Adamczyk, M., Perez-Mon, C., Gunz, S., & Frey, B. (2020). Strong shifts in microbial community structure are associated with increased litter input rather than temperature in High Arctic soils. Soil Biology and Biochemistry, 151, 108054 (14 pp.). https:/

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Soil Biology and Biochemistry 151 (2020) 108054

Available online 21 October 2020

Strong shifts in microbial community structure are associated with increased litter input rather than temperature in High Arctic soils

Magdalene Adamczyk, Carla Perez-Mon, Samuel Gunz, Beat Frey

^*

Rhizosphere Processes Group, Swiss Federal Research Institute WSL, Zürcherstrasse 111, 8903, Birmensdorf, Switzerland

A R T I C L E I N F O Keywords:

Global warming High arctic Plant litter Microbiome Transplant Temperature increase

A B S T R A C T

Rising temperatures in the Arctic and the expansion of plants to higher latitudes will significantly alter belowground microbial communities and their activity. Given that microbial communities are major producers of greenhouse gases, understanding the magnitude of microbial responses to warming and increased carbon input to Arctic soils is necessary to improve global climate change models. In this study, active layer and permafrost soils from northern Greenland (81^◦N) were subjected to increased carbon input, in the form of plant litter, and temperature increase, using a combined field and laboratory approach. In the field experiment, unamended or litter-amended soils were transplanted from the permafrost layer to the top soil layer and incubated for one year, whereas in the laboratory experiment active layer and permafrost soils with or without litter amendment were incubated at 4 ^◦C or 15 ^◦C for six weeks. Soil microbial communities were evaluated using bacterial 16S and fungal ITS amplicon sequencing and respiration was used as a measure of microbial activity. Litter amendment resulted in similar changes in microbial abundances, diversities and structure of microbial communities, in the field and lab experiments. These changes in microbial communities were likely due to a strong increase in fast- growing bacterial copiotrophic taxa and basidiomycete yeasts. Furthermore, respiration was significantly higher with litter input for both active layer and permafrost soil and with both approaches. Temperature alone had only a small effect on microbial communities, with the exception of the field-incubated permafrost soils, where we observed a shift towards oligotrophic taxa, specifically for bacteria. These results demonstrate that alterations in High Arctic mineral soils may result in predictable shifts in the soil microbiome.

1. Introduction

The Arctic is changing rapidly with climate warming at a rate of more than twice the global average (IPCC, 2019), resulting in the retreat of glaciers and ice sheets, as well as the accelerated thawing of permafrost (Camill, 2005; Jorgenson et al., 2010). Permafrost is covered by an active layer, soil that undergoes seasonal freeze-thaw cycles. The depth of this active layer has increased in many locations over the last few decades, at the expense of the underlying permafrost (Gruber and Haeberli, 2009).

Despite the harsh conditions High Arctic environments present for microbial life, such as low temperatures, freeze-thaw cycles and low water availability, microbes (i.e. bacteria and fungi) are able to grow and metabolise there. In fact, active layer and permafrost soils in this region harbour a very diverse, yet largely unknown, microbial life (Gilichinsky et al., 2008; Jansson and Tas¸, 2014; Margesin and Collins,

2019; Nikrad et al., 2016; Vishnivetskaya, 2009).

In the past few decades, research on Arctic permafrost has increased considerably due to emerging concerns about the impacts of rising at- mospheric temperatures. Arctic permafrost soils are known to contain large organic carbon (C) stocks (Donhauser and Frey, 2018; Schuur et al., 2015; Tarnocai et al., 2009). As permafrost thaws, more liquid water becomes available and microbial activity increases. This can result in increased decomposition of previously frozen soil organic matter and in turn can increase the production of greenhouse gases such as CO2, CH4 and N2O (Knoblauch et al., 2013; Schuur et al., 2015).

Recent research has also documented shrub expansion in Arctic ecosystems in response to ongoing climate warming (Elmendorf et al., 2012; Myers-Smith et al., 2011; Naito and Cairns, 2011; Tape et al., 2006). This shrub expansion, also called shrubification, is manifested by an increase in shrub biomass, cover and abundance. An advancement of plants to higher latitudes will cause new interactions between plants and

* Corresponding author.

E-mail addresses: magdalene.adamczyk@wsl.ch (M. Adamczyk), carla.perezmon@wsl.ch (C. Perez-Mon), samuel.gunz@wsl.ch (S. Gunz), beat.frey@wsl.ch (B. Frey).

Contents lists available at ScienceDirect

Soil Biology and Biochemistry

journal homepage: http://www.elsevier.com/locate/soilbio

https://doi.org/10.1016/j.soilbio.2020.108054

Received 17 July 2020; Received in revised form 14 October 2020; Accepted 18 October 2020

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microbes and may lead to changes in ecosystem functions. Furthermore, it has been suggested that deep plant roots can invade newly thawed permafrost, where they interact with C and nitrogen released during thawing (Blume-Werry et al., 2019).

Root biomass provides essential C input into Arctic soils, and shrub expansion can therefore be expected to increase their C content. How- ever, root exudation and litter input can enhance soil C loss by stimu- lating microbial activity and may lead to increased mineralisation of organic material, a process known as the priming effect (Fontaine et al, 2003, 2007; Pegoraro et al., 2019; Walker et al., 2016).

An increase in C and nutrient availability as a result of shrub expansion in Arctic tundra ecosystems is expected to stimulate microbial activity overall but may benefit some taxa more than others, which could lead to shifts in taxonomic and functional traits of microbial communities (Eilers et al., 2010; Ridl et al., 2016). Following the concept of copiotrophic/oligotrophic lifestyles proposed by several au- thors (Fierer et al., 2007; Koch, 2001; Lauro et al., 2009; Roller and Schmidt, 2015), microbes with copiotrophic traits, such as fast growth and metabolic versatility, respond relatively quickly to an increase in C and nutrient availability. In contrast, oligotrophic taxa are characterised by an inability to grow and thrive in nutrient-rich environments, and they exhibit slower growth rates and more stress tolerance.

Both changes in plant cover and rising temperatures are expected to lead to fundamental shifts in microbial communities in Arctic soils.

