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Table of Contents
Abstract
Introduction
Results And Discussion
Isotopic Signatures Indicate The Biological Origin Of Nitrate
Unique Microbial Nitrogen Cycle In Coastal Antarctica
Highly Diversity Of Microbiomes And Their Roles In Microbial N-Cycling Processes
Abundant And Novel Nitrification Drivers In Coastal Antarctica
Metabolic Potential And Surviving Strategy Of Clade B Comammox Nitrospira In Antarctica
Nitrification And N 2 O Production Activities Driven By Comammox Nitrospira
Implications And Outlook
Methods
Sample Collection And Treatment
Physiochemical Analysis
Isotopic Analysis Of Nitrate
DNA Extraction, PCR, QPCR, Sequencing And Phylogenetic Analysis
Metagenomic Sequencing
Assembly And Binning Of Metagenomes
Phylogenomic Analysis And Genome Annotation
DNA-Stable Isotope Probing (SIP) Microcosm Incubations, Gradient Fractionation, Quantification, And Sequencing
Data Availability
Supporting Information
Author Contributions
Competing Interests
Competing Interest Statement
Competing Interest Statement
Abstract
Although microbial nitrogen (N) cycling plays a pivotal role in Antarctic ecosystems, its underlying mechanisms are largely uncharted. In this study, we unravel the biological origin of nitrate via triple oxygen isotopic composition analysis and systematically profile functional N-cycling genes within soil and lake sediment samples from the ice-free areas of East Antarctica. We successfully reconstruct 1,968 metagenome-assembled genomes (MAGs) spanning 29 microbial phyla, enabling the analysis of the presence or absence of 52 diverse metabolic marker genes. Consistent with quantitative data, our metagenomic analyses confirm the active processes of microbial nitrogen fixation, nitrification, and denitrification. We find no detectable anaerobic ammonium oxidation (anammox) processes, underscoring a unique microbial N-cycling dynamic in the region. Notably, we identify the predominance of complete ammonia-oxidizing (comammox) Nitrospira , a recently discovered bacterial guild capable of performing the entire nitrification process within a single organism. Further genomic investigations reveal their adaptive strategies in the Antarctic environment. These strategies likely involve the synthesis of trehalose to counteract cold stress, high substrate affinity to efficiently utilize available resources, and alternative metabolic pathways to adapt to nutrient-scarce conditions. Their significant role in the nitrification process is validated through 13 C-DNA-based stable isotope probing (DNA-SIP). This research provides a comprehensive illustration of nitrification’s crucial contribution to the nitrogen budget in coastal Antarctica, highlighting comammox Nitrospira clade B as a novel nitrifying agent and shedding new light on the complex biogeochemical processes of nitrogen cycling in coastal Antarctica.
Introduction
Antarctica, a region largely insulated from anthropogenic nitrogen (N) deposition, exhibits notably low N concentrations 1 . These diminished N levels play an integral role in sustaining ecological equilibrium in the continent’s barren, ice-free terrains 2 , 3 . Consequently, elucidating the intricacies of microbial nitrogen cycling pathways in Antarctic ecosystems—including diazotrophy (N-fixation), nitrification, anaerobic ammonium oxidation (anammox) and denitrification—is paramount 4 . Diazotrophic Cyanobacteria are the primary agents of biologically accessible ammonium (NH 4 + ) synthesis in these environments 5 . Alongside NH 4 + , nitrate (NO 3 − ) constitutes a significant inorganic nitrogen pool in polar biomes 6 , 7 . The genesis of NO − in Antarctic ecosystems can be attributed to both abiotic atmospheric deposition, particularly within mineral-rich Antarctic soils 8 , and to biotic nitrification processes. Still, the discrete contributions of these sources to the coastal Antarctic NO 3 − reservoir remain inadequately characterized, necessitating further investigation. Nitrification plays a pivotal role in the global biogeochemical nitrogen cycle and contributes significantly to the emissions of nitrous oxide (N 2 O), a potent greenhouse gas. This process encompasses the oxidation of ammonia (NH 3 ) to nitrate (NO 3 − ) via the intermediate nitrite (NO 2 − ). Traditionally, chemolithoautotrophic nitrification has been attributed to two distinct microbial guilds: ammonia-oxidizing bacteria (AOB) and archaea (AOA), responsible for converting NH 3 to NO 2 − , and nitrite-oxidizing bacteria (NOB), which facilitate the subsequent oxidation of NO 2 − to NO 3 − . Beyond these canonical nitrifiers, the recently identified complete ammonia oxidizers (comammox) within the genus Nitrospira , specifically lineage II, perform full nitrification as a singular 9 , 10 . These organisms are increasingly recognized for their significant role in nitrification across diverse ecosystems, including both engineered systems 11 and natural environments 12 . The contribution of comammox bacteria to nitrification and subsequent NO 3 − production in Antarctic ecosystems, however, remains an enigma. Phylogenetically, comammox Nitrospira bifurcate into clades A and B 9 , distinguished by their respective ammonia monooxygenase genes ( amoA ). Physiological assessments of clade A comammox Nitrospira have revealed an exceptionally high ammonia affinity, suggestive of an oligotrophic niche adaptation 13 , 14 . In stark contrast, clade B comammox Nitrospira , despite their widespread distribution in various habitats, including those characterized by low temperatures such as the Tibetan Plateau 15 , 16 and Arctic permafrost 17 , have yet to be cultured for physiological scrutiny. This study aims to unravel the complexities of the microbial nitrogen cycle in the ice-free regions of East Antarctica, identifying the primary sources of NO 3 − and examining the potential roles of novel microbes in the nitrification process. Our research is strategically centered on the Larsemann Hills (LH) 18 , a vast ice-free rocky landscape in East Antarctica, dotted with over a hundred oligotrophic lakes shrouded in ice. Through a comprehensive approach, we discover that (i) the origin of NO 3 − is primarily the biological nitrification process; (ii) the microbial nitrogen cycle is distinct, encompassing most microbial N-cycling processes except for the anammox pathway; and (iii) the comammox Nitrospira clade B serves as a novel, abundant, and active driver of nitrification, possessing unique survival strategies against the cold and oligotrophic conditions of the coastal Antarctic environment.
