Development of archaeal communities in biological soil crusts along a revegetation chronosequence in the Tengger Desert, north central China

Soil & Tillage Research 196 (2020) 104443

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Development of archaeal communities in biological soil crusts along a revegetation chronosequence in the Tengger Desert, north central China

T

Lina Zhaoa,b,d, Yubing Liua,b,⁎, Shiwei Yuanc,d, Zhaohuan Lia,d, Jingyao Suna,d, Xinrong Lia,b,⁎ a

Shapotou Desert Research & Experiment Station, Northwest Institute of Eco-Environment and Resources, Chinese Academy of Sciences, Lanzhou, 730000, China Key Laboratory of Stress Physiology and Ecology in Cold and Arid Regions of Gansu Province, Northwest Institute of Eco-Environment and Resources, Chinese Academy of Sciences, Lanzhou, 730000, China c Key Laboratory of Remote Sensing of Gansu Province, Northwest Institute of Eco-Environment and Resources, Chinese Academy of Sciences, Lanzhou, 730000, China d University of Chinese Academy of Sciences, Beijing, 100049, China b

A R T I C LE I N FO

A B S T R A C T

Keywords: Arid and semi-arid areas Archaeal diversity Archaeal function Successional stage of BSCs

Archaea are major contributors to biogeochemical cycles and energy metabolism among soil microorganisms under extremely acidic and high-temperature conditions, however, the biodiversity and ecological function of archaea in biological soil crusts (BSCs) of desert ecosystems are not fully understood. Here, we used Illumina MiSeq sequencing and microbial functional gene array (GeoChip 5.0) to test the following hypotheses: (1) the composition and function related to biogeochemical cycles of the archaeal community would change significantly in the development process of BSCs; and (2) the key factors driving these changes may be soil biogeochemical properties. The results showed that the diversity, abundance, and functional potential of the archaeal community showed their highest levels in 5-year-old BSCs. The dominant phyla were Thaumarchaeota, Euryarchaeota, and an unclassified phylum in the archaeal community during BSC succession. The functional genes of the archaeal community were mainly involved in carbon (C) and nitrogen (N) cycles, and the functions of the three dominant phyla were complementary in these cycles. Moreover, redundancy analysis showed that soil biogeochemical properties were negatively related to the composition and function of the archaeal community during BSC succession, and the soil C:N ratio might be the major limiting factor. These results provided evidences for our hypotheses and revealed that the archaeal community played an important ecological role in the early development stage of BSCs, and might be pioneer species of soil microbial communities during BSC succession.

1. Introduction Biological soil crusts (BSCs) are assemblages of microscopic (cyanobacteria, algae, fungi, and bacteria) and macroscopic (lichens, mosses, and micro arthropods) organisms combined with surface soil particles, and make up to 40% of the living cover of desert ecosystems and can even exceed 70% in some unique soil habitats (Belnap and Lange, 2003; Eldridge and Greene, 1994; Weber et al., 2016). Being considered as the desert ecosystem engineers, BSCs have many important ecological functions in arid and semi-arid areas, such as fixing dune surfaces (Li et al., 2010a, b, 2012), reducing soil erosion (Warren and Eldridge, 2001), enhancing soil fertility (Belnap and Gardner, 1993; Li, 2005), influencing soil nutrient cycling (Belnap and Lange, 2003; Li et al., 2002), regulating soil moisture (Belnap and Lange, 2003; George et al., 2003), affecting the germination and establishment of vascular plants (Godínez-Alvarez et al., 2012; Li et al., 2005), and ⁎

establishing a favorable living environment for other organisms (Neher et al., 2009). The formation of BSCs is generally regarded as a process in which the dominance of pioneer cyanobacteria is subsequently replaced by lichens and mosses with corresponding improvement in soil environments (Neher et al., 2009). As environmental changes can result in rapid variation in the microbial communities in BSCs (Cong et al., 2015; Han et al., 2007; Harris, 2009; López-Lozano et al., 2013), microbial communities are often used as sensitive bio-indicators for predicting the degradation and restoration of BSCs (Emmerling et al., 2001; Fernandes et al., 2005; Liu et al., 2017a, b). With the recent development of high-throughput sequencing technology, research on the composition and function of bacterial and fungal communities in BSCs has become in-depth and comprehensive (Bates and Garcia-Pichel, 2009; Bates et al., 2010a; Green et al., 2008; Gundlapally and Garcia-Pichel, 2006; Maier et al., 2014; Wang et al., 2015). It has been reported that the dominant bacteria in BSCs are

