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The frigid environment under glaciers is inhospitable to all but the most intrepid of microscopic life. To eke out a living, these microbes must do without sunlight and the photosynthetically fixed carbon that fuels most other ecosystems on Earth. Instead, such ecosystems are likely supported by chemosynthetic primary production that capitalizes on energy from inorganic reactions to produce biomass, but the exact mechanisms enabling such chemosynthetic life under the ice are unknown.

Eric Dunham, from Montana State University, USA, and colleagues collected sediments from a glacial system in Iceland that overlays a silicate mineral-rich basaltic catchment, conditions that are prevalent across glacial systems. High concentrations of the reductant hydrogen (H2) were detected, which likely formed when silicate minerals pulverized by the glacier reacted with water. In microcosms seeded with the sediments and amended with H2 and 14CO2, subglacial microbes could oxidize H2, using the resulting energy for chemosynthetic carbon fixation. Metagenomic sequencing from enrichment cultures revealed two prominent autotrophic hydrogenotroph populations, one likely restricted to H2-based chemoautotrophy and one with genomic potential for mixotrophy. The populations exhibited rates of H2 oxidation and carbon fixation approximately tenfold higher than those taken from a Canadian glacier overlying carbonate and shale, suggesting specialization to H2-rich conditions in basalt-glacier systems.

Credit: Natthawat/Getty Images

Interactions between glaciers and rock that can turn an otherwise inhospitable environment into a home for microbes could have implications beyond present-day Earth. Icy H2-dependent primary production could have sustained life during Snowball Earth episodes in our planet’s distant past, or could pave the way for life to evolve on Saturn’s frozen moon Enceladus. More

Stage development and survival rates
Egg duration of G. molesta was not affected by fruit species (F = 0.54, df = 2, 261, P = 0.581), by collection date (F = 0.06, df = 2, 261, P = 0.941), or by fruit species by collection date interaction (F = 0.24, df = 4, 261, P = 0.914) (Table 1). Durations of other life stages were all significantly affected by fruit species (larva F = 28.16, df = 2, 144, P  More

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Sample collection
Algae were sampled from August 27th to September 21st (2017) from seven populations also collected for Bonthond et al. [28], including three native populations; Akkeshi (Japan), Soukanzan (Japan), Rongcheng (China); and four non-native populations; Pleudihen-sur-Rance (France), Nordstrand (Germany), Cape Charles Beach (Viriginia) and Tomales Bay (California, Fig. 1, Table S1). Individuals fixed to hard substratum (see [30]) were sampled at least a meter apart from one another and stored in separate plastic bags. As A. vermiculophyllum has a complex, haplodiplontic life-cycle only diploids were included in the experiment. Life-cycle stages were identified in the field with a dissecting microscope or post-hoc by microsatellite genotyping [31]. After transport in coolers and storage at 4 °C in the lab, bags with algae were shipped to Germany, arriving within 4–6 days after collection. In the climate room (15 °C), individuals were transferred to separate transparent aquaria with transparent lids, containing 1.75 L artificial seawater (ASW) prepared from tap water and 24 gL−1 artificial sea salt without CaCO3 (high CaCO3 concentrations increase disease risk, Weinberger data unpublished) and exposed to 12 h of light per day (86.0 µmol m−2s−1 at the water surface). Aquaria were moderately aerated with aeration stones. Per population, four diploid individuals were acclimated over 31–32 days to climate room conditions prior to starting the experiment. Water was exchanged weekly with new ASW enriched with 2 mL Provasoli-Enrichment Solution (PES; [32]). At the start of the experiment, wet weight was recorded and individuals were divided into two parts of ~10 g each and placed into two plastic tanks with 1.75 L water and 2 mL PES (Fig. 1).
Fig. 1: Schematic overview of the sampling design and experimental process.

Algae were collected from native populations Rongcheng (ron), Soukanzan (sou) and Akkeshi (akk) and non-native populations Tomales Bay (tmb), Cape Charles Beach (ccb), Pleudihen-sur-Rance (fdm) and Nordstrand (nor). In the climate room algae were acclimated for 5 weeks and divided into two thalli. One of the thalli was treated for three days with an antibiotic mixture after which both groups were monitored for six weeks, during which the treated algae received inoculum with each water change. Microbiota samples were taken in the field (tfield), directly after disturbance (t0) and after 1, 2, 4 and 6 weeks (t1, t2, t4 and t6).