However, studies on soil microbial communities and their responses to substrate input such as litter are scarce, particularly in remote High Arctic regions. Most research has focussed on the decomposition of soil organic matter in response to substrate input in Arctic soils (Pegoraro et al., 2019; Wild et al., 2014). The effects of substrate input on microbial communities have been studied, for example litter in subarctic heath soils (Rinnan et al., 2008) and fertiliser in moist acidic Arctic tundra soils (Campbell et al., 2010; Koyama et al., 2014), yet bacterial and fungal communities in the High Arctic polar desert have received little attention. More research has been conducted on the effects of temperature on microbial communities in Arctic soils (Ballhausen et al., 2020; Deslippe et al., 2012; Feng et al., 2020; Johnston et al., 2019). In a 16-year-long experiment Lamb et al. (2011) found that microbial communities in the High Arctic appear to be resistant to nutrient input and warming, and that such alterations also have little impact on greenhouse gas fluxes. To date, little is known about how climate change will impact belowground communities and their activity in active layer and permafrost soils, or the subsequent effects on soil C dynamics, particularly in remote High Arctic regions.

In this study, we aimed to deepen the understanding of the effects of litter input and temperature on microbial communities in active layer and permafrost soils from a remote site in northern Greenland (81^◦N).

Our objectives were to examine: (i) how litter input and temperature affect microbial activity; (ii) how bacterial and fungal abundances, diversities and structure of microbial communities are affected; and (iii) which taxonomic groups are favoured or hindered as a result of these treatments. We conducted two experiments, one in the field and one in the laboratory. For both experiments, active layer and permafrost soils were amended with dried leaves of the plant Ledum groenlandicum (Ericaceae), which is native in southern Greenland as well as in the Canadian Arctic. In the lab experiment, active layer and permafrost soils were incubated at 4 ^◦C or 15 ^◦C over a period of six weeks, during which CO₂measurements were taken once a week. For the field experiment, control and litter-amended active layer and permafrost soils were transplanted to the uppermost soil layers and incubated for one year. To determine the effect of temperature, a comparison was performed between transplanted permafrost unamended soils after one year of incubation and the same soils at the start of the incubation period. Microbial communities were analysed using bacterial 16S and fungal ITS amplicon sequencing.

2. Materials and methods

2.1. Site characteristics and soil sampling

The study site was located in northern Greenland at the Villum Research Station (VRS, 81^◦36^′N, 16^◦40^′W, 24 m a. s. l.). Hourly soil temperature measurements made on site at a depth of 5 cm (two Geo- Precision sensors, GeoPrecision GmbH, Ettlingen, Germany) during the one-year field incubation showed minimum and maximum temperatures of − 16.3 ^◦C and 19.0 ^◦C. Mean soil temperatures were 5.8 ±3.8 ^◦C in August 2018 and 10.2 ±3.7 ^◦C in July 2019 (Table S1; Fig. S1). Soils were covered by snow from beginning of October 2018 until beginning of July 2019 (based on HOBO light data loggers, Onset Computer Cor- poration, Pocasset, MA, USA) (Fig. S1). Mean annual precipitation (MAP) in the region is 188 mm (https://eu-interact.org/field-sites/villu m-research-station/). Permafrost in the area is continuous and it is assumed to occur below a depth of 20 cm (https://eu-interact.org/field-s ites/villum-research-station/). The bedrock is described as quaternary, undifferentiated cover and dominated by carbonate minerals (htt ps://data.geus.dk). The terrain features patterned ground forms (i.e.

ice-wedged polygons) with patchy occurrences of biological soil crusts and scattered vascular plants, in particular Saxifraga oppostifolia (family Saxifragaceae), Papaver radicatum (family Papaveraceae) and Draba spp (family Brassicaceae).

Soils were collected in July 2018 during a field campaign at VRS.

Three soil profiles located 200 m apart from each other were excavated with shovels down to a depth of 50 cm. Soils were >90% sand and slightly alkaline (pH =7.4), with total C ranging from 0.7 to 0.8% and total N (nitrogen) values of 0.1% (Table S2). In line with the previous information about the permafrost depth (https://eu-interact.org/field-s ites/villum-research-station/), temperature measured along the soil profiles during the excavation showed values below 0 ^◦C at a depth of 30 cm (data on request). Accordingly, bulk soil samples were collected from each of the profiles at depths of 5–10 cm (active layer, AL) and 35–45 cm (permafrost, PF), and sieved with a 4 mm mesh. To minimise the con- founding effects linked to the spatial heterogeneity of the terrain (e.g.

variations in vegetation cover and soil moisture between the soil profiles), pools of AL and PF (approx. 20 kg in total for each soil type) were prepared by homogenising the respective samples collected from the three different soil profiles. Pre-sterilised equipment was used during the collection and processing of the soil samples. Samples were directly used in the field for the soil transplant experiment or transported frozen to the WSL laboratory facilities, where they were stored at − 2 ^◦C before being used for the incubation experiment (see below).

2.2. Soil transplantation and litter amendment set-up (field study) Aliquots of about 1 kg of the AL and PF pools were used to establish the in situ incubations. The soil aliquots were either amended with dried leaves of Ledum groenlandicum (C: 52%; N: 1.5%, TrancePlants, Quebec, Canada) or left unamended (controls). The substrate amounts were adjusted to final concentrations of 5 mg C g⁻¹DW, which adds to the original soil C concentrations (7 mg C g⁻¹DW). The soils were then transferred to stainless steel containers (12 cm diameter x 13 cm height, catalogue no. 300.118.32, IKEA, Almhult, Sweden). Four experimental ¨ replicates were established for each soil type (AL, PF) and treatment (with or without plant litter), producing a total of 16 containers filled with the soil samples. The containers were perforated at the bottom to allow exchange of gases and water with the surroundings while at the same time preventing waterlogging (Zumsteg et al., 2013a). The 16 soil samples were placed in a randomised arrangement, at an unvegetated land area approx. 2 km away from the VRS. The containers were buried in the surface, leaving a rim of approx. 1 cm to avoid the runoff of external materials into the containers. A pit of approx. 1 m²was excavated down to a depth of 40 cm. Containers filled with amended (4x) and unamended (4x) PF soils (8 containers in total) were buried at the

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bottom of the pit, together with two GeoPrecision sensors. The pit was closed again using the excavated soil materials. The containers were covered with stones to prevent the surrounding soil from mixing with the experimental soils. These containers will serve as long-term incubation controls (e.g. resampling after 5 years). Aliquots from the AL and PF pools were collected at the start of the experiment (July 19, 2018).