Results and Discussion
Isotopic signatures indicate the biological origin of nitrate
Depending on how NO 3 − is produced, the composition of its oxygen isotopes differs, allowing differentiation between abiotic and biotic sources 19 . For biologically produced NO 3 − , one oxygen atom (O) is expected from atmospheric oxygen (O 2 ) and two from the surrounding water (H 2 O) 19 . Biologically produced NO 3 − would have a δ 18 O of ∼0.6‰, since δ 18 O of atmospheric O 2 is 23.9‰ 20 and the measured δ 18 O of LH lake water is –12.7±1.5‰ (Supplementary Table 1). Atmospheric NO 3 − in Antarctica is mainly produced by the reaction of nitrogen oxides (NO x ), ozone (O 3 ), and hydroxyl radicals (OH·) 21 . In addition, while biologically produced NO 3 − has a Δ 17 O of 0‰, which is identical to that of O 2 and H 2 O, atmospheric NO 3 − is usually characterized by high oxygen isotopic ratios 21 , 22 and can reach Δ 17 O ≥35‰ in the atmosphere in Antarctica 22 . The mean δ 18 O of NO 3 − in lake sediments from LH is 4.6‰. The annual mean δ 18 O of NO 3 − in snow and the atmosphere at the coastal Zhongshan station, situated in LH, is approximately 73‰ 21 . From these, the contributions of atmospheric deposition and biological production to sedimentary NO 3 − pools can be quantified by isotope mass balancing, indicating that the nitrification process constitutes 93% of the NO 3 − in sediments. In addition, limited data of sediment NO 3 − Δ 17 O produced a mean of approximately 1.3‰, which strengthens the observed >90% contribution of nitrification considering the very high Δ 17 O of atmospheric NO 3 − at Zhongshan station (approximately 35‰). Similarly, the isotope mass balance of δ 18 O and Δ 17 O suggested ∼96% of soil NO 3 − in the two study areas is from nitrification. Thus, we demonstrated here that sediment and soil NO 3 − predominantly originated from biological production in and nearby the investigated lakes.
Unique microbial nitrogen cycle in coastal Antarctica
The primary production of NO 3 − is largely attributed to the microbial nitrification process, rather than atmospheric precipitation. Consequently, the origin of NH 4 + , the substrate for nitrification, can be traced back to biological N-fixation. Based on the qPCR-based quantification of functional N-cycling genes ( Fig. 1 ), we noticed a comparable amount of N-fixation ( nifH, encoding nitrogenase) to nitrification genes ( amoA and nxrB , encoding ammonia monooxygenase and nitrite oxidoreductase, respectively), with quantities ranging from 10 2- 10 4 copies ng − 1 DNA ( Fig. 1 ). The relatively low quantity of the gene encoding hydroxylamine dehydrogenase ( hao ) can be attributed to the low coverage of the applied primers and the absence of the bacterial hao gene in the genomes of AOA 23 , which are of notable abundance in the studied samples. In addition to N-fixation and nitrification processes, we also observed significant activity of denitrifier, as indicated by the high abundances of genes encoding the sequential reduction of NO 3 − ( nar, nap ), NO 2 − ( nir ), nitric oxide (NO, nor ), and N 2 O ( nos ) ( Fig. 1 ). The relatively higher abundances of nirS and nirK genes can likely be attributed to their presence not only in denitrifiers but also in nitrifiers. Interestingly, we found substantial quantities of the functional gene for dissimilatory nitrate reduction to ammonium (DNRA, nrfA ) and assimilatory nitrite reduction (ANR, nasA ) ( Fig. 1 ). The potential for DNRA and ANR is widespread among phylogenetically diverse microorganisms and these processes ensure the retention of inorganic nitrogen in an ecosystem. Strikingly, we detected minuscule amount of the hzo gene (encoding hydrazine oxidoreductase), a biomarker for the anammox process, in all tested sediment and soil samples. Anammox bacteria catalyze the anaerobic oxidation of NH 4 + using NO 2 − as electron acceptor, producing N 2 as final product. These bacteria were first identified in wastewater treatment systems 24 and were subsequently discovered in various environments, including marine 25 , coastal 26 , terrestrial 27 and engineering systems 28 . The absence of anammox functional markers in coastal Antarctica suggests unique microbial nitrogen cycling properties in this remote region.
