Summary
Increasing frequency and intensity of drought and periodic precipitation events caused by global climate change can lead to varying soil moisture which directly and radically affect plants and soil microbial communities. The soil microbial biomass-C significantly reduce, and the microbial composition turns to be more fungi dominant in respond to drying-rewetting cycles. As root exudations communicating roots and their soil microbes, the change on associated soil community cause indirect impacts on plant growth and the resistance and resilience of soil microbial communities exist in response to short-term and long-term drought. Due to root exudation varied in plant species and growth stages, different growth strategies of plant species and the same species at different growth stages also lead to more bacteria-dominant or fungi-bacteria-dominant microbial communities in response to drought. Here, we aimed at quantifying changes in soil microbial community composition driven by drought under two different plant strategies: fast and slow growing. Therefore, we sampled soil under fast-growing and slow-growing plant species and under C. vulgris at different growth stages with control and drought. We extracted DNA from these soil samples and will characterized the microbial communities based on amplicon sequencing (16S for bacteria and ITS for fungi). To explore the variation in microbial community composition, we will calculate alpha and beta diversity indices for each community and will do taxonomical classifications based on existing databases. We hypothesise that under a long-term drought, the microbial composition varies under plant species with different growth strategies and the soil microbial community under slow-growing species change less drastically than the community of fast-growing species when facing drought due to the slow-growing species invest more in defense to drought. Furthermore, we expect that under the drought condition, the microbial composition will differ between growth stages of C. vulgaris. More specifically, the microbial community of younger Calluna change more drastically than the older plants under drought condition since the younger Calluna invest more in plant growth which leads them more vulnerable to drought. Thus, soil microbial communities indicate plants with different growth strategies to cope with drought stress and changing soil moisture during their whole lifespan. And during a long period of drought, plants mediate their root microbial composition and evolve by selecting more drought-durable communities. Theoretical framework
With climate change, drought events continue to occur, reducing the total water availability on land, especially in most parts of the Southern Hemisphere, the United States, most of Europe and the Mediterranean (Pokhrel et al., 2021). Most grassland ecosystems are suffering from the increasing frequency and intensity of drought events, mixed with periodic and extreme precipitation events (Jansson & Hofmockel, 2020). Changes in terrestrial water availability in grassland ecosystems can have an influence not only on plant growth, but also on the composition and functions of soil microbial communities (Jansson & Hofmockel, 2020). After a long-term drought effect, there could be an increase in the abundance of more drought-tolerant species in plant communities, and subsequently different root-associated microorganisms would be selected with the shift of plant species (de Vries et al., 2018; de Vries & Shade, 2013). Bacteria have been shown to be more sensitive to drought than fungi in grassland ecosystems through network analyses in mesocosms (Upton et al., 2018) and multi-year field experiments (de Vries et al., 2018). Thus, fungi potentially has a more prominent role than bacteria in keeping the carbon and nitrogen cycling when soil moisture is insufficient (Treseder et al., 2018). Furthermore, for the bulk microbiomes, fungal hyphae may help to make up for the spatially dispersed resources, in the case of drought when the diffusion of microorganisms in soil pores is limited (Jansson & Hofmockel, 2020). Although some soil microorganisms have evolved various physiological strategies to cope with drought stress, members of some bacterial groups, such as actinomycetes and bacilli, can survive in drought-affected soil (Bouskill et al., 2016; Naylor et al., 2017) because they can conserve activity and become dormant under dry
conditions. However, a subsequent increase in water availability may cause enhanced connectivity, more dispersal, more anaerobic niches in microbiomes and a sudden increase in nutrients, resulting in an increase in anaerobic taxa and a decrease in diversity (Jansson & Hofmockel, 2020). To imitate the natural precipitation and drought events, drought and re-wetting in laboratory can show how soil moisture variation influences soil microbial communities.