After one year (August 10, 2019), soil aliquots of 25 g were collected from the middle of the experimental containers (i.e. field-incubated soils), at a depth of 0–5 cm, using ethanol-sterilised shovels. Samples were transported on ice and stored in the lab at − 20 ^◦C until analyses were performed.

2.3. Incubation experiment under controlled conditions

A 35-day incubation experiment was set up similarly to in previous laboratory-based temperature studies (Donhauser et al., 2020; Lul´akov´a et al., 2019; Perez-Mon et al., 2020). An 80 g aliquot of wet soil from each layer (AL and PF) was weighed into 250 ml glass jars and covered with a cotton wool permeable lid to allow for aeration. The glass jar samples were amended with dried leaves of Ledum groenlandicum or left unamended (controls). The amount of added plant leaves was the same as in the field study (adjusted to 5 mg C g⁻¹soil). The glass jars were incubated in growth chambers set to 4 ^◦C or 15 ^◦C. There were four replicates for each soil layer (AL, PF), treatment (with or without plant litter) and temperature (4 ^◦C, 15 ^◦C), resulting in a total of 32 glass jars.

At the end of the incubation period, the lab-incubated soils were analysed for DNA content and the structure of microbial communities, microbial abundance and respiration. The temperatures of 4 ^◦C and 15 ^◦C were chosen based on historical data (>10 years) of daily air temperature measurements at VRS (https://www2.dmu.dk/asiaqmet/Default.as px). The temperature of 4 ^◦C is common at daytime during July and August, whereas 15 ^◦C represents the maximum temperature that is reached at the site during this period.

2.4. DNA extraction, PCR amplification and Illumina MiSeq sequencing Total genomic DNA was extracted from approximately 0.5 g of soil per sample using the DNeasy PowerSoil Kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions. DNA was quantified with PicoGreen (Invitrogen, Carlsbad, CA, USA). The V3 –V4 region of the bacterial small-subunit (16S) rRNA gene and the internal transcribed spacer region 2 (ITS2) of the eukaryotic (fungal groups and some groups of protists and green algae) ribosomal operon were PCR amplified from 10 ng of DNA template, using primers and conditions previously described (Adamczyk et al., 2019; Frey et al., 2016). Negative controls were included for the DNA extractions (extraction buffer without soil) and PCR amplifications (high purity water without DNA template).

Triplicate PCR reactions were pooled, purified using Agencourt Ampure XP (Beckman Coulter, Beverly, USA) and sent to the Génome Québec Innovation Centre at McGill University (Montréal, Canada) for paired-end sequencing on the Illumina MiSeq v3 platform (Illumina Inc., San Diego, CA, USA). Raw sequences were deposited in the NCBI Sequence Read Archive under the BioProject accession number PRJNA645283.

2.5. Sequence quality control, OTU clustering and taxonomic assignments Quality filtering, clustering into operational taxonomic units (OTUs) and taxonomic assignment were performed as previously described (Adamczyk et al., 2019; Donhauser et al., 2020; Frey et al., 2016) using a customised pipeline based on UPARSE (Edgar, 2013).The pipeline is based on clustering sequences into OTUs at 97% identity. Prokaryotic centroid sequences were queried against the SILVA database v. 132 (Quast et al., 2013). Eukaryotic centroid sequences were first curated using a custom-made ITS2 database retrieved from NCBI GenBank, and sequences assigned to fungi were classified to finer taxonomic levels

using the UNITE database v. 8.0 (Abarenkov et al., 2010). Prokaryotic sequences identified as originating from organelles (chloroplast, mito- chondria), as well as eukaryotic sequences identified as originating from soil animals (Metazoa) or plants (Viridiplantae), or of unknown eukaryotic origin, were removed from downstream analyses. A brief description of the overall microbial community composition as well as complete lists of all archaeal, bacterial and fungal OTUs including taxonomic assignment, the number of sequences and abundance information can be found in Supplementary Results and Data S1.

2.6. Microbial parameters

Soil basal respiration was measured once per week during the laboratory incubation period. The respiration measurements were performed 7 days after the start of the incubation, to avoid potential artefacts in microbial activity estimations created by the soil manipu- lation during experiment set-up. Five measurement series were performed, each approx. one week apart. During each measurement series, the mesocosms were gas-tight sealed, and CO2 concentrations accumu- lating in the flasks were measured at two different time points (0 h and 24 h for controls, 0 h and 2 h for litter-amended samples, with the exception of litter-amended AL 4 ^◦C and PF 4 ^◦C, which were also measured after 24 h during the first two measurements series) using an infrared absorption CO2 analyser (EGM-4 Environmental Gas Monitor;

PP systems, Amesbury, MA, USA) as previously described (Lul´akov´a et al., 2019; Perez-Mon et al., 2020). The gas-tight seals were replaced by permeable cotton wool lids after each CO2 measurement series (maximum duration of 24 h). Similar gas measurements were performed on the one-year in situ incubated soils at 4 ^◦C, directly in the lab facilities of VRS. To express respiration rates per soil dry weight, the final respiration rates were converted from the headspace CO2 change in ppm to μmol C–CO2 g⁻¹day⁻¹.

Bacterial and fungal abundances were determined by quantitative PCR (qPCR) on a 7500 Fast Real-Time PCR System (Thermo Fisher Scientific, Waltham, Massachusetts, USA). qPCR reactions were prepared using the universal primer pairs 27F/519 R amplifying the V1 – V3 region of the 16S rRNA gene (prokaryotes) and ITS3/ITS4 amplifying the internal transcribed spacer region 2 (ITS2) of the eukaryotic ribosomal operon (fungi). qPCR programs were run as described by Rime et al. (2016a).

2.7. Data analysis

All statistical analyses were performed using R (v. 3.6.0; R Core Team, 2019) and all graphs were generated with the ggplot2 package (v.

3.2.0; Wickham, 2016), unless specified otherwise.

For analysis of microbial alpha diversity, observed richness (number of OTUs) and Shannon diversity index were estimated based on OTU abundance matrices rarefied to the lowest number of sequences per sample (lab experiment: bacteria 13,919 and fungi 7,633 sequences;

field experiment: bacteria 17,012 and fungi 8,905 sequences) using the R package phyloseq (v. 1.28.0; McMurdie and Holmes, 2013). To assess the main and interactive effects of litter input and incubation temperature on alpha diversities of the AL and PF lab-incubated soils, a two-way analysis of variance (ANOVA) was performed. Correspond- ingly, a one-way ANOVA was performed to assess the effects of litter and temperature (permafrost soil transplanted to the top soil layer) on microbial alpha diversity in field-incubated soils. Pairwise comparisons of significant effects were conducted using Tukey’s HSD post-hoc tests.

