Methanogens are strictly anaerobic archaea capable of energy conservation by methane production, yet their presence in oxic and arid environments challenges existing paradigms. In this study, we enriched and genomically characterized seven methanogenic cultures from desert biocrusts, affiliated with the genera Methanobacterium, Methanosarcina, and Methanocella. Six of these new enrichment cultures represent new species. Nonetheless, phylogenomic analyses revealed close genetic relationships with organisms from anoxic environments, indicating the absence of an evolutionary distinction. Comparative genomics exposed diverse though non-unique repertories of antioxidant (e.g. catalase, superoxide dismutase and desulfoferrodoxin), and desiccation-resistance genes (including genes for maintaining osmotic pressure and repair of cell wall and membrane), with Methanobacterium spp possessing the lowest gene abundance and diversity for oxygen and desiccation tolerance. Nevertheless, the occurrence of a Class I methanogen such as Methanobacterium in arid soils challenges the notion that members of this class are less oxygen tolerant than Class II. Pangenome analysis further uncovered unique genes enriched in membrane-associated functions and potentially non-functional stress-related genes. Via a global metagenomic survey we find that methanogens are underdetected in dryland soils, likely due to sequencing depth limitations. Our findings highlight previously overlooked methanogen diversity and ecological plasticity in oxic and desiccated habitats, and emphasize the need for further studies to elucidate their survival strategies.

Efficient tertiary effluent desalination is hindered by membrane biofouling, leading to plant downtime, shortened membrane lifespan, heightened energy consumption, and reduced permeability. While the composition of these biofouling layers is studied, little is known about bacterial succession and function. To address this, we used a benchtop reverse osmosis (RO) system to process synthetic tertiary effluent with diluted sludge. System flux monitoring tracked biofouling layer development, and RO biofilm samples were collected at early and mature stages (48, 72, 120 h). Scanning electron microscopy visualized the samples, and their 16S rRNA genes were sequenced. DNA-stable isotope probing with labeled glucose identified growing taxa in early and late biofouling stages (48, 120 h). Over time, biofouling layer biomass increased, with shifts in bacterial diversity and composition. Proteobacteria, notably oligotrophic genera, dominated early stages along with Bacteriodota, while Actinobacteria increased in mature biofilms. Functional changes included a shift from biosynthesis of cellular components like DNA, peptidoglycan, membrane lipids, and antimicrobials, to production of extracellular polysaccharides and reactive oxygen scavenging agents. In conclusion, our research enhances the understanding of biofouling dynamics within tertiary effluent desalination processes, providing insights that could improve biofouling management strategies in RO desalination systems, potentially applicable to larger-scale operations.

Biological soil crusts (biocrusts) are key contributors to desert ecosystem functions, therefore, biocrust restoration following mechanical disturbances is imperative. In the Negev Desert hyperarid regions, phosphate mining has been practiced for over 60 years, destroying soil habitats and fragmenting the landscape. In this study, we selected one mining site restored in 2007, and we used DNA stable isotope probing (DNA-SIP) to identify which bacteria grow in post-mining and adjacent natural biocrusts. Since biocrust communities activate only after wetting, we incubated the biocrusts with (H2O)-O-18 for 96 h under ambient conditions. We then evaluated the physicochemical soil properties, chlorophyll a concentrations, activation, and functional potential of the biocrusts. The DNA-SIP assay revealed low bacterial activity in both plot types and no significant differences in the proliferated communities' composition when comparing post-mining and natural biocrusts. We further found no significant differences in the microbial functional potential, photosynthetic rates, or soil properties. Our results suggest that growth of hyperarid biocrust bacteria after wetting is minimal. We hypothesize that due to the harsh climatic conditions, during wetting, bacteria devote their meager resources to prepare for the coming drought, by focusing on damage repair and organic compound synthesis and storage rather than on growth. These low growth rates contribute to the sluggish recovery of desert biocrusts following major disturbances such as mining. Therefore, our findings highlight the need for implementing active restoration practices following mining.

