Global warming facilitated environmental change effects on CO2 releasing microbes in Antarctic sediments

doi:10.21203/rs.3.rs-5441636/v1

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Global warming facilitated environmental change effects on CO₂ releasing microbes in Antarctic sediments

https://doi.org/10.21203/rs.3.rs-5441636/v1

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Rapid melting of the Western Antarctic Peninsula (WAP) glaciers is a compelling piece of evidence of how climate change affects our planet. This study investigated the impact of global warming-facilitated environmental change on microbial community structure and function by subjecting sediments sampled near the Fourcade Glacier in Potter Cove, WAP, to a temperature gradient and supply of metabolic nutrients relevant for the fate of carbon in marine ecosystems. We found that (i) temperature as a key environmental change driver will significantly impact microbial community structure, but ecological functions supported by fresh supply of nutrients from glacial meltwater will prevail; (ii) keystone species responsible for specialized functions are metabolically flexible, persisting from 2°C to 25°C; and (iii) in addition to keystone species, global warming will activate certain hitherto inactive but endogenous microorganisms in response to either changes in temperature or nutrient flux to sustain ecosystem functions. Our study presents evidence of sediment microbiome resilience in response to strong temperature or nutrient flux shifts, thereby adding another layer of evidence of nature’s adaptability to global warming.

global warming

Antarctica

marine sediment

RNA stable isotope probing

organic matter mineralization

acetate

The marine biosphere contributes about 45 percent (global net primary productivity = ~ 49 Pg C year^− 1) to the amount of organic matter (OM) degraded or preserved in the earth system¹. The terminal steps in the degradation of organic matter in marine sediments involve coupling fermentation intermediate (H₂/acetate) oxidation to manganese, Fe(III), sulfate, and CO₂ reduction. Therefore, microbes that respire sulfate and metal oxides as terminal electron acceptors are important for the final step of OM mineralization to CO₂ ². A previous study³ showed that these specialists microbes represent keystone species shaping the microbial community composition and function in permanently cold sediments, irrespective of their abundance⁴. Environmental change, such as increased temperature due to global warming, might affect the rates and pathways of OM degradation, and possibly causes a shift in compositions of communities involved in determining the final fate of OM in the environment. It is therefore important to study the effects of environmental change on abundance and activity of sulfate and metal-reducing microbes to enhance knowledge of the terminal fate of organic matter. Understanding the mechanistic control of organic matter degradation and preservation during transport and burial is essential for assessing the impacts of past, present, and future climate and, thus, environmental change on the marine carbon cycle².

The Western Antarctic Peninsula (WAP) is one of the fastest-warming places on earth⁵. As a result, terrestrial and marine-terminating glaciers in the area have melted at unprecedented rates. For example, the Fourcade Glacier (Fig. 1), a polythermal glacier draining into Potter Cove, King George Island/Isla 25 de Mayo, WAP, has receded by at least 1.5 km² between 1956 and 2018 ⁶, creating several newly ice-free areas and transforming from an initial tidewater glacier to land-terminating glacier^7,8. Consequently, glacial meltwater transports easily reducible Fe (oxyhydr)oxides either directly, via subglacial groundwater discharge, or via icebergs that are known to contain reactive Fe nanoparticles, into the sediments⁹. The implications that this environmental change brought about by global warming for sediment microbial communities, especially regarding how they respond to and mineralize OM in Antarctica, is barely explored.

It is difficult to detect extreme temperature effects in situ on organisms and their biology within realistic climate relevant experimental timeframes (about 20–25 years¹⁰). Therefore, previous studies evaluating climate change effects on responses of (Ant)arctic biotic communities have either used laboratory cultures to simulate long-term climate change effects^11,12 or network analysis of clustering similarities of metagenomic data¹³. The slow growth rates of marine sediment microbes present another constraint for designing such studies¹⁴. This constraint has been mitigated by the development of stable isotope probing of DNA and RNA. RNA-stable isotope probing (RNA-SIP) confers the advantage that microbial physiological activity can be studied within a relatively short incubation period under conditions that closely resemble those of the natural environment, thereby avoiding the enrichment of artificial communities that do not reflect those relevant conditions¹⁵.

Here, we combine stable isotope and molecular biology techniques to investigate the survival strategies of microorganisms that influence the fate of organic matter in marine sediments under current global warming scenarios. We also discuss the implications of microbial adaptation to global warming for the release of CO₂ from marine sediments.

