Observations on two microbial life strategies in soil: Planktonic and biofilm-forming microorganisms are separable

Soil Biology and Biochemistry 136 (2019) 107535

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Observations on two microbial life strategies in soil: Planktonic and biofilmforming microorganisms are separable

T

Lukáš Bystrianskýa, Martina Hujslováb, Hana Hršelováb, Veronika Řezáčováb, Lenka Němcováa, Jana Šimsovác, Hana Gryndlerováb, Olga Kofroňovád, Oldřich Benadaa,d, Milan Gryndlera,* a

Faculty of Science, Department of Biology, J. E. Purkyně University in Ústí nad Labem, České mládeže 8, CZ40096, Ústí nad Labem, Czech Republic Laboratory of Fungal Biology, Institute of Microbiology CAS, v.v.i, Vídeňská 1083, CZ14220, Prague 4, Czech Republic c Faculty of Social Economics, Department of Mathematics and Informatics, J. E. Purkyně University in Ústí nad Labem Moskevská 54, CZ40096, Ústí nad Labem, Czech Republic d Laboratory of Molecular Structure Characterization, Institute of Microbiology CAS, v.v.i, Vídeňská 1083, CZ14220, Prague 4, Czech Republic b

ARTICLE INFO

ABSTRACT

Keywords: Biofilm Plankton Glass fiber Microbial community Microbiome

Biofilms, the communities of sessile microbial forms, are hotspots of biological activity that coexist in soil together with free-living (planktonic) biota. Sessile and planktonic microbial communities may constitute functionally different groups of organisms with different roles in interactions with organic matter and plants. Nevertheless, soil microbiomes are regularly analyzed without distinguishing biofilm inhabitants and planktonic organisms. Consequently, basic information regarding taxa constituting both communities is severely lacking, which limits the understanding of the basic structure of soil microbiome and consequently also of the microbiome functioning. In this study, we tested the hypothesis that soil biofilm and planktonic microbial communities are different. Glass fiber filters were exposed to three different field soils for 12 weeks and biofilms arose on their surfaces. The biofilms were further separated from the planktonic forms by washing the latter out of the filters and both communities were analyzed using next generation sequencing. The results revealed significant differences between biofilm and planktonic communities of bacteria and eukaryotic organisms. Our data indicate common production of motile microbial cells in the soil and specialization of some taxa (Legionella spp.) to planktonic life mode. We also noted an association between the abundance of some bacterial taxa and eukaryotic grazers suggesting a trophic interaction. Tillage, as a cause of soil disturbance, did not result in a significant increase in the abundance of most abundant biofilm associated microbial taxa. This is the first analysis of separated sessile (biofilm) and motile (planktonic) communities of soil microorganisms.

1. Introduction Soils contain microbial communities that are characterized by highly complex interactions between their components, resulting in the formation of biofilms. This is strongly suggested by the ordered and patchy distribution of microbial cells existing in clusters associated with organic particles (Nunan et al., 2003; Redmile-Gordon et al., 2014) or with organic nutrition available in the rhizosphere (Burns and Stach, 2002). Biofilm is defined as a bacterial (microbial) community surrounded by a self-produced polymeric matrix, and attached to an inert or biotic surface (Costerton et al., 1995; Flemming et al., 2016), unlike planktonic (pelagic) microbial cenosis which remains unattached to a physical surface. A wider concept of biofilm involves also microbial

*

aggregates, microcolonies and floccules (Rinaudi and Giordano, 2010). Biofilms may affect the soil structure stability and functioning as they are constituted by the extracellular polymeric matrix that may work as a flocculating agent or protective surface coating stabilizing soil aggregates (Lentz, 2015). Information regarding the composition and behaviour of soil biofilms may be the key factor for better management of biologically mediated nutrient turnover and soil health in general (Burns et al., 2013). Many beneficial rhizobacteria are found in biofilms (Kasim et al., 2016), and occurrence of biofilms in the rhizosphere allows the plants to survive environmental stresses and provides protection against pathogens (Xu et al., 2013). Plant cultures have been many times experimentally inoculated by a beneficial microorganism or complex

Corresponding author. E-mail address: [email protected] (M. Gryndler).

https://doi.org/10.1016/j.soilbio.2019.107535 Received 14 September 2018; Received in revised form 28 June 2019; Accepted 9 July 2019 Available online 10 July 2019 0038-0717/ © 2019 Elsevier Ltd. All rights reserved.

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inocula (Compant et al., 2010; van de Voorde et al., 2012), aiming at the improvement of the plant culture performance. However, these experiments were not always successful (Trabelsi and Mhamdi, 2013). The reason for these variable results may consist in the inability of the inoculant to persist in the soil, which might be caused (among other causes) by an inability to incorporate itself into the pre-existing soil biofilms. From the viewpoint of agricultural practices, this phenomenon may be the most important biofilms-connected topic. As the beneficial bacterial components of biofilms may be protected by the biofilm structure against antimicrobial compounds, they may be less susceptible to the action of preparations targeted to invading (i. e. motile, planktonic) phytopathogens. On the other hand, if the phytopathogen itself is able to live in biofilm (Koczan et al., 2009), this will influence the development of plant protection strategy by the phytopatologist. Basic understanding regarding the composition and behaviour of soil biofilms is thus a subject of great practical importance and merits serious attention. It is evident that the studies of microbial life in nature (soil biofilms being a particular example) must involve the multispecies aspect (Røder et al., 2016). At the same time, the studies of soil biofilms encounter several difficulties and there are numerous questions to be answered. Information concerning the dynamics of the biofilm establishment and development, the effects of plant/soil system properties on its development, its responsivity to external stimuli, and even the basic information with respect to the biological diversity and quantity of biofilms naturally formed in the soil is lacking. In fact, soil microbial communities are taken as a black box in many studies which do not account for living in biofilm and in plankton as two particular life strategies. The goal of this research is to contribute to understanding of the composition of microbial communities forming soil biofilms. We supposed that two types of the soil microbiota exist: first, organisms that are attached to physical surfaces (biofilm communities) and second, soil inhabitants that are freely moving in the soil environment or floating in the soil solution (communities of planktonic organisms). We tested the hypothesis that the composition of the two communities is different, though the majority of organisms may be present in both of them. Our strategy was to expose inert physical surfaces to the field soil for a period enabling soil biota to produce biofilm, and then partially separate attached microorganisms from unattached individuals by a washing procedure (Fig. 1). This way, we obtained a fraction enriched in attached organisms and a fraction enriched in planktonic forms (Fig. 1B). Having the data on prokaryotic (mainly bacterial) and eukaryotic biofilm communities at hand, we attempted to detect association of components of both communities in different samples. This association may indicate a trophic interaction between biofilm inhabitants reported by others (Huws et al., 2005; Seiler et al., 2017). Since the stability of soil structure is affected by the formation of biofilm extracellular polymeric matrices (Costa et al., 2018; Lentz, 2015), it was further aimed to assess the effects of soil tillage (a cause of mechanical soil disturbance) on biofilm communities in three different soils. Hypothetically, soil microbiome may tend to compensate for mechanical disturbance effects by increasing the activity of microbial taxa forming the biofilm. This activity increase may be accompanied by increased abundance of the active taxa. To confirm this compensation effect, we tried to detect microbial taxa that are associated simultaneously with biofilm and soil tillage/disturbance.

Fig. 1. Principle of the separation of biofilm (white circles) and planktonic (black circles) microbial communities. Mixed community (A) is formed on glass fiber filter exposed to the soil environment. During the separation procedure, the streaming of a liquid through the filter causes washing off of the majority of planktonic cells whereas biofilm structures mostly remain attached to surfaces (B). This procedure is performed in a glass filtration apparatus where the filter is fixed with its side originally exposed to the soil oriented to the bottom (C). The washed part of the filter is then aseptically cut off (D) and further processed.

the porosity of the filter enables easy flow-through of water, which is necessary for washing off the planktonic microbial cells. However, the filter itself is a fragile material and must be mechanically protected by a solid holder (trap) that can be safely immersed into the soil, as described further. Circular glass microfiber filters (Whatman, grade GF/F) with 45 mm diameter were placed into the ad-hoc fabricated polyamide traps with the one-sided opening (Supplementary Fig. S1) and covered by PVCcoated glass-fiber protective netting (1.4 mm mesh). The trap opening diameter was 40 mm. The traps were immersed into the soil, the filter surface being oriented vertically with the center located 8 cm below the soil surface. In total, 24 traps were immersed into the soils at 3 sites in the Czech Republic: Žalhostice (county Litoměřice, soil 1), Soběnice (county Liběšice, soil 2) and Ruzyně (county Prague 6, soil 3). The soil characteristics are presented in Supplementary Table S1. Eight traps were immersed per site: four in uncultivated and four in cultivated soil. The arrangement of traps at the site is shown in Supplementary Fig. S2. The traps were incubated in soils for 12 weeks (April to June 2017) and then collected. Immediately after the collection, the samples were stored on ice in a wet chamber and further processed within 24 h.

