, and is predicted to occur most rapidly at high latitudes. Here we show that, in the laboratory, adult krill are able to survive, grow, store fat, mature, and maintain respiration rates when exposed to near-future ocean acidification (1000 – 2000 μatm pCO2) for one year. Despite differences in seawater pCO2 incubation conditions, adult krill are able to actively maintain the acid-base balance of their body fluids in near-future pCO2, which enhances their resilience to ocean acidification.

This data set was collected from a ocean acidification minicosm experiment performed at Davis Station, Antarctica during the 2014/15 summer season. It includes: - description of methods for all data collection and analyses. - environmental data logged throughout the experiment; nutrients, temperature, light climate. - carbonate chemistry data; pH (on Total scale), fugacity of CO2, dissolved inorganic carbon concentration, practical alkalinity, Omega calculations for both araganite and calcite. - product datasheet (including transmission spectra) of Osram 150W HQI-TS/NDL metal halide lamps.

Experimental Set-up: An unreplicated, 6-level, dose-response experiment was conducted on a natural microbial community over a range of pCO2 levels (343, 506, 634, 953, 1140 and 1641 micro atm). Seawater was collected on the 19th November 2014 approximately 1 km offshore from Davis Station, Antarctica (68 degrees 35' S, 77 degrees 58' E) from an area of ice-free water amongst broken fast-ice. The seawater was collected using a thoroughly rinsed 720L Bambi bucket slung beneath a helicopter and transferred into a 7000 L polypropalene reservoir tank. Six 650 L polyethene tanks (minicosms), located in a temperature-controlled shipping container, were immediately filled via teflon lined house via gravity with an in-line 200 micron Arkal filter to exclude metazooplankton. The minicosms were simultaneously filled to ensure they contained the same starting community. The ambient water temperature at time of collection was -1.0 degrees C and the minicosms were maintained at a temperature of 0 degrees C plus or minus 0.5 degrees C. At the centre of each minicosm there was an auger shielded for much of its length by a tube of polythene. This auger was rotated at 15 rpm to gently mix the contents of the tanks. Each minicosm tank was covered with an acrylic air-tight lid to prevent pCO2 off-gasing outside of the minicosm headspace. The minicosm experiment was conducted between the 19th November and the 7th December 2014. Initially, the contents of the tanks were given a day to equibrate to the minicosms. This was followed by a five day acclimation period to increasing pCO2 at low light (0.8 plus or minus 0.2 micro mol m-1 s-1), allowing cell physiology to acclimated to the pCO2 increase (days 1-5). During this period the pCO2 was progressively adjusted over five days to the target level for each tank (343 - 1641 micro atm). Thereafter pCO2 was adjusted daily to maintain the pCO2 level in each treatment (see carbonate chemistry section below). Following acclimation to the various pCO2 treatments light was progressively adjusted to 89 plus or minus 16 micro mol m-2 s-1 at a 19 h light:5 h dark cycle. The community was incubated and allowed to grow for a further 10 days (days 8-18) with target pCO2 adjusted back to target each day (see carbonate chemistry section below). 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, 2017. Light microscopy sampling and analysis: Samples from each minicosm were collected on days 1, 3, 5, 8, 10, 12, 14, 16 and 18 for microscopic analysis to determine protistan identity and abundance. Approximately 960 mL were collected from each tank, on each day. Samples were fixed with 20 40 mL of Lugol's iodine and allowed to sediment out at 4 degrees C for greater than or equal to 4 days. Once cells had settled the supernatant was gently aspirated till approximately 200 mL remained. This was transferred to a 250 mL measuring cylinder, again allowed to settle (as above), and the supernatant gently aspirated. The remaining 20 mL. This final 20 mL was transferred into a 30 mL amber glass bottle. All samples were stored and transported at 4 degrees C to the Australian Antarctic Division, Hobart, Australia for analysis. Lugols-fixed and sedimented samples were analysed by light microscopy between July 2015 and February 2017. Between 2 to 10 mL (depending on cell-density) of lugols-concentrated samples was placed into a 10 mL Utermohl cylinder (Hydro-Bios, Keil) and the cells allowed to settle overnight. Due to the