Corresponding author at: Donggang West Road 320, Lanzhou, 730000, China. E-mail addresses: [email protected] (Y. Liu), [email protected] (X. Li).

https://doi.org/10.1016/j.still.2019.104443 Received 13 July 2018; Received in revised form 22 August 2019; Accepted 5 October 2019 0167-1987/ © 2019 Elsevier B.V. All rights reserved.

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following decades. The appearance of BSCs following stabilization of sand dunes is considered an important indicator of desertification reversion. Since then, the fenced revegetated areas from different establishment years have provided ideal experimental sites by a “space-fortime” approach for exploring related ecological succession and environmental changes following revegetation, especially in the formation and development of BSCs.

Bacilli (mainly Bacillus, Enterococcus, and Lactococcus), Alphaproteobacteria, and Sphingobacteria (mainly Chitinophagaceae and Cytophagaceae), which account for the highest proportion of microorganisms in BSCs from the Tengger Desert (Liu et al., 2017a, b), Tabernas Desert (Maier et al., 2014), and Sandia Mountains (Moquin et al., 2012), and Ascomycota were noted to be the dominant fungi of BSCs in desert ecosystems (Abed et al., 2013; Bates et al., 2010a, b). Bacterial and fungal communities in BSCs contribute to improving soil fertility as decomposers, providing nutrients to plants and other organisms, and promoting carbon (C), nitrogen (N), sulfur (S), and phosphorus (P) cycling in desert ecosystems (Abed et al., 2013; Bates and Garcia-Pichel, 2009; Bates et al., 2010a, 2012; Grishkan and Kidron, 2013; Grishkan et al., 2015; Maier et al., 2014; Moquin et al., 2012; Zhang et al., 2016). Moreover, fungal communities play critical roles in maintaining the BSC structure through their mycelia (Barger et al., 2006; Meadows et al., 1994). Therefore, bacterial and fungal communities have important ecological functions and are potential indicators of the BSC succession process. However, few studies have been conducted on the biodiversity and function of archaeal communities in BSCs. Soule et al. (2009) investigated the light-colored, dark-colored, lichen-type, and moss-type BSCs in arid land of North America using denaturing gradient gel electrophoresis technology, and found that all detected phylotypes belonged to Crenarchaeota and the archaeal community composition was constant across different BSC types and climatic regions. However, Du et al. (2014) reported that the structure of the archaeal community in light-colored BSCs significantly varied with seasonal changes, and the major archaeal groups belonged to Thaumarchaeota (a novel archaeal phylum) in the Hunsandake Desert of China. Due to these inconsistent results and lack of other supporting evidence, it is necessary to study the changes of biodiversity and function in the archaeal community during BSC succession. The archaeal community can promote biogeochemical cycles and energy metabolism in the ocean and other ecosystems (Brochier-Armanet et al., 2008; Dumont et al., 2011; Guy and Ettema, 2011; Offre et al., 2013; Williams et al., 2013); however, research on the archaeal functions of BSCs in desert ecosystems is in the initial stage, and only the functional group of ammoniaoxidizing archaea (AOA) in the N cycle has been studied (Marusenko et al., 2013). In the present study, archaeal community composition and ecological function related to biogeochemical cycles (C, N, S, and P cycles) in BSCs were studied along the chronosequence of over 50-year-old revegetation in the Shapotou revegetated area using Illumina MiSeq sequencing and microbial functional gene array (GeoChip 5.0). Our aim is to test the following hypotheses: (1) the composition and function related to biogeochemical cycles of the archaeal community will change significantly during BSC succession; and (2) the key factors driving these changes are soil biogeochemical properties.