Similarly, the main and interactive effects of soil type, temperature and litter input on respired CO2 were tested by ANOVA. Furthermore, the main and interactive effects of litter input and temperature (lab experiment) and of litter only (field experiment) on microbial abundance (total DNA content as a proxy) along with 16S and ITS copy numbers were tested for each soil type using ANOVA. In a similar manner, we examined the effect of transplanting permafrost to the top

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soil layer on microbial abundance. Assumptions regarding homosce- dasticity and normality of residuals were tested and, where necessary, transformation (Tukey’s ladder of powers) of response variables was performed.

Bray-Curtis dissimilarities were calculated based on square-root- transformed relative abundances of OTUs. The effect of soil type on the structure of microbial communities (beta diversity) was assessed by conducting a permutational ANOVA (PERMANOVA, number of permutations =9,999) with the function ‘adonis’ implemented in the R package vegan (v. 2.5.5; Oksanen et al., 2019). Similarly, for each soil type the main and interactive effects of litter and incubation temperature (lab experiment) and litter alone (field experiment) on the structure of microbial communities were analysed. In the same way, the effect of transplanted permafrost (field experiment) on the structure of microbial communities was examined. Principal coordinate analysis (PCoA) or- dinations of the structure of microbial communities were calculated using the ‘ordinate’ function implemented in the R package phyloseq.

To identify microbial phyla and genera that were significantly different between litter-amended and unamended samples, we first agglomerated OTUs to the phylum and genus level, and generated subsets for each soil type. To avoid problems that can arise from ana- lysing compositional data, we performed differential abundance analysis by applying a negative binomial generalised linear model to the OTU count data using the R package DESeq2 (v. 1.24.0; Love et al., 2014). For the lab experiment, the effect of litter amendment on microbial taxa was assessed by controlling first for incubation temperature.

We applied normal shrinkage to log2 fold changes using the ‘lfcShrink’ function implemented in DESeq2. Phyla and genera were considered significantly different (Wald test) between litter-amended and unamended samples if the false discovery rate (adjusted p-value) was

<0.01. Only the ten most abundant phyla and genera with significant log2 fold changes were plotted. The relative abundances of microbial taxa displayed in the plots include data from both unamended and litter-amended samples for each soil type and were grouped into cate- gories. The same analysis was used for examining significant differences in microbial taxa of permafrost soil incubated for one year in the top soil layer (temperature treatment).

3. Results

3.1. Changes in respiration

During the six-week lab incubation of active layer and permafrost soils, we measured the respired CO₂of litter-amended and unamended soils once a week (Fig. 1). Litter-amended active layer and permafrost soils incubated at 15 ^◦C had similar respiration rates, with the highest

initial levels across all samples (>0.45 μmol C–CO2 g⁻¹day⁻¹), which then dropped steeply over the measurement period. On the other hand, respiration rates of active layer and permafrost soils incubated at 4 ^◦C were initially close to zero, then rose quickly within the first 8 days.

From approximately 16 days onwards the respiration rates of both 4 ^◦C and 15 ^◦C samples settled to comparable levels (between 0.06 and 0.2 μmol C–CO2 g⁻¹day⁻¹). In contrast, none of the unamended soils exhibited respiration rates of greater than 0.0045 μmol C–CO2 g⁻¹day⁻¹ over the testing period. We found a statistically significant relationship between respiration rates and litter amendment, whereas temperature alone had a relatively minor effect despite the large difference during the first two measurements. Furthermore, no significant differences were found between the active layer and permafrost soils either with or without litter addition (Table S3). Respiration rates of active layer and permafrost soils incubated in the field for one year with or without litter input also showed a significant increase in respired CO2 in litter- amended soils (Fig. S2).

3.2. Microbial abundance

Total DNA content, used as a proxy for microbial biomass, as well as bacterial 16S and fungal ITS copy numbers, used as proxies for bacterial and fungal abundance, respectively, significantly increased in litter- amended active layer and permafrost soils (Table 1; Fig. S3). We observed the same patterns for both the lab and field experiment, with the exception of bacterial abundance in active layer soils during the one- year field incubation, in which litter addition resulted in a decrease in DNA content. Moreover, we found a significant effect of temperature alone on 16S copy numbers and on total DNA content in lab-incubated permafrost soils (Table 1).

The ratio of fungal ITS to bacterial 16S copy numbers, i.e. the F:B ratio, increased five-fold in litter-amended soils (F:B 0.0041) relative to unamended soils (F:B 0.0008) in the lab experiment. Field-incubated soils exhibited an even greater, nine-fold increase in the F:B ratio in litter-amended soils (F:B 70.1) compared with unamended soils (F:B 7.5).

Bacterial 16S copy numbers did not change significantly after the one-year incubation of permafrost soils transplanted to the top soil layer to simulate an increase of temperature. In contrast, fungal ITS copy numbers and the total DNA content exhibited a significant increase in transplanted permafrost (Table 1; Fig. S3).

3.3. Changes in microbial alpha diversities in response to litter amendment and temperature

We observed a decrease in microbial diversity, richness and Shannon index with litter amendment in both the field and the laboratory experiment (Fig. 2). In the lab experiment, bacterial and fungal richness were significantly lower (bacteria: F(1,24) =219.6, p <0.001; fungi:

F(1,24) =40.1, p <0.001) in permafrost soils than in the active layer.

Shannon diversity was less affected (bacteria: F(1,24) =17.1, p <0.001;

fungi: F(1,24) =5.4, p =0.03) by soil type. Litter amendment was the strongest driver of changes in bacterial diversity, as shown for individ- ually analysed soils in Table 2. Bacterial richness and Shannon diversity decreased by more than 50% in response to litter amendment in 4 ^◦C- and 15 ^◦C-lab-incubated active layer and permafrost soils (Fig. 2A, C).

Fungal diversity was less affected by litter amendment (Fig. 2B, D;

Table 2). Incubation temperature alone had only a marginal effect on microbial diversities compared with the effect of litter input (Table 2).

