Bacterial communities and soil chemistry from ten established invasions of Lupinus polyphyllus in southwestern Finland, 2020
Data files
Oct 24, 2024 version files 14.54 MB
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Bacteria.root.nodule.csv
14.41 MB
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Bacteria.soil.csv
124.26 KB
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README.md
5.45 KB
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Soil.chemistry.csv
821 B
Abstract
Plants host microorganisms that can facilitate their success in becoming invasive. Established plant invasions might thus provide useful insights into potential changes in plant-associated microbiomes over the course of the invasion process. Here, we investigated the endophytic bacterial communities of the invasive herbaceous legume Lupinus polyphyllus, which is able to form mutualistic associations with N-fixing bacteria. More specifically, we examined the alpha diversity (observed bacterial taxa richness and Shannon diversity) and composition of bacterial communities in roots and nodules sampled from core and edge locations within 10 established invasion sites (>10 years old) in southwestern Finland. Moreover, we compared the alpha diversity and structure of bacterial communities in the rhizosphere and bulk soil between core and edge locations within these invasion sites. We found that roots and nodules had distinctive endophytic bacterial communities, with roots having 24% higher bacterial alpha diversity (Shannon diversity) than nodules. In nodules, the dominant bacteria were assigned to the family Bradyrhizobiaceae, which includes N-fixing bacteria. Soil bacterial communities, instead, were shaped by soil type, with bulk soil hosting up to 27% higher alpha diversity (richness and Shannon diversity) than rhizosphere soil; however, there was no apparent difference in their community composition. Soil bacterial communities were only weakly associated with soil chemistry. Endophytic and soil bacterial communities did not differ between core and edge locations within the established invasions. Our findings suggest that L. polyphyllus may not induce dramatic changes in the bacterial communities with which it associates over the course of the local invasion process.
README: Bacterial communities and soil chemistry from ten established invasions of Lupinus polyphyllus in southwestern Finland, 2020
[Access this dataset on Dryad] doi:10.5061/dryad.qz612jmrc
Content: This dataset consists of three data files (Bacteria.root.nodule.csv, Bacteria.soil.csv, Soil.chemistry.csv) that contain all variables used for the statistical analyses presented in Ramula, Mousavi, and Vesterinen "Root, nodule, and soil bacterial communities associated with the invasive nitrogen-fixing Lupinus polyphyllus", submitted for Ecology and Evolution.
Experimental procedures: To examine the alpha diversity and composition of endophytic and soil bacterial communities in core and edge locations within established invasions, samples were collected within ten sites invaded by the herbaceous Lupinus polyphyllus (>10 years old) in southwestern Finland. The distances between the ten sample sites varied from 1.7 to 32.7 km. In each invasion site, 10 haphazardly chosen flowering individuals were sampled from the core and edge locations, respectively, resulting in a total of 200 root samples (10 plants × 2 locations × 10 invasion sites). In addition, soil samples were collected from the rhizosphere of three flowering plants (at a depth of 10–15 cm) and bulk soil (at a depth of >20 cm) in the core and edge locations within invasion sites; these samples were pooled within each location per invasion site.
Bacterial DNA was identified by sequencing the gene 16S rRNA, resulting in a total of 13 487 ZOTUs (zero-radius operational taxonomic units) for the root and nodule samples, and 1 141 ZOTUs for the soil samples after rarefying the data. To characterise soil bacterial diversity, bacterial richness (i.e., number of bacterial ZOTUs per sample) and Shannon diversity index were calculated based on the bacterial ZOTUs.
To characterise soil chemistry, separate soil samples were collected from the rhizosphere of L. polyphyllus in the core and edge locations of each invasion site (no bulk soil was sampled). Again, three flowering plants were sampled per location and their rhizosphere soil was pooled (0.5 L soil in total), resulting in 20 soil samples (2 locations × 10 invasion sites). The following soil properties were analysed: total nitrogen (N), nitrate (NO3-), ammonium (NH4+), calcium (Ca), phosphorus (P), potassium (K), magnesium (Mg), and pH. The soil analysis was carried out by Eurofins Agro in Finland (https://www.eurofins.fi/agro).