Highly diversity of microbiomes and their roles in microbial N-cycling processes
From an extensive dataset exceeding 280 gigabases of sequencing data, we managed to reconstruct a dereplicated collection of 724 high-quality and 1244 medium-quality 29 metagenome-assembled genomes (MAGs) (Supplementary Data 1). The recovered genomes encompass 29 distinct phyla (Supplementary Data 2), marking, to the best of our knowledge, the most comprehensive inventory of Antarctica coastal soil and sediment genomes to date. On average, we obtained 50 to 60 MAGs per sample site studied ( Fig. 2a ). The most abundant phyla included Actinobacteriota , Pseudomonadota , Bacteroidota , Chloroflexota , Verrucomicrobiota , Acidobacteriota , Patescibacteria , Planctomycetota , and Gemmatimonadota ( Fig. 2b , Supplementary Data 1), aligning with findings from other Antarctic surveys 5 , 30 , 31 . Cyanobacteriota also featured among the top ten most abundant MAGs (Supplementary Data 1), a deviation from the Mackay Glacier Region, where they were largely absent in most soil samples 31 . We identified only five archaeal MAGs within the Thermoproteota phylum, all of which were further classified as ammonia-oxidizing archaea (AOA) within Group I.1b. To comprehend the metabolic strategies that sustain the abundant bacterial life in these extremely nutrient-poor sediments and soils, we examined the distribution and affiliation of 52 marker genes conserved across different energy conservation and carbon acquisition pathways in the MAGs we retrieved. As predicted, genes for aerobic organotrophic respiration were encoded by nearly all community members ( Fig. 2c , Supplementary Fig.1). Consistent with observations from the Mackay Glacier Region 31 , a significant number of the MAGs appear to fix carbon via the Calvin-Benson-Bassham (CBB) cycle or the 3-hydroxypropionate cycle ( Fig. 2c ). These processes provide a pathway for biomass generation that is independent of photoautotrophy, a function primarily performed by Cyanobacteriota (Supplementary Fig.1). Genomic analysis revealed that the most abundant and widespread community members encoded trace gas oxidation genes. In a pattern similar to previous discoveries 31 , 32 , carbon monoxide (CO) dehydrogenases (CoxL) were exclusive to Actinobacteriota and Chloroflexota ( Fig. 2c , Supplementary Data 3). Interestingly, uptake hydrogenases were encoded by MAGs from Acidobacteriota , Chloroflexota , and Verrucomicrobiota ( Fig. 2c , Supplementary Data 3), which aligns with observations from temperate soil where Acidobacteriota are known to be active atmospheric H 2 consumers 33 , albeit with slight differences. In the realm of N-metabolism, a comprehensive array of functional genes was identified within the MAGs we analyzed ( Fig. 2c ). Nitrogen fixation appears to be predominantly carried out by taxa within Acidobacteriota , Cyanobacteriota , Desulfobacterota, Myxococcota, and Verrucomicrobiota ( Fig. 2c ). Consistent with the quantitative analyses, the hzs gene, indicative of the anammox process, was not present in the retrieved MAGs. While a total of 96 Planctomycetota MAGs were obtained, subsequent verification through taxonomic classification to check for affiliation with the Brocadiaceae confirmed that none of these MAGs belong to anammox bacteria (Supplementary Data 2). The absence of anammox process may be attributed to the discrepancy between the optimal temperature range of 12–17°C, which is characteristic of anammox process found in similarly cold Arctic fjord sediments 34 , suggesting a limited tolerance to low temperatures. This assertion is further reinforced by the absence of anammox bacteria in the McMurdo Dry Valleys (an Antarctic desert) 35 and in Arctic regions 36 as well. The pervasive distribution of denitrification genes across nearly all phyla underscores the robust and widespread microbial denitrification activity in Antarctica, corroborating previous research that highlights the ubiquity of cold-adapted denitrifiers across diverse Antarctic ecosystems 37 . Regarding the nitrification process, the marker genes- amoA (associated with ammonia oxidation) and nxrB (linked to nitrite oxidation) were found exclusively in the Thermoproteota and Nitrospirota phyla ( Fig. 2c ). Although Antarctic soil nitrification was previously reported back in 1997 38 , attributed to AOB genera Nitrosospira and Nitrosomonas , as well as AOA within group I.1b 39 – 42 , no investigation has yet been conducted into the presence and function of the recently discovered comammox bacteria within the Nitrospira genus 9 , 10 . Here, we present data showing the prevalence of the comammox Nitrospira amoA gene in Antarctica lake sediments and soils, strongly suggesting the activity of comammox Nitrospira in this region.