In the field experiment, litter amendment also resulted in a decrease in microbial diversities (Fig. 2E – H); however, the overall response was weaker than the results for lab-incubated soils. In contrast to bacteria, for which richness and Shannon diversity decreased significantly in active layer and permafrost soils, fungal diversity was only significantly affected by litter input in permafrost soils (Table 2; Fig. 2E – F). In permafrost soils transplanted to the top soil layer, mimicking a Fig. 1.Respiration rates (μmol C–CO2 g⁻¹dry soil day⁻¹) of active layer and

permafrost soils incubated at either 4 ^◦C or 15 ^◦C, with or without litter amendment, during a five-week period. The inset depicts a detailed view of the control (unamended) samples. Mean values with standard errors are shown for each measurement series starting 7 days after initial temperature incubation.

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temperature increase of approx. 4 ^◦C, bacterial diversity slightly increased during the one-year field incubation; however, this effect was only significant for bacterial Shannon diversity (Fig. 2G; Table 2).

Fungal diversity was not significantly altered in the transplanted permafrost soils (Fig. 2F, H; Table 2).

3.4. Changes in the structure of microbial communities in response to litter amendment and temperature

Principal coordinate analysis (PCoA) revealed similar patterns of change for bacterial and fungal communities in the lab and field incubation experiments (Fig. 3). Bacterial communities in lab-incubated active layer and permafrost soils of unamended samples clustered closely together (Fig. 3A), although in the field experiment significant differences were found among communities between unamended samples of active layer and permafrost soils (after the one-year incubation) (F(2,10) =2.2, p =0.009; Fig. 3C). The fungal community appeared to be more heterogeneous across unamended samples (Fig. 3B, D). As for bacteria, significant differences in fungal community structure in active layer and permafrost soils were only found for unamended samples in the field experiment (F(2,10) =2.9, p <0.001).

The greatest changes in the structure of bacterial and fungal communities, which were highly significant, were found between litter- amended soils and unamended ones. Lab-incubated soils clustered by soil type and incubation temperature (Fig. 3A and B), but the only significant differences in the structure of microbial communities were between permafrost soils incubated at 4 ^◦C and 15 ^◦C (Table 3). After one year of field incubation, litter-amended active layer and permafrost soils had distinct microbial communities compared with unamended samples (Fig. 3C and D; Table 3).

Bacterial community structure shifted in permafrost soils that were transplanted to the top soil layer and incubated for one year in the field.

The community became more similar to that in active layer soils, yet still differed significantly (Table 3; Fig. 3C). For fungi, the shift in community structure that occurred when permafrost soils were transplanted was less pronounced but still significant (Table 3; Fig. 3D).

3.5. Differential abundance of bacterial and fungal phyla in response to litter amendment

We further analysed which taxa, in particular among the most

abundant phyla and genera, drove the changes in the structure of bacterial and fungal communities in soils with and without litter input.

Using differential abundance analysis (Figs. 4 and 5), we observed similar patterns of change for lab- and field-incubated soils. As expected, changes in abundance at the phylum level were less prominent than at the genus level. In active layer and permafrost soils in both the lab and field experiment, we observed an increase in Proteobacteria, Actino- bacteria and Bacteriodetes. The phylum Patescibacteria also increased as a result of litter amendment, except in lab-incubated permafrost soils.

Firmicutes increased in lab-incubated soils, predominantly in permafrost soils, yet this phylum in general had a much lower abundance compared with the aforementioned phyla. Responses of phyla to litter addition were less clear between lab and field experiments. Of the most abundant phyla, Verrucomicrobia, Acidobacteria and Gemmatimona- detes decreased in abundance in lab-incubated active layer and permafrost soils. For samples that were incubated in the field, the only significant decrease in active layer soils was found for the less abundant phylum Fibrobacteres (<1%). Furthermore, field-incubated permafrost soils exhibited a decrease in the more abundant Acidobacteria and Gemmatimonadetes, similar to the findings for lab-incubated soils.

Within the fungal kingdom, the only significant increase in abundance in lab-incubated soils in response to litter input was found for the phylum Basidiomycota in active layer soils. Phyla that significantly decreased in active layer soils were only the less abundant (<1%) Rozellomycota, Basidiobolomycota and Mortierellomycota. In lab- incubated permafrost soils only Mortierellomycota and Mucrocomy- cota decreased in response to litter addition. In the field experiment, the same phyla in both active layer and permafrost soils showed significant responses; Basidiomycota increased in abundance, while Mortier- ellomycota strongly decreased. No significant changes were found for the highly abundant (82.6%) phylum Ascomycota.

3.6. Differential abundance of bacterial and fungal genera in response to litter amendment

We examined the responses of the ten most abundant bacterial and fungal genera to litter input (Figs. 4 and 5). In almost all cases genera belonging to the phyla Proteobacteria, Actinobacteria and Bacteriodetes significantly increased in abundance in litter-amended active layer and permafrost soils during lab and field incubations. Of particular note was the genus Pseudomonas (phylum Proteobacteria), which increased Table 1

Main and interactive effects of litter input and incubation temperature on bacterial 16S and fungal ITS copy numbers and total DNA content in lab-incubated active layer and permafrost soils (upper section); and main effects of litter input and one-year incubation in the top soil layer (permafrost soil transplant) during the field experiment (lower section).

16S copy numbersâ ITS copy numbersâ Total DNAâ

F-value p-value F-value p-value F-value p-value

Lab incubation Active layer

Litter 83.56(1,12) < 0.001 154.00(1,12) < 0.001 1.25(1,12) 0.29

Temperature 0.89(1,12) 0.36 2.68(1,12) 0.13 1.46(1,12) 0.25

Litter ✕ Temperature 5.39(1,12) 0.04 4.12(1,12) 0.07 2.57(1,12) 0.14

Permafrost

Litter 113.89(1,12) < 0.001 55.52(1,12) < 0.001 104.52 < 0.001

Temperature 5.85(1,12) 0.03 2.80(1,12) 0.12 8.95(1,12) 0.01

Litter ✕ Temperature 5.54(1,12) 0.04 0.15(1,12) 0.71 0.24(1,12) 0.63

Field incubation Active layer

Litter 13.41(1,6) 0.01 53.68(1,6) < 0.001 27.6(1,6) 0.002

Permafrost

Litter 64.88(1,6) 0.005 36.74(1,6) < 0.001 13.24(1,6) 0.01

Top soil transplant 0.30(1,5) 0.61 9.27(1,5) 0.03 35.98(1,5) 0.002

The F-statistics of ANOVAs are shown. Subscripts denote the degrees of freedom and residuals for each factor. Significant effects (p-value <0.05) are given in bold.

a16S/ITS copy numbers: 16S/ITS copies μ_g⁻¹DNA, Total DNA: μ_{g DNA g}⁻¹_{dry soil.}

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considerably in litter-amended active layer and permafrost soils in both the lab and field incubations. Pseudomonas was also by far the most abundant genus in lab-incubated soils. For the active layer and permafrost soils in the field experiment, we also observed significant increases in the genera Caulobacter, Massilia, Rhizobium and Sphingomonas, which are also members of the phylum Proteobacteria. The genera Galbitalea and Pseudarthrobacter (both Actinobacteria) significantly increased in litter-amended active layer and permafrost soils in the field experiment.