Soil moisture and re-wetting events affect the size, composition and ecosystem processes of microbial communities directly and radically (Rousk et al., 2013). In a series of experiments near Santa Barbara, CA, USA, Fierer et al. (2003) measured the influence of soil water potential on the microbial mineralization of native soil organic carbon (C). This study found that carbon mineralization rates increased with soil water potential (Fierer et al., 2003). Although comparing the results describing the relationship between soil water potential and microbial respiration with published studies is difficult, lower relative respiration rates indicates that soil moisture can directly affect microbial growth. When re-wetted, soil respiration increases (Kieft et al., 1987), and after several cycles of dry-rewetting around 40% of CO2 can come from the
mineralization of organic C (Wu & Brookes, 2005). Together, these results show how soil water potential affects C mineralization rates, but the underlying mechanisms of which microorganisms are driving these patterns across different ecosystems are still not well understood. However, Evans and Wallenstein (2012) found that respiration and microbial biomass-C were relatively unchanged by moisture pulses and indicated that environmental history can affect contemporary rates of biogeochemical processes. When the situation moves from incubations to grassland ecosystems, the dry-rewetting event significantly reduced microbial biomass-C (Gordon et al., 2008). Not only the microbial biomass C, but also the composition of microbial communities can vary with soil moisture. Soil bacterial and archaeal communities respond to drought stress diversely and for different soils, response strategies can be different (Placella et al., 2012). In the soil with drought history, the effects related to the induction of laboratory water stress were predominantly associated with a decrease in the putative fungal biomarker with little to no change in the bacterial biomarkers (M. A. Williams, 2007). This was also proved in the result of microorganisms comparing unimproved and improved grassland. The dry-rewetting stress leads to greater nutrient leaching from improved than from unimproved grassland soils, which have a greater microbial biomass and abundance of fungi relative to bacteria (Gordon et al., 2008). In the comparison of grassland and agriculture soil, actinomycetes and Gram-negative bacteria had relatively higher abundances before rewetting (Steenwerth et al., 2005). These studies indicate that the effect of drought events on soil microbial composition also varies with the different use of soil. Additionally, drought events have impacts on soil microbial ecosystem (Rousk et al., 2013). Respiration and bacterial growth tend to increase with soil moisture contents (Iovieno & Bååth, 2008). However, the pattern is not the same for all bacteria, with archaeal communities having a distinct response to moisture (Placella et al., 2012). This indicates that specific groups of microorganisms may be poised to respond to the wet-up event in various soil types. In a sand and a sandy loam which were pre-incubated at optimal water content for microbial activities and dried to different water contents, respiration was measured over 14 days (Chowdhury et al., 2011). In both soils, cumulative respiration decreased with decreasing water potential and microbial community composition changed with water potential. Two strategies by which microbes respond to water potential were found, a decrease in water potential up to 2 MPa kills a proportion of the microbial community, however, at lower water potential, the microbes survive and their activity per unit biomass is reduced (Chowdhury et al., 2011). In forest soil, the pulse of microbial respiration following re-wetting was investigated. Compared with the control soil, the respiratory response in drought-exposed soil was slower and reached a lower rate, which was transformed into less C mineralization one week after rewetting (Göransson et al., 2013). Moreover, the study on resistance and resilience of microbial communities in soil after drought and drying-rewetting cycles implies that drying-rewetting cycles structure microbial communities to respond quickly and efficiently and microbial communities can adapt to changing climatic conditions (de Nijs et al., 2019). Although it is shown that soil microbial
communities respond to imitated natural drought and precipitation events efficiently, their response to drought is specific to the affected period.