Abundant and novel nitrification drivers in coastal Antarctica
The quantitative evaluation of nitrification-related functional genes, including amoA and nxrB (Supplementary Fig. 2), established that AOA and comammox bacteria were the predominant nitrifiers in the analyzed soils and sediments (Supplementary Fig. 3). Our metagenomic analysis corroborates these findings, having identified only AOA and Nitrospira MAGs among the various nitrifying groups ( Fig. 2c ), suggesting their relatively high abundances (Supplementary Table 3). Furthermore, a more detailed investigation into the abundance and community structure of all identified nitrifying organisms using amplicon sequencing and subsequent phylogenetic assessment indicated an unexpected dominance of clade B comammox Nitrospira in this environment (Supplementary Results and Discussion). Among the de-replicated MAGs, five were classified as Group I.1b-AOA within the genera Nitrosocosmicus and Nitrospharea of Thermoproteota phylum (Supplementary Data 2). This categorization stemmed from phylogenomic assessments and average nucleotide identity (ANI) analyses (Supplementary Fig. 4). These findings are in agreement with the data obtained from amplicon sequencing, as discussed in the Supplementary Results and Discussion section.). Together with their previous detection in Arctic soils 43 , 44 , this suggests a capacity of Nitrosocosmicus -like AOA to thrive in cold and nutrient-deficient conditions. Of the seven Nitrospirota MAGs analyzed, four were verified as comammox Nitrospira , and the remaining three belonged to the exclusively nitrite-oxidizing lineages II and IV of Nitrospira . This classification was substantiated by phylogenomic and ANI analyses ( Fig. 3a and 3b , Supplementary Fig. 5) and these results concur with the phylogenetic assessment based on the amoA and nxrB gene of Nitrospira (Supplementary Fig. 8 & Fig. 9). The notably high abundance of the AOA and these Nitrospira MAGs in the soil and sediment metagenomes studied (reaching up to 0.173%, Supplementary Table 3) further emphasizes their significant roles in nitrification within these ecosystems. Two high-quality comammox MAGs, Nitrospira sp. La1-X1 (hereafter referred to as “La1”) and Nitrospira sp. La3-X1 (hereafter referred to as “La3”), had genome sizes of 4.26 Mb and 3.78 Mb, respectively (Supplementary Table 3). The amoA gene sequences from La1 and La3 align closely with the dominant amoA OTUs and with previously published sequences from clade B comammox ( Fig. 4c and Supplementary Fig. 8). Consistent with the amoA -based phylogenetic findings, phylogenomic analysis also demonstrates a distinct clustering of these comammox MAGs within clade B ( Fig. 3a ). For La1, the ANI was highest with MAG Nitrospira sp. palsa1310, which was identified from Arctic permafrost soil 17 . The ANI between these genomes is 96.67% ( Fig. 3b ), surpassing the species threshold of 95% 45 , suggesting that these genomes represent different strains of the same Nitrospira species, potentially adapted to cold environments. In contrast, La3 is grouped within a clade B subset that includes MAGs from drinking water treatment systems and glacier surface soil 46 , but it did not share high ANI values with any other clade B genomes (≤85%, Fig. 3b ). This suggests that La3-bin1 represents a unique so far undetected lineage within clade B.
Metabolic potential and surviving strategy of clade B comammox Nitrospira in Antarctica
The clade B comammox MAGs Nitrospira sp. La1-X1 and Nitrospira La3-X1 harbor the complete genetic machinery for NH 3 and NO 2 − oxidation, the respiratory chain, and the reduced tricarboxylic acid (rTCA) cycle, which is the conserved CO 2 fixation pathway in Nitrospira ( Fig. 3c and 3d ). These core metabolic features are highly conserved in comammox Nitrospira as reported in previous studies 47 , 48 . Similar to other Nitrospira genomes 46 , La1 and La3 do not encode nitric oxide reductase (NOR), which is crucial for enzymatic N 2 O production 49 . They do carry genes for urea transport and hydrolysis by the urease, indicating the use of urea as an alternative ammonia source, as shown for other comammox Nitrospira 9 , 10 , 50 . In addition, like some Nitrospira that have the confirmed ability to use hydrogen and formate as alternative energy sources 51 , 52 , the comammox MAGs La1 and La3 contain genes for formate and hydrogen oxidation. Although the formate dehydrogenases of comammox are similar to their nitrite-oxidizing counterparts, comammox bacteria possess a 3b-type [NiFe] hydrogenase. This hydrogenase is rarely identified in canonical Nitrospira and its physiological role remains unclear 48 , 53 . While the capacity for formate oxidation is widely distributed in clade B, the 3b-type [NiFe] hydrogenase has been mainly identified in clade A comammox species 48 ( Fig. 3d ). However, the presence of this hydrogenase type in a few clade B genomes 46 , 48 including La3 challenges the clade specificity of this