Date of data collection: June-July 2020 (range: 2020-07-29 - 2020-07-02)
Description of the data and file structure:
Bacteria.root.nodule.csv
Description: This data file contains bacterial communities in roots and nodules sampled within 10 invasion sites of the legume Lupinus polyphyllus in southwestern Finland, 2020.
Definitions of all variables in the file:
Site: ten invasion sites of L. polyphyllus in alphabetics (range: A-J)
Location: sampling location within each invasion site with two levels (Core = core location, Edge = edge location)
Tissue.type: plant tissue used for the extraction of bacterial DNA with two levels (Root, Nodule)
Replicate: a replicate code per location at each site (range: 1-10 L. polyphyllus individuals sampled)
Sample.ID: a unique sample code used for genetic libraries (380 samples in total)
Richness: number of ZOTUs oberved per sample
Shannon: Shannon deiversity index (H’) calculated from ZOTUs
ZOTU1-ZOTU9491: columns containing the relative abundances of bacterial zero-radius operational taxonomic units (ZOTUs), 13478 ZOTUs in total. Note that the ZOTUs are not shown in consecutive numbers from smallest to largest.
Bacteria.soil.csv
Description: This data file contains soil bacterial communities sampled in core and edge locations within 10 invasion sites of the legume Lupinus polyphyllus in southwestern Finland, 2020.
Definitions of all variables in the file:
Site: ten invasion sites of L. polyphyllus in alphabetics (range: A-J)
Location: sampling location within each invasion site with two levels (Core = core location, Edge = edge location)
Soil.type: soil type used for the extraction of bacterial DNA with two levels (Bulk, Rhizosphere)
Sample.ID: a unique sample code for genetic libraries (38 samples in total)
Richness: number of ZOTUs per sample
Shannon: Shannon diversity index (H’) calculated from ZOTUs
ZOTU1-ZOTU9513: columns containing the relative abundances of bacterial zero-radius operational taxonomic units (ZOTUs), 1141 ZOTUs in total. Note that the ZOTUs are not shown in consecutive numbers from smallest to largest.
Soil.chemistry.csv
Soil chemistry in core and edge locations within 10 invasion sites of the legume Lupinus polyphyllus (n=20 soil samples) in southwestern Finland, 2020.
Definitions of all variables in the file:
Site: ten invasion sites of L. polyphyllus in alphabetics (range: A-J)
Location: sampling location within each invasion site with two levels (Core = core location, Edge = edge location)
pH: soil pH
Ca: soil calsium content (mg/L)
P: soil phosphorus content (mg/L)
K: soil potassium content (mg/L)
Mg: soil magnesium content (mg/L)
NO3: soil nitrate content (mg/L)
NH4: soil ammonium content (mg/L)
N.total: soil total nitrogen content (mg/L)
Software
Microsoft Excel can be used to view Bacteria.root.nodule.csv, Bacteria.soil.csv, and Soil.chemistry.csv. The data files can be used independently.