In the lab-incubated soils only Galbitalea increased in both soil types, while Pseudarthrobacter only become more abundant in active layer soils. Flavobacterium (Bacteriodetes) exhibited a significant increase in active layer and permafrost soils in both the lab and field experiment.

The abundances of RB41 (Acidobacteria) and Chthoniobacter (Verruco- microbia) significantly declined in response to litter input in the soils of both experiments. Overall, we observed similar patterns for bacterial genera between the lab and field experiments, as well as between active

layer and permafrost soils in response to litter addition.

Among fungal genera, the majority that exhibited changes in response to litter input, either increasing or decreasing in abundance, were of the phylum Ascomycota. Lichenised fungi, such as Verrucaria and lichenicolous Rhinocladiella, decreased in field-incubated active layer and permafrost soils, while Cephalotrichum showed the greatest increase. Similarly, in lab-incubated soils Rhinocladiella decreased, whereas Verrucaria only increased in active layer soils. In both experiments we observed an increase in Pseudogymnoascus, which was notably abundant in field-incubated permafrost soils. In lab-incubated active layer and permafrost soils, the basidiomycete yeast Mrakia exhibited a particularly large increase, but the yeast Naganishia also significantly increased as a result of litter input. In field-incubated soils, there was also an increase in basidiomycete yeast genera, such as Mrakia, Mrakiella and Bullera in active layer soils and Vishniacozyma, Mrakia and Naga- nishia in permafrost soils. In both the lab and field experiment the genus

Bacteria Fungi

Lab experimentField experiment

Active Layer 4°C Active Layer 15°C Permafrost 4°C Permafrost 15°C Control Litter Control Litter Control Litter Control Litter 0

50 100 150 200

Richness

B *

1 2 3 4

Shannon Index

D _* _**

500 1000 1500

Richness

A *** *** *** ***

2 4 6

Shannon Index

C _*** _*** _*** _***

Active Layer Permafrost

Year 1 Control Year 1

Litter Year 0

Control Year 1 Control Year 1

Litter 0

500 1000 1500 2000 2500

Richness

E *** **

Litter Year 0

Litter 0

2 4 6

Shannon Index

G *** ** ***

Litter Year 0

Litter 0

50 100 150 200

Richness

F ***

Litter Year 0

Litter 0

1 2 3

Shannon Index

H ***

Fig. 2. Variation in richness and Shannon index of microbial communities in lab- and field-incubated active layer and permafrost soils with and without litter amendment. (A) Bacterial and (B) fungal richness; (C) bacterial and (D) fungal Shannon index in lab- incubated soils. (E) Bacterial and (F) fungal richness; (G) bacterial and (H) fungal Shan- non index in field-incubated soils. Mean values (with standard errors) of four replicates are shown (except in E − H for Permafrost Year 0 Control ⌠unamended⌡ with three replicates). Asterisks represent the significance of differences between litter- amended and control (unamended) samples.

Significance levels: * p <0.05, ** p <0.01,

*** p <0.001.

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Mortierella decreased greatly.

3.7. Differential abundance of bacterial and fungal taxa in transplanted permafrost soils

Bacterial phyla in unamended permafrost soils transplanted to the top soil layer responded in the opposing direction as in litter-amended permafrost soils. Phyla such as the relatively abundant Proteobacteria and Actinobacteria decreased in transplanted permafrost soils, whereas Acidobacteria, Chloroflexi, Verrucomicrobia and Planctomycetes increased in abundance (Fig. S3). Overall, however, the responses of bacterial phyla in the transplanted permafrost soils were weak. For bacterial genera, the strongest responses were observed for Pseudomonas and Rhodoferax (both Proteobacteria) as well as Aeromicrobium (Acti- nobacteria), which decreased. The Planctomycetes genera Pir4 Lineage, Gemmata and Pirellula all increased in abundance, albeit these increases,

in relation to the observed decreases, were only marginal.

None of the fungal phyla exhibited a significant response in transplanted permafrost soils (Fig. S4). Only a few less abundant genera (<1%) were found to have a significant response: Botrytis and Venturia (both Ascomycota) as well as Dioszegia (Basidiomycota) increased, while Didymella (Ascomycota) decreased in abundance.

4. Discussion

4.1. Litter input enhances microbial activity

In this study, we aimed to elucidate the effects of plant litter (L. groenlandicum) input and temperature increase, in both lab and field conditions, on the soil microbiome of active layer and permafrost soils from northern Greenland (81^◦N). Litter addition to both active layer and permafrost soils resulted in significantly enhanced soil respiration, a Table 2

Main and interactive effects of litter input and incubation temperature on bacterial and fungal alpha diversities in lab-incubated active layer and permafrost soils (upper section); and main effects of litter input and one-year incubation in the top soil layer (permafrost soil transplant) during the field experiment (lower section).