feature. In addition to the canonical F 1 F 0 H + -driven ATPase, La1 and La3 encode a potentially Na + -pumping F 1 F 0 ATPase, previously detected in the haloalkalitolerant nitrite-oxidizing Ca. Nitrospira alkalitolerans 54 and clade A comammox Ca. N. kreftii 14 . Similar to these genomes, La3 also possesses a Na + -translocating NADH:ubiquinone oxidoreductase (NQR), a feature missing in La1( Fig. 3c ). These Na + -pumping enzyme complexes might represent an adaption of La3 to saline or haloalkine conditions similar to other Nitrospira . The disaccharide trehalose is one of several solutes known to protect bacteria against cold stress, and may also protect against other harmful environmental conditions, such as osmotic stress 55 . La1 possesses a trehalose-6-phosphate synthase (OtsA) and the corresponding phosphatase (OtsB) to potentially produce the non-reducing disaccharide in two steps from UDP-glucose, as shown for E. coli 56 . In contrast, La3 encodes several other trehalose synthesis enzymes in one gene cluster, indicating that this species might convert NDP-glucose (trehalose synthase) and maltose (trehalose synthase/amylase) to trehalose. However, all these trehalose synthesis pathways have also been identified in Nitrospira from mesophilic environments and might thus not be a distinguishing feature for cold adaptation. In addition to trehalose synthesis, La1 and La3 possess other features for coping with different stresses. Whereas a gene cluster encoding a nitrile hydratase has been detected in La1 and La3, the genomes lack an amidase to degrade the produced amides further to the corresponding carboxylic acid and ammonia, which could serve as an energy source. An example of a potential amidase is a putative formamidase identified in a clade B MAG obtained from the Rifle aquifer 46 . Other detoxification mechanisms include catalases and Fe orMn superoxide dismutases (SOD) for reactive oxygen defence identified in both clade B genomes, and a periplasmic Cu-Zn SOD present in La3 and many other clade B genomes, but absent in La1 ( Fig. 3c and 3d ). An adaptation to oligotrophic conditions (Supplementary Table 1) might be the methionine salvage pathway (MSP) of La1 ( Fig. 3c ). This pathway recycles the sulphur-containing intermediate 5’-methyl-thioadenosine back to methionine, thus allowing the use of reduced sulphur compounds under sulphur limitation. However, similar to the high-quality genome of the nitrite-oxidizing Nitrospira lenta and other clade B genomes, the mtnE gene encoding the last step in this pathway is missing in La1. Whether other enzymes might complete the MSP in these Nitrospira species remains to be determined.
Nitrification and N 2 O production activities driven by comammox Nitrospira
We have demonstrated that comammox Nitrospira potentially serve as crucial drivers of nitrification in coastal East Antarctica, and further investigated their survival strategies from a genomic perspective. Additionally, we confirmed the nitrification activity of comammox Nitrospira using DNA-SIP on two lake sediment samples (LA1 and LA2) and one soil sample (LS4). DNA-SIP incubations were conducted at 4°C (for LA1) and 10°C (for LA1, LA2, and LS4) and the production of NO 3 − served as indicator of nitrification activity ( Fig. 4a , Supplementary Fig. 11). During the incubations, comammox Nitrospira were actively growing, as evidenced by the peak shifts of their DNA the 13 C-treatment ( Fig. 4b , Supplementary Fig. 4). Subsequent sequencing of the amoA genes from the labelled DNA revealed clade B comammox Nitrospira amoA OTUs, including those of comammox MAGs La1-X1 and La3-X1, in both the 4°C and 10°C incubations ( Fig. 4c ). Collectively, these data bolster the evidence for the active role of clade B comammox Nitrospira in nitrification. The addition of NH 4 + did increased the nitrate production, however, did not stimulate higher N 2 O production ( Fig. 4a ). These can be explained by the fact that the major active nitrifiers-comammox Nitrospira- are low N 2 O producers 49 . The application of chlorate, a potential comammox-specific inhibitor 57 , slightly reduced the production of NO 3 − and N 2 O ( Fig. 4a ). However, given that chlorate can also influence other nitrite-oxidizing Nitrospira species as well as denitrifiers, it is not feasible to ascribe the observed reductions in NO 3 − and N 2 O levels solely to the activity of comammox Nitrospira . Nevertheless, prior research has shown the resilience of clade B comammox Nitrospira to freeze-thaw cycles, which underscores the endurance of this particular nitrifier group in environments with low and even subfreezing temperatures 58 . Raising the temperature (10°C) stimulated the activity of AOA (in LA2) and AOB (in LA2 and LS4) (Supplementary Fig.4, Supplementary Results and Discussion), indicating that these nitrifiers are more competitive under elevated temperature conditions, an important finding to predict ecosystem changes in response to global warming.