Drought can have short-term (up to 3 years) and long-term (10 years or more) legacy effects on plant and soil microbial communities (Rousk et al., 2013). In the short-term drought, plant growth can increase in soils with previous drought history compared to soils without drought, and fungal-based microbial communities have been shown to be more resistant to drought (de Vries, Liiri, Bjørnlund, Bowker, et al., 2012; de Vries, Liiri, Bjørnlund, Setälä, et al., 2012). Soil microbial composition and functions respond to short-term legacy effect of drought. Plant growth (de Vries, Liiri, Bjørnlund, Setälä, et al., 2012) and the resistance and resilience (de Vries, Liiri, Bjørnlund, Bowker, et al., 2012) to drought of soil food webs in two contrasting land-use systems: intensively managed wheat with a bacterial-based soil food web and extensively managed grassland with a fungal-based soil food web. After repeated drought in 2.5 months, in soils with a drought legacy effect, plant growth increased in both grasslands and wheat systems (de Vries, Liiri, Bjørnlund, Setälä, et al., 2012) and the more resistant grassland ecosystem adapted to drought better than the bacterial-based food web of wheat soil (de Vries, Liiri, Bjørnlund, Bowker, et al., 2012). As drought can induce a higher microbial C-use efficiency during the burst of microbial activity induced by the rewetting of dry soil, this can lead to a negative feedback to climate warming (Göransson et al., 2013). These drought legacy effects might be driven by declines in microbial abundance and possible changes in microbial community composition after drought treatments and this indicated that environmental changes can affect rates of ecosystem processes indirectly through microbial abundances and communities (Allison et al., 2013). However, in the long-term, drought can decrease plant growth and cause a shift in microbial communities. the flow of C to fungal and bacterial communities in the laboratory water stress treatment is diverse but not related to the 11 years of field irrigation (M. A. Williams, 2007). Not only the different time period of drought legacy effect influences the performance of soil microbial communities, but also the root exudates of different plant species can mediate the growth and composition of soil microbial communities. The ecological strategies of plants can lead to different responses to drought (de Vries et al., 2018). Slow-growing species invest more in defence than in growth and are therefore least sensitive to water availability and more resistant to drought than fast-growing species, the latter being more vulnerable but with a faster recovery (Ouédraogo et al., 2013). Therefore, A. Williams and de Vries (2020) suppose that fundamental differences in the response to root exudation between plant species with more conservative traits and those with acquisitive traits drive the responses of their associated microbial communities to drought, which in turn feeds back into their own regrowth and ultimately affects the form and function of ecosystems. Fast-growing plant species respond to drought rapidly with reduced growth and total root exudation and increased relative root exudation while slow-growing species don’t reduce their exudation, continuing to grow and exudate and both strategies show various dependence on bacteria and fungi networks through root exudates. Drought-induced changes in root exudates of fast-growing plants might mobilize more bacterial-dominated microbial communities (de Vries et al., 2018) and also increase the respiration per unit root exudate C (de Vries et al., 2019), in turn facilitating plant regrowth. Contrary to fast-growing species, slow-growing species with thicker, coarser roots generally rely more on associations with mycorrhizal fungi (Ma et al., 2018) and are associated with fungal-dominated soil microbial communities that function better under drought conditions (de Vries et al., 2018). Slow-growing plant species can be expected to be less vulnerable to drought than fast-growing species, but the latter has been shown to even increase in abundance in the presence of drought (de Vries et al., 2018). Changes in the quantity and quality of root exudates after drought may also result in increased soil organic C decomposition and contribute to CO2 production peaking when rewetting ("birch effect"); Birch,
1958). Such a mechanism, in the case of repeated droughts, would result in severe fluctuations in community biomass and ecosystem C flux, an overall increase in fast-growing species, and soil C loss, which could eventually erode resilience and lead to a regime shift (de Vries et al., 2018; A. Williams & de Vries, 2020). Microbial communities resistant to drought stress vary in
composition under plant species with two different growth strategies, but even under the same strategy, differences in drought resistance can be found when comparing different growth stages. As the direct study on the microbial composition variation of C. vulgaris plants, a slow-growing species, under natural drought and rewetting is scarce, the C stock of plants can offer a hint to explore the potential microbial variation. Aboveground C stock of plants reached its peak at approximately 18 years of age by detecting the concentration of C and nitrogen (Kopittke et al., 2013). In the aboveground C pools, it is believed that the belowground C stock of the plant community increases with their community age and phase of development, and peaks at about 18 years of age and then remain stable. When the C accumulates in plants and plants grow older, root exudates can increase in quality and quantity which might lead to increasing gram-positive bacteria and stable gram-negative bacteria and fungi under natural drought and re-wetting than in younger plants (Fuchslueger et al., 2016). Additionally, the concentration of N in plants with drought history decreases in shoots and increases in fine root (Fuchslueger et al., 2016) and younger Calluna vulgris plants tend to invest more in shoots than roots (Meyer‐Grünefeldt et al., 2015). Therefore, we hypothesize that older plants might have a more specific microbial community that they may be able to feed with root exudates even under drought stress, since they have deeper roots which allow themselves to access more water and suffer less from drought than younger plants. Thus, drought affects more strongly younger C. vulgris than older and we expect that this will also influence how their associated microbial communities respond to drought.
