Ectomycorrhizal fungi live in close association with their host plants and form complex interactions with bacterial/archaeal communities in soil. We investigated whether abundant or rare ectomycorrhizal fungi on root-tips of young beech trees (Fagus sylvatica) shape bacterial/archaeal communities. We sequenced 16S rRNA genes and fungal internal transcribed spacer regions of individual root-tips and used ecological networks to detect the tendency of certain assemblies of fungal and bacterial/archaeal taxa to inhabit the same root-tip (i.e. modularity). Individual ectomycorrhizal root-tips hosted distinct fungal communities associated with unique bacterial/archaeal communities. The structure of the fungal-bacterial/archaeal association was determined by both, dominant and rare fungi. Integrating our data in a conceptual framework suggests that the effect of rare fungi on the bacterial/archaeal communities of ectomycorrhizal root-tips contributes to assemblages of bacteria/archaea on root-tips. This highlights the potential impact of complex fine-scale interactions between root-tip associated fungi and other soil microorganisms for the ectomycorrhizal symbiosis.
The mycorrhizal symbiosis is an ancient association between plants and soil fungi, which facilitates the growth of both partners by transferring benefits, such as reciprocal nutrient acquisition, increased stress resistance and protection from pathogens1,2. Fungal hyphae emanating from mycorrhizal root-tips expand the surface area of the host root system for enhanced nutrient uptake and potential tripartite interactions with other soil microorganisms3. The mycorrhizosphere, i.e., the soil in direct contact with mycorrhizal hyphae influenced by root and hyphal exudates, provides an attractive habitat for soil bacteria and archaea. The presence of readily available carbon sources enhances fungal and bacterial abundance, diversity and their interplay4,5. However, interactions taking place at fine-scale resolution remain unclear.
Ectomycorrhizal fungi form symbioses mainly with trees in temperate and boreal forests. Numerous fungal species coexist on single host plants6,7,8,9,10,11,12,13,14, such as 80–90 fungal taxa on roots of old-growth Fagus sylvatica trees15. The diversity of ectomycorrhizal fungi colonizing a plant is reflected in the morphological diversity of ectomycorrhizal root-tips, which differ in shape, color and other visual properties. Based on the assumption that the fungal tissue of one ectomycorrhizal root-tip is formed by one fungal species, morphological properties have been used, supported by microscopy and catalogued information, to identify the fungal species involved in its formation16,17. Studies investigating ectomycorrhizal fungi via sequencing of the internal transcribed spacer (ITS) region, however, often found more than one ectomycorrhizal fungal species present on mycorrhizal root-tips10,12,13,18,19,20,21. This raises the question if and how the additionally present ectomycorrhizal species may contribute to the formation of a specific morphotype and to its functionality. As fungal-specific root-tip morphologies create special microenvironments for soil organisms such as bacteria or archaea5,22,23, the fungi building an ectomycorrhizal root-tip may influence the associated bacterial and archaeal communities.
The development of ectomycorrhizal root-tips is complex and the fungal inoculation of the root initiates the formation of the fungal mantle which can precede the association of bacteria24. The ectomycorrhizal mantle surface of Fagus sylvatica harbors high numbers of bacteria25, and the thin interface between the host root and adhered fungal mantle is considered to represent a particular hotspot for bacterial colonization26. Many studies demonstrated that mycorrhizosphere-associated microbial communities can vary according to their host plants, the present soil and environmental conditions21,25,27,28,29,30,31. A rare number of studies, however, investigated fungal and/or bacterial communities at the level of individual ectomycorrhizal root-tips18,31,32 and none has, to the best of our knowledge, explored the complex linkages between the diverse fungal and bacterial/archaeal communities associated with single root-tips. Interesting attempts have been made in this direction in the studies by Marupakula33,34, who showed that the dominant fungal species on root-tips of Pinus sylvestris were associated with distinct bacterial communities33. Until now, however, research has mainly focused on the dominant fungal species on ectomycorrhizal root-tips, which is likely responsible for morphotype forming. As such, the influence of less abundant or even rare members of the fungal root-tip communities on bacterial associations remains unknown.