2.2. Sample collection and preparation The BSC and underlying soil (0–5 cm) samples were collected in early November 2015 from the Shapotou revegetated area: established in 1964 (51 years of revegetation, 51YR), 1981 (34 years of revegetation, 34YR), 1987 (28 years of revegetation, 28YR), 2000 (15 years of revegetation, 15YR), and 2010 (5 years of revegetation, 5YR). Furthermore, as representation of initial status, soil samples from nearby mobile sand (MS) were used. In each revegetated area, five sample cores (3.5 cm diameter) from a diagonal intersection and four vertices of each plot were sampled. To compensate for spatial heterogeneity, each sample was collected from six individual plots (20 m × 20 m quadrat separated by at least 20 m). Thus, 30 samples (5 cores × 6 plots) were obtained and immediately mixed together to form one composited sample. Triplicate composite samples for each revegetation area were collected using the same method. Composited samples were stored in an ice box and transported to the laboratory, then immediately homogenized and sieved through a 1-mm screen to remove stones and plant roots. Approximately 500 g of each composited BSC sample was stored at −80 °C for nucleic acid-based molecular analysis. Approximately 1000 g of each composited soil sample was airdried for soil biogeochemical properties. 2.3. Soil characteristics analysis Soil pH was determined using a 1:5 soil–water mixture. Silt and clay contents were determined by the pipette method after salts removal with sodium acetate, organic matter destruction with H2O2 and chemical dispersion using sodium hexametaphosphate (Gee and Or, 2002). Organic C (OC) was determined by the dichromate oxidation method (Nelson and Sommers, 1982). Total N (TN) was measured using dry combustion in a Vario Macro Cube Elementar (Elementar Analysensysteme, Hanau, Germany). The soil C:N ratio was calculated using OC and TN (Aitkenhead and McDowell, 2000). Soil total P (TP) was determined by perchloric acid digestion according to Sommers and Nelson (1972). Urease (UE) activity was measured by the amount of NNH4+ released from urea hydrolysis at 578 nm (Yang et al., 2007). Dehydrogenase (DHA) activity was measured by the reduction of 2,3,5triphenyl tetrazolium chloride to triphenyl formazan (TPF) at 485 nm, following the method described by Casida et al. (1964). Invertase (IT) activity was measured by the amount of glucose released from sucrose hydrolysis at 508 nm (Jin et al., 2009). Catalase (CAT) activity was measured by following the reduction of hydrogen peroxide (H2O2) at 240 nm (Beers and Sizer, 1952). One unit of enzyme activity of UE, DHA, IT, and CAT was defined as the amount of enzyme that decomposed 1 μg of N-NH4+, 1 μg of TPF, 1 μg of glucose, and 1 μmol H2O2 per gram dry soil sample per hour, respectively. All soil physicochemical characteristics and enzyme activities are shown in Table S1.

2. Materials and methods 2.1. Site description The study site is located in the Shapotou revegetated area (37°32′N, 105°02′E) on the southeast edge of the Tengger Desert in north central China. The natural landscape of this area comprises large, dense, and reticulate dune chains. The annual average temperature is 9.6 °C, average annual rainfall is ˜180 mm, with most precipitation occurring during July–September, and annual average pan evaporation is ˜2900 mm (Li et al., 2010a; Liu et al., 2006). The soil is wind-borne sand (according to Eutric Arenosols in the World Reference Base, FAOISRIC-ISSS, 1998), saline and alkaline, with pH 7.5–11.2 (He et al., 2016). The dominant plants are xerophytic shrubs including Artemisia ordosica Krasch and Caragana korshinskii Kom (Wang et al., 2005; Zhang et al., 2013). The non-irrigated revegetation protection system in this area was mainly established in 1956, and expanded several times in the