Sediment samples were collected from the closest possible distance to the Fourcade Glacier, and used for an analysis of how keystone microbes for the terminal mineralization of organic matter in sediments adapt to changes in temperature (5°C, 10°C, 15°C, 20°C, 25°C and 30°C) versus the baseline in situ temperature of 2°C ¹⁶. While some of the temperatures the sediments were subjected to for the study are experimental, the WAP remains one of the fastest warming places on earth, and continues to experience unprecedented and extreme warming events with certain areas recording up to 17.1°C to 18.3°C recently¹⁷. To avoid the enrichment of target microbes and promote the possibility of mimicking environmental conditions, sediment incubations were supplied with traceable acetate in low concentrations (500 µM ¹³C) as carbon source. The incubations were also amended with or without fresh electron acceptors to optimize conditions for anaerobic respiration (iron and sulfate reduction). A similar set-up was used with unlabeled acetate (¹²C) as control. The sediments were incubated in total for 17 days or less depending on rates of CO₂ release in the incubations (suppl. Fig. S1) as indicator for microbial activity, and then subjected to RNASIP. The results show that (i) ecological functions in the sediments prevail over the applied temperature range albeit at different rates, despite the change in active microorganisms performing those ecological functions; (ii) keystone microorganisms are more metabolically flexible than expected; certain species persisted from psychrophilic (2°C) to mesophilic (25°C) conditions; and (iii) as warming and glacier retreat continues to affect the nutrient balance of Antarctic sediments, microbial communities will develop adaptation strategies that may reactivate hitherto inactive or low activity microbial populations.

Temperature and nutrients have a significant effect on microbial community structure

Recession of the Fourcade Glacier results in increased glacial meltwater discharge, supplying more suspended particulate matter including iron oxides to coastal waters and underlying sediments¹⁸, thereby changing the dynamics of carbon mineralization. Therefore, the experimental design aimed to mimic the prevailing environmental conditions in the sediments using acetate and lepidocrocite (a low crystallinity, bioavailable γ-FeO(OH)) to stimulate iron reduction as a consequence of a fresh supply of iron oxides as nutrients to the sediments⁹. Subglacial meltwater can supply more labile iron, which is more bioavailable to microorganisms performing iron reduction with OM degradation intermediates such as acetate⁹. However, the glacier has recently started to recede to land⁸, which may change the supply and characteristics of iron oxide with predicted possible decrease in suspended particulate matter¹⁹. This scenario potentially favors other processes such as sulfate reduction over iron reduction²⁰, since the sediment is replete with up to 28 mM sulfate²¹. Therefore, a separate set of experiments was set up with acetate and sulfate to mimic this scenario of reduced iron supply to the sediments. These conditions are optimal for sulfate reduction or sulfur cycling, which likely becomes more important in an emerging environmental change scenario of very low iron input due to receding glaciers to land²⁰. As a baseline control for both scenarios, “acetate only” treatments were also set up. Acetate was selected as a representative volatile fatty acid intermediate of fermentation²², which is obligatorily used by anaerobically respiring microorganisms as a terminal electron donor and is also suitable for anabolic respiration by organotrophic organisms. In all treatments across all temperatures, we obtained indicators for ¹³CO₂ release from microbial catabolic consumption of ¹³C-acetate (suppl. Fig. S1) and incorporation of ¹³C-carbon from acetate into the biomass of active organisms (suppl. Fig. S2). The experiments were stopped at different days, based on the pace of build-up of ¹³CO₂ in the incubations i.e., after 10 days at 2°C and 5°C, 11 days at 10°C and 15°C, 15 days at 30°C and 17 days at 20°C and 25°C. Sulfide was not detected in the incubations and a minor net sulfate concentration decrease was detected, mainly between 10°C and 20°C (suppl. Fig. S3a). A qualitative measurement of Fe²⁺ build up in the relevant iron amended treatments was used as proxy for iron reduction rates. Iron reduction rates were optimal at 5°C and 10°C but decreased consistently as the temperature gradient became more mesophilic (suppl. Fig. S3b).