This study focused on microbial colonization patterns of ectomycorrhizal root-tips, and particularly on the question whether diverse ectomycorrhizal fungal communities influence the association of bacteria/archaea. More precisely, we hypothesized that the whole fungal community, rather than only the dominant ectomycorrhizal fungi is responsible for patterns of bacterial/archaeal communities across root-tips. To test this we used modularity, a network property that determines the presence of assemblies of bacterial/archaeal taxa that tend to inhabit the same root-tip as a given set of fungal taxa. No changes in modular structure among networks that gradually consider less abundant fungal taxa would suggest that mainly the highly abundant fungi shape the assembly of bacterial/archaeal communities on the root-tips, while gradually contrasting network structures would suggest that the consideration of rare fungi generate co-occurrence patterns, and thus also contribute to shaping the bacterial assemblies. We sequenced the 16S rRNA gene and the ITS1 region from 62 individual mycorrhizal root-tips of 13 young European Beech (Fagus sylvatica) trees, in addition to corresponding rhizosphere soil and root-distant soil (bulk soil). We aimed to get a comprehensive picture of the dominant and rare fungal community members on individual root-tips and to assess the associated bacterial/archaeal communities. By investigating which fungal-bacterial/archaeal taxa tend to inhabit the same mycorrhizal root-tip, we assessed tendencies in the associations of fungal and bacterial/archaeal communities. In combination with a unique conceptional framework, this study provides insights into the fungal-bacterial interplay on individual root-tips.
Screening roots of the 13 sampled trees confirmed that 80–90% of root-tips in each root system were mycorrhized (see Gorka et al.35 for details). Mycorrhizal root-tips of sampled Fagus sylvatica roots showed a variety of fungal morphological appearances, differing in color, branching pattern and extensive mycelium (Fig. 1, a1–6). In a pre-experiment, we also confirmed bacterial colonization on the ectomycorrhizal fungal mantle (built from ectomycorrhizal fungal hyphae surrounding the root-tips) of Fagus sylvatica root-tips via scanning electron microscopy (Fig. 1, section b).
Fagus sylvatica ectomycorrhizal root-tips harbor specific fungal OTUs and many unique bacterial/archaeal OTUs
Interestingly, among all investigated habitats (rhizosphere, bulk soil and mycorrhizal root-tips), mycorrhizal root-tips harbored most of the unique bacterial/archaeal OTUs (1000), representing 15% of all OTUs (Fig. 2a). In contrast, bulk soil harbored most of the unique fungal OTUs (227), accounting for 17% of all OTUs (Fig. 2b).
Analyzing the 100 most abundant shared and unique fungal and bacterial/archaeal OTUs across the three investigated habitats revealed that a large group of OTUs was solely present on mycorrhizal root-tips, while bulk and rhizosphere shared OTUs (Fig. 2c, e). Within the 100 most abundant bacterial/archaeal OTUs, OTUs almost entirely (to 90-100%) associated with mycorrhizal root-tip habitats belonged to members of bacterial orders Rhizobiales, Streptomycetes and Burkholderiales. Within the 100 most abundant fungal OTUs, OTUs with higher relative abundances on mycorrhizal root-tips than in bulk soil and rhizosphere habitats belonged to fungal orders Thelephorales, Sebacinales, Pezizales, Agaricales. (Fig. 2c, e). As expected, ectomycorrhizal and other symbiotrophic fungi tended to be associated more with mycorrhizal root-tips than with the other habitats, whereas saprotrophic and pathotrophic fungi tended to occur more in bulk soil and rhizosphere habitats (Fig. 2d). However, a large part of the fungal taxa found on root-tips was also found in the soil (Fig. 2b, d). When considering all OTUs (Supplementary Fig. 1), the patterns of OTU distribution were similar to the ones observed for the 100 most abundant OTUs. Fungal and bacterial/archaeal communities on mycorrhizal root-tips showed clear differences in their taxonomic composition compared to bulk and rhizosphere-inhabiting communities (Supplementary Figure 2). Moreover, the fungal and bacterial/archaeal community compositions on mycorrhizal root-tips were significantly less diverse (Shannon diversity index; fungal dataset: ANOVA, Tukey HSD, p < 0.001; bacteria/archaea: Dunn test, Kruskal-Wallis multiple comparison, p < 0.001; Supplementary Fig. 3) and significantly different from bulk and rhizosphere soil (Bray-Curtis dissimilarity; PERMANOVA, p = 1e−04; Supplementary Fig. 3). However, dispersion between groups was not homogenous (ANOVA, F = 21.99, p > 0.001), most likely due to the high group variance dispersion in mycorrhizal root-tip samples (Permdist, p = 1e−04; Supplementary Fig. 4).