Implications and outlook
In this study, we elucidated the unique microbial N-cycle in pristine and oligotrophic coastal Antarctic soil and lake sediments, identifying the microbial nitrification process as the primary pathway for NO 3 − production. Our metagenomic and quantitative functional gene-targeted analyses both revealed clade B comammox Nitrospira to be key nitrification drivers in these environments. This finding not only expands our understanding of microbial diversity but also underscores the pivotal role of specific microbial groups in biogeochemical cycling in extreme environments. Our results also revealed fascinating patterns of niche differentiation between clade A and B comammox Nitrospira and canonical nitrifiers. The genomic data strongly suggest that clade B species might have superior affinity for their substrate ammonia, and are potentially best adapted to survive and thrive in cold and oligotrophic environments. This niche differentiation could have significant implications for N-cycling in cold environments and it will be important to determine how the nitrifier and overall microbial communities will react to global warming. In addition, we have successfully obtained in total 1968 MAGs, significantly enriching the existing microbial genomic data for Antarctica. These MAGs provide invaluable insights into the microbial diversity and their metabolic capabilities in the extreme and unique environment of coastal East Antarctica. The availability of these MAGs will undoubtedly facilitate further research into the intricate relationships between microbial communities, biogeochemical cycles, and climate change in polar regions and beyond. This study underscores the importance of understanding the unique microbial N-cycle in coastal East Antarctica. We have uncovered clade B comammox Nitrospira to be a novel nitrification driver in low-temperature environments, and investigated their survival strategy and potential impact on climate change through the production of greenhouse gas N 2 O 49 , 59 . It is crucial to further investigate how comammox Nitrospira evolve and survive in Antarctic ecosystems, meeting one of the six priorities for Antarctic science 60 . Recognizing the climatological significance of N-cycling and biogeochemical processes, future research should continue to monitor these microbes, as they could hold the key to understanding the broader implications of microbial activity on our planet’s climate.
Methods
Sample collection and treatment
Samples were collected in Larsemann Hills (LH), the second-largest ice-free land in East Antarctica with an area of ∼50 km 2 . LH has a cold and dry continental climate, with an annual mean temperature of ∼-10 °C and the temperatures occasionally above 0 °C in summer 61 , leading to this region typically free of snow cover in summer. As a result of seasonal snow cover and the glacier melting, a number of landlocked lakes are developed in this region. Surface sediment (the upper ∼5 cm) and surface water samples were collected from six lakes (LA1-LA6) in LHs in February 2020 ( Fig. 1a ). The surface sediment samples were procured using a stainless-steel spade on the shore of lakes, with typical water depths of ∼150-200 cm. Approximately 1L of surface water near the lake shore was also sampled with clean polyethylene (PE) bottles, a portion of which was used to measure temperature, pH, salinity, and conductivity utilizing a multi-probe water quality meter (YSI Professional Plus series; Supplementary Table 1). The remainder of the water samples was filtered using 0.22-μm polytetrafluoroethene (PTFE) filters for chemical analyses. Additionally, a surface soil sample (the top ∼5 cm) was collected near each lake, ∼100 m from the lake shore, using a clean stainless spatula. At each sampling site, larger gravels were removed firstly, and five soil sub-samples (four corners and the center of a square) were collected at a distance of 5–10 m and then mixed to obtain a representative sample (LS1-LS6). For a comprehensive understanding of N cycling processes in the ice-free areas in East Antarctica, two surface soil samples (SVL1 and SVL2) were also collected in February 2022 at Inexpressible Island, South Victoria Land where the climate is comparable to that of LH, following the same sampling protocols. All of the sediment and soil samples were stored in sealed PE bags, and all samples were transported to the laboratory at temperatures of ∼-20 °C for subsequent analysis.
Physiochemical analysis
In the laboratory, approximately 50 g of the sediment and soil samples were freeze-dried in 50-mL clean centrifuge tubes (ALPHA 1-4/LD, Martin Christ Inc.). After drying, the samples were homogenized using an agate mortar and pestle, and subsequently passed through a 1 mm sieve for further chemical analysis. For determining total organic carbon (TOC) content, approximately 5 g of samples were digested with 10% HCl ( v / v ) to remove carbonate. Then, TOC was measured with an automatic element analyser (Elementar, VARIO EL III), with acetanilide used as the external standard. The detection limit (DL) of the TOC was estimated to be approximately 0.005%. All samples were measured in triplicate, yielding a relative standard deviation (1σ) of <10% for each sample. For chemical ion analysis, approximately 5 g of freeze-dried samples were placed in sterile 50-mL centrifuge tubes and suspended in 25 mL Milli-Q water (18.2 MΩ). The solution was then ultrasonicated for 40 min. The supernatant was first centrifuged for 15 min at 3000g, and then filtered through 0.22-μm PTFE filters for nutrient determination. Nutrient concentrations (NH 4 + , NO 3 − , and PO 4 3− ) in the sediment, soil extracts, and lake water samples were determined using an Aquion RFIC ion chromatograph (IC, Thermo Scientific, USA), equipped with the analytical columns CS12A (2×250 mm), AS11-HC (2×250 mm), methanesulfonic acid (MSA), and potassium hydroxide (KOH) as eluents for cations and anions, respectively. In addition, the concentrations of SiO 3 2− were determined using an automated QuAAtro™ nutrient analyser (Seal Analytical Ltd., UK). During sample analysis, replicate determinations ( n = 5) were performed, and 1σ for all species was <5%.