Including information about soil microbial communities into global decomposition models is critical for predicting and understanding how ecosystem functions may shift in response to global change. Here we combined a standardised litter bag method for estimating decomposition rates, the Tea Bag Index (TBI), with high-throughput sequencing of the microbial communities colonising the plant litter in the bags. Together with students of the Federal College for Viticulture and Fruit Growing, Klosterneuburg, Austria, acting as citizen scientists, we used this approach to investigate the diversity of prokaryotes and fungi-colonising recalcitrant (rooibos) and labile (green tea) plant litter buried in three different soil types and during four seasons with the aim of (i) comparing litter decomposition (decomposition rates (k) and stabilisation factors (S)) between soil types and seasons, (ii) comparing the microbial communities colonising labile and recalcitrant plant litter between soil types and seasons, and (iii) correlating microbial diversity and taxa relative abundance patterns of colonisers with litter decomposition rates (k) and stabilisation factors (S). Stabilisation factor (S), but not decomposition rate (k), correlated with the season and was significantly lower in the summer, indicating a decomposition of a larger fraction of the organic material during the warm months. This finding highlights the necessity to include colder seasons in the efforts of determining decomposition dynamics in order to quantify nutrient cycling in soils accurately. With our approach, we further showed selective colonisation of plant litter by fungal and prokaryotic taxa sourced from the soil. The community structures of these microbial colonisers differed most profoundly between summer and winter, and selective enrichment of microbial orders on either rooibos or green tea hinted at indicator taxa specialised for the primary degradation of recalcitrant or labile organic matter, respectively. Our results collectively demonstrate the importance of analysing decomposition dynamics over multiple seasons and further testify to the potential of the microbiome-resolved TBI to identify the active component of the microbial community associated with litter decomposition. This work demonstrates the power of the microbiome-resolved TBI to give a holistic description of the litter decomposition process in soils.

Including information about soil microbial communities into global decomposition models is critical for predicting and understanding how ecosystem functions may shift in response to global change. Here we combined a standardised litter bag method for estimating decomposition rates, Tea Bag Index (TBI), with high-throughput sequencing of the microbial communities colonising the plant litter in the bags. Together with students of the Federal College for Viticulture and Fruit Growing, Klosterneuburg, Austria, acting as citizen scientists, we used this approach to investigate 30 the diversity of prokaryotes and fungi colonising recalcitrant (rooibos) and labile (green tea) plant litter buried in three different soil types and during four seasons with the aim of (i) comparing litter decomposition [decomposition rates (k) and stabilisation factors (S)] between soil types and seasons, (ii) comparing the microbial communities colonising labile and recalcitrant plant litter between soil types and seasons (iii) correlating microbial diversity and taxa relative abundance patterns of colonisers with litter decomposition rates (k) and stabilisation factors (S). Stabilisation factor (S), but not 35 decomposition rate (k), correlated with the season and was significantly lower in the summer. This finding highlights the necessity to include colder seasons in the efforts of determining decomposition dynamics in order to quantify nutrient cycling in soils accurately. With our approach, we further showed selective colonisation of plant litter by fungal and prokaryotic taxa sourced from the soil. The community structures of these microbial colonisers differed most profoundly between summer and winter, and rooibos litter was generally a stronger selector than green tea litter. Moreover, this study 40 indicates an equal, if not higher, importance of fungal vs prokaryotic degraders for recalcitrant and labile plant litter decomposition. Our results collectively demonstrate the importance of analysing decomposition dynamics over multiple seasons and isolating the effect of the active component of the microbial community. https://doi.org/10.5194/soil-2021-110 Preprint. Discussion started: 21 October 2021 c © Author(s) 2021. CC BY 4.0 License.

The following protocol is intended as a downstream application for our Purification of RNA from a DNA/RNA Extract protocol. This protocol describes how to synthesise a first-strand non-specific complementary DNA (cDNA) from a purified RNA extract using SuperScript IV Reverse Transcriptase. The second strand synthesis is usually not required for most downstream applications. This protocol is a simplified and condensed version of the full protocol provided by the manufacturer.

Phenol is a common chemical used for nucleic acid extraction. Phenol can be a component in a commercial reagent (e.g. QIAzol, TRIzol) or prepared as part of a mixture in the laboratory (e.g. chloroform:phenol). Because phenol solutions are an integral part of routine life science applications, their hazards may be taken for granted. Phenol can be very dangerous, and the hazards are not just those of a typical corrosive. The hazards of phenol are 2 fold: it is both corrosive (can cause severe burns) and toxic (absorbed phenol acts as a systemic toxin). In one case, death resulted from ingestion of as little as 15 ml. Liquid phenol can penetrate the skin with efficiency approximately equal to that of inhalation. Deaths have been reported for exposures of 25% or more of body surface area. Phenol has an anaesthetic effect and can cause severe burns that may not be immediately painful or visible. The threshold concentration of human skin damage from phenol is 1.5%. It can cause permanent eye injury and blindness. Prevention Anyone working with phenol should be familiar with its risks, chemical properties, and ways of handling spillages or exposure. Please read the MSDS and this protocol before starting to work with phenol. Work with phenol is only allowed in a working chemical hood, and a phenol decontamination kit should be at reach. The user must wear protective equipment described in this protocol at all times. First aid for dermal (skin) exposures It is recommended to use polyethylene glycol 300 or 400 (PEG-300 or PEG-400) rather than water for immediate first aid treatment of dermal exposures. First aid for spills For phenol spill on a floor or bench (non-body case), use an absorbent (e.g. Vermiculite). After it soaks the liquid, collect them into a plastic bag and store them together with other toxic/phenol waste below the fume hood in a ventilated cabinet.