Methods
Root and soil sampling
We carried out root and soil sampling in 10 sites with established invasions of L. polyphyllus over four days (29 June–2 July) in the summer of 2020 when the species was flowering. All of these sites were close to Turku in southwestern Finland (latitude: 60.357–60.521, longitude: 22.168–22.740; distances 1.7–32.7 km apart). They featured sandy moraine or clay soil and were located in wastelands, road verges, former fields, or forest understoreys. The exact age of the invasions is unknown, but the species has been present in each location since at least 2010 when we visited the populations for the first time (Ramula, 2014). The size of these invasions vary between 65-2400 m2 (mean ± SD = 617.4 ± 736.4 m2) and currently, they contain hundreds of individuals (mean cover = 59.7%, range: 2%–96%). In each invasion site, we haphazardly sampled 10 flowering individuals from the core and edge locations, respectively, with plant density typically being lower in the latter (mean ± SD = 3.3 ± 1.1 for core and 2.0 ± 0.9 for edge). We dug up each sampled plant, used a spade to gently remove soil close to its root system, cut a ca. 10-cm piece of root containing nodules, and placed it in a plastic ziplock bag, resulting in a total of 200 root samples (10 plants × 2 locations × 10 invasion sites). In addition, we collected soil samples from the ectorhizosphere (hereafter rhizosphere) of three plants (at a depth of 10–15 cm) and bulk soil (at a depth of >20 cm) in the core and edge locations; these samples were pooled within each location per invasion site and stored in a plastic bag. Altogether, we had 40 soil samples (2 locations (core and edge) × 2 soil types (rhizosphere and bulk) × 10 invasion sites). To avoid cross-contamination, the spade and secateurs were wiped with ethanol after each sampling event. We transported all samples in a cold box to the laboratory and preserved the soil samples at -80oC for later use. In the laboratory, we rinsed the roots with tap water to remove the soil, and cut about 2 cm of each sample and some nodules for further analysis. These root and nodule samples were surface-sterilised in 96% ethanol for 1 min and 3% NaClO for 3–5 min and rinsed with deionised water three times. The specimens were preserved in Eppendorf tubes at -80oC for DNA isolation.
To characterise soil chemistry, we collected separate soil samples from the rhizosphere of L. polyphyllus in the core and edge locations of each invasion site (no bulk soil was sampled). Again, we sampled three flowering plants per location and pooled their rhizosphere soil (0.5 L soil in total), resulting in 20 soil samples (2 locations × 10 invasion sites). The following soil properties were analysed: total nitrogen (N), nitrate (NO3-), ammonium (NH4+), calcium (Ca), phosphorus (P), potassium (K), magnesium (Mg), and pH. The soil analysis was carried out by Eurofins Agro in Finland (https://www.eurofins.fi/agro).
DNA extraction
We cut the root samples into pieces 2–4 mm in length using sterilised (in 70% ethanol) blades. We then crushed the nodules and root pieces (up to 150 mg) using tweezers sterilised in 70% ethanol for DNA extraction. For the soil samples, we used 300 mg of each sample for DNA extraction. For all samples (roots, nodules, soil), we extracted DNA with the NucleoSpin Soil Kit (Macherey-Nagel GmbH Co. KG, Duren, Germany) following the provided protocol. To improve the DNA yield, root and nodule samples were placed in TissueLyser II (Qiagen) for the sample lysis step. The extracted DNA samples were preserved at -20oC for PCR and sequencing.
Polymerase chain reaction (PCR)
Locus-specific PCR and library preparation was done by the company Bioname Ltd, Finland (www.bioname.fi). The ribosomal bacterial 16S v8–v9 gene region was amplified using nested PCR and two primer pairs (799F [AACMGGATTAGATACCCKG; Chelius & Triplett, 2001] and 1492R [GGYTACCTTGTTACGACTT; Lane 1991]), and (1062F [GTCAGCTCGTGYYGTGA; modified from Ghyselink et al., 2013] and 1390R [ACGGGCGGTGTGTRCAA; modified from Zheng et al., 1996]). The secondary pair of the primers included a linker-tag to enable the subsequent attachment of NGS adapters. To increase amplicon library diversity, the secondary primer pair was used as four different versions so that they included heterogeneity spacers between the linker-tag and the locus-specific oligo. A blank PCR control was added to each PCR batch to evaluate the purity of reagents and the level of cross-contamination. In this project, the PCR controls were done in parallel in separate tubes because the DNA plates contained no empty wells. Initially, we carried out a pre-PCR using the primers 799F and 1492R to amplify a longer fragment of the bacterial 16S region. This primer pair is used to exclude plant chloroplast amplification (Chelius & Triplett, 2001). The reaction setup followed Kankaanpää et al. (2020) and included 5 μL of 2×MyTaq HS Red Mix (Bioline, UK), 2.4 μL of H2O, 150 nM of each primer, and 2 μL of DNA extract per sample in a 10-μL volume. Cycling conditions for 799F and 1492R were as follows: 3 min at 95°C, 35 cycles of 45 sec at 95°C, 45 sec at 54°C, and 1 min at 72°C, ending with 5 min at 72°C. The nested PCR reaction included the locus-specific primer pair 1062F and 1390R. The PCR mixture from the pre-PCR round was used as a DNA template in this round of PCR. All PCR reactions after the pre-PCR were carried out in two technical replicates, in which each replicate contained two heterogeneity versions of each primer.