2.4. Illumina MiSeq sequencing and data processing The total DNA was extracted from 1 g of the BSC sample using an E.Z.N.A. Soil DNA Kit (Omega Bio-Tek, Norcross, GA, USA), according to the manufacturer’s instructions. To avoid low DNA concentration, the DNA of a sample was extracted thrice and pooled. The obtained DNA was checked for purity and concentration using NanoDrop 2000c (Thermo Fisher Scientific, Waltham, MA, USA). The archaeal 16S rRNA gene was amplified by PCR using primer set 524F10-extF 2

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(5′-TGYCAGCCGCCGCGGTAA-3′) and Arch958R-modR (5′-YCCGGCGTTGAVTCCAATT-3′) (Pires et al., 2012). PCR was performed using a 20-μL reaction mixture containing 5 μL of 2 × Taq PCR Master Mix (Sangon Biotech Shanghai Co., Ltd., Shanghai, China), 1 μL of each primer (10 μM), 1 μL of total DNA (10 ng), and sterilized ultrapure water. The PCR conditions were as follows: 94 °C for 4 min, followed by 35 cycles of 94 °C for 30 s, 50 °C for 30 s, and 72 °C for 30 s, and a final extension at 72 °C for 10 min. The amplicons were separated by 1% agarose gel electrophoresis, purified using SanPrep Column DNA Gel Extraction Kit (Sangon Biotech Shanghai Co. Ltd.) and quantified with QuantiFluor™-ST (Promega Corporation, Madison, WI, USA). The purified amplicons were sequenced using Illumina MiSeq platform (Majorbio Bio-Pharm Technology Co. Ltd., Shanghai, China; http://www. majorbio.com/). All sequence data were submitted to the NCBI Sequence Read Archive (SRA) under accession number SRP118745. The raw FASTQ files were demultiplexed and filtered using QIIME (version 1.17) with the following criteria (Liu et al., 2017a): (i) The 300-bp reads were truncated at any site receiving an average quality score < 20 over a 50-bp sliding window, discarding the truncated reads shorter than 50 bp; (ii) exact barcode matching, two nucleotide mismatch in primer matching, reads containing ambiguous characters were removed, and (iii) only sequences that overlapped > 10 bp were assembled according to their overlap sequence. Reads that could not be assembled were discarded. Subsequently, the sequences were clustered to operational taxonomic units (OTUs) at 97% sequence identity using UPARSE (version 7.1 http://drive5.com/uparse/). Alpha diversity analysis was used to reflect the diversity and richness of the archaeal community (Table S2). The overall differences in archaeal community composition among BSC samples were evaluated by principal co-ordinate analysis (PCoA) and permutational multivariate analysis of variance (PERMANOVA) performed with R (http://www.r-project.org/) based on the Bray–Curtis distance.

the detection signal was obtained at 72 °C for 10 s and analyzed. All the experiments were performed on Stratagene Mx3000 P system (Stratagene, Agilent Technologies Inc.). PCR products were gel-purified and cloned into pUCm-T vector (Sangon Biotech Shanghai Co. Ltd.), and the resulting ligation products were transformed into Escherichia coli DH5α competent cells (Sangon Biotech Shanghai Co. Ltd.). The standard curve was constructed from serial dilution of linearized plasmids containing the target gene fragment of interest that was cloned from the amplified pure culture DNA. Amplification efficiencies of 82.4% were obtained with r-squared values of 0.988. The melting curve was obtained to confirm that the amplified products were of appropriate size as well as to ensure the absence of primer dimer. The gene copy numbers of the target samples for each reaction were calculated from the standard curves. qPCR was replicated at least six times for each sample.