2. Materials and methods 2.1. Exposure of glass filter surfaces to the soil The glass microfiber filter is highly porous and provides inert surfaces that can be colonized by soil microorganisms. At the same time, 2

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2.2. Separation of attached and free microorganisms

Washed glass filter pieces (1 cm2 per trap, see above) were fixed in 3% glutaraldehyde in 50 mM cacodylate buffer, pH 7.2, at 4 °C overnight. After that, the filters were extensively washed with 50 mM cacodylate buffer, pH 7.2, and dehydrated through ethanol series (25, 50, 75, 90, 96, 100 and 100%). Dehydrated filters were directly dried from 100% ethanol in the critical-point-dryer (K850, Quorum Technologies Ltd, Laughton, UK) using liquid CO2 as transitional medium. Finally, the samples were sputter coated with 3 nm of platinum in a high resolution sputter-coater (Q150T, Quorum Technologies Ltd, Laughton, UK) and observed in an FEI Nova NanoSEM scanning electron microscope (FEI, Brno, Czech Republic) at 5 kV using SE, TLD and CBS detectors. Beam deceleration mode of the microscope was used for minimization of charging artifacts.

Eukaryotic SSU rRNA gene fragment (V4 region) was also amplified in triplicate in a reaction mixture containing 12.5 μl 2 × Combi PPP Master Mix (hot-start type, Top-Bio, Prague), 0.5 μl tagged primer TAReuk454FWD1 (10 μM), 0.5 μl tagged primer TAReukREV3 (10 μM), 1 μl of the same template DNA as that used for amplification of prokaryotic DNA, and 10.5 μl water. Primers were adopted from Stoeck et al. (2010) with tags according to Lejzerowicz et al. (2015). The cycling conditions published by Mahé et al. (2014) were modified as follows: 94 °C for 4 min, followed by 40 cycles of 94 °C for 30 s, 47 °C for 45 s, and 72 °C for 1 min, after which a final elongation step at 72 °C for 5 min was performed. A fragment of the ITS region of fungal communities was first amplified in triplicate from each sample. To this purpose, a reaction mixture was composed of 12.5 μl 2 × Combi PPP Master Mix, 0.5 μl primer ITS1F (10 μM), 0.5 μl primer ITS4 (10 μM), 1 μl of the same template as used in the amplification of prokaryotic and eukaryotic DNA, and 10.5 μl water. Primers were adopted from White et al. (1990). The mixture was first denatured at 94 °C for 4 min and then subjected to 35 cycles of denaturation at 94 °C for 45 s, annealing at 52 °C for 45 s and elongation at 72 °C for 90 s. Final elongation at 72 °C lasted for 10 min. The three replicates per sample were then pooled, purified by Qiaquick PCR purification kit (Qiagen) with elution volume 30 μl, diluted 1:1000 and used as a template in a second PCR amplification. A single PCR amplification was performed per sample, the reaction mixture is the same as in the first amplification described above except that tagged primers gITS7_T (Ihrmark et al., 2012) and ITS4 (White et al., 1990), targeting the ITS2 region, were used. The reaction mixture was first denatured at 94 °C for 4 min and then subjected to 25 cycles of denaturation at 94 °C for 30 s, annealing at 56 °C for 30 s and elongation at 72 °C for 30 s. Final elongation at 72 °C lasted 10 min. The final PCR product was purified by Qiaquick PCR purification kit. Products of amplification of prokaryotic, eukaryotic and fungal DNA were quantified using Picogreen fluorescence, diluted to a concentration of 20 ng/μl and combined to prepare a single sequencing library that was sequenced using Mi-Seq Illumina platform.

2.4. DNA extraction from microbial communities and sequencing

2.5. Data analysis

Whole samples of free microflora fraction (solids concentrated by 5 min/10000×g centrifugation from 10 ml of flow-through liquid, described above) or 1 cm2 washed glass paper (described above) were used as sources of DNA for molecular analysis of free and attached microbial communities, respectively. Before the DNA extraction, a volume of 4 μl of 100 μM homopolymeric carrier DNA (deoxythymidine 130-mer) was always added to samples. DNA was extracted from the samples using a glassmilk method as described by Gryndler et al. (2014). The resulting DNA extracts were further amplified to produce prokaryotic, eukaryotic and fungal PCR amplicons per each sample of attached and free microbial community. DNA fragments originating from different samples were distinguished by tag combinations of the primers. For primer sequences see Supplementary Table S2. A fragment of V4 region of SSU rRNA gene of bacterial communities was amplified in triplicate using reaction mixture containing 12.5 μl 2 × TPHS DNA-free Master Mix (hot start type, Top-Bio, Prague), 0.5 μl modified tagged primer 515Fd_T (10 μM), 0.5 μl modified tagged primer 806Rd_T (10 μM), 1 μl template DNA (5–50 ng/μl) and 10.5 μl water. The primers were modified from Apprill et al. (2015). The authors indicate that the primers are directed to prokaryotic DNA but, in reality, they amplify mainly V4 region of bacterial SSU rRNA gene. The amplicons obtained using these primers are thus further referred to as both „bacterial” and „prokaryotic“. The cycling conditions were: initial denaturation for 4 min at 94 °C, followed by 35 cycles of 45 s denaturation at 94 °C, annealing for 60 s at 50 °C, and elongation for 75 s at 72 °C. Final elongation at 72 °C lasted 10 min. The three replicates per each sample were pooled and purified by Qiaquick PCR purification kit.

Sequencing data were treated by Seed software (Větrovský and Baldrian, 2013), version 2.0.4, as follows: After the creation of contigs and removing low-quality sequences, sequences shorter than 150 base pairs (bp) or longer than 400 bp (Prokaryota) or sequences shorter than 40 bp (fungi) were discarded. ITS sequences were extracted from the fungal amplicons. The contigs were chimaera-cleaned and clustered by the Usearch tool (version 8.1.1861, Edgar and Flyvbjerg, 2015) at the similarity level of 97%. The most abundant sequences representing sequence clusters were compared (blastn algorithm, Blast tool, version 2.2.26+, Altschul et al., 1990) with GenBank database (environmental sequences, metagenomes, and unidentified organisms excluded). The sequences originating from amplicons of prokaryotic communities that were assigned to mitochondria, plastids or eukaryotes were excluded from further analyses. Similarly, the sequences of Streptophyta, Bacteria and Craniata were excluded from eukaryotic data. The remaining sequences were randomly resampled to retain 3000 prokaryotic, 2000 eukaryotic and 500 fungal sequences per sample. This procedure unified the number of sequence reads per sample for the purposes of permutation multivariate analysis of variance (PERMANOVA). The sequences have been deposited in Sequence Read Archive (NCBI) under accession number SRP149198. Operational taxonomic units (OTUs) corresponding to clusters at the level of the genus were named according to the best GenBank hit genus name and OTUs with the same genus assignment were summed and further treated as compound OTUs. Effects of three environmental factors (life strategy, tillage, soil) on microbial communities were tested on Bray-Curtis distance using PERMANOVA (Anderson, 2001), available for R programming

The traps were disassembled and protective nettings were removed from the surface of glass filters. If a root reached the surface of the filter, it was removed by fine forceps. Each filter was then superficially washed with 50 ml of sterile 0.1% MgSO4.7H2O, and aseptically fixed in glass filtration apparatus (Fig. 1C) with the side originally exposed to the soil oriented to the bottom. As it was shown in preliminary testing, this washing solution does not disturb the biofilms formed on glass surfaces exposed to soil. A 10 ml volume of autoclaved 0.1% MgSO4.7H2O solution was further left to spontaneously pass through the filter without the application of any vacuum or pressure, and collected in a sterile vessel. This liquid contained mainly fine soil particles and free (planktonic) microbial cells (see Supplementary Fig. S3). The washed part of the filter containing mainly biofilm communities was then aseptically cut off and two 1 cm2 pieces were prepared: the first was further used for electron microscopic observations and the second was used for molecular analysis of the attached (biofilm) microbial community. 2.3. Scanning electron microscopy

3

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Fig. 2. Scanning electron microscopy images of microorganisms forming biofilms on glass fiber filters immersed to soil 1 (A, B), soil 2 (C, D) or soil 3 (D, E). Attachment of bacterial cells (b) to glass fibers (g) with scarce (A, B) or rich (C–E) reticulate extracellular structures (black arrows) that connect the cells to the support is visible. The images also depict the amorphous clay particles (c) clung to glass fibers (mainly A, B). Extracellular fibrillar structures that strongly resemble nanowires (open arrows) are displayed (mainly C, D).