To study the legacy effect of drought on soil microbial communities under plant species with different growth strategies and under conservative plants at different growth stages, we sampled soil under two different plant species at a long-term drought experiment. Furthermore, for the slow-growing species C. vulgaris, we sampled soil underneath individuals at different growth stages. Specially, we asked the following questions:
1) Is there a drought legacy effect on soil microbial communities under fast growing plant species and under slow-growing plant species? Does the effect differ between the two growth strategies?
2) Is there a difference on the drought legacy effect on soil microbial communities under C. vulgaris at different growth stages?
We hypothesized that under a long-term drought, the growth rate of soil microbial communities decreases, and the microbial composition varies under plant species with different growth strategies and community age. More specifically, we expect that the soil microbial community under slow-growing species change less drastically than the community of fast-growing species when facing drought due to the slow-growing species invest more in defense to drought. And for slow-growing plant C. vulgaris, the microbial community of younger Calluna change more drastically than the older plants under drought condition since the younger Calluna invest more in plant growth which leads them more vulnerable to drought.
Material and Methods Study Site
The soil samples came from a heathland site which is one of five shrublands in Europe of the experimental field-manipulations of summer drought, located in Oldebroek, the Netherlands (52°24'N, 05°55'E) (Rousk et al., 2013), with a mean annual precipitation of 1,005 mm(de Nijs et al., 2019). This site is characterized by sandy podzol and mainly covered with Calluna vulgaris shrubs, a slow-growing plant species and Deschampsia flexuosa and Mollinia caerulea grasses, both fast-growing plant species. Since 1998, three 20 m2 plots were subjected to drought-like conditions by using a retractable transparent and waterproof cover to prevent precipitation from entering the plot of the drought treatment during the growing season, thereby
reducing the annual rainfall reaching the ground by 30% and causing average soil moisture to be reduced by 60% (Beier et al., 2004; Rousk et al., 2013). Three other same size control plots were left under ambient conditions with uncovered scaffolding.
Soil sample collection
Soil samples were collected from 6 different plots, 3 control and 3 drought plots. Each plot has had parts of it disturbed (mowed) at different time points: 2009, 2013 and the “old” not mowed part from at least 1998. These different mowing treatments allowed us to explore different growth stages of C. vulgaris, respectively representing pioneer, building, mature and degenerate at age of 7, 11 and 30 years based on its life history (Barclay-Estrup & Gimingham, 1969). In each plot, soil samples were collected underneath three individuals of C.vulgaris (from all mowing
treatments) and underneath three individuals of one of the two fast-growing grasses (only from the 2013 mowed part), all randomly selected (Figure 1). We collected a total of 69 samples as in one control plot there were no grass plants present at the time of sampling. Each soil sample consisted of several cores (up to 10 cm of soil depth) that together amounted to at least 90g of fresh soil. Upon return in the lab, the fresh soil samples were homogenized through 2 mm sieves and stored in a -20°C fridge until microbial analyses.