2.5. GeoChip 5.0 analysis and data processing

3. Results

GeoChip 5.0 is a high-throughput, microarray-based metagenomic tool to analyze the functional composition and structure of microbial communities. The purified DNA (500 ng) of each BSC sample was labeled with the fluorescent dye Cy 3 (GE Healthcare, CA, USA) as described previously (Tu et al., 2014), and hybridized with GeoChip 5.0 at 67 °C in an Agilent hybridization oven (Agilent Technologies Inc., Santa Clara, CA, USA) for 24 h. After hybridization, the slides were washed using Agilent Wash Buffers at room temperature to remove unbound DNA. The arrays were then scanned with NimbleGen MS200 Microarray Scanner (Roche NimbleGen Inc., Madison, WI, USA). To avoid false positives, the spots with weak signals and signal-to-noise ratio of less than 2.0 were removed before statistical analysis. Data normalization was based on logarithm transform, calculating signal intensity in each sample, then scaled up by average (Mean Ratio). Genes were considered positive if its corresponding probes were positive in at least 2/3 of replicate sets. The sum of normalized signal intensity for gene subcategory was used to represent the total normalized signal intensity, and the normalized average signal intensity was obtained from the triplicate (Bai et al., 2013).

3.1. Archaeal richness and abundance in BSCs of different stages

2.7. Statistical analysis One-way analysis of variance and Tukey’s honestly significant difference analysis were carried out using SPSS 16.0 software (SPSS Inc., Chicago, IL, USA) to determine the differences of the Chao Indies, the relative proportion of the classifications, and the intensity of function of archaeal communities in BSCs at different stages. The homogenization of microbial Chao index was carried out according to Min-Max Normalization (Jain and Bhandare, 2011). Redundancy analysis (RDA) was used to test the effects of soil biogeochemical characteristics on the compositional and functional structures of the archaeal community, the statistical significance of the RDA was assessed by the Monte Carlo Permutation test with 499 permutations, and the analytical program CANOCO 4.5.1 was used (Ter Braak, 1986). Figures were generated with Origin 8.0 (Origin Lab Corporation, Northampton, MA, USA).

A total of 666,327 archaeal trimmed sequences were retrieved with average lengths of 447 bp, and 56–85 OTUs of archaea were generated using a threshold of 97% (Table S2). The PCoA and PERMANOVA tests showed that the six different stages of BSCs were divided into two groups: MS and 5YR belonging to group 1; and 15YR–51YR belonging to group 2 (R2 = 0.43654, F = 12.40, p < 0.01; Fig. 1). The archaeal community richness (Fig. 2) and 16S rRNA gene copies (Table 1) significantly increased in the first 5 years of BSC development (p < 0.05) and then decreased with succession.

2.6. Quantitative PCR Quantitative PCR (qPCR) was performed to determine the abundance of archaeal 16S rRNA gene using the primers Arch349 F (5′-GYGCASCAGKCGMGAAW-3′) and Arch806R (5′-GGACTACVSGGGTATCTAAT-3′) (Takai and Horikoshi, 2000). The 20-μL qPCR reaction mix contained 10 μL of 2 × SYBR Green PCR Master Mix (Takara Bio, Shiga, Japan), 0.4 μL of each primer (10 μM), 1 μL of total DNA template (diluted to 1 ng μL−1), and molecular-grade H2O. The reaction was conducted as follows: 50 °C for 2 min and 94 °C for 10 min, followed by 45 cycles of 94 °C for 1 min, 54 °C for 1 min, and 72 °C for 30 s;

Fig. 1. PCoA of the archaeal communities in BSCs of six different stages at the OTU level based on 97% similarity. MS, 5YR, 15YR, 28YR, 34YR, and 51YR represent mobile sand, 5-, 15-, 28-, 34-, and 51-year-old BSCs, respectively. 3

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Fig. 2. Chao index of archaeal communities at the OTU level in BSCs of six different stages. Different lowercase letters denote significant differences between samples (p < 0.05). MS, 5YR, 15YR, 28YR, 34YR, and 51YR represent mobile sand, 5-, 15-, 28-, 34-, and 51-year-old BSCs, respectively.

Fig. 3. Archaeal community composition in BSCs of six different stages at the phylum level. Different lowercase letters represent a significant difference between samples (p < 0.05). MS, 5YR, 15YR, 28YR, 34YR, and 51YR represent mobile sand, 5-, 15-, 28-, 34-, and 51-year-old BSCs, respectively.