environment, package Vegan 2.5–3, function Adonis. As the use of PERMANOVA assumes homogeneity of dispersions, complementary multivariate homogeneity of group dispersions analysis (full model, Anderson, 2006) was applied to evaluate the within-group variations (package Vegan, function betadisper). The similarity of the communities in the samples at different levels of environmental factors was presented by Non-metrics Multidimensional Scaling (NMDS, function metaMDS) on Bray-Curtis distance. The sequencing data (without resampling) obtained from biofilm or planktonic microbial communities was further subjected to differential count analysis using DESeq2 method (R-package DESeq2, Love et al., 2014). This method is based on a generalized linear model assuming negative binomial distribution of the count data. Its main outputs are the logarithm of fold change (i. e. the logarithm of the ratio of sequence counts observed in the two treatments compared) and padj-value. In our

case, the latter parameter represents the Wald test p-value as a measure of statistical significance of the fold change, adjusted for multiple testing by the false discovery rate method. The values of padj≤0.05 are taken as statistically significant. The soil/sampling site (as an experimental factor) was left in the analysis design as a covariate. If an OTU tended to reach significantly higher sequence counts in biofilm samples than in planktonic community (fold change > 1), it was designated a „biofilm” or „biofilm associated” OTU. If an opposite trend was noted (fold change < 1), the OTU was further referred to as „planktonic” OTU, but see the notes in the first and fifth paragraphs of section 4.1. Similarly, if an OTU showed significantly higher sequence counts in samples from tilled soil compared to the samples from untilled one (fold change > 1), it was taken as associated with tilled soil. The fold change < 1 indicates an association with untilled soil. Interactions of the life mode (biofilm versus planktonic) and soil exploitation (tilled 4

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versus untilled) was further evaluated after the addition of the interaction term to the experimental design formula. OTU counts are available in the supplementary file “Supplementary_data_S1.xlsx”. To evaluate the relationships between Prokaryota and Eukaryota in biofilms (which suggest potential trophic interactions), we used a normalized version of variation matrix T (Aitchison, 1986) calculated from merged prokaryotic, eukaryotic and fungal datasets. The elements of this matrix τij meet the form ij

= var

1 x ln i 2 xj

sequences (1913 clusters) assigned to fungi, forming 292 compound OTUs (excluding unidentified sequences). 3.3. High-level microbial taxa in biofilm and planktonic communities of soils In all the three soils, Actinobacteria were more abundant in biofilm samples compared to samples of planktonic communities (Supplementary Fig. S4, merged data from tilled and untilled samples). Similarly, the bacterial group Planctomycetia also tended to be more abundant in biofilm communities, though lower sequence counts of this group were generally observed compared to Actinobacteria. Another less abundant bacterial group, Chitinophagia, was always more abundant in planktonic communities. Similar obvious patterns were not obtained for eukaryotic high-level taxa. Notably, this is also true for fungi constituting only a minor part of all eukaryotic sequences (mean abundance value 8.5%).

(1)

where the components xi and xj represent transformed (y = 100x+1) abundances of two different OTUs being analyzed. The OTU abundances were calculated as fractions of sequence numbers of these OTUs within the total sequence count in the sample. Values τij are close to zero where the ratio xi is almost constant throughout the samples, inxj

dicating a strong positive relation between the components xi and xj. To reach the analysis output formally similar to the correlation coefficient, the transformation using the formula y = e ( ij) was applied to the matrix elements. This converts the values to fall into the interval ⟨0, 1⟩. The transformed values close to one indicate a strong positive relationship between the two components. We arbitrarily set the τij≥0.8 as an indication of perceptible association. The values close to zero indicate that there is either no relationship between the components or there is a relationship of inverse proportions. The transformed variation matrix is available in the supplementary file “Supplementary_data_S1.xlsx”, sheet “Transformed variation matrix”. Only biofilm (i. e. surface-attached) microbiota was analyzed in this way. Before calculating the variation matrix, the abundance of each OTU was divided by the mean abundance of this OTU in the corresponding sampling site. This treatment excluded the influence of different abundances of the OTUs in different sampling sites and eliminated potential false results. Only the OTUs represented by at least 100 sequences at each sampling site (resampled biofilm data) were included in this analysis.

3.4. Statistical analysis at the level of genus The result of PERMANOVA (Supplementary Table S3) indicates highly significant effects of all the three explanatory factors (life mode, tillage, soil) on the composition of communities of Prokaryota, Eukaryota and Fungi in our samples. The significance of main factors was always higher than the significance of interactions. The test of the dispersion homogeneity (Supplementary Table S3), however, rejected the null hypothesis in the case of the dispersion within the eukaryotic data. In the case of prokaryotic and fungal data subsets, the null hypothesis was not rejected. NMDS ordination diagrams (Supplementary Fig. S5) clearly revealed a separation of biofilm and planktonic samples according to dissimilarities of prokaryotic, eukaryotic and fungal communities. The dissimilarities of the communities (in terms of centroid separation) were less pronounced in the case of soil as an explanatory factor. The effect of tillage was weakest and was best visible in the case of fungal communities. When the effects of life mode (biofilm associated versus planktonic OTUs) and tillage were tested using DESeq2 method, highly significant results were obtained (Table 1 and Supplementary_data_S2.xlsx). Twenty five prokaryotic OTUs showed significant association with biofilm life mode while 14 showed significant association with planktonic life mode, which constitutes, respectively, 13.9 and 7.8% of all prokaryotic OTUs with the padj value available. The numbers of OTUs associated with biofilm were higher than the numbers of OTUs associated with planktonic life mode also in the case of Fungi and Eukaryota (Table 2). Eleven prokaryotic OTUs were associated with untilled soils whereas only 3 with the tilled ones. Still lower numbers of eukaryotic OTUs showed significant association with any level of tillage as experimental factor and no fungal OTU was significantly affected by this factor. Large numbers of prokaryotic, fungal and eukaryotic OTUs were excluded from evaluation by DESeq2 method due to either independent filtering (OTUs with very low normalized sequence counts, mainly the singleton OTUs) or due to the presence of outliers within the data (Table 2). For these OTUs, the padj value is not available. DESeq2 analysis reveals that higher numbers of prokaryotic, fungal and eukaryotic OTUs tended to associate with biofilm life mode than with planktonic life mode (Table 2), but all the responsive OTUs constitute only a small fraction of the total numbers of the OTUs whose association with a particular life mode has been tested (i. e. whose with available padj value). Examples of 10 most abundant OTUs (the OTUs with highest base mean value) per each group of organisms are presented in Table 1. Among them, a number of responsive OTUs can be found. For the purposes of this work, the responsive OTUs are defined as those identified by DESeq2 method as significantly affected by an environmental factor. According to the DESeq2 results, the highly abundant prokaryotic

3. Results 3.1. Electron-microscopic observations of washed filters Electron microscopic images show that biofilms attached to the glass fibers produce extracellular structures in different amounts (Fig. 2). These structures are of reticulate character and connect cells directly to glass surfaces or surfaces of clay particles clung to glass fibers. They may represent artifacts resulting from dehydration of extracellular polymeric matrix produced by microorganisms in the specimen. Nevertheless, some of these structures (mainly visible in Fig. 2C–D) strongly resemble bacterial nanowires. 3.2. Sequencing Sequencing of prokaryotic amplicons resulted in 369199 sequences (reads) grouped in 10649 clusters. After removing sequences assigned to Eukaryota (mostly mitochondrial or plastid rDNA), 368336 sequences forming 10453 clusters remained in the dataset. In total, 901 compound prokaryotic OTUs were formed at the level of genus, excluding unidentified sequences. Out of 188235 sequences obtained from amplicons produced by Eukaryota-specific PCR (grouped in 3149 clusters), 169141 sequences (constituting 3021 clusters) were assigned to eukaryotic microorganisms whereas the remaining sequences, assigned to Bacteria, Streptophyta, and Craniata, were excluded from the dataset. When the remaining eukaryotic clusters with the same genus identification (i. e. with the same GenBank best hit genus) were summed, 529 compound eukaryotic OTUs were formed, excluding unidentified sequences. Sequencing the amplicons obtained in fungalspecific PCR produced, after in-silico extraction of ITS region, 134983 5

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Table 1 Results of DESeq2 method of differential sequence read count analysis of prokaryotic, eukaryotic and fungal OTUs in communities associated with biofilm or planktonic life mode in tilled and untilled soils. The fold change for the factor Life mode higher than 1 indicates that the OTU shows higher abundance in biofilm communities than in plankton and vice versa. In analogy, the fold change for the factor Tillage higher than 1 indicates the higher abundance of the OTU in tilled soil samples, the fold change lower than 1 indicates that this OTU is more abundant in untilled soil samples. padj represents the Wald test p values adjusted for multiple testing by the false discovery rate method. The values of padj≤0.05 are given in bold. The base mean value is the average of the normalized count values taken over all samples. The analysis of 10 most abundant OTUs is presented, for complete analysis output see Supplementary_data_S2. NA – the value is not available. OTU

