Inorganic carbon addition stimulates snow algae primary productivity

Earth has experienced glacial/interglacial oscillations accompanied by changes in atmospheric CO2 throughout much of its history. Today over 15 million square kilometers of Earth’s land surface is covered in ice including glaciers, ice caps, and ice sheets. Glaciers are teeming with life and supraglacial snow and ice surfaces are often darkened by the presence of photoautotrophic snow algae, resulting in accelerated melt due to lowered albedo. Few studies report the productivity of snow algal communities and the parameters which constrain their growth on supraglacial surfaces—key factors for quantifying biologically induced albedo effects (bio-albedo). We demonstrate that snow algae primary productivity is stimulated by the addition of inorganic carbon. Our results indicate a positive feedback between increasing CO2 and snow algal primary productivity, underscoring the need for robust climate models of past and present glacial/interglacial oscillations to include feedbacks between supraglacial primary productivity, albedo, and atmospheric CO2.

by the Laboratory for Isotopes and Metals in the Environment at the Pennsylvania State University.

CO2 photoassimilation and stable isotope signals
A microcosm-based approach was employed to assess in situ inorganic carbon uptake through the addition of NaH 13 CO3 as we have described previously (Hamilton and Havig, 2017). While we are aware of debate regarding validity of microcosm amendment studies for studying microbial activities in snow, particularly dry snow (i.e., Carpenter et al., 2000), we targeted areas of active melt with snow of high water content and samples were collected during the Summer months when daytime ambient temperatures were well above freezing (~12-14ºC). In our previous study (Hamilton and Havig, 2017), we observed inorganic carbon uptake following a 60-minute incubation time in ~15-mL assays amended with 100 µM NaH 13 CO3. Here we employed a similar method: The surface layer of wet snow where phototrophic populations were visibly apparent (red or orange colored snow) was collected using a sterile spatula and placed into a clean / sterile container. The snow was allowed to melt to a snow-slush slurry (~15 minutes) and was homogenized with the spatula. Approximately 20-mL of the snow-slush slurry was then transferred to clear 40-mL acid-washed screw-cap polypropylene bottles. Assays were then initiated by amending with NaH 13 CO3 (Cambridge Isotope Laboratories, Inc., Andover MA). NaH 13 CO3 stock solutions were prepared with 18.2 MΩ/cm deionized water such that 100 µL was added to each microcosm to reach the final concentration of amendment (50 µM, 100 µM, 500 µM, and 1 mM).
NaH 13 CO3 was added using a sterile syringe. Following amendment, bottles were briefly and gently shaken to distribute the label. Bottles were wrapped in foil for dark assays. Bottle control assays and assays for natural abundance received unlabeled NaH 13 CO3 prepared as described above. All assays were performed in triplicate. Bottles were pushed into areas of soft snow near the collection site such that approximately 50% of the bottle was directly exposed to sunlight and incubated for approximately 2 hours in situ. To stop the assays, bottles were flash-frozen on dry ice, transported on dry ice, and stored at -20°C until processing.
For processing, assays were thawed and biomass was washed with HCl (1 M) to remove any carbonate minerals and residual NaH 13 CO3, then washed with 18.2 MΩ/cm deionized water and dried (60°C for three days). Dried biomass was ground/homogenized with a clean mortar and pestle. Dried and ground samples for determination of carbon concentration and stable isotope signal were weighed and placed into tin boats and sealed. Samples were analyzed via a Costech Instruments Elemental Analyzer (EA) periphery connected to a Thermo Scientific Delta V Advantage Isotope Ratio Mass Spectrometer (IR-MS) in the Department of Geology at the University of Cincinnati. Linearity corrections were made using NIST Standard 2710. δ 13 C values were calibrated using reference standards USGS-40 and USGS-41 and checked with a laboratory standard (glycine). All carbon stable isotope results are given in delta formation expressed as per mil (‰). Carbon stable isotopes were calculated according to the following equation: where Ra is the 13 C/ 12 C ratio of the sample or standard, and are reported versus the Vienna Pee Dee Belemnite (VPDB) standard. Reported values of DIC uptake (carbon fixation rates) were calculated using the difference in absolute 13 C/ 12 C ratios between the labeled assays and unlabeled controls. 16S rRNA sequences were targeted using the primers (515f) (Caporaso et al., 2012) and 806rB (Apprill et al., 2015).  (Schloss et al., 2009) following the MiSeq SOP (Kozich et al., 2013). Read pairs were assembled and resulting contigs with ambiguous bases were removed. Contigs were trimmed to include only the overlapping regions and unique sequences were aligned against a SILVA-based reference alignment and classified using a Bayesian classifier within Mothur against the against the Silva (v128) reference taxonomy. Chimeras were identified and removed using UCHIME (Edgar et al., 2011).

DNA extraction and sequence analysis
Sequences were classified Operational taxonomic units (OTUs) were assigned to all classified sequences at a sequence similarity of 97.0% for archaea and bacteria and 98% for eukarya using the averageneighbor algorithm. Rarefaction was calculated within mothur and based on rarefaction analysis, >95% of the predicted 16S and 18S rRNA gene diversity was sampled at this depth of sequencing (data not shown