Bacteria Fungi

Richness Shannon Richness Shannon

F-value p-value F-value p-value F-value p-value F-value p-value

Lab incubation Active layer

Litter 1823.22(1,12) < 0.001 3294.43(1,12) < 0.001 15.15(1,12) 0.002 12.58(1,12) 0.004

Temperature 6.18(1,12) 0.03 29.33(1,12) < 0.001 0.20(1,12) 0.66 3.31(1,12) 0.09

Litter ✕ Temperature 0.43(1,12) 0.52 18.50(1,12) 0.001 1.05(1,12) 0.33 0.65(1,12) 0.43

Permafrost

Litter 757.84(1,12) < 0.001 1329.15(1,12) < 0.001 0.79(1,12) 0.39 16.73(1,12) 0.002

Temperature 1.41(1,12) 0.26 59.86(1,12) < 0.001 3.20(1,12) 0.10 0.01(1,12) 0.91

Litter ✕ Temperature 2.66(1,12) 0.13 7.26(1,12) 0.02 7.11(1,12) 0.02 3.50(1,12) 0.09

Field incubation Active layer

Litter 59.92(1,6) < 0.001 178.80(1,6) < 0.001 2.58(1,6) 0.16 2.58(1,6) 0.16

Permafrost

Litter 30.72(1,6) 0.001 154.90(1,6) < 0.001 187.80(1,6) < 0.001 186.40(1,6) < 0.001

Top soil transplant 3.18(1,5) 0.14 32.25(1,5) 0.002 2.46(1,5) 0.18 1.78(1,5) 0.24

The F-statistics of ANOVAs are shown. Subscripts denote the degrees of freedom and residuals for each factor. Significant effects (p-value <0.05) are given in bold.

Fig. 3. Variation in the structure of microbial communities in response to litter input to active layer and permafrost soils. The upper row shows the structure of (A) the bacterial and (B) the fungal communities in lab-incubated soils and the lower row shows the structure of (C) bacterial and (D) fungal communities in field-incubated soils. Principal coordinate analyses (PCoA) are based on Bray-Curtis distance matrices.

Distances between symbols on the ordination plots reflect relative dissimilarities in the community structure. The variation in the structure of microbial communities explained by each PCoA axis is given in parentheses.

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finding also observed in several other studies on soils from the Alaskan tundra (Lynch et al., 2018), Siberian tundra (Walz et al., 2017), subarctic tundra heath (Jonasson et al., 2004), alpine glacier forefields (Zumsteg et al., 2013b) and switchgrass fields (De Graaff et al., 2010).

This increase in respiration rate was, however, short-lived and slowly decreased over time.

The decline in respiration after the initial surge was likely due to the depletion of energy-rich C sources, as suggested by Hartley et al. (2008).

Similar observations were previously made by Zumsteg et al. (2013b), who noted a peak in respiration in nutrient-poor soils from a glacier forefield, followed by a decrease before stabilising 13 days after substrate addition. In particular, the substrate depletion was likely the result of a rapid utilisation of easily available C substrates, leaving behind more complex C compounds.

Incubation temperature alone (4 ^◦C vs. 15 ^◦C) only had a marginal effect on respiration rate, and differences were mainly present at the start of the six-week lab incubation. During this period, the respiration rates of 4 ^◦C-incubated soils were considerably lower than those of the 15 ^◦C-incubated soils. This is likely explained by increased microbial activity at higher temperatures (Jansson and Tas¸, 2014; Lul´akov´a et al., 2019), indicating that additional energy-rich inputs to High Arctic soils that are becoming warmer under climate change will likely cause higher CO₂emissions. Furthermore, research suggests that C input into soils in which energy- and nutrient-poor conditions prevail for microbes may increase soil respiration as a result of positive priming (Fontaine et al., 2004). Increased carbon and nutrients availability in the soils, as well as higher temperatures also favour plant growth, which potentially coun- teracts these effects. Further investigation is required into the soil microbial-plant interactions in the High Arctic to determine their long-term effect on soil C stocks.

4.2. The microbial community structure shifts with litter input and temperature

In both the lab and field experiment, litter amendment led to a significantly reduced microbial alpha diversity. The decrease in both richness and Shannon diversity was particularly pronounced for bacteria in lab-incubated soils. Previous studies have shown similar findings, with bacterial diversity declining considerably in response to glucose addition to soils from a ponderosa pine ecosystem (Mau et al., 2015) and

to plant residue input to temperate soils (Pascault et al., 2013). We also observed a strong increase in bacterial abundance with litter amendment, suggesting that the decrease in bacterial diversity was likely due to fast-growing populations profiting from the newly added plant litter, as previously suggested by Bernard et al. (2007). Fungal richness and Shannon diversity were less affected by the addition of litter compared with bacterial diversity, yet fungi also decreased significantly in diversity. As observed for bacteria, litter amendment led to a higher fungal abundance, indicating an increase of opportunistic fungal taxa with enhanced resource utilisation in response to litter input. The incubation temperatures of 4 ^◦C and 15 ^◦C used in the lab experiment played only a minor role in microbial alpha diversities. Similar observations for bacterial alpha diversity and temperature effects were previously reported by Donhauser et al. (2020) and Lul´akov´a et al. (2019) for alpine soils.

The decrease in microbial alpha diversities in litter-amended soils was further accompanied by strong shifts in the structure of bacterial and fungal communities in both the lab and field experiment. The observed shifts in the structure of microbial communities likely resulted from the expansion of fast-growing bacteria and fungi, which are known to respond quickly to added labile substrates as found in various other studies (Blagodatskaya and Kuzyakov, 2008; Stenstrom et al., 2001; ¨ Trivedi et al., 2018; Wild et al., 2014). These taxa likely also profited from the easily degradable compounds of the added litter to the examined soils. The strong increase of the bacterial 16S copy numbers further indicates a shift towards a copiotrophic community. Copiotrophs possess a higher copy number of the 16S rRNA gene which presumably enable them to capitalize rapidly on sudden availability of carbon (Klappen- bach et al., 2000). Large shifts in the structure of bacterial communities in response to fertiliser addition (nitrogen and phosphorous) and C substrates of varying chemical recalcitrance have been reported previously for terrestrial cryoenvironments such as Arctic tundra soils (Koyama et al., 2014) and glacier forefields (Rime et al., 2016b; Zumsteg et al., 2013b). Moreover, in lab-incubated soils, we found that temperature also affected the structure of microbial communities; however, this effect was only apparent for permafrost soils. An increase in soil temperature has been shown to induce changes in the structure of microbial communities in Arctic soils (Deslippe et al., 2012; Feng et al., 2020).

In transplanted permafrost (with no litter addition) incubated over a period of one year in the top soil layer, no significant changes were found for microbial diversities. Correspondingly, only marginal Table 3

Main and interactive effects of litter input and incubation temperature on the structure of bacterial and fungal communities in lab-incubated active layer and permafrost soils (upper section); and main effects of litter input and one-year incubation in the top soil layer (permafrost soil transplant) during the field experiment (lower section).