Prokaryota Pseudomonas Rhizobium Acidovorax Arthrobacter Brevundimonas Bosea Pelomonas Pseudoxanthomonas Legionella Hydrogenophaga Fungi Fusarium Cladosporium Cylindrocarpon Ilyonectria Paraphoma Pyrenochaeta Plectosphaerella Neonectria Alternaria Paratricharina Eukaryota Acrobeloides Colpoda Fusarium Chiloplacus Pedospumella Cercomonas Heteromita Paracercomonas Mesorhabditis Oscheius

Factor

Life mode (biofilm vs.planktonic)

Tillage (tilled vs. untilled)

Interaction (Life mode vs.Tillage)

Base mean

Fold change

padj

Fold change

padj

padj

454 250 200 184 177 170 162 159 159 155

2.48 2.69 1.62 6.13 0.671 2.54 2.04 1.68 0.0637 1.40

0.002 < 0.001 0.167 < 0.001 0.242 < 0.001 0.320 0.153 < 0.001 0.479

0.827 1.99 1.35 1.59 2.74 1.12 0.657 1.70 1.46 1.14

0.681 0.052 0.504 0.372 0.004 0.683 0.674 0.231 0.395 0.805

0.994 0.994 0.994 0.994 0.994 0.994 0.994 0.994 0.994 0.994

1192 270 159 156 93.6 85.9 54.1 47.3 41.5 36.5

12.6 0.302 58.0 16.5 234 248 5.14 240 0.141 1.89 × 108

< 0.001 0.014 < 0.001 < 0.001 < 0.001 < 0.001 0.095 < 0.001 0.017 NA

2.06 0.818 2.12 2.31 3.72 0.429 0.195 0.721 1.55 0.00216

0.991 0.991 0.991 0.991 0.991 0.991 0.862 0.991 0.991 NA

0.997 0.997 0.997 0.997 0.997 0.997 0.997 0.997 0.997 NA

350 143 138 128 128 126 123 99.0 86.5 85.7

0.121 0.462 21.0 0.0174 1.00 6.20 14.6 9.79 0.0945 0.00972

0.001 0.063 < 0.001 NA 1.000 < 0.001 < 0.001 < 0.001 0.020 NA

1.46 0.937 1.96 0.436 3.37 2.45 0.552 0.589 0.0150 0.00103

0.635 0.867 0.096 NA 0.086 0.052 0.274 0.382 < 0.001 NA

0.991 0.991 0.991 NA 0.991 0.991 0.991 0.991 0.991 NA

OTUs corresponding to genera Pseudomonas, Rhizobium, Arthrobacter and Bosea are associated with biofilm and the highly abundant OTU corresponding to the genus Legionella shows significant association with planktonic life mode. The significant association of fungal OTUs with biofilm life mode may merely indicate that the corresponding fungal mycelia are entangled into glass fiber filter used as the filling of traps and are not necessarily physically attached to the surfaces. Taking in mind this reservation, we noted 6 highly abundant fungal OTUs (Fusarium, Cylindrocarpon, Ilyonectria, Paraphoma, Pyrenochaeta and Neonectria)

presented in Table 1 as significantly correlating with „biofilm life mode“ and 2 OTUs (Cladosporium and Alternaria) tending to avoid it. Significant preference for tilled or untilled soil was not observed among the 10 most abundant fungal OTUs. DESeq2 method also identified 4 highly abundant eukaryotic OTUs listed in Table 1 as significantly associated with biofilm life mode: Fusarium, Cercomonas, Paracercomonas and Heteromita. Two other OTUs, Acrobeloides and Mesorhabditis, were associated with planktonic life mode. Among the prokaryotic, fungal and eukaryotic OTUs presented in

Table 2 Overview of results of DESeq2 analysis. The table shows the numbers of OTUs with significant fold change values > 1 (OTUs associated with biofilm life mode or tilled soil) and the numbers of OTUs with significant fold changes < 1 (OTUs associated with planktonic life mode or untilled soil). Further, the numbers of OTUs that were excluded from the analysis (padj not available) due to the presence of outliers within the data (Outlier cut) or due to the action of independent filtering (Independent filter cut) are presented. The numbers in brackets indicate percentages of OTUs showing significant fold change within the number of all OTUs with padj available. Factor

Life mode

Tillage

Group of organisms

Prokaryota

Fungi

Eukaryota

Prokaryota

Fungi

Eukaryota

Significant fold change > 1 Significant fold change < 1 Independent filter cut Outlier cut padj not available Total OTU number

25 (13.9) 14 (7.8) 692 30 722 902

9 (6.7) 4 (3.0) 107 51 158 293

19 (15.4) 7 (5.7) 335 72 407 530

3 (1.8) 11 (6.7) 707 30 737 902

0 (0) 0 (0) 0 51 51 293

1 (3.6) 5 (17.9) 430 72 502 530

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Fig. 3. Mean abundance of selected bacterial (A–D) and eukaryotic (E–H) OTUs in planktonic (white columns) and biofilm (black columns) communities. The abundance of an OTU has been calculated as a fraction of count of sequences belonging to this OTU within all sequences in the sample. „T” and „U” symbols in the column description indicate tilled and untilled soil, respectively. The numerals 1, 2 and 3 indicate the sampling site. Vertical error bars represent standard error of the mean, n = 4.

Table 1, only 2 were affected by tillage. Bacterial OTU Brevundimonas was significantly associated with tilled soils while eukaryotic OTU Mesorhabditis tended to prefer untilled ones. None of the abundant OTUs presented in Table 1 shows a significant interaction between life mode and tillage. As seen in Supplementary_data_S2.xlsx, this interaction was very rare and occurred only in the case of 1 prokaryotic (Ralstonia) and 2 eukaryotic OTUs (Rhabditis, Acanthamoeba). No significant interaction was observed for fungal OTUs.

family Rhizobiaceae, Bosea and Rhizobium, were also observed to associate with biofilm life mode (Table 1). The sequences belonging to the OTU Bosea (44 clusters) showed mostly 97% or higher similarity to GenBank hits LT934154, KJ878628, and MF525612 (Bosea sp.). Rhizobium OTU was composed of 52 clusters and the vast majority of sequences belonged to clusters with best GenBank best hits KY292475 (Rhizobium sp.) and MG461568 (Rhizobium giardinii), with sequence similarity values at least 98.6%. Legionella OTU was very rare in the biofilm and its abundance among planktonic bacteria (though relatively high) was burdened by high variation (Fig. 3C). This compound OTU was composed of 280 sequence clusters. It seems, however, that it is well defined. The most abundant clusters were highly similar to GenBank hits LN614827 (99%) and KU508792 (97.3%), belonging to Legionella falloni and Legionella neerlandica, respectively, and even the majority of less abundant clusters showed similarities to the members of the Legionella genus with the mean best hit similarity being equal to 94.3%. Only 10 rare clusters of this OTU showed the best sequence similarity to the medicinally most important species Legionella pneumophila. Other planktonic OTU, corresponding to the genus Aquicella (Fig. 3D), was represented by 106 sequence clusters with the similarity to GenBank best hit mostly not exceeding 95%. The vast majority of the clusters was most similar to GenBank sequence NR_025764 (Aquicella siphonis). Two examples of eukaryotic OTUs showing high fold change value indicating association with the biofilm were those corresponding to

3.5. Examples of responsive abundant OTUs The OTU showing the highest fold change among highly abundant bacterial OTUs is Arthrobacter (Fig. 3A). This OTU was composed of 69 sequence clusters similar mainly to GenBank best hits MG461455 (Arthrobacter pascens), MG196048, MG519282 and KF870415 (Arthrobacter sp.) with similarity of at least 99%. Other biofilm associated bacterial OTU, Geodermatophilus, tended to occur mainly in untilled soils (Fig. 3B). This OTU (composed of 17 sequence clusters) is well defined, mostly showing the similarity higher than 97% with best GenBank hit LT608342 (Geodermatophilus sp.). A highly abundant OTU significantly associated with biofilm life mode is that corresponding to the genus Pseudomonas (Table 1). This OTU was constituted by 202 sequence clusters, the 3 most abundant clusters showing highest similarity (at least 99%) to Genbank sequence MG516208 (Pseudomonas mohnii). Abundant representatives of the 7