After sequencing 16S rRNA in the labeled and unlabeled fractions obtained from RNA-SIP experiments, distance-based redundancy analyses (dbRDA), which combines a distance matrix calculation and principal coordinate analysis, were performed on the sequence data to evaluate community composition. The results show significant influence of temperature and nutrient (sulfate and iron as electron acceptors) on microbial communities (Fig. 2; suppl. Table S1, Fig. S4). Temperature had a significant effect (F(1,205) = 53.4; p < 0.001) on the microbial community composition across all treatments (Fig. 2a). The further the experimental temperature differed from the in-situ temperature, the more significant was the shift in microbial community composition across all treatments (suppl. Table S1). The addition of electron acceptors had a similar, but less pronounced significant effect (F(2,205) = 3.9; p < 0.001) on the microbial community compositions (Fig. 2; suppl. Table S1). The influence of electron acceptors was, however, more pronounced than temperature when comparing the communities at 2°C and 5°C across all treatments and within the ¹³C-labelled treatment fractions reflecting active use of ¹³Cacetate for anabolic function (heavy + ultra-heavy fractions; suppl. Table S1, treatment F(1,8) = 3.27; p = 0.012; temperature p > 0.05). This finding demonstrates a probable scenario in situ. The Northern and Northwestern Antarctic Peninsula have experienced a warming trend of 0.46°C ± 0.96°C per decade between 1951 and 2018 or of 3.12°C ± 1.02°C in total over the same period⁵. These warming rates are alarming as they represent a 2.5 times increase when compared to the rest of the world’s warming rate at currently 0.18°C to 0.19°C per decade over the last 50year period^{23, 24}. Given the current scenarios, a 3°C warming over the next 50–100 years might occur in the WAP^{25, 26}. Our study predicts that in the likelihood of increased warming, microbial community structures will be impacted, thereby triggering community adaptation to sustain ecosystem function. As the warming is accompanied by a change in nutrient composition or an initial increase and subsequent cease in the supply of a particular nutrient (such as iron), microbial community composition will also change, selecting for new dominant members better suited to the shifted environmental conditions.

Iron reducers and sulfur-cycling organisms dominate the incubations

The active microorganisms identified in the microbial communities across all treatments (Fig. 3; suppl. Fig. S5-S15) use acetate for anabolic processes and are well known for their distinct ecosystem function. We classified them to sequence level (ASV: amplicon sequence variant) to define the ecology of each organism under environmental change scenarios. The dominant microorganisms included iron reducing bacteria such as (i) Sva1033 (Fig. 3a), previously identified in permanently cold sediments^3,27,28; (ii) Desulfuromusa (Fig. 3b), a well-known metal oxide reducer able to switch between manganese and iron reduction²⁹; (iii) unclassified Geopsychrobacteraceae (Fig. 3c), very close relatives of Desulfuromusa, which harbors many species previously described as iron reducers³⁰; (iv) unclassified members of the order Desulfuromonadales (Fig. 4d, l), the overarching order of iron reducers³¹; (v) Desulfuromonas (Fig. 3e), a well-studied dissimilatory iron reducing genus often identified in multiple environments from temperate to permanently cold areas^29,32; and (vi) Trichloromonas (formerly Desulfuromonas³³; Fig. 3f) from the Desulfuromonadaceae family (according to used silva taxonomy release 138.1³⁴) of iron reducers³⁵.

Other active organisms included those well studied for their role in sulfur cycling such as (i) Sulfurimonas (Fig. 3g)³⁶, (ii) Sulfurospirillum (Fig. 3h)³⁷; (iii) Desulfobacter (Fig. 3i), a well-known sulfate reducer³⁸ that was only stimulated at 20°C with sulfate addition; and (iv) unclassified Arcobacteraceae (Fig. 3j), detected only at 2°C and known for its sulfur cycling or metal reduction capabilities^28,39. Certain identified taxa were stimulated at specific temperatures but not previously connected to sulfur or iron metabolisms inter alia; (i) Colwellia (Fig. 3k), an extremely psychrophilic heterotrophic bacteria found in cold-polar sediments, sea ice and the deep sea^28,40 and (ii) Hoeflea (Fig. 3m), previously isolated from temperate halophilic marine settings⁴¹.