Isotopic analysis of nitrate
The isotopic composition of NO 3 − was determined using the bacterial denitrifier method at the Environmental Stable Isotope Laboratory of East China Normal University (ECNU-ESIL). Briefly, the denitrifying bacterium Pseudomonas aureofaciens , which lacks the N 2 O reductase enzyme, quantitatively transforms NO 3 − into gaseous N 2 O 62 , 63 . The δ 15 N and δ 18 O of the generated N 2 O were measured in duplicates using isotope-ratio mass spectrometry (IRMS, Thermo Scientific Delta V). The Δ 17 O of NO 3 − was separately analyzed through the thermal decomposition of N 2 O into to N 2 and O 2 64 , followed by measurements at m/z 32, 33, and 34 on the IRMS. The pooled standard deviation (1σ p ) was employed to ascertain the measurement precision of the overall denitrifier method 65 , 66 . The 1σ p of all duplicate samples executed in at least two different batches was 0.6‰ for δ 15 N (n=10), 0.3‰ for δ 18 O (n=10), and 0.8‰ for Δ 17 O (n=8). However, due to the limited amounts of NO 3 − in the samples, only 3 sediment and 5 surface soil samples were analyzed for Δ 17 O of NO 3 − . In addition, lake water stable isotopes (δ 18 O and δ 2 H) were analyzed using laser absorption spectrometry (TIWA-45EP, Los Gatos Research, Inc.). To ensure quality control, replicate analyses (n=5) were performed, yielding relative standard deviations of 0.05‰ and 0.2‰ for δ 18 O and δ 2 H, respectively.
DNA extraction, PCR, qPCR, sequencing and phylogenetic analysis
Total DNA was extracted from 0.5 g sediment/soil samples using the Fast DNA SPIN kit (MP Biomedicals, Santa Ana, CA) according to the manufacturer’s protocols. The final DNA quality and quantity were determined using Quant-iT PicoGreen dsDNA Assay Kit (ThermoFisher Scientific, China). A detailed description of the Quantitative PCR (qPCR) analysis for functional nitrogen cycling genes was provided in the Supplementary Methods section. It also elaborates on the procedures for PCR and high-throughput sequencing, as well as the subsequent phylogenetic analysis of nitrification genes.
Metagenomic sequencing
The total DNA from the original sediment and soil samples was sequenced on the Illumina HiSeq Xten platform using a 150-bp paired-end library at Beijing Novogene Biotech Co., Ltd. (Beijing, China). The NEXTFLEX Rapid DNA-Seq Library Prep Kit 2.0 (Bioo Scientific, Austin, TX, USA) was used for DNA library preparation with an insert size of ∼300 bp according to the manufacturer’s recommendations. DNA was sheared using a Covaris S220 Focused Ultrasonicator to create 150 bp fragments. Subsequently, metagenomic sequencing was performed on a cBot Cluster Generation System according to the manufacturer’s standard protocols. we acquired a data range between 20-40 gigabases sequencing data for each individual sample.
Assembly and binning of metagenomes
Raw reads were processed using fastp v0.19.7 67 for adapter trimming, quality filtering, and per-read quality trimming. The seven quality-controlled metagenomes were individually assembled and co-assembled using MEGAHIT v1.1.3 68 with default parameters (k-mers: 21,29,39,59,79,99,119,141). Each of the fourteen assemblies was initially binned using the binning module (– metabat2 –maxbin2 –concoct; –metabat2 for co-assembly) in the metaWRAP pipeline 69 v1.3.2, and were consolidated using DAS Tool v1.1.2 70 with default parameters. After dereplication check using dRep v3.0.0 71 (-comp 50 -con 10) and completeness and contamination evaluation using CheckM v1.1.3 72 , 1968 MAGs were obtained, and the taxonomy of each MAG was assigned using GTDB-Tk v1.5.0 73 with the Genome Taxonomy Database (Release 06-RS202). In the process of annotating metabolic functions, the genomes that were extracted were examined using DIAMOND 74 against 52 tailor-made protein databases, which consisted of representative metabolic marker genes. To confirm the existence of crucial metabolic genes in the MAGs, phylogenetic trees were generated using the maximum-likelihood method. The dereplicated MAGs had their percentage relative abundances determined by aligning each sample’s clean paired-end reads to the MAGs utilizing CoverM (version 0.6.1, available at https://github.com/wwood/CoverM ) in genome mode, applying the default configurations. Moreover, the trimmed reads were incorporated into CLC Genomics Workbench version 20.0 (CLCBio, Qiagen, Germany), and the de novo assembly algorithm of CLC was employed to search for amoA and nxrB sequences. Contigs that contained either amoA or nxrB genes were selected for phylogenetic analysis, in conjunction with amplicon sequences.