Mycorrhizal root-tips were characterized by one to three dominating fungal OTUs and a more even bacterial/archaeal distribution
Most root-tips fungal communities were dominated by a few fungal OTUs, which constituted more than 50% of the relative read abundance. Most mycorrhizal root-tips harbored one to three dominant OTUs with relative abundance ranging from 45% up to 97% and many rare fungi (relative read abundance <5%, on average 904 OTUs per root-tip) (Fig. 3). OTUs that dominated the community of individual root-tips were mostly associated with the fungal orders Thelephorales, Agaricales, Pezizales and Sebacinales (Fig. 3). The dendrogram demonstrates that the most similar fungal communities were derived from root-tips stemming from different plants.
We identified 19 highly abundant fungal OTUs (relative abundance >50% on individual root-tips), which likely contribute to the formation of the mantle tissue and the morphological structure of each individual root-tip (labeled OTUs in Supplementary Fig. 5). They consisted mainly of ectomycorrhizal members of Thelephorales, Pezizales, Sebacinales and Agaricales (see Supplementary Table 2 for detailed taxonomic classification). Seven of these OTUs (OTU_1, 2, 4, 7, 11, 15, 23), encompassing members of the genera Inocybe, Tomentella, Peziza and Laccaria, were prevalent in more than 90% of all root-tips and thus part of the shared fungal community among all investigated root-tips (see labeled OTUs in Supplementary Fig. 5).
The bacterial/archaeal community composition was more even among the investigated root-tips (Supplementary Fig. 2), containing less dominant OTUs (relative average abundance of 8% across all root-tips) that were mainly members of Proteobacteria and Actinobacteria. Reads assigned to Proteobacteria represented more than 50% on 35 root-tips, specifically Alpha- and Gammaproteobacteria, while reads assigned to Actinobacteria were predominant on 23 root-tips and represented more than 50% of the reads in 16 samples. Reads assigned to Acidomicrobia, Verrucomicrobiae, Bacteroidia, Chloroflexia, Cyanobacteria, Planctomycetes, Firmicutes and Gemmatimonadetes were less abundant, but present on the majority of the sequenced mycorrhizal root-tips (Supplementary Fig. 2). When analyzing all possible combinations of shared fungal or bacterial/archaeal OTUs across the investigated mycorrhizal root-tip samples, we were able to identify four fungal OTUs (belonging to fungal orders Thelephorales and Agaricales) and 15 bacterial OTUs (belonging to bacterial orders Bacillales, Burkholderiales, Chthoniobacterales, Corynebacteriales, Frankiales and Rhizobiales) that co-occurred on all investigated mycorrhizal root-tips (Supplementary Fig. 6, see Supplementary Table 3 for detailed taxonomic classification of co-existing fungal and bacterial OTUs).
Bacterial/archaeal communities on ectomycorrhizal root-tips are shaped by the entire fungal community including less abundant fungi
The non-random community structure on ectomycorrhizal root-tips indicates a link between the distribution of associated fungi and bacteria/archaea. This was shown by the significant difference of the observed modularity in most bipartite networks when compared to their respective null models (Table 1). In addition, we saw a gradual increase of significance in modularity when comparing network “10%” (restricted network including only fungal OTUs with >10% relative abundance in individual samples) to network “0.1” (most complete network including fungal OTUs with relative abundance >0.1% in individual samples; Tables 1 and 2). The individual networks encompassed various numbers of modules, each of which consisted of different fungal and bacterial OTUs (Table 2).