The high relative abundance and capacity to thrive over a wide range of temperature suggest that iron and sulfur cycling organisms dominate and shape the ecosystem functioning of the microbial communities in our incubations (Fig. 3), as similarly found in sub-Antarctic South Georgia sediments³. The addition of fresh iron oxide promoted the predominance of seven different organisms (i.e., ASVs) of Sva1033 over a wide temperature range, the most of any taxon stimulated in this study. This result suggests that Sva1033 will remain the dominant key microorganism for the terminal steps of OM degradation in Antarctic sediments if the current scenario of glacial recession continues to provide fresh iron to the sediments as part of glacial meltwater. Sva1033 was less competitive in the treatments without adding fresh iron oxide, providing the opportunity for other iron reducers such as Desulfuromonas, Trichloromonas, and unclassified Geopsychrobacteraceae to thrive competitively in those treatments (Fig. 3). The observation that Sva1033 was not competitive when fresh iron oxide supply was limiting and giving rise to the dominance of other recognized iron reducers, especially Desulfuromonas, is interesting. It predicts the adaptation pattern of the microbial communities in the sediments to the recently emerging scenario of the Fourcade Glacier receding to land that may eventually result in a decrease of fresh iron supply. Nutrient supply was not the only factor determining the dominance of the activity of one microbe over the other in these experiments as temperature had a similar effect. At 30°C, we observed that all the previous iron reducers that competed favorably and were active, regardless of the nutrient conditions in the sediments, were absent, except for Desulfuromusa, with two distinct organisms that were only stimulated when the conditions became mesophilic (20°C to 30°C). At 30°C, these Desulfuromusa organisms were the only iron reducers stimulated to retain iron reducing function in these microbial communities, maintaining 50–55% relative abundance of the entire active populations in all treatments. In another study done at the same study site, Desulfuromusa was the main active metal oxide reducer at the in situ temperature of 2°C metabolizing acetate with manganese as the preferred nutrient instead of iron oxides as electron acceptor⁴². Here, we provide further insight into the ecosystem function of Desulfuromusa in cold environments, that they are likely outcompeted for growth on acetate by microbes such as Sva1033 and Desulfuromonas when the dominant electron acceptor is iron, and the conditions are optimally psychrophilic. However, they can become competitive when manganese is the main nutrient available to support acetate consumption or the sediments are subjected to warmer environmental conditions.

The alternating abundance of Sulfurimonas and Sulfurospirillum as active populations, mainly in the sulfate amended and the control treatments was interesting (Fig. 3g, h). Both play a similar ecosystem function facilitating sulfur cycling, yet they were well differentiated in proliferation between psychrophilic and mesophilic conditions. Sulfurimonas was more adapted to psychrophilic metabolism and was barely detected when the temperature increased to 20°C (Fig. 3g). Sulfurospirillum was detected with a high abundance as Sulfurimonas at low temperatures, but only at 20°C and 25°C (Fig. 3h). Despite the clear stimulation of microbial activity with sulfate (Fig. 3; suppl. Fig. S1, S3), the only known sulfate reducer identified amongst the active communities was Desulfobacter which displayed a 10% relative abundance with sulfate addition at 20°C (Fig. 3i). Nevertheless, we tested for the prevalence of sulfate reduction activity by quantitative polymerase chain reaction (qPCR) on the messenger RNA (mRNA) transcripts using primers specific for the detection of the dissimilatory sulfite reductase gene alpha subunit (dsrA) which encodes the activity for sulfite reduction to sulfide (suppl. Fig. S16). We found that up to 6x10¹² copies of dsrA transcripts were present per ng of complementary DNA (cDNA) across treatments except at 30°C. A qPCR targeting the bacterial 16S rRNA transcript detected up to 6x10¹⁵ copies of bacterial 16S rRNA per ng of cDNA in the same samples, thereby confirming that the lack of detection of the dsrA transcript at that temperature was not due to experimental detection limits. This finding clarifies that sulfate reduction is ongoing in the sediments at in situ conditions and up to 25°C as part of the cryptic sulfur cycle, even as sulfate reducers are not competitive for use of acetate for biomass production. At 30°C, the absence of sulfate reducers in the heavy fractions, and the non-detection of the dsrA mRNA transcript as a signature for sulfate reduction suggests sulfate reduction as a microbial process in the sediments stopped thriving.

Distinct keystone species showing metabolic flexibility from psychrophilic to mesophilic temperature

We identified five distinct keystone species – based on physiological capabilities rather than network analysis-based co-occurrence – among the specific organisms (ASVs) that adapted to the effects of increased temperature with great metabolic flexibility, transitioning from psychrophilic to mesophilic metabolism to sustain the key ecosystem function of iron reduction. These include Sva1033, Desulfuromusa, Desulfuromonas, an unclassified Geopsychrobacteraceae, and Trichloromonas (Fig. 3). Amongst the seven observed organisms of Sva1033 (Fig. 3a), two displayed the metabolic flexibility to survive from 2°C to 20°C. The first, labeled for identification purposes as ASV 1, was most active at 2°C (23%) and remained enriched, displaying metabolic flexibility up to 20°C (10%). ASV 10, although growing optimally at 20°C (13%), was active down to 2°C (5%) while ASV 43 and ASV 7, which grew optimally at 10°C, remained psychrophilic. There were three organisms that grew only at mesophilic temperatures and were not stimulated at psychrophilic ones: ASV 12 (37% only at 25°C), ASV 44 (9% at 25°C) and ASV 93 (6%) only at 20°C. Family Sva1033 was previously identified as an iron reducer only in permanently cold sediments^3,28. The findings here suggest that there are certain Sva1033 organisms that prefer mesophilic temperatures, and that this iron reducer is metabolically more flexible than previously thought, being able to thrive from 2°C to 25°C in our experiments, but not thriving at 30°C.