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genera Telaepolella and Heteromita. OTU Telaepolella (Fig. 3E) is not equally distributed among the three soils, being particularly abundant in the biofilm of tilled soil 1 and very rare in the biofilm of the same untilled soil. This OTU (constituted by 53 clusters) was highly uniform, with almost all GenBank best hits being identical: KP86409 (Telaepolella sp.). The mean similarity was, however, a mere 87.6%. OTU Heteromita (24 clusters, Fig. 3F) was abundant in all the three soils and contained two large clusters constituting together 94% of all sequences of the OTU. The most abundant cluster was highly (98.6%) similar to GenBank best hit U42447 (Heteromita globosa) whereas the second most abundant cluster showed 99.5% similarity with best GenBank hit HM536169 (Heteromita sp.). The OTUs Mesorhabditis and Acrobeloides represent organisms associated with planktonic life mode. OTU Mesorhabditis (Fig. 3G), which was composed of 20 sequence clusters, showed the similarity - mostly higher than 98% - mainly to GenBank best hits EU040138 and KY119802. Both best hits are identified as Mesorhabditis sp. Acrobeloides OTU (35 sequence clusters, Fig. 3H) was similar to GenBank hits KX669638 (A. nanus) and EU543176 (A. thornei) with similarity mostly higher than 99%.

particles. These properties of the trap material can cause a selection effect acting during the biofilm formation and development (Renner and Weibel, 2011), which may result in establishment of biofilm microbial communities that are different, to some extent, from those formed in the soil. Also, the unnatural surfaces of the glass fibers may hypothetically represent less hospitable environment for biofilm forming microorganisms, compared to true soil particles, which may lead to a shift of their life strategy from the biofilm to planktonic one. Our results thus should not be taken as an exact description of microbial communities constituting soil biofilm and planktonic communities but rather as the approximative reflection of their existence. The development of biofilms is a type of microbial growth and, as such, it demands energy in the form of organic nutrition. At the same time, energy accessibility strongly affects the composition of the biofilm community composition and biomass accumulation (Martins et al., 2018; Truu et al., 2019). As the purpose of our work is to provide the view of soil biofilms that would be as realistic as possible, the energy availability in the soil surrounding the trap should correspond to that in the inside of the trap itself. From this viewpoint, the use of a planar filter as a trapping medium is advantageous. The thickness of the filter is 0.3 mm only, which enables easy penetration of diffusible organic compounds into the trap volume, even without taking into account the massive flux of the liquid between the trap volume and surrounding soil. Also, the particulate organic matter gets into close superficial contact with the filter and can affect the development of biofilm here. Thus, the energy availability in the trap and in surrounding soil should be comparable and should not heavily distort the results. Electron microscopic study of the biofilms developed on glass filters exposed to soil enabled us to distinguish bacterial cells attached to the glass surfaces or to surfaces of small clay particles that clung to the surface of the filter. The attachment seems to be secured by extracellular structures associated with bacterial cells. Though the reticular character of these structures suggests that they represent artifacts produced during the dehydration of the extracellular polymeric material, it is possible that a portion of the reticulate material represents nanowires, electroconductive bacterial appendages that might mediate unique cell-to-cell or cell-to-surface interactions (Reguera et al., 2005). Interestingly, the OTU corresponding to bacterial genus Geobacter, which contains the frequently studied nanowires producing organism Geobacter sulfurreducens (Sure et al., 2016), has been identified in some of our samples. Two bacterial OTUs, Geodermatophilus (biofilm inhabitant) and Legionella (planktonic) are demonstrative examples of organisms with two contrasting life modes as defined for the purposes of this work. With these OTUs, the fold change (biofilm versus planktonic life mode) reaches the value 25.7 and 0.064, respectively. Though the fold changes have been calculated from the sets of compositional data obtained from independent PCR reactions, the sequence counts in biofilm and planktonic fractions can be compared in a relative manner. Significant results of this comparison indicate, at least for the cases of prokaryotic and fungal OTUs, that the washing procedure is effective. Testing of the dispersion homogeneity showed a significant difference in this parameter within the data obtained using Eukaryota-specific PCR. This suggests that the eukaryotic communities are more heterogeneous than the other communities studied here. At the same time, this fact prevents us to make strong conclusions about the behaviour of eukaryotic OTUs in our study on the basis of the PERMANOVA output alone. Nevertheless, the separation of centroids corresponding to eukaryotic biofilm and planktonic samples in NMDS ordination diagram suggests that the communities are different. This notion is supported by the significant results of the downflow independent DESeq2 analysis of the life mode preferences of individual eukaryotic OTUs. A significant separation of OTUs with two different life mode preferences was observed in spite of the fact that quite feeble stream of the washing liquid was applied. This flux was powered just by gravity,

3.6. Relationships between eukaryotic/fungal and prokaryotic OTU abundances in biofilm communities In total, the τij parameter reached the threshold value of 0.8 in the case of 12 pairs of identified eukaryotic/bacterial OTUs and never exceeded the value of 0.9 (see the supplementary file “Supplementary_data_S1.xlsx”, sheet “Transformed variation matrix”). The highest τij value equal to 0.873 was observed for the Bosea/Colpoda OTU pair. Using this criterion, eukaryotic OTU Colpoda associated with 7 bacterial OTUs involved in the analysis and was the most active one in this regard. Further, OTU Heteromita was associated with 3 bacterial OTUs and OTU Cercomonas showed association with a single bacterial OTU. No association with bacterial OTUs was noted among fungal OTUs (i. e. the OTUs detected in amplicons produced in PCR with fungal specific primers). Comparing the fungal data obtained using Eukaryota-specific PCR and Fungi-specific PCR, the τij parameter reached the threshold value in a single case of the OTU pair Cylindrocarpon (Fungi-specific PCR)/Fusarium (Eukaryota-specific PCR). 4. Discussion 4.1. Effectivity of the washing procedure and trap functioning The washing of the porous biofilm carrier material (glass fibers in our case) cannot be, in principle, absolutely effective in separating planktonic organisms and biofilm inhabitants because some planktonic organisms, though fully motile, can be caught in dead end pores of the filter. On the other hand, pieces of the authentic biofilm could be snatched by liquid streaming during the washing procedure. We suppose that these effects would, to some extent, deteriorate the separation of planktonic and biofilm OTUs. We thus define the two fractions of organisms as enriched in planktonic organisms and as enriched in biofilm inhabitants. Consequently, the life mode preferences (biofilm versus planktonic) may be underestimated in our work. Not only the washing procedure itself but also the character of the trap material can influence the results (Hsu et al., 2013). Though the porous structure of the glass fiber filters used in our work provides microspaces where microbes can find the locations suitable for attachment, the dimensions of glass fibers and their spatial organization substantially differ from the microstructure of natural soils. Thus, the access of the air, as well as the streaming of water, are different there. Furthermore, the chemical composition of glass fibers that serve as a supporting material for developing biofilms is different from that of soil 8

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resembling the situation when the rainfall water infiltrates into the soil under natural conditions. This supports the hypothesis that bacteria are easily released and transported by soil solution movement from upper to deeper horizons, forming a depth-dependent development of bacterial communities with ecological and biogeochemical consequences (Lehmann et al., 2018).

aggregates with a diameter lower than 20 μm (Constancias et al., 2014). This further supports the character of Rhizobia as important inhabitants of soil biofilms. Legionella OTU was detected among the organisms that produce cells constituting an important fraction of planktonic microbial community. It is interesting that these bacteria may colonize existing biofilms and grow vigorously there (Rogers et al., 1994). This is probably due to their ability to parasitize protozoans, which commonly graze on biofilm communities (Greub and Raoult, 2004; Kuiper et al., 2004; Declerck et al., 2005, 2007). The formation of mono-species biofilms by members of this bacterial genus have been described as well (for review see Hilbi et al., 2011). Nevertheless, the very low abundance of this OTU in the biofilm community suggests that it does not massively parasitize sessile biofilm grazers in our soil samples and may be specialized to planktonic protists instead. Interestingly, the other OTU with notable affinity to planktonic life mode, Aquicella, corresponds to a bacterial genus that also infects protozoans (Santos et al., 2003). Eukaryotic OTU Telaepolella associated with biofilm life mode corresponds to a genus of Amoebozoa containing naked amoebae with fine pseudopodia. This genus is considered a group of aquatic organisms but was also reported from a rice field (Takenouchi et al., 2016). Similarly, the OTU Heteromita, corresponding to the genus of the same name which belongs to Cercozoa (Ekelund et al., 2004), was consistently very abundant in biofilm communities observed in all the three soils. A representative of this genus, Heteromita globosa, has been reported as a potent consumer of bacterial biomass clogging porous materials (Mattison et al., 2002). This species is probably identical with an important portion of organisms constituting OTU Heteromita in our work. It represents one of the most commonly recorded species of small flagellates that are capable of living in thin water layer on surfaces of soil particles (Nisbet, 2012, p. 44) where it may regulate the growth of biofilms. The eukaryotic OTUs showing significant association with planktonic life mode correspond to genera Acrobeloides and Mesorhabditis. These genera belong to Nematoda and their numerous members feed on bacteria (Bird and Ryder, 1993). As these organisms are common in soils, they may represent highly motile planktonic regulators of soil bacterial communities.