3.2. Archaeal community composition in BSCs of different stages There were three dominant phyla detected in BSCs (Fig. 3). Thaumarchaeota was the dominant phylum across all samples, and its relative abundance decreased during BSC succession (from 72.65% in MS to 57.39% in 51YR). Euryarchaeota also showed similar changes (maximum 19.20% in MS and minimum 2.94% in 51YR) to Thaumarchaeota. In contrast, an unclassified phylum increased from 7.93% in MS to 39.67% in 51YR. At the genus level (Fig. 4), the dominant genera norank_Soil_Crenarchaeotic_Group_SCG (SCG) and norank_Marine_Group_II (MG II) decreased; however, an unclassified genus and Candidatus Nitrososphaera (Ca. N.) increased in the development process of BSCs.

and denitrification were mainly in Euryarchaeota, genes involved in assimilatory N reduction were focused in the unclassified phylum, and genes involved in nitrification mainly occurred in Thaumarchaeota and the unclassified phylum.

3.4. Relationships between archaeal community structure and soil biogeochemical properties The RDA showed that the cumulative variation in the compositional and functional structures of the archaeal community explained by the first four axes (as constrained by the measured soil environmental variables) was 92.5% and 97.0%, respectively, with the first axis explaining 59.5% and 76.1%, and with the corresponding second axis explaining 18.1% and 12.0% (Fig. 6; Table S3). According to a Monte Carlo permutation test, the first canonical axis was highly significant in both the compositional (F = 5.072, p < 0.01) and functional (F = 10.742, p < 0.01) structures (Table S3). Archaeal compositional structure along the soil texture (Silt + Clay), nutrient contents (C:N ratio and TP), and enzyme activities (UE, DHA, CAT, and IT) gradient were reflected on axis 1, where the highest correlated variable was Silt + Clay (–0.8996) and C:N ratio (–0.8827) (Fig. 6a; Table S4). For archaeal functional structure, the C:N ratio (–0.8493) and IT (–0.8593) were most strongly related to the archaeal community with the highest correlated variables in axis 1, showing that these two characteristics were the prime determinants of archaeal functional structure (Fig. 6b; Table S4). Because the highest levels of richness and function of the archaeal community were in the early successional stage of BSCs, there was a close negative correlation between soil factors and archaeal compositional and functional structures, except for soil pH.

3.3. Archaeal functions in biogeochemical cycle in BSCs of different stages The normalized average signal intensities of C-cycling genes were much higher than those of other biogeochemical cycles (Fig. 5a), and gene subcategories were most abundant in N-cycling genes, with the normalized average signal intensities of assimilatory N reduction and nitrification higher than others. Compared with C and N cycles, there were fewer genes involved in S and P cycling in subcategories and they had lower normalized average signal intensity. The normalized average signal intensity of genes involved in C fixation, methane, ammonification, assimilatory N reduction, N fixation, sulfite reduction, polyphosphate synthesis, and polyphosphate degradation was significantly higher in 5YR than for 15YR–51YR (p < 0.05). Moreover, the functional gene structures in different dominant phyla of the archaeal community significantly differed (Fig. 5b). Genes of Thaumarchaeota were only involved in C-, N-, and S-cycling, but genes of Euryarchaeota were found in all biogeochemical cycles. Genes involved in C fixation were mainly in Thaumarchaeota, and those involved in C degradation and methane were focused in Euryarchaeota. In N-cycling, all genes involved in dissimilatory N reduction and N fixation belonged to Euryarchaeota, genes involved in ammonification

Table 1 Absolute abundances of archaea (16S rRNA gene copies per gram of BSCs) in BSCs quantified by qPCR. MS, 5YR, 15YR, 28YR, 34YR, and 51YR represent mobile sand, 5-, 15-, 28-, 34-, and 51-year-old BSCs, respectively. Dominant Archaeal abundance

MS

5YR 4

5.83 × 10 ± 1.36 × 103e

15YR 8

4.49 × 10 ± 3.82 × 107a

28YR 8

3.13 × 10 ± 4.04 × 106b

34YR 8

2.11 × 10 ± 1.97 × 107c

51YR 8

1.20 × 10 ± 3.36 × 107d

3.16 × 107 ± 2.24 × 106e

Data were expressed as means ± standard deviation (n ≥ 6); different lowercase letters represent a significant difference between samples (p < 0.05). 4

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Fig. 4. Archaeal community composition in BSCs of six different stages at the genus level. Different lowercase letters represent a significant difference between samples (p < 0.05). MS, 5YR, 15YR, 28YR, 34YR, and 51YR represent mobile sand, 5-, 15-, 28-, 34-, and 51-year-old BSCs, respectively.