Full size image

Experimental setup
To rigorously disturb the microbial community, one of each of the pairs of aquaria containing the same algal individual was treated with a combination of antibiotics, aiming to increase the effectivity (10 mgL−1 ampicillin, 10 mgL−1 streptomycin, 10 mgL−1 chloramphenicol) and the other (control) remained untreated. All experimental work was conducted with disposable gloves and sterilized equipment, to minimize contamination. After three days, the water was removed from all tanks (treated and control) and the wet weight was recorded for all algae. All individuals were rinsed with one 1.75 L volume ASW and re-incubated in 1.75 L ASW. Subsequently, both groups received new ASW with 2 mL PES weekly and individuals treated with antibiotics received also 2 mL inoculum. The inoculum was prepared from individuals of all 7 populations, following the procedure to remove epibiota as described in Bonthond et al. [28]. Briefly, apical fragments of 1 g were separated from the thallus and transferred to 50 mL tubes containing 15 ± 1 glass beads (3 mm) and 15 mL ASW and vortexed for 6 min to separate epibiota from the algal tissue. In total, 8 samples were prepared from one individual per population. The resulting suspensions were pooled and mixed with glycerol (20% final glycerol concentration), aliquoted in 50 mL tubes and stored at −20 °C. For each water exchange, a new aliquot was defrosted at room temperature and added to the water of treated algae. Wet weight was recorded weekly with water exchanges. Before weighing the individual on aluminum foil, it was dipped twice on a separate aluminum foil sheet, to reduce attached water in a systematic way. Endo- and epiphytic microbiota were sampled in the field (tfield, [28]), at the start of the experiment (t0), after one week (t1), two weeks (t2), four weeks (t4) and six weeks (t6, Fig. 1). To equalize acclimation times across populations the experiment was stacked into five groups (Table S2). At each sampling moment, 0.5 or 1 g of tissue was separated from all individuals with sterilized forceps and epibiota were extracted similarly to the preparation of the inoculum. The resulting suspension was filtered through 0.2 µm pore size PCTA filters. Both the filters and the remaining tissue were preserved at −20 °C.
DNA extraction and amplicon sequencing
Tissue samples were defrosted, rinsed with absolute ethanol and DNA free water to remove hydro- and moderately lipophilic cells and molecules from the surface and cut to fragments with sterilized scissors. DNA was then extracted from these fragments (endobiota) and from preserved filters (epibiota) using the ZYMO Fecal/soil microbe kit (D6102; ZYMO-Research, Irvine, CA, USA), following the manufacturer’s protocol. Although this method to separate endo- and epibiota was shown to resolve distinct communities [28], tightly attached epiphytic cells may not be completely removed from the surface and detectable in endophytic samples as well. Two 16S-V4 amplicon libraries, over which the samples were divided in a balanced manner, were prepared as in Bonthond et al. [28], following the two-step PCR strategy from Gohl et al. [33], using the same set of 16S-V4 target primers and indexing primers. The libraries were sequenced on the Illumina MiSeq platform (2×300 PE) at the Max-Planck-Institute for Evolutionary Biology (Plön, Germany), including four negative DNA extraction controls and four negative and positive PCR controls (mock communities; ZYMO-D6311). The fastq files were de-multiplexed (0 mismatches). Relevant field samples from Bonthond et al. [28] were combined with the new dataset and assembled, quality filtered and classified altogether with Mothur v1.43.0 [34] using the SILVA-alignment release 132 [35]. Sequences were clustered within 3% dissimilarity into OTUs using the opticlust algorithm. Mitochondrial, chloroplast, eukaryotic and unclassified sequences were removed. To prepare the community matrix we discarded singleton OTUs (in the full dataset), samples with More

Department of Geography, Texas A&M University, College Station, TX, USA
J. Loisel

Department of Geography, University of Exeter, Exeter, UK
A. V. Gallego-Sala, M. J. Amesbury, D. J. Charman & T. P. Roland

Ecosystems and Environment Research Programme, University of Helsinki, Helsinki, Finland
M. J. Amesbury, A. Korhola, M. Väliranta, S. Juutinen, K. Minkkinen & S. Piilo

Department of Geography and Geotop Research Center, University of Quebec at Montreal, Montreal, Quebec, Canada
G. Magnan & M. Garneau

Magister of Environment and Soil Science Department, Tanjungpura University, Pontianak, Indonesia
G. Anshari

Department of Geography and Environment, University of Hawaii at Manoa, Honolulu, HI, USA
D. W. Beilman

Department of Ecology and Territory, Pontificial Xavierian University, Bogota, Colombia
J. C. Benavides

Organic Geochemistry Unit, School of Chemistry, and School of Earth Sciences, University of Bristol, Bristol, UK
J. Blewett & B. D. A. Naafs

Environmental Studies Program and Earth and Oceanographic Science Department, Bowdoin College, Brunswick, ME, USA
P. Camill

Department of Geology, Chulalongkorn University, Bangkok, Thailand
S. Chawchai

Department of Geography, University of California, Los Angeles, Los Angeles, CA, USA
A. Hedgpeth

Max Planck Institute for Meteorology, Hamburg, Germany
T. Kleinen & V. Brovkin

Faculty of Engineering, Chemical and Environmental Engineering, University of Nottingham, Nottingham, UK
D. Large