4.2. Biofilm inhabitants versus planktonic organisms Relatively low numbers of OTUs that are significantly associated with a biofilm life mode may indicate that a high number of the soil microbial taxa exist in the form of both biofilm and planktonic cells and do not necessarily need to prefer the biofilm life mode suggested by other authors (Costerton et al., 1987; Stoodley et al., 2002; Flemming et al., 2016) as the prevailing microbial life strategy. However, this notion should be taken with reservation regarding the limitation of our methodology that must be taken into account: the predisposition of organisms to choose between the two distinct life modes - biofilm versus planktonic - may be underestimated in our work (see above the discussion on the efficiency of the washing procedure) and, at the same time, the direct quantitative comparison of the abundance of an OTU in biofilm and in the planktonic community is not possible because of the sets of compositional data are compared. The production of extracellular polymeric substances is a prerequisite for stable biofilm formation. In accordance with this, the bacterial genus Pseudomonas, corresponding to the most abundant biofilm associated bacterial OTU detected in our samples, is known not only as a predominant bacterial biofilm forming genus but also as an efficient producer of extracellular polymers (Mann and Wozniak, 2012). Some members of the other abundant biofilm associated genus, Arthrobacter, have a significant ability to produce copious extracellular polymeric substances as well (Quintelas et al., 2011). Actinobacteria of the genus Geodermatophilus produce motile cells („zoospores“, Hayakawa et al., 2000). This indicates that they should be detectable in both biofilm and plankton. Though we observed this OTU mainly in biofilm, it was detectable in small amounts among planktonic organisms, suggesting a possible release of motile cells. These bacteria are members of the family Geodermatophilaceae that involves stone dwelling actinobacteria forming biofilms resisting harsh environmental conditions (Sghaier et al., 2016). The genus Geodermatophilus was also frequently detected in biofilms on highly superficially altered stones (Urzi et al., 2001), which suggests its role in mineral bedrock weathering. Bacterial genus Bosea (Rhizobiaceae) has been reported as a component of biofilms produced in reverse osmosis waste water cleaning device (Belgini et al., 2014). It was also detected as an abundant inhabitant of biological soil crusts (Liu et al., 2017). Both the above reports are consistent with the presence of the corresponding OTU in the biofilm in our samples. Similarly, the genus Rhizobium is known to form biofilms, though its attachment to the root and not to inert surfaces is mainly discussed (Rinaudi and Giordano, 2010). The ability of Rhizobium to colonize roots is probably closely connected with their ability to form biofilms (Fujishige et al., 2008). Free living planktonic bacteria can spread easier than taxa that live as components of biofilms. As a result, planktonic organisms should show the vertical stratification less marked than sessile biofilm inhabitants. Similarly, planktonic taxa should be distributed more homogeneously in soil environments. Yang et al. (2018) analyzed vertical distribution of bacterial taxa in three soil horizons till the depth of 55 cm and found a significant stratification. In accordance with our notion of Rhizobium OTU as a non-motile biofilm component, they found these bacteria almost exclusively in the middle horizon (depths 25–35 cm). In addition to this, large quantities of biofilm inhabiting bacteria should not be detectable in the finest soil particles because biofilms are larger structures containing many microbial cells. Interestingly, Rhizobium bacteria were almost absent from very small soil

4.3. Effect of soil tillage In general, tillage is reported as a significant factor affecting soil microbial communities (Mathew et al., 2012; Wang et al., 2012) but this is not reflected in our data. Among the highly abundant OTUs, only 2 were significantly affected by tillage. This indicates that the effect of tillage, as a factor affecting microbial communities, is much weaker than the effect of separation of biofilm and planktonic communities. It is suggested that tillage enlarges numerous ecological niches that are less available in untilled soils, as documented for example by the results of Constancias et al. (2014). The reason why we were not able to document this even for the bacterial group Planctomycetes, reported as highly responsive to soil management history and amendment (Buckley et al., 2006), is unknown. No significant simultaneous association of highly abundant OTUs with biofilm and tilled soil was observed. This does not support our hypothesis suggesting the stimulatory effect of tillage induced soil disturbance on the abundance of biofilm associated bacterial OTUs. It seems that the compensation of the effect of disturbance on the soil structure by stimulation of the biofilm constituting organisms (producing soil protective extracellular macromolecular matrix) either does not exist or is not detectable by the experimental method used in our study. 4.4. Potential trophic interactions In our work, we tried to detect signs of possible trophic interactions 9

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between biofilm prokaryotic community („lawn“) and protozoan grazers. Such interactions can be anticipated on the basis of already published data. However, the existence of an association between two organisms still does not necessarily indicate the trophic interaction and must be taken as an impetus for detailed analysis. Different grazers probably have different abilities to feed on a particular biofilm component (Seiler et al., 2017). This predicts some specificity in biofilm/ grazer interaction and, interestingly, stimulates the formation of the biofilm by the consumed fraction of the prokaryotic community. In addition, complex interactions between biofilm components may affect grazing intensity (Raghupathi et al., 2018). In our data, the interactions between biofilm bacteria and protozoan grazers were weak and infrequent. The OTU corresponding to eukaryotic genus Heteromita, found in biofilms in our soil samples, was associated with only 3 out of 23 different bacterial OTUs tested. This suggests the existence of a specific trophic interaction of this OTU with bacterial taxa that can serve as food. If the association really represents a sign of trophic interaction, the OTU corresponding to the genus Colpoda, associated with 7 bacterial OTUs, is probably more generalistic in its food preferences than Heteromita. This OTU was more abundant in plankton but was sufficiently frequent in biofilm samples to be included in the analysis of potential trophic interactions. It is thus possible that Colpoda individuals float in the soil solution seeking the prey, immerse into the biofilm whenever the prey is found and reduce the biofilm thickness by grazing (Huws et al., 2005). Further, our data indicate that several fungal OTUs are significantly more abundant in amplicons obtained from washed glass fiber filters, which could indicate a „biofilm life mode“. However, the filamentous character of the typical fungal thallus may result in its entanglement into the glass fiber filter without any other kind of attachment to the surfaces. The persistence of fungi in the washed filters thus does not necessarily mean that they constitute part of a biofilm. Nevertheless, these mycelia may represent physical surfaces that can be attractive for bacteria and may be colonized by bacterial biofilms, i. e. they can serve as biofilm bearers. The existence of fungal hyphae as novel hospitable microhabitats has been reported as a prerequisite to specific biofilm formation (Warmink and van Elsas, 2009). Several bacteria-fungi interactions have been reported in the literature, but mycorrhizal fungi mainly appeared as fungal organisms involved (Nazir et al., 2010). Warmink et al. (2009) showed the selective proliferation of bacteria in the proximity of mycelia (mycosphere) of different fungi and introduced the concept of fungiphilic bacteria (‘fungiphiles’), i.e. bacteria adapted to the exploitation of hyphal exudates as a carbon source. In this study, however, the existence of a perceptible association of bacterial groups with fungi was not confirmed. This might be due to the fact that fungi constituted only a minor part of eukaryotic biomass (in terms of sequence numbers) in our samples and potential association of bacterial OTUs with fungal hyphae might be hidden by natural variation in bacterial OTUs abundance in the field. It is possible that, should such an observation be performed under more homogeneous artificial conditions in microcosms, the association will be revealed.