4. Discussion

related to biogeochemical cycles during BSC succession in the revegetation of the Tengger Desert, to test our hypotheses that archaeal composition and function shifted due to the improvement of soil biogeochemical properties during BSC succession. The BSCs of different successional stages were divided into two groups according to the archaeal community structure. Group 1 was dominated by the physical crusts which can be identified as the early successional stage of BSC development, and group 2 was dominated by

In temperate desert ecosystems, bacteria and fungi play important roles in maintaining the structural and functional stability of BSCs (Belnap and Lange, 2003; Bowker, 2007; Yeager et al., 2004). However, there has been very limited study of the composition and function of BSC archaeal community in desert ecosystems. The present study provided an overview of archaeal community composition and function

Fig. 5. Changes in functional genes involved in biogeochemical cycles (carbon, nitrogen, sulfur, and phosphorus cycles) of archaeal communities in BSCs of six different stages. The normalized signal intensity in overall gene subcategories (a) and in the dominant phyla (b) were compared among BSCs. Different lowercase letters represent a significant difference between samples (p < 0.05). 5YR, 15YR, 28YR, 34YR, and 51YR represent 5-, 15-, 28-, 34-, and 51-year-old BSCs, respectively.

5

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Fig. 6. RDA of soil biochemical characteristics in relation to archaeal compositional (a) and functional (b) structures. Each circle in the RDA represents the archaeal community of each sample. The arrow represents the direction and magnitude of the soil factors associated with the archaeal community structure, and the length of the arrow denotes the intensity of the correlation. MS, 5YR, 15YR, 28YR, 34YR, and 51YR represent mobile sand, 5-, 15-, 28-, 34-, and 51-year-old BSCs, respectively.

2009; Pester et al., 2012; Spang et al., 2010). The Euryarchaeota, with the dominant genus of MG II, are mainly involved in C degradation and methane metabolism. Genes of carbohydrate/lipid transport and metabolism are rich in MG II and play important roles in energy metabolism (Berg et al., 2010; Deschamps et al., 2014). In addition, anaerobic archaea belonging to Euryarchaeota are capable of methanogenesis and anaerobic methane oxidation, the two key steps in the C cycle (Offre et al., 2013). Although the relative abundance of Thaumarchaeota and Euryarchaeota decreased during BSC succession, the normalized average signal intensity of genes involved in the C and N cycles was highest for 5YR. Furthermore, an unclassified phylum increased significantly during the BSC succession, and its genes were mainly involved in nitrification and assimilatory N reduction. The functions of the three dominant phyla were redundant, but their major functions were complementary and played an important role in the C and N cycles, especially during the BSC early successional stage. These results further revealed the function of pioneer species of archaea during BSC development. However, compositional and functional structures of the microbial community in a specific condition are closely related to their habitat environmental factors (Angel et al., 2010; Fierer and Jackson, 2006). Development of bacterial and fungal communities of BSCs were positively correlated with soil texture and nutrient concentration during BSC succession (Abed et al., 2012, 2013; Bates et al., 2010a; Li et al., 2010a; Liu et al., 2017a, b); however, this differed to results for the archaeal community which were not consistent with the development of soil characteristics. The main reason for this was that the negative feedback of the archaeal community to soil C or N inputs (Bates et al., 2011; Nielsen et al., 2010), and their diversity and function, decreased with higher soil C:N ratio and available element concentration in the later successional stage of BSCs. Soil C:N ratio was the main determinant factor for both archaeal community composition and function. Our results further confirmed that archaea inhabited a far more restricted ecological niche in soils than bacteria, perhaps because of competitive interactions with nitrifying bacteria (Bates et al., 2011). The activities of IT and UE in soil involved in C and N metabolism increased with the higher C and N concentrations. Accordingly, CAT and DHA activities, as indicators of soil fertility and soil microbial activity, respectively (Kaczynski et al., 2016; Trasar-Cepeda et al., 2007), were negatively correlated with the archaeal community.