Network “10%” was significantly modular (z = 56.52) compared to 100 randomized networks serving as a null model and consisted of four modules comprising 40 fungal OTUs (see Supplementary Table 4 for detailed taxonomic classification). Similarly, network “5%” was highly modular (z = 95.47) and was divided into six modules encompassing 54 fungal OTUs (see Supplementary Table 5 for detailed taxonomic classification). The significance of modularity peaked in network “0.1%” (z = 851.24), which included less abundant fungal OTUs with >0.1% relative abundance (in total 416 fungal OTUs; Tables 1, 2). Within all three networks, each module differed in its fungal and bacterial composition (Fig. 4). While some modules were dominated by ectomycorrhizal members of Agaricales, Sebacinales, Pezizales, Russulales, Thelephorales, other modules showed a more diverse distribution of various members of ectomycorrhizal and non-ectomycorrhizal fungal guilds (Fig. 4). Bacterial community composition was dominated by members of Proteobacteria, Acidobacteria, Actinobacteria, Bacteroidota and Planctomycetota, but their relative abundances changed across the different modules of each network (Fig. 4).
To assess whether the whole fungal community, and not only the most abundant ectomycorrhizal fungi, shape the bacterial/archaeal communities present on each root-tip, we evaluated the consistency of modules’ composition across networks with different fungal OTU abundance cut-offs. We compared the bacterial/archaeal and fungal taxa assignments into modules between network “10%” and network “0.1%”, as well as network “5%” and network “0.1%” (Fig. 5).
Including rare fungi altered the structure of the network allowing the emergence of specific bacterial/archaeal associations with rare fungi. We observed two patterns in the structure of the networks that explain the effect of rare fungi on the fungal-bacterial/archaeal assemblies. We show statistical evidence for those patterns that relate to two non-exclusive ecological processes. First, comparing the module patterns across networks showed that network “0.1%” (comprising also rare fungi >0.1% relative abundance) contained two modules (M1, M4) solely composed of rare fungal taxa (with a relative abundance of less than 5% and more than 0.1%), which were not included in the restricted network “10%” or “5%” (consisting of only abundant fungal OTUs with relative abundance >10% and >5%, respectively; Fig. 5). This pattern was clearly visible in the comparison with network “10%” (Fig. 5a) and less strong, but still distinct in the comparison with network “5%” (Fig. 5c, M1 and M4 of network “0.1%” contain only a few abundant fungi and mostly rare fungi). Second, including the rare fungi also changed which abundant fungi tended to be associated to the same bacteria/archaea. Comparing networks revealed that highly abundant fungal taxa change their module’s partners when also considering rare fungi; taxa belonging to the same module in the restricted networks did not share the same module in the most complete network (network “0.1%”). The proportion of highly abundant fungal taxa of networks “10%” and “5%” (>10% and >5% relative abundance) grouping together within the same module in the compared network “0.1%” was significantly smaller than 1 for both comparisons, network “10%” and “0.1%” (mean = 0.28, t = −7.99, df = 6, p value < 0.001) and network “5%” and “0.1%” (network “5%”; mean= 0.20, t = −13.97, df = 13, p value < 0.001; a proportion of 1 would indicate that 100% of the fungal taxa belonging to the same module in one network also share a module in the other network).
The bacterial/archaeal taxa were distributed across all modules of both network pairs, although a bigger proportion of these taxa was shared with M2 and M3 of network “5%” and “10%” (Fig. 5b, d) in comparison with network “0.1%”.
In this study we showed that the unique microenvironment of established Fagus sylvatica ectomycorrhizal root-tips harbors specific microbial communities with particular co-occurrence patterns. Our data suggest that mycorrhizal root-tips host diverse fungal communities as well as unique bacteria/archaea. These specific bacterial/archaeal communities associated to mycorrhizal root-tips are selected by the fungal community present on each root-tip. Furthermore, our results demonstrate that not only the most abundant fungi, but also less abundant fungi are associated with the bacterial/archaeal community within a root-tip and alter the structure of the bipartite network.
Each mycorrhizal root-tip was inhabited by one to three dominant ectomycorrhizal OTUs, which comprised more than 50% of the reads. It is likely, that the fungal mantle tissue of root-tips is mostly formed by those highly abundant fungi, which subsequently shape the morphological structure of each individual root-tip. As such, they constitute preferable microhabitats for bacterial/archaeal, as well as (lower abundant) fungal colonization.