biomass or rRNA gene copy numbers) of different microbial taxa between planktonic and biofilm communities, we were able to demonstrate that some microbial OTUs are detectable almost exclusively in planktonic communities. This indicates that living outside biofilm is not forbidden for soil microorganisms and that a significant portion of soil microbiota produces motile forms. Direct comparison of absolute abundances of motile and sessile forms of soil microorganisms would be highly desirable to reach conclusions of higher strength and precision. The tool that would enable this comparison is under development. For the purposes of our research, we used traps containing glass fibers as inert mechanical support of developing biofilms. It proved to be an excellent material that is sufficiently persistent in soil, can be easily manipulated in the laboratory and allows easy washing procedure. It is possible, however, that the character (roughness, chemical composition) of this material may show some selection effect to microorganisms during the development of biofilms. Our future work will thus involve the testing of the effects of different support materials on biofilm development in soil. We believe that the use of inert materials used as trap fillings represents a new and valuable tool for studies of soil biofilms. This tool, in combination with advanced molecular methods of the microbial community analysis, might provide new insight into the behaviour and diversity of soil microorganisms with potential practical consequences for soil management. Acknowledgements The work was financially supported by the Czech Science Foundation [grant number 17-09946S]. Highly valuable and insightful comments, kindly provided by the two anonymous peer reviewers and the editor, are gratefully acknowledged. The authors are further indebted to dr. K. Pranaw for language correction. Appendix A. Supplementary data Supplementary data to this article can be found online at https:// doi.org/10.1016/j.soilbio.2019.107535. References Aitchison, J., 1986. The Statistical Analysis of Compositional Data. Monographs on Statistics and Applied Probability (Reprinted in 2003). Chapman and Hall, London, pp. 416 (1986). Altschul, S.F., Gish, W., Miller, W., Myers, E.W., Lipman, D.J., 1990. Basic local alignment search tool. Journal of Molecular Biology 215, 403–410. Anderson, M.J., 2001. A new method for non-parametric multivariate analysis of variance. Austral Ecology 26, 32–46. Anderson, M.J., 2006. Distance-based tests for homogeneity of multivariate dispersions. Biometrics 62, 245–253. Apprill, A., McNally, S., Parsons, R., Weber, L., 2015. Minor revision to V4 region SSU rRNA 806R gene primer greatly increases detection of SAR11 bacterioplankton. Aquatic Microbial Ecology 75, 129–137. Belgini, D.R.B., Dias, R.S., Siqueira, V.M., Valadares, L.A.B., Albanese, J.M., Souza, R.S., Torres, A.P.R., Sousa, M.P., Silva, C.C., De Paula, S.O., Oliveira, V.M., 2014. Culturable bacterial diversity from a feed water of a reverse osmosis system, evaluation of biofilm formation and biocontrol using phages. World Journal of Microbiology and Biotechnology 30, 2689–2700. Bird, A.F., Ryder, M.H., 1993. Feeding of the nematode Acrobeloides nanus on bacteria. Journal of Nematology 25, 493–499. Buckley, D.H., Huangyutitham, V., Nelson, T.A., Rumberger, A., Thies, J.E., 2006. Diversity of Planctomycetes in soil in relation to soil history and environmental heterogeneity. Applied and Environmental Microbiology 72, 4522–4531. Burns, R.G., Stach, J.E.M., 2002. Microbial ecology of soil biofilms: substrate bioavailability, bioremediation and complexity. In: Violante, A., Huang, P.M., Bollag, J.M., Gianfreda, L. (Eds.), Soil Mineral-Organic Matter-Microorganism Interactions and Ecosystem Health. Developments in Soil Science 28/2. Elsevier Science, Amsterdam, pp. 17–42. Burns, R.G., DeForest, J.L., Marxsen, J., Sinsabaugh, R.L., Stromberger, M.E., Wallenstein, M.D., Weintraub, M.N., Zoppini, A., 2013. Soil enzymes in a changing environment: current knowledge and future directions. Soil Biology and Biochemistry 58, 216–234. Compant, S., Clément, C., Sessitsch, A., 2010. Plant growth-promoting bacteria in the rhizo- and endosphere of plants: their role, colonization, mechanisms involved and prospects for utilization. Soil Biology and Biochemistry 42, 669–678. Constancias, F., Prévost-Bouré, N.C., Terrat, S., Aussems, S., Nowak, V., Guillemin, J.P., Bonnotte, A., Biju-Duval, L., Navel, A., Martins, J.M.F., Maron, P.A., Ranjard, L.,

4.5. Conclusions and future prospects Our observations clearly indicate substantial differences in biofilm and planktonic microbial communities in soil. These two distinct communities contain organisms specialized to living in biofilm or to planktonic life mode but most probably also the organisms that exist in soil as sessile forms producing copious planktonic cells that can migrate and colonize new spaces. It is probable that the last mentioned life mode is very frequent in soils because many OTUs detected in our soils did not show significant association with biofilm or planktonic life mode. Though the currently available methodology does not allow precise quantitative comparison of abundances (in absolute amounts of 10

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columns. Applied and Environmental Microbiology 68, 4539–4545. Nazir, R., Warmink, J.A., Boersma, H., van Elsas, J.D., 2010. Mechanisms that promote bacterial fitness in fungal-affected soil microhabitats. FEMS Microbiology Ecology 71, 169–185. Nisbet, B., 2012. Nutrition and Feeding Strategies in Protozoa. Springer, Netherlands. Nunan, N., Wu, K.J., Young, I.M., Crawford, J.W., Ritz, K., 2003. Spatial distribution of bacterial communities and their relationships with the micro-architecture of soil. FEMS Microbiology Ecology 44, 203–215. Quintelas, C., da Silva, V.B., Silva, B., Figueiredo, H., Tavares, T., 2011. Optimization of production of extracellular polymeric substances by Arthrobacter viscosus and their interaction with a 13X zeolite for the biosorption of Cr(VI). Environmental Technology 32, 1541–1549. Raghupathi, P.K., Liu, W., Sabbe, K., Houf, K., Burmølle, M., Sørensen, S.J., 2018. Synergistic interactions within a multispecies biofilm enhance individual species protection against grazing by a pelagic protozoan. Frontiers in Microbiology 8, 2649. Redmile-Gordon, M.A., Brookes, P.C., Evershed, R.P., Goulding, K.W.T., Hirsch, P.R., 2014. Measuring the soil-microbial interface: extraction of extracellular polymeric substances (EPS) from soil biofilms. Soil Biology and Biochemistry 72, 163–171. Reguera, G., McCarthy, K.D., Mehta, T., Nicoll, J.S., Tuominen, M.T., Lovley, D.R., 2005. Extracellular electron transfer via microbial nanowires. Nature 435, 1098–1101. Renner, L.D., Weibel, D.B., 2011. Physicochemical regulation of biofilm formation. MRS Bulletin 36, 347–355. Rinaudi, L.V., Giordano, W., 2010. An integrated view of biofilm formation in rhizobia. FEMS Microbiology Letters 304, 1–11. Røder, H.L., Sørensen, S.J., Burmølle, M., 2016. Studying bacterial multispecies biofilms: where to start? Trends in Microbiology 24, 503–513. Rogers, J., Dowsett, A.B., Dennis, P.J., Lee, J.V., Keevil, C.W., 1994. Influence of temperature and plumbing material selection on biofilm formation and growth of Legionella pneumophila in a model potable water system containing complex microbial flora. Applied and Environmental Microbiology 5, 1585–1592. Santos, P., Pinhal, I., Rainey, F.A., Emphadinhas, N., Costa, J., Fields, B., Benson, R., Veríssimo, A., da Costa, M.S., 2003. Gamma-proteobacteria Aquicella lusitana gen. nov. sp. nov. and Aquicella siphonis sp. nov. infect protozoa and require activated charcoal for growth in laboratory media. Applied and Environmental Microbiology 69, 6533–6540. Seiler, C., van Velzen, E., Neu, T.R., Gaedke, U., Berendonk, T.U., Weitere, M., 2017. Grazing resistance of bacterial biofilms: a matter of predators' feeding trait. FEMS Microbiology Ecology 93, fix112. Sghaier, H., Hezbri, K., Ghodhbane-Gtari, F., Pujic, P., Sen, A., Daffonchio, D., Boudabous, A., Tisa, L.S., Klenk, H.-P., Armengaud, J., Normand, P., Gtari, M., 2016. Stonedwelling actinobacteria Blastococcus saxobsidens, Modestobacter marinus and Geodermatophilus obscurus proteogenomes. The ISME Journal 10, 21–29. Stoeck, T., Bass, D., Nebel, M., Christen, R., Jones, M.D., Breiner, H.W., Richards, T.A., 2010. Multiple marker parallel tag environmental DNA sequencing reveals a highly complex eukaryotic community in marine anoxic water. Molecular Ecology 19, 21–31. Stoodley, P., Davies, D.G., Costerton, J.W., 2002. Biofilms as complex differentiated communities. Annual Review of Microbiology 56, 187–209. Sure, S., Ackland, M.L., Torriero, A.A.J., Adholeya, A., Kochar, M., 2016. Microbial nanowires: an electrifying tale. Microbiology 162, 2017–2028. Takenouchi, Y., Iwasaki, K., Murase, J., 2016. Response of the protistan community of a rice field soil to different oxygen tensions. FEMS Microbiology Ecology 92, fiw104. Trabelsi, D., Mhamdi, R., 2013. Microbial inoculants and their impact on soil microbial communities: a review. BioMed Research International 863240, 1–11. Truu, M., Oopkaup, K., Krustok, I., Koiv-Vainik, M., Nolvak, H., Truu, J., 2019. Bacterial community activity and dynamics in the biofilm of an experimental hybrid wetland system treating greywater. Environmental Science and Pollution Research 26, 4013–4026. Urzi, C., Brusetti, L., Salamone, P., Sorlini, C., Stackebrandt, E., Daffonchio, D., 2001. Biodiversity of Geodermatophilaceae isolated from altered stones and monuments in the Mediterranean basin. Environmental Microbiology 3, 471–479. van de Voorde, T.F.J., van der Putten, W.H., Bezemer, T.M., 2012. Soil inoculation method determines the strength of plant–soil interactions. Soil Biology and Biochemistry 55, 1–6. Větrovský, T., Baldrian, P., 2013. Analysis of soil fungal communities by amplicon pyrosequencing: current approaches to data analysis and the introduction of the pipeline SEED. Biology and Fertility of Soils 49, 1027–1037. Wang, J.J., Li, X.Y., Zhu, A.N., Zhang, X.K., Zhang, H.W., Liang, W.J., 2012. Effects of tillage and residue management on soil microbial communities in North China. Plant, Soil and Environment 58, 28–33. Warmink, J.A., van Elsas, J.D., 2009. Migratory response of soil bacteria to Lyophyllum sp. strain Karsten in soil microcosms. Applied and Environmental Microbiology 75, 2820–2830. Warmink, J.A., Nazir, R., van Elsas, J.D., 2009. Universal and species-specific bacterial ‘fungiphiles’ in the mycospheres of different basidiomycetous fungi. Environmental Microbiology 11, 300–312. White, T., Bruns, T., Lee, S., Taylor, J., 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: Innis, M.A., Gelfand, D.H., Sninsky, J.J., White, T.J. (Eds.), PCR Protocols: A Guide to Methods and Applications. Academic Press, San Diego, pp. 315–322. Xu, Z., Shao, J., Li, B., Yan, X., Shen, Q., Zhang, R., 2013. Contribution of bacillomycin D in Bacillus amyloliquefaciens SQR9 to antifungal activity and biofilm formation. Applied and Environmental Microbiology 79, 808–815. Yang, J.K., Liang, J.F., Xiao, L.M., Yang, Y., Chao, Q.F., 2018. Vertical stratification of bacteria and the chemical compounds in crude oil-contaminated soil layers of the semi-deserted Dzungharian Basin. PLoS One 13, e0203919.