Library preparation and sequencing
Library PCR followed, with minor modifications, the procedure of Vesterinen et al. (2016). We used a dual indexing strategy, in which each reaction (including technical replicates) was prepared with a unique combination of forward and reverse indices. All index sets were balanced perfectly so that each nucleotide position included either T/G or A/C, which ensures base calling for each channel during sequencing. For a reaction volume of 10 μL, we mixed 5 μL of MyTaq HS RedMix, 500 nM of each tagged and indexed primer (i7 and i5), and 3 μL of locus‐specific PCR product from the first PCR phase. For library preparation PCR, we used the following protocol: initial denaturation for 3 min at 98°C, then 12 cycles of 20 s at 95°C, 15 s at 60°C, and 30 s at 72°C, followed by 3 min at 72°C. All the indexed reactions were then pooled and purified using magnetic beads (Vesterinen et al., 2018). Sequencing was done at the Turku Centre for Biotechnology (Turku, Finland), on an Illumina NovaSeq6000 SP FlowCell v1.5 PE 2x250 apparatus (Illumina Inc., San Diego, California, USA), run together with other samples, including a PhiX control library.
Bioinformatics and diversity measures
Our bioinformatics pipeline closely followed that in Kaunisto et al. (2020) as summarised below. In this study, we processed a total of 44 489 226 raw reads (25 151 901 in PCR replicate 1 and 19 337 325 in PCR replicate 2). Paired-end reads were merged and trimmed for quality using 64-bit VSEARCH version 2.14.2 (Rognes et al., 2016), and the primers were removed from the merged reads using the software CUTADAPT version 2.7 (Martin, 2011) with 20% rate for primer mismatches and strict length parameters: minimum length 285 bp, maximum length 320 bp (longer sequences clipped from 3’ end). After removing singletons and chimers, the unique reads were clustered into ZOTUs (zero-radius operational taxonomic units) with 32-bit USEARCH version 11 (Edgar, 2010). ZOTUs were mapped back to the primer-trimmed reads to construct a zotu table with the ‘usearch_global’ algorithm in VSEARCH (Rognes et al., 2016). The resulting sequence variants were assigned to taxa using custom databases with SINTAX (Edgar, 2010) in VSEARCH (Rognes et al. 2016) The databases were downloaded from https://drive5.com/usearch/manual/sintax_downloads.html with the following reference: 16S RDP training set rdp_16s_v18 (21k seqs). The reads from both PCR replicates were pooled ifreads were detected in both of them. We removed singletons (read count <2) and only retained reads assigned as Bacteria.
Bioinformatic processing resulted in 15 003 ZOTUs for subsequent analyses. Since the sampling design for soil differed from that for roots and nodules, we analysed these two data sets separately. After rarefying (8 104 sequences per sample for roots and nodules and 88 sequences for soil), we continued our analyses with a total of 13 478 ZOTUs for the root and nodule samples, and 1 141 ZOTUs for the soil samples. Low sequence counts for soil were due to the inadequate dilution of the samples. To evaluate the diversity of these bacterial communities, we calculated two alpha diversity measures (observed ZOTU richness and Shannon index (H’)) separately for the root and nodule data and for the soil data using the R (R Core Team, 2023) packages “phyloseq” (McMurdie & Holmes, 2013) and “microeco” (Liu et al., 2021).
References
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