The following protocol describes how to perform an RNA-Stable Isotope Probing experiment. The scope of this protocol only covers the parts involving separating labelled RNA from unlabelled RNA using ultracentrifugation in a caesium trifluoroacetate density gradient and downstream quantification to evaluate whether the labelling and separation of the RNA were successful. Total RNA should be extracted from an environmental sample or an enrichment culture that was incubated with an isotopically-labelled substrate. Labelling can be of the carbon, oxygen or nitrogen in the RNA (or any combination of the 3). For environmental samples, we recommend extracting RNA using our protocol Total Nucleic Acids Extraction from Soil and purifying it using the Purification of RNA from Crude NA Extract protocol. This protocol is based on the following papers: Whiteley et al. (2007); Dumont et al. (2011); Angel and Conrad (2013). For a comprehensive discussion on how to design a SIP experiment and how to analyse the resulting data, we recommend referring to the recent book on the subject: Stable Isotope Probing: Methods and Protocols, especially chapters: 1-3 and 9-18.

The following protocol is intended for the quantification of double-stranded DNA using Quant-iT™PicoGreen® dsDNA Assay Kit (ThermoFisher). This protocol is a simplified and condensed version of the full protocol from the manufacturer. The procedure described here is for 96 reactions. If samples are run in duplicates, then this should allow quantifying 40 samples.

Universal 16S rRNA probe-based-qPCR assay for bacteria. The primers target the V4 region of the 16S rRNA gene and were specifically designed for Illumina amplicon sequencing. The original primers were designed by Caporaso et al. (2012) and modified by Walters et al. (2015). For barcoding, we use the Fludigm Access Array for barcoding the sample and therefore the primers are synthisized with the CS1 and CS2 regions.

Millipedes are among the largest and most important invertebrates, with over 12,000 identified and 80,000 expected species worldwide. Millipedes are detritivores living on leaf litter, deadwood, or soil. Because of the poor nature of their diets, millipedes compensate through high food consumption. Thanks to this, they are keystone species in many terrestrial ecosystems. In fact, in tropical and temperate zones, they rank the third most essential macrodetritivores after termites and earthworms and consume 10-36% of the annual litter. Thus, they contribute to soil formation and are essential forest ecosystem engineers. Despite their ecological importance, it remains unclear what role does their microbiome play in their diet. We studied the gut microbiota of 11 millipede species and measured key physicochemical conditions (redox, pH and O2 levels). We found that the bacterial and archaeal communities were phylogenetically conserved while the fungi matched the diet. Methanogenic millipedes had a distinct community dominated by fermenting and syntrophic microorganisms. Follow-up experiments on the methanogenic and non-methanogenic species Epibolus pulchripes and Glomeris connexa, respectively, showed that both could survive prolonged antibiotic treatment, although with some disruption of their digestion. Antibiotics treatment significantly reduced the faecal bacterial colony counts after seven days in both species. Additionally, methane production dropped by 74% in the group treated with antibiotics and 52%, in the group that received sterile feed without antibiotics. Microbiome analysis of these groups showed major shifts of the community composition in response to antibiotics, but less so with sterile feed. Apart from the presence of methanogens, high methane production correlated with a high relative abundance of Bacteroidia, while Gammaproteobacteria dominated the guts of millipedes with low, or no, methane production. By supplementing the millipedes' diet with BES, methane production could be suppressed entirely within 21 days. Microscopic analysis of the faeces (using CARD-FISH) revealed methanogens from the orders Methanobacteriales and Methanomassiliicoccales associated with ciliates. These methanogens persisted even in the absence of methane production. Our results indicate a significant gut microbiome activity in cellulolytic, fermentative and methanogenic litter decomposition processes, however, unlike in ruminants and termites with a limited nutritional contribution to the host.