2014. Microscale evidence for a high decrease of soil bacterial density and diversity by cropping. Agronomy for Sustainable Development 34, 831–840. Costa, O.Y.A., Raaijmakers, J.M., Kuramae, E.E., 2018. Microbial extracellular polymeric substances: ecological function and impact on soil aggregation. Frontiers in Microbiology 9, 1639. Costerton, J.W., Cheng, K.J., Geesey, G.G., Ladd, T.I., Nickel, J.C., Dasgupta, M., Marrie, T.J., 1987. Bacterial biofilms in nature and disease. Annual Review of Microbiology 41, 435–464. Costerton, J.W., Lewandowski, Z., Caldwell, D.E., Korber, D.R., Lappin-Scott, H.M., 1995. Microbial biofilms. Annual Review of Microbiology 49, 711–745. Declerck, P., Behets, J., Delaedt, Y., Margineanu, A., Lammertyn, E., Ollevier, F., 2005. Impact of non‐Legionella bacteria on the uptake and intracellular replication of Legionella pneumophila in Acanthamoeba castellanii and Naegleria lovaniensis. Microbial Ecology 50, 536–549. Declerck, P., Behets, J., Van Hoef, V., Ollevier, F., 2007. Detection of Legionella spp. and some of its amoeba hosts in floating biofilms from anthropogenic and natural aquatic environments. Water Research 41, 3159–3167. Edgar, R.C., Flyvbjerg, H., 2015. Error filtering, pair assembly and error correction for next-generation sequencing reads. Bioinformatics 31, 3476–3482. Ekelund, F., Daugbjerg, N., Fredslund, L., 2004. Phylogeny of Heteromita, Cercomonas and Thaumatomonas based on SSU rDNA sequences, including the description of Neocercomonas jutlandica sp. nov., gen nov. European Journal of Protistology 40, 119–135. Flemming, H.C., Wingender, J., Szewzyk, U., Steinberg, P., Rice, S.A., Kjelleberg, S., 2016. Biofilms: an emergent form of bacterial life. Nature Reviews Microbiology 14, 563–575. Fujishige, N.A., Lum, M.R., De Hoff, P.L., Whitelegge, J.P., Faull, K.F., Hirsch, A.M., 2008. Rhizobium common nod genes are required for biofilm formation. Molecular Microbiology 67, 504–515. Greub, G., Raoult, D., 2004. Micro‐organisms resistant to free‐living amoebae. Clinical Microbiology Reviews 17, 413–433. Gryndler, M., Černá, L., Bukovská, P., Hršelová, H., Jansa, J., 2014. Tuber aestivum association with non-host roots. Mycorrhiza 24, 603–610. Hayakawa, M., Otoguro, M., Takeuchi, T., Yamazaki, T., Iimura, Y., 2000. Application of a method incorporating differential centrifugation for selective isolation of motile actinomycetes in soil and plant litter. Antonie van Leeuvenhoek International Journal of General and Molecular Microbiology 78, 171–185. Hilbi, H., Hoffmann, C., Harrisson, C.F., 2011. Legionella spp. outdoors: colonization, communication and persistence. Environmental Microbiology Reports 3, 286–296. Hsu, L.C., Fang, J., Borca-Tasciuc, D.A., Worobo, R.W., Moraru, C.I., 2013. Effect of micro- and nanoscale topography on the adhesion of bacterial cells to solid surfaces. Applied and Environmental Microbiology 79, 2703–2712. Huws, S.A., McBain, A.J., Gilbert, P., 2005. Protozoan grazing and its impact upon population dynamics in biofilm communities. Journal of Applied Microbiology 98, 238–244. Ihrmark, K., Bödeker, I.T.M., Cruz-Martinez, K., Friberg, H., Kubartova, A., Schenck, J., Strid, Y., Stenlid, J., Brandström-Durling, M., Clemmensen, K.E., Lindahl, B.D., 2012. New primers to amplify the fungal ITS2 region – evaluation by 454-sequencing of artificial and natural communities. FEMS Microbiology Ecology 82, 666–677. Kasim, W.A., Gaafar, R.M., Abou-Ali, R.M., Omar, M.N., Hewait, H.M., 2016. Effect of biofilm forming plant growth promoting rhizobacteria on salinity tolerance in barley. Annals of Agricultural Sciences 61, 217–227. Koczan, J.M., McGrath, M.J., Zhao, Y., Sundin, G.W., 2009. Contribution of Erwinia amylovora exopolysaccharides amylovoran and levan to biofilm formation: implications in pathogenicity. Phytopathology 99, 1237–1244. Kuiper, M.W., Wullings, B.A., Akkermans, A.D., Beumer, R.R., Van Der Kooij, D., 2004. Intracellular proliferation of Legionella pneumophila in Hartmannella vermiformis in aquatic biofilms grown on plasticized polyvinyl chloride. Applied and Environmental Microbiology 70, 6826–6833. Lehmann, K., Schaefer, S., Babin, D., Kohne, J.M., Schluter, S., Smalla, K., Vogel, H.J., Totsche, K.U., 2018. Selective transport and retention of organic matter and bacteria shapes initial pedogenesis in artificial soil - a two-layer column study. Geoderma 325, 37–48. Lejzerowicz, F., Esling, P., Pillet, L., Wilding, T.A., Black, K.D., Pawlowski, J., 2015. Highthroughput sequencing and morphology perform equally well for benthic monitoring of marine ecosystems. Scientific Reports 5, 13932. Lentz, R.D., 2015. Polyacrylamide and biopolymer effects on flocculation, aggregate stability, and seepage in a silt loam. Geoderma 241–242, 289–294. Liu, L., Liu, Y., Zhang, P., Song, G., Hui, R., Wang, Z., Wang, J., 2017. Development of bacterial communities in biological soil crusts along a revegetation chronosequence in the Tengger Desert, northwest China. Biogeosciences 14, 3801–3814. Love, M.I., Huber, W., Anders, S., 2014. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biology 15, 550. Mahé, F., Mayor, J., Bunge, J., Chi, J., Siemensmeyer, T., Stoeck, T., Wahl, B., Paprotka, T., Filker, S., Dunthorn, M., 2014. Comparing high-throughput platforms for sequencing the V4 region of SSU-rDNA in environmental microbial eukaryotic diversity surveys. Journal of Eukaryotic Microbiology 62, 338–345. Mann, E.E., Wozniak, D.J., 2012. Pseudomonas biofilm matrix composition and niche biology. FEMS Microbiology Reviews 36, 893–916. Martins, S.J., Medeiros, F.H.V., Lakshmanan, V., Bais, H.P., 2018. Impact of seed exudates on growth and biofilm formation of Bacillus amyloliquefaciens ALB629 in comon bean. Frontiers in Microbiology 8, 2631. Mathew, R.P., Feng, Y., Githinji, L., Ankumah, R., Balkcom, K.S., 2012. Impact of notillage and conventional tillage systems on soil microbial communities. Applied and Environmental Soil Science 2012, 548620. Mattison, R.G., Taki, H., Harayama, S., 2002. The bacterivorous soil flagellate Heteromita globosa reduces bacterial clogging under denitrifying conditions in sand-filled aquifer

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