While one Desulfuromusa organism (ASV 36, Fig. 3b) was stimulated from 2°C until 20°C mainly in the iron amended treatment, the other three stimulated Desulfuromusa organisms constituted the majority (20–55%) of the active communities from 20°C to 30°C (Fig. 3b) but could not compete at psychrophilic conditions. Three Desulfuromonas organisms were identified (Fig. 3e). Particularly ASV 6 was the most metabolically flexible organism identified from all incubations throughout the experiment. This organism was detected with 2% relative abundance at 2°C and remained detectable up to 25°C (13%). The other two Desulfuromonas organisms, ASV 26 and ASV 3, were only stimulated by psychrophilic conditions reaching up to 13–22% relative abundance. All identified Desulfuromonas organisms were better stimulated in the absence of fresh iron oxide supply. One organism (ASV 56) of unclassified Geopsychrobacteraceae (Fig. 3c) was detected at 5°C and was stimulated more abundantly in the absence of fresh iron supply, until 25°C. One Trichloromonas organism (Fig. 3f) was detected in our incubations, its growth stimulated from 2°C to 20°C, mainly by sulfate addition and in the control treatments.

By maintaining metabolic flexibility and surviving multiple temperature shifts, these microbes demonstrated that, despite the effect of warming on sediment communities, certain keystone species will adapt over wide temperature ranges to sustain relevant ecosystem functions supported by prevailing environmental conditions. In the case of our site, this function is the release of CO₂ driven by iron reduction as the final sink of OM degradation. The fresh supply of iron oxides from the glacial meltwater supports the activity of iron reducers, ensuring that iron reduction remains a relevant metabolism in the sediments. Therefore, iron reducers, mainly Sva1033, act as keystone species in this specific environment, adapting to increased temperature and sustaining OM degradation. However, in the event of a limited iron supply in the distant future scenario, threatening the activity of Sva1033 in the environment, other iron-reducing microorganisms will replace such ecosystem function acting as keystone species.

Implication for the fate of organic matter on an increasingly warming planet

The Antarctic Peninsula experienced an extremely warm event and record-high surface melt in February 2022 ¹⁷. On February 7-8th, 2022, extreme record-high near-surface temperatures (13.6°C to 13.7°C) were recorded on King George Island/Isla 25 de Mayo, including the Potter Cove¹⁷ where sediment near the Fourcade Glacier was sampled for our study three years earlier. Similar events occurred in Western Antarctica in February 2020, featuring an unprecedented regional temperature anomaly of + 4.5°C during a six-day period⁴³. In the face of global warming and glacier retreat, warming events will become frequent, causing increased discharge of glacial meltwater into underlying sediments. The discharge of nutrients-containing glacial meltwater from the Fourcade Glacier ice is expected to alter the physical and chemical properties of marine sediments with impact on biological communities⁴⁴, and ultimately the fate and rate of degradation of organic matter. One of such impacts is the significant change in the structure and function of microbial communities as comprehensively demonstrated by our study. We observed that just 10 days of significant warming of 3°C and above is sufficient to impact the microbial community structure and ecosystem function of active populations in marine sediments from the Antarctic Peninsula (Fig. 2; suppl. Table S1, Fig. S4). The detection of certain microorganisms such as Colwellia, Desulfuromusa, Arcobacteraceae and Hoeflea at specific temperature windows is a key finding from our study. It validates the age-old Baas-Becking hypothesis⁴⁵ of spatial distribution that “everything is everywhere, but the environment selects”. Becking alluded to the remarkable spatial distribution potential of microbes, but that only specifically adapted organisms will thrive and proliferate under specific environmental guides^46,47. Climate change-driven warming in Antarctica will activate hitherto inactive or low-activity microbial populations as a response to changes in temperature or nutrient flux, helping microbial communities sustain ecosystem functions. For example, Desulfuromusa stepped up at 30°C to ensure iron reduction ecosystem function continues in these sediments when other more successful relatives at lower temperatures could not thrive.