The following protocol describes how to perform an RNA-Stable Isotope Probing experiment. The scope of this protocol only covers the parts involving separating labelled RNA from unlabelled RNA using ultracentrifugation in a caesium trifluoroacetate density gradient and downstream quantification to evaluate whether the labelling and separation of the RNA were successful. Total RNA should be extracted from an environmental sample or an enrichment culture that was incubated with an isotopically-labelled substrate. Labelling can be of the carbon, oxygen or nitrogen in the RNA (or any combination of the 3). For environmental samples, we recommend extracting RNA using our protocol Total Nucleic Acids Extraction from Soil and purifying it using the Purification of RNA from Crude NA Extract protocol. This protocol is based on the following papers: Whiteley et al. (2007); Dumont et al. (2011); Angel and Conrad (2013). For a comprehensive discussion on how to design a SIP experiment and how to analyse the resulting data, we recommend referring to the recent book on the subject: Stable Isotope Probing: Methods and Protocols, especially chapters: 1-3 and 9-18.

The following protocol describes how to perform an RNA-Stable Isotope Probing experiment. The scope of this protocol only covers the parts involving separating labelled RNA from unlabelled RNA using ultracentrifugation in a caesium trifluoroacetate density gradient and downstream quantification to evaluate whether the labelling and separation of the RNA were successful. Total RNA should be extracted from an environmental sample or an enrichment culture that was incubated with an isotopically-labelled substrate. Labelling can be of the carbon, oxygen or nitrogen in the RNA (or any combination of the 3). For environmental samples, we recommend extracting RNA using our protocol Total Nucleic Acids Extraction from Soil and purifying it using the Purification of RNA from Crude NA Extract protocol. This protocol is based on the following papers: Whiteley et al. (2007); Dumont et al. (2011); Angel and Conrad (2013). For a comprehensive discussion on how to design a SIP experiment and how to analyse the resulting data, we recommend referring to the recent book on the subject: Stable Isotope Probing: Methods and Protocols, especially chapters: 1-3 and 9-18.

The following protocol is intended for the quantification of RNA using Quant-iT™ RiboGreen™ RNA Assay Kit (ThermoFisher). This protocol is a simplified and condensed version of the full protocol from the manufacturer. The procedure described here is for 96 reactions. If samples are run in duplicates, then this should allow quantifying 40 samples.

The protocol was designed to quantify microbial eukaryotic fungi using ITS region copy number evaluation by Droplet Digital PCR technology (ddPCR) from Bio-Rad company. For the assay, we are using a universal fungal ITS rRNA primer pair: ITS1f 5'- CTT GGT CAT TTA GAG GAA GTA A -3', 38 bp upstream of ITS1 from White et al., 1990 ITS2 5'- GCT GCG TTC TTC ATC GAT GC -3', identical to ITS2 from White et al., 1990until now because of its extreme length variability among the fungi species. In fact, this variability introduced a bias to final copy numbers estimated by qPCR,. Amplicon size: ~250–600 bp (Bokulich & Mills, 2013; Hoggart et al., 2018) The same ITS1f and ITS2 primers are recommended for use for fungi identification by the Earth Microbiome Project.

Universal 16S rRNA probe-based-qPCR assay for bacteria. The primers and probe are taken from Yu et al. (2005).

The following protocol is intended as a downstream application for our Total Nucleic Acids Extraction from Soil protocol. This protocol describes how to purify RNA from a DNA and RNA extract using TURBO™ DNase and GeneJET RNA Cleanup and Concentration Micro Kit. This protocol is a simplified and condensed version of the full protocols provided by the manufacturers.

The protocol is dedicated to evaluation of 18S rRNA fungal gene copy number using Droplet Digital PCR technology (ddPCR) from Bio-Rad company. This is the up-to-date modification and improvement of clasical probe based qPCR assay. For the assay, we are using universal fungal 18S rRNA primers and a probe that showed broad-coverage and favourable quantitative parameters (adapted from Liu et al., 2012): FungiQuantF 5'- GGR AAA CTC ACC AGG TCC AG -3' , targets S. cerevisiae 1199-1218, FungiQuantPrb* 5'- TGG TGC ATG GCC GTT -3', targets S. cerevisiae 1269-1283, FungiQuantR 5'- GSW CTA TCC CCA KCA CGA -3', targets S. cerevisiae 1532-1549. * Probe must be dual-labelled either with 5’-6-FAM, 3’-BHQ1 or any other valid combination. The expected amplicon length is 351bp.

The following protocol is intended for the quantification of double-stranded DNA using Quant-iT™PicoGreen ® dsDNA Assay Kit (ThermoFisher). This protocol is a simplified and condensed version of the full protocol from the manufacturer. The procedure described here is for 96 reactions. If samples are run in duplicates, then this should allow quantifying 40 samples.