biological crusts identified as the later successional stage (Li et al., 2007a, 2010b; Liu et al., 2017a). Unexpectedly, the richness and diversity of the archaeal community did not increase similarly to bacterial or fungal communities during BSC succession (Fig. S1, Table S2). Our previous studies showed that bacterial diversity increases rapidly in the first 15 years and then remains relatively stable, whereas it may take decades or even hundreds of years to achieve such a stable state in fungal diversity of BSCs in revegetation areas (Liu et al., 2017a,b). These results showed that archaeal, bacterial, and fungal communities reached their highest richness level successively during BSC development, and the cause of this difference may be related to the adaptability of different microbial communities to soil environments (Fries et al., 1997; Ishii et al., 2000; Nemergut et al., 2007). Archaea can live in extreme environments and have strong adaptation to the poor soil texture and nutrient environment in the early successional stage of BSCs, and can effectively utilize the limited nutrients and low water content for growth and reproduction in temperate desert ecosystems (Chapin et al., 2002; Grime, 1979; Konings et al., 2002; MartensHabbena et al., 2009). However, in the later successional stage of BSCs, the richness (Fig. S1), diversity (Table S2), and abundance (Table 1) of the archaeal community decreased and their ecological functions may have been replaced by bacteria, fungi, or cryptogamic species to further promote BSC succession (Bates et al., 2010a; Belnap, 2002; Belnap and Lange, 2003; Li et al., 2007b, 2010a; Liu et al., 2017a, b). The possible reason was competition between different microbial communities due to limited internal resources (Feng et al., 2017; Rothrock and GarciaPichel, 2005), and the archaeal community may be at a disadvantage in competition with bacteria or fungi (Bates et al., 2011). Thus, the development of the archaeal community was in the early successional stage of BSCs, meaning that archaea might act as pioneer species in soil microbial communities during BSC development. Among the archaeal community, Thaumarchaeota, Euryarchaeota, and an unclassified phylum were the dominant phyla during BSC development. This result was not consistent with research on different types of BSCs in arid land of North America (Soule et al., 2009) and light-colored BSCs in the Hunsandake Desert (Du et al., 2014), revealing that the archaeal compositional structure of BSCs differed according to soil environments. We found that Thaumarchaeota, which contains two dominant genera of SCG and Ca. N., played a major role in C fixation and nitrification. The SCG and Ca. N. belong to the AOA group (Brochier-Armanet et al., 2008; Buckles et al., 2013; Jiang et al., 2015; Prosser and Nicol, 2008; Zhalnina et al., 2014), and can use carbon dioxide as a C source for autotrophic growth (Zhang et al., 2012). Additionally, AOAs oxidize ammonia to nitrite with further oxidation to nitrate, which is generally considered to be the rate-limiting step of the nitrification process in the N cycle (Gubry-Rangin et al., 2010; Jung et al., 2011; Leininger et al., 2006; Martens-Habbena et al.,

5. Conclusions The results provided strong evidence for our hypotheses that the composition and function related to biogeochemical cycles of the archaeal community changed significantly during BSC succession, and that the key factors driving these changes were soil biogeochemical 6

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properties, especially the soil C:N ratio. Archaea might serve as a pioneer species of microbial communities in BSC development and played an important role in biogeochemical cycles in the early successional stage of BSCs, as their diversity, abundance, and functional potential showed their highest levels at that period. Moreover, the dominant phyla Thaumarchaeota, Euryarchaeota and an unclassified phylum were complementary in their ecological functions. This study supplemented knowledge gaps of archaeal diversity and function in BSCs of desert ecosystems.

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