Phylogenomic analysis and genome annotation
All available genomes and MAGs classified by the GTDB-Tk database R202 75 as Nitrospiracea , with estimated genome completeness ≥70% and contamination ≤10%, were downloaded from NCBI. Dereplication was performed using the drep v2.4.2 71 dereplicate workflow with cut-offs for estimated genome completion ≥70% and contamination ≤10%, but otherwise default settings. In addition to these 95 Nitrospiracea genomes, seven Nitrospira lineage IV and two clade B genomes were included in the phylogenetic analysis using the UBCG pipeline for the extraction and concatenated alignment of 91 single-copy core genes 76 . In addition, two Leptospirillum genomes (GCF_000284315.1, GCF_000299235.1) were included as the outgroup. A maximum-likelihood phylogenetic tree was constructed using IQ-TREE v1.6.12 77 with 1000 ultrafast bootstrap replications and the GTR+F+I+G4 model identified by the implemented Modelfinder 78 . ANI analysis of all clade B genomes was performed using the OrthoANI 79 . Gene calling and automatic genome annotation were performed using the MicroScope platform, and annotations of selected features were manually checked and refined. For analyzing the distribution patterns of selected key features, 17 comammox clade B genomes were annotated using prokka v.1.14.6 80 with the “--gcode 11” and “--metagenome” options to obtain all protein sequences for generating a BLAST database. The distribution of selected key proteins within clade B was analyzed by conducting a BLASTp search against this database with default settings, except for an e-value cutoff of 1e-6. Only hits with an identity ≥35% (pident) and a query coverage ≥80% (qcovs) were reported as present. Phylogenomic and ANI analyses were carried out on the retrieved AOA MAGs. The process followed was as previously described, using representative genomes (which include Group I.1a and Group I.1b AOA) and Group I.1b Nitrosocosmicus AOA genomes as references, respectively.
DNA-stable isotope probing (SIP) microcosm incubations, gradient fractionation, quantification, and sequencing
Lake sediments (LA1 and LA2), and the soil sample (LS4) were selected for DNA-SIP microcosm incubation experiments (Supplementary Fig. 10) to mimic a low-temperature oligotrophic environment under in situ flooding conditions. Microcosms were constructed in 120 mL serum bottles containing 20 g sediments at 60% of maximum water-holding capacity and were incubated for 56 days at 10 °C in the dark. For each sample, two different treatments were established in triplicate microcosms. The 13 CO 2 microcosms were amended with 5% ( v/v ) 13 CO 2 (99 atom%; Sigma-Aldrich Co., St. Louis, MO, USA) plus approximately 5 µg 15 N-NH 4 Cl-N g −1 dry weight soil/sediment (d.w.s.), while the 12 CO 2 control treatments received 5% ( v/v ) 12 CO 2 plus approximately 5 µg 14 N-NH 4 Cl-N g −1 d.w.s.. The water content was restored weekly over an 8-week incubation period by opening the bottles. The sediment/soil samples were destructively sampled after 56 days of incubation and transferred immediately to –80°C for subsequent molecular analysis. The remaining ∼2 g of sediments were used for end-point quantification of NH 4 + , NO 2 − , and NO 3 − concentrations. The LA1 sediment sample was additionally chosen for an incubation at 4 °C, adhering to the same procedure previously outlined. In addition to the ammonium substrate, 50 μM chlorate, a specific comammox inhibitor 57 , was introduced to study the activity of comammox Nitrospira . Moreover, the production of nitrous oxide (N 2 O) 57 was actively monitored during these 4°C DNA-SIP incubations. The fractionation of DNA post DNA-SIP incubations, along with the subsequent quantification analysis of functional nitrification groups (detailed in the Supplementary Methods).
Data availability
All sequences of AOA-, AOB-, and comammox Nitrospira-amoA and Nitrospira-nxrB obtained in this study were deposited in GenBank, with accession numbers MZ956347 - MZ956585 . All amplicon sequencing data, unprocessed metagenomes, metagenomic assemblies, and metagenome-assembled genomes (MAGs) have been submitted to the Sequence Read Archive of the National Center for Biotechnology Information (NCBI), under the BioProject accession No. PRJNA855145. The sequences and associated annotations of the five AOA and seven Nitrospira MAGs are publicly available at MicroScope ( https://mage.genoscope.cns.fr/microscope ).
Supporting information
Author contributions
P.H., G.S. and M.L. conceived the study. P.H. and X.T. conducted the lab work. P.H. and H.K. performed the comparative genomic analyses. X.D. and P.H. analyzed the metagenomic data. D.W. and Z.L. collected the samples. Q.Z. did the physiochemical analyses. L.H. and S.L. helped with the data interpretation. P.H. and G.S. wrote the manuscript, with contributions from all co-authors.
Competing interests
The authors declare no competing interests.
Competing Interest Statement
Competing Interest Statement
The authors have declared no competing interest.
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