Our study contrasts previous observations in the water column of the Arctic Ocean where ecosystem function under warming scenarios is being performed by new species, introduced to the environment by dispersion from temperate environments^48,49. The introduction of new better-adapted organisms to sustain ecosystem function in the Arctic Ocean is facilitated by the influence of the Atlantic waters leading to increased warming and saltiness, otherwise called Atlantification^48,50. Our contrasting observations are in line with the geomorphology of Antarctica, given that Antarctic waters are isolated from the rest of the global oceans by the Antarctic Circumpolar Current (ACC)⁵¹. Besides, introduction of temperate organisms by dispersion is more likely to occur in pelagic settings than in sediments. Consequently, indigenous distinct microbial populations, instead of newly introduced microbes as reported for the Arctic, respond to warming to conserve ecosystem function in Antarctica as our study reveals. As demonstrated in Fig. 3, conservation of ecosystem function is achieved either by keystone species with incredible metabolic flexibility to adapt to changes in temperature and nutrient conditions (such as Sva1033 – ASVs 1 and 10 or Desulfuromonas – ASVs 6 and 35) or by previously inactive species better suited to the changing environmental guides (such as Desulfuromusa – ASVs 5, 18, and 19).

These findings have implication for the biogeochemical cycle of elements and for the ocean’s fluxes and control of CO₂ release to the atmosphere. The types of microbes our study targeted utilize iron and sulfate as electron acceptors to support their ecosystem function of mineralizing a significant portion of the organic matter in marine sediments. A global warming-induced change in microbial community composition (Fig. 2), which could often be accompanied by the loss of certain specialist microbes or the stimulation of microbes better adapted to the environmental change (Fig. 3), will also affect the rate of CO₂ release from the sediments as simulated by the CO₂ release trajectory in our study (suppl. Fig. S1). Although the temperature ranges that we tested are largely experimental, we argue that the findings are relevant for predicting current and future impacts of climate change on organic matter degradation and associated microbial communities in both permanently cold and temperate environments.

Sediment sampling

Sediment samples were collected from Potter Cove close to the Fourcade Glacier during the Antarctic summer in 2019. The sediments were collected from the safest spatial distance to the Fourcade Glacier where it was still possible to sample without risk. The geochemical setting of Potter Cove and the site sampled (STA 13; S62°13'31.4''/W58°38'28.2'') were previously described⁹. Because of the shallow water depth at the site (15 m), SCUBA divers from the Argentinian Diving Division could retrieve short sediment cores. The cores were immediately processed in the Dallmann laboratory (AWI- DNA/IAA cooperation) of the Argentine Carlini Station. In detail, the 29 cm core (designated as STA13-04) used for this study was sliced into 5 cm sections and stored at 2°C in 500 mL hermetically closed Schott bottles, under N₂ headspace (99.999% purity, Linde, Germany). After three months in transit back to the laboratory in Germany under constant storage in the dark at near in situ conditions, the entire core was pooled to have sufficient material for incubation experiments.

Incubation experiments

To study temperature responses of respiring organisms in glacial meltwater-influenced sediments from Potter Cove, short-term incubation experiments were set up. Anoxic sediment slurries were prepared using 5 g sediment to a final volume of 30 mL (1:6 w/v) in 60 mL serum bottle with anoxic, sterile sulfate-free artificial sea water (ASW; composition [L^− 1]: 26.4 g NaCl, 11.2 g MgCl₂ ∙ 6 H₂O, 1.5 g CaCl₂ ∙ 2 H₂O and 0.7 g KCl). The ratio 1:6 w/v was used because several bottles (n = 126) were required for the study; however, the sediment available was insufficient for making slurries with larger sediment input. Slurries were made anoxic under N₂ headspace. Thereafter, 18 slurry bottles per temperature were pre-incubated at the various experimental temperatures (2°C, 5°C, 10°C, 15°C, 20°C, 25°C, 30°C) for 7 days to acclimatize the sediment communities to the incubation temperatures before substrate addition. Afterward, three treatment types were prepared in biological triplicates per temperature: I. acetate (500 µM) II. acetate and lepidocrocite (5 mM) and III. acetate and sulfate (5 mM) with either natural abundance acetate (¹²C) or ¹³C-labelled acetate. After substrate addition, all treatments were subsequently sampled for dissolved iron (Fe²⁺) and sulfate measurement by collecting 1 mL slurry under anoxic conditions. Thereafter, all treatments were subjected to static incubation in the dark at the respective temperatures. The incubations were run between 10 and 17 days depending on the evolution of ¹³C-CO₂ in the headspace of incubations at the respective temperatures (see analytical methods section).