DNA extraction and amplification
Microbial DNA extraction was carried out using the DNeasy PowerSoil extraction kit from Qiagen by following the instructions of the kit. Initially, DNA was extracted from 0.25 g of fresh soil. After the extraction, DNA concentration was measured using the Qubit dsDNA HS Assay kit. Samples that could not be detected in an agarose gel after being amplified for 16S and ITS genes, were extracted again from one gram of soil.
Polymerase chain reactions were conducted with 25 μL assays. For the fungal ITS2
amplification, 25.0 μL of Platinum Hot Start PCR Master Mix (2x) from ThermoFisher®, 0.5 μLof each primer (10 μM), 13.0 μL of PCR-grade water, and 1.0 μL of a genomic DNA template (5 ng μL−1) were mixed in a 75 μL PCR tube for each sample. The following thermal profile was used for the fungal PCR: an initial denaturation and enzyme activation step of 95 °C for 3 min, followed by 30 cycles of 95 °C for 20 s, 55 °C for 30 s, and 72 °C for 30 s, with a final extension of 72 °C for 5 min. For the 16S amplification, 10 μL of Platinum Hot Start PCR Master Mix (2x) from ThermoFisher®, 0.5 μL of each primer (10 μM), 13.0 μLof PCR-grade water, and 1.0 μL of a genomic DNA template (5 ng μL−1) were mixed in a 75 μL PCR tube for each sample. The following thermal profile was used for bacterial PCR: an initial denaturation and enzyme activation step of 95 °C for 3 min, followed by 25 cycles of 95 °C for 20 s, 55 °C for 30 s, and 72 °C for 30 s, with a final extension of 72 °C for 5 min. Qualities of PCR products were evaluated by agarose gel electrophoresis.
Data analysis
To investigate the effects of plants, growth stage and drought treatment on soil microbial composition, I will use the QIIME2 platform (Bolyen et al., 2019). Specifically, raw sequencing reads will be demultiplexed and quality filtered, from which I will identify and tabulate amplicon sequencing variants (ASVs) using the DADA2 and deblur-denoise plugins. After
Fig. 1 Soil samples from each plot with 3 control and drought treatment.
constructing a table with relative abundance of ASVs per sample and representative sequences per ASV, I will import SILVA 16S rRNA database and UNITE ITS rRNA database separately with their trimmed sequences and train the classifier to get 16S and ITS rRNA taxonomic classification and bar plots which allows to see which taxonomic groups change under drought and under each plant species. Meanwhile, I will generate a tree for phylogenetic diversity analysis and then compute alpha and beta diversity metrics. After conducting alpha and beta diversity analysis, I will choose Shannon’s diversity index metrics which represent the abundance of soil microbial communities for alpha diversity analysis and unweighted UniFrac distance metrics which represent the dissimilarity of microbial communities for beta diversity analysis. Then, stats and plots of alpha and beta diversity will be gained and the plots can show (1) the richness and diversity of microbe species under fast- and slow-growing plant species and (2) under C.vulgaris at different growth stages with or without drought stress and (3) the differences in taxonomic abundance profiles from different soil samples. After I form the metrics of Shannon’s diversity index, unweighted UniFrac distance and taxonomic classification, I will use a linear mixed effect model in R to see if these indices differ between treatments.
Time schedule
This project will be started in March and analysis in QIIME2 and R will take more than a month respectively. Literature review will last during the whole project and thesis writing will be done from July to August. Totally, this project will take 6 months.
Funding
The project is supported by European Research Council (ERC) Start Grant to Franciska de Vries and the main cost in this project is Illumina Sequencing which is £6,876.00.
Table. 1 Time schedule for master thesis
March April May June July August
Literature review
QIIME2 for 16S rRNA
QIIME2 for ITS rRNA
Statistical analysis in R
Thesis writing
Table. 2 Funding in this project
Cost category Total in Euro
Illumina sequencing 8028.87
Travel to the field 100
Material to collect samples 20
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