Bacteria Fungi

F-value p-value R² F-value p-value R²

Lab incubation

Active layer 24.4(1,12) 0.0001 59.8% 10.8(1,12) 0.0001 42.2%

Litter 2.2(1,12) 0.128 5.3% 1.5(1,12) 0.135 6.0%

Temperature 2.2(1,12) 0.142 5.4% 1.3(1,12) 0.155 5.0%

Litter ×Temperature

Permafrost 32.1(1,12) 0.0001 63.0% 7.6(1,12) 0.0001 32.1%

Litter 3.4(1,12) 0.039 6.7% 2.0(1,12) 0.037 8.4%

Temperature 3.4(1,12) 0.034 6.7% 2.1(1,12) 0.028 8.8%

Litter ×Temperature Field incubation

Active Layer 8.2(1,6) 0.031 57.8% 3.8(1,6) 0.032 39.9%

Litter

Permafrost 10.1(1,6) 0.028 62.6% 7.3(1,6) 0.030 55.0%

Litter 3.1(1,5) 0.032 38.0% 1.8(1,5) 0.029 26.0%

Top soil transplant

The pseudo F-statistics of PERMANOVAs are shown (number of permutations 9,999, except for top soil transplant 5,039). Subscripts denote the degrees of freedom and residuals for each factor. Significant effects (p-value <0.05) are given in bold.

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differences were observed in bacterial and fungal abundances between the start of the experiment and after the one-year field incubation. We observed however, a shift in bacterial communities in permafrost soils towards that of active layer soils. This suggests that warmer permafrost will resemble the community structure in active layer soils, a finding which was also observed for bacterial communities in a long-term permafrost thawing experiment (Monteux et al., 2018). Similarly, other research on permafrost soils has shown that bacterial communities change significantly with increased temperatures, while fungi are not

affected due to the larger variability in the structure of fungal communities (Feng et al., 2020), a finding that we also observed in this study.

Changes in community structure in soil transplant experiments have been reported to be particularly pronounced when soil from lower temperatures was transferred to locations with higher temperatures (Zumsteg et al., 2013a). However, our findings should be interpreted cautiously, as we cannot exclude that time (one-year incubation) contributed to the shift in community structure. Control samples taken from the permafrost zone may lead to further insights.

Fig. 4. Differential abundant bacterial taxa in response to litter amendment in (A–D) lab-incubated soils and (E–H) field-incubated soils. The left-hand plots depict (A, E) phyla and (C, G) genera in active layer soils, whereas the right-hand plots depict (B, F) phyla and (D, H) genera in permafrost soils. Only significant log2 fold changes between litter-amended and control (unamended) samples of the ten most abundant phyla and genera are shown, based on a significance level of p <0.01 after false discovery rate correction. Error bars represent standard error. The relative abundance values for each phylum and genus are calculated for all samples within either active layer or permafrost soil. The genus Allorhizobium-Neorhizobium-Pararhizobium-Rhizobium was shortened to Rhizobium for display- ing purposes.

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4.3. Fast-growing taxa benefit from litter addition

The shifts in the structure of microbial communities in response to litter amendment were driven by various bacterial and fungal taxa. In general, it has been proposed that substrate addition to soils will initially benefit fast-growing copiotrophic taxa that take advantage of readily degradable C compounds. Slow-growing oligotrophic taxa subsequently develop on the remaining recalcitrant C compounds (e.g. lignin), due to their greater catabolic capacities compared with the taxa stimulated earlier (Fierer et al., 2007; Koch, 2001; Lauro et al., 2009; Pascault et al., 2013; Rime et al., 2016b). Applying this concept to microbial phyloge- netic groups, however, is a subject of debate. As physiological traits of microbial groups are context dependent, the same phylum can be dominated by copiotrophic taxa in one environment and oligotrophic taxa in another (Hartmann et al., 2017; Morrissey et al., 2016; Senechkin et al., 2010). Thus, to avoid misinterpretations at the phylum level, it is important to analyse taxa at lower taxonomic levels.

In the present study, we observed that copiotrophic bacterial phyla were particularly stimulated by the addition of litter to active layer and permafrost soils in both the lab and field experiment. This included the highly abundant Proteobacteria and the slightly less abundant Actino- bacteria and Bacteroidetes. Previous research on microbial community responses to C substrates with varying recalcitrance in soils from various

ecosystems, such as glacier forefields (Rime et al., 2016b), Arctic tundra (Koyama et al., 2014), grasslands (Ali et al., 2018; Leff et al., 2015) and forests (Eilers et al., 2010), likewise revealed increases in copiotrophic taxa. Overall, active layer and permafrost soils exhibited similar responses of microbial taxa. The global increase in Proteobacteria, in particular in lab-incubated soils, was mainly due to the stimulation of the genus Pseudomonas. Pseudomonas appears to be abundant in Arctic soils, as a large number of Pseudomonas strains have been isolated from the Canadian High Arctic (Marcolefas et al., 2019). Furthermore, Pseu- domonas has a putative role as a copiotroph (Bittman et al., 2005; Tardy et al., 2015) and has been shown to respond strongly to glucose addition to soils (Jenkins et al., 2010). A question that may arise, however, is if Pseudomonas was already present in large amounts in the litter added to the soils. Yet, preliminary experiments we conducted with sterilised litter showed that the abundance of Pseudomonas still increased in the examined active layer and permafrost soils (unpublished data). In field-incubated soils, Pseudomonas also increased considerably but was overall less abundant, possibly as a result of environmental influences.

Massilia, another copiotrophic group belonging to Proteobacteria (Ofek et al., 2012) and commonly found in terrestrial cryoenvironments (Lul´akov´a et al., 2019; Perez-Mon et al., 2020; Rime et al., 2016b), also increased in abundance with litter input. This genus utilises multiple C sources and is known to degrade a wide variety of organic compounds, Fig. 5.Differential abundant fungal taxa in response to litter amendment in (A–D) lab-incubated soils and (E–H) field-incubated soils. The left-hand plots depict (A, E) phyla and (C, G) genera in active layer soils, whereas the right-hand plots depict (B, F) phyla and (D, H) genera in permafrost soils. Only significant log2 fold changes between litter-amended and control (unamended) samples of the ten most abundant phyla and genera are shown, based on a significance level of p <0.01 after false discovery rate correction. Error bars represent standard error. The relative abundance values for each phylum and genus are calculated for all samples within either active layer or permafrost soil.