Centro de Investigación GAIA Antártica, University of Magallanes, Punta Arenas, Chile
C. A. Mansilla

Climate and Environmental Physics, Physics Institute and Oeschger Centre for Climate Change Research, University of Bern, Bern, Switzerland
J. Müller & F. Joos

Consortium Érudit, Université de Montréal, Montreal, Quebec, Canada
S. van Bellen

Department of Ecology and Conservation Biology, Texas A&M University, College Station, TX, USA
J. B. West

Department of Earth and Environmental Sciences, Lehigh University, Bethlehem, PA, USA
Z. Yu

Institute for Peat and Mire Research, School of Geographical Sciences, Northeast Normal University, Changchun, China
Z. Yu

Department of Environmental Studies, Mount Holyoke College, South Hadley, MA, USA
J. L. Bubier

Department of Geography, McGill University, Montreal, Quebec, Canada
T. Moore

Department of Physical Geography, Stockholm University, Stockholm, Sweden
A. B. K. Sannel

School of Geography, Geology and the Environment, University of Leicester, Leicester, UK
S. Page

Department of Earth and Environmental Sciences, KU Leuven, Leuven, Belgium
M. Bechtold & W. Swinnen

School of Geography & Sustainable Development, University of St Andrews, St Andrews, UK
L. E. S. Cole

Department of Earth, Ocean & Atmospheric Science, Florida State University, Tallahassee, FL, USA
J. P. Chanton

Department of Bioscience, Aarhus University, Roskilde, Denmark
T. R. Christensen

Department of Earth Sciences, University of Toronto, Toronto, Ontario, Canada
M. A. Davies & S. A. Finkelstein

Instituto Franco-Argentino para el Estudio del Clima y sus Impactos, Buenos Aires, Argentina
F. De Vleeschouwer

Institute for the Study of Earth, Oceans, and Space, University of New Hampshire, Durham, NH, USA
S. Frolking & C. Treat

Department of Geobotany and Plant Ecology, University of Lodz, Lodz, Poland
M. Gałka

Laboratoire d’Ecologie Fonctionnelle et Environnement, UMR 5245, CNRS-UPS-INPT, Toulouse, France
L. Gandois

Cranfield Soil and Agrifood Institute, Cranfield University, Cranfield, UK
N. Girkin

Department of Renewable Resources, University of Alberta, Edmonton, Alberta, Canada
L. I. Harris

Stockholm Environment Institute, University of York, York, UK
A. Heinemeyer

Max Planck Institute for Biogeochemistry, Jena, Germany
A. M. Hoyt

Lawrence Berkeley National Laboratory, Berkeley, CA, USA
A. M. Hoyt

Florence Bascom Geoscience Center, United States Geological Survey, Reston, VA, USA
M. C. Jones

Department of Marine and Coastal Environmental Science, Texas A&M University at Galveston, Galveston, TX, USA
K. Kaiser

Department of Biology, University of Victoria, Victoria, British Columbia, Canada
T. Lacourse

Faculty of Geographical and Geological Sciences, Climate Change Ecology Research Unit, Adam Mickiewicz University, Poznań, Poland
M. Lamentowicz

Natural Resources Institute Finland (Luke), Helsinki, Finland
T. Larmola

Agroscope, Zurich, Switzerland
J. Leifeld

Institute for Atmospheric and Earth System Research, University of Helsinki, Helsinki, Finland
A. Lohila

Finnish Meteorological Institute, Climate System Research, Helsinki, Finland
A. Lohila

Department of Geography, Royal Holloway, University of London, Egham, UK
A. M. Milner

Department of Forest Sciences, University of Helsinki, Helsinki, Finland
K. Minkkinen

School of Earth and Environmental Sciences, University of Queensland, Brisbane, Queensland, Australia
P. Moss

Lamont-Doherty Earth Observatory, Palisades, NY, USA
J. Nichols

National Park Service, Washington DC, WA, USA
J. O’Donnell

Department of Environment & Geography, University of York, York, UK
R. Payne

Department of Chemistry, and Department of Geological and Environmental Science, Hope College, Holland, MI, USA
M. Philben

Department of Geography and Environmental Science, University of Reading, Reading, UK
A. Quillet

Department of Applied Earth Sciences, Uva Wellassa University, Badulla, Sri Lanka
A. S. Ratnayake

School of Biosciences, University of Nottingham, Nottingham, UK
S. Sjögersten

Département de Géographie, Université de Montréal, Montréal, Québec, Canada
O. Sonnentag & J. Talbot

Geography, School of Natural and Built Environment, Queen’s University Belfast, Belfast, UK
G. T. Swindles

Department of Environmental Science, Policy, and Management, University of California, Berkeley, Berkeley, CA, USA
A. C. Valach

Department of Environment and Sustainability, Grenfell Campus, Memorial University, Corner Brook, Newfoundland, Canada
J. Wu More