Analytical methods

As a proxy for iron reduction in these short-term incubations, Fe²⁺ was measured at day 0 and the end time point of the incubations at the various temperatures, i.e., day 10 at both 2°C and 5° C, day 11 at both 10°C and 15°C, day 15 at 30°C and day 17 at both 20°C and 25°C. To arrive at a probable comparative end time point for the incubations at the various temperatures, the evolution of ¹³C-CO₂ in the headspace of incubations was measured over time (suppl. Fig. S1). Incubations were stopped when δ¹³C-CO₂ values, expressed in per mille (‰) relative to the Vienna Peedee belemnite (VPDB), were similar at the end time points across the different temperatures. The evolution of δ¹³C-CO₂ in the headspace was measured by injecting either 100 µL or 500 µL of the gas sample into a Thermo Finnigan Trace GC ultra connected to a Finnigan MAT DELTA Plus IRMS via a Thermo Finnigan GC Combustion III interface using a chromatographic and temperature set-up as previously described⁵².

Dissolved ferrous iron accumulation rates were calculated from the slope of the difference in Fe²⁺ concentrations between day 0 and the end point for each temperature. To determine Fe²⁺ concentrations, 1 ml slurry was collected into 1.5 mL reaction tubes under anoxic conditions from the respective time points using N₂ pre-flushed 1 mL syringes. Reaction tubes were centrifuged (15,300 x g, 5 min at 4°C) to obtain supernatants. 100 µL of supernatant were used for dissolved Fe²⁺ measurements as previously described⁵³ with modifications⁵⁴. Dissolved Fe²⁺ measurement was used as a proxy to calculate iron reduction rates because of the difficulty of detecting total Fe(II) produced in the sediment incubations due to the reaction complex of most of the produced Fe(II) with sediment carbonate system within the sediment matrix⁵⁴. To serve as a proxy for sulfate reduction, sulfate concentrations between the start and end point of the incubations were measured as described elsewhere³. The slope of the sulfate concentrations was similarly used as proxy for sulfate reduction rates. Evidence for sulfate reduction via sulfide production was not obtained from all treatments, as characteristic sulfide smell was not detected during the incubations. This could be due to the abiotic reaction of produced sulfide with Fe(II) in the slurries.

RNA stable isotope probing

It is highly recommended for successful RNA-SIP density centrifugations to use between 0.5–1 µg RNA. This amount of RNA is difficult to obtain from marine sediment due to low biomass. Therefore, to obtain sufficient RNA input for SIP, sediment slurry was pooled from triplicates of the same treatments before nucleic acid extraction. Nucleic acids were extracted following an established protocol⁵⁵ with modifications⁵⁶. To obtain DNA-free RNA from nucleic acids, DNA was removed using the RQ1 DNase kit (Promega, Wisconsin, USA). RNA was subjected to a 2% gel electrophoresis to ensure DNA bands were absent followed by a fluorometric quantification of RNA using the Quant-iT RiboGreen kit (Invitrogen, Thermo Fisher Scientific). Given the different RNA concentrations obtained from RNA extraction, isopycnic density centrifugation was performed on an 8-sample rotor using samples with similar RNA concentrations per run. Isopycnic density centrifugation and gradient fractionation were done following a previously described protocol⁵⁵ with modifications⁵⁶. After gradient fractionation, RNA gradients were quantified using the Quant-iT RiboGreen kit and immediately stored at -80°C until further processing. Subsequently, 10 of the 14 obtained fractions per sample (suppl. Fig. S2) were pooled in the following format: fraction 3 and 4 (Ultra-heavy, density: 1.806–1.832 g/ml), fraction 5 and 6 (Heavy, density: 1.797–1.820 g/ml), fraction 7 and 8 (Midpoint, density: 1.786–1.809 g/ml), fraction 9 and 10 (Light, density: 1.774–1.797 g/ml), fraction 11 and 12 (Ultra-light, density: 1.766–1.786 g/ml). To evaluate the enriched microorganisms in the different pooled fractions, reverse transcription of RNA to cDNA was performed using the GoScript reverse transcriptase kit (Promega). The resulting cDNA, derived from 5 pooled fractions per sample, with 6 samples representing the 6 treatments from each temperature, totaling in 210 cDNA samples, served as template for amplicon sequencing. PCR was performed using KAPA HiFi HotStart PCR kit (KAPA Biosystems) with barcoded bacterial 16S rRNA primers (Bac515F (5′-GTGYCAGCMGCCGCGGTAA-3′)⁵⁷; Bac805R (5′-GACTACHVGGGTATCTAATCC-3′)⁵⁸. DNA amplification, PCR product purification and sequencing library preparation were performed as described previously⁵⁴. Amplicon sequencing was performed by Nogovene (Cambridge, UK) using a Novaseq6000 platform (2x 250 bp). Raw reads were processed with the DADA2 sequence analysis pipeline⁵⁹ as described previously^42,60. Resulting amplicon sequence variants (ASVs) were taxonomically assigned using the SILVA database (SSU Ref NR 99 release 138.1³⁴) and sequences assigned outside Bacteria or as mitochondria or chloroplasts were removed. Sufficient sequencing depth per sample was checked by rarefaction curves as described previously⁴² (suppl. Fig. S17-S19). For identifying the ASVs that were clearly labeled by ¹³C-acetate, the following calculation was performed: the calculated average of the relative abundance per ASV in heavy and ultra-heavy fractions of ¹³C-labeled treatments had to be higher than the calculated average in light and ultra-light fractions of ¹³C-labeled treatments.

The transcripts of sulfate reduction marker gene subunit dsrA, encoding dissimilatory sulfite reductase, were quantified by quantitative PCR (qPCR) (primer DSR1-F+ (5′ACSCACTGGAAGCACGGCGG-3′), DSR-R (5′-GTGGMRCCG TGCAKRTTGG-3′)⁶¹) in cDNA samples also used for amplicon sequencing as described previously³. As template, 2 µL cDNA were used and after quantification of cDNA concentration using the Quanti-it PicoGreen kit (Invitrogen, Thermo Fisher Scientific), quantified copies per ng cDNA were calculated. For samples below detection limit an additional qPCR targeting the bacterial 16S rRNA was performed (primer Bac8Fmod (5′-AGAGTTTGATYMTGGCTCAG-3′), Bac338Rmod (5′-GCWGCCWCCCGTAGGWGT3′)³) using a similar procedure, as described previously³.

Statistical analysis

A distance-based redundancy analysis (dbRDA) was performed on a Bray-Curtis dissimilarity distance matrix of the bacterial 16S rRNA gene relative abundance data using treatment and incubation temperature as explanatory variables. The significance (p < 0.05) of the model was tested with an ANOVA like permutation test⁶² (anova.cca function in vegan R package⁶³, version 2.6.6.1). All analyses and plots were made within the R environment⁶⁴ version 4.4.1.

Data availability

The raw sequence data were submitted to European Nucleotide Archive (ENA) under accession number PRJEB82428. All codes used for data analysis were submitted to the Github repository https://github.com/Microbial-Ecophysiology/tempSIP-PotterCove.

Competing Interests

The authors declare no competing interests.

Funding

This work was supported by the Deutsche Forschungsgemeinschaft (DFG) in the framework of the priority program “Antarctic Research with Comparative Investigation in Arctic Ice Areas” SPP 1158 in the project “Environmental Controls of Iron-Reducing Microorganisms in Antarctic marine sediments (ECIMAS)” (project number 404648014) and the University of Bremen. Graciana Willis-Poratti was funded by individual fellowships supported by the Deutscher Akademischer Austauschdienst [German Academic Exchange Service (DAAD)]: grant numbers 57440915, 57507442 and 57681226.

Author Contributions

DAA, MWF and GWP designed the study. DAA and GWP conducted the field sampling. DAA, GWP, LCW and CO performed the lab experiments. DAA, LCW, XY, GWP, and TRH analyzed the data. LCW and CN produced the figures. GWP, WMC, SV, SH and MWF facilitated the field trip to the WAP. MWF secured funding for the research. DAA led the study and wrote the manuscript with LCW. All co-authors contributed to the manuscript.

Acknowledgments

The authors thank the Instituto Antártico Argentino (IAA) – Dirección Nacional del Antártico (DNA), the crew at Carlini Station, Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET-Res. N◦ 4252/116), and the Alfred Wegener Institute Helmholtz Centre for Polar and Marine Research (AWI) for logistics support during the field campaign to the King George Island/Isla 25 de Mayo in the Antarctic Peninsula. The authors acknowledge Principal Corporal Javier Alvarez from the crew at Carlini Station and Argentinian Navy (Armada Argentina) and the Argentinian Army (Ejército Argentino) divers of CAV 2018–2019 for their support during the sampling at Potter Cove.

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Global warming facilitated environmental change effects on CO₂ releasing microbes in Antarctic sediments

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