We observed vastly differing fungal communities amongst single root-tips of the same plant, which can be explained by competition, including priority effects, facilitation or parasitism among species10,36,37,38. In particular, positive interactions and competition between ectomycorrhizal fungal species determine root colonization39. Especially multi-host ectomycorrhizal fungi colonizing different host trees, such as members of the genera Inocybe, Cenoccocum, Laccaria and Russulales, are known to be strong competitors17,36. They were also among the most prevalent fungi in our system, which confirms their consistent presence across the investigated root-tips.
Based on scanning electron microscopy (Fig. 1, section b), we detected dense bacterial colonization on the ectomycorrhizal mantle surface of Fagus sylvatica, which is in accordance with previous research25. Mycorrhizal fungi themselves exude carbon compounds and their varying quality and quantity influences mycorrhizosphere bacterial/archaeal community composition33,40,41,42,43. In addition, root exudates can be altered by fungi, leading to quantitative and qualitative changes in root exudates between rhizosphere and mycorrhizosphere, selectively shaping the associated microbial communities, the so called mycorrhizosphere effect4,27,44. The significant difference in community richness, evenness and diversity between mycorrhizal root-tip and soil-related bacterial/archaeal communities indicates a strong mycorrhizosphere effect in our system4. However, since the dispersion among group was not homogenous, the results must be interpreted with care.
Dominant bacteria on the root-tips, such as members of Burkholderiales, Streptomycetales and Rhizobiales were previously found in ectomycorrhizal systems: Burkholderiales, the most abundant colonizers of soil fungi45, have been detected on mycorrhizal root-tips29,33,46,47 and endophytes of Fagus sylvatica25. Members of Rhizobiales have been found on ectomycorrhizal root-tips of Pinus muricata46, and peridia of E. granulatus and truffles have been found to provide a suitable niche for Bradyrhizobia48,49,50. Many bacterial strains have been reported to enhance ectomycorrhizal formation45,51,52 and improve plant nutrition and health. Members of Streptomycetales, for instance, are known as important symbiosis modulators by inhibiting growth of pathogenic fungi and promoting ectomycorrhizal growth formation53. In crop plant systems, such as soybean roots, rice roots or sugarcane rhizosphere, it has been reported that Burkholderiales, Streptomycetales and Rhizobiales act as plant growth promoting bacteria and are the core-responsive bacteria under drought conditions, mediating drought tolerance54,55,56. However there is still a lack of knowledge about the role of non-easily culturable bacteria living in the mycorrhizosphere.
The large group of unique bacterial/archaeal OTUs present on mycorrhizal root-tips indicates that ectomycorrhizal fungi create preferred microhabitats for distinct bacteria/archaea26. The non-random community structure of the investigated mycorrhizal root-tips suggests that they are determined by the fungal taxa present. Fungal hyphae can select for certain bacteria and determine their community composition57,58 and vice-versa, mycorrhizosphere bacteria can act in a fungi-specific manner and stimulate mycorrhizal formation in combination with certain fungi, but inhibit the symbiosis with others53,59.
The colonization of mycorrhizal root-tips is complex, triggered by a multitude of signaling pathways in the symbiotic interfaces of host and fungi, preceding fungal inoculation of the roots24,60. Bacterial colonization follows the formation of the initial fungal mantle. By analyzing the fungal patterns in module assignments among the different networks, we identified potential underlying processes contributing to shifts in the structure of co-occurring bacterial/archaeal communities due to fungal-bacterial/archaeal interactions. We showed that not only highly abundant, but also rare fungal OTUs shape the fungal-bacterial/archaeal interactions, highlighting the importance of comprehensive analysis of fungal communities on individual root-tips, that is not based solely on the characterization of dominant fungi. However, we do not claim to have covered the entire rare fungal community associated to mycorrhizal root-tips, as this is dependent on factors such as the used extraction method and sequencing depth. Based on the distribution of abundant and rare fungi among the modules (Fig. 5), we propose a framework (Fig. 6) to assess two non-exclusive processes contributing to shifts in the structure of the resulting network when all fungi (including the rare ones) are considered compared to only the abundant ones. The consideration of rare fungi can result in (a) the emergence of specific bacterial/archaeal associations to the rare fungi, and (b) changes in the similarity patterns of bacterial/archaeal associations between the highly abundant fungi. On the one hand, if changes are induced by the emergence of specific bacterial/archaeal associations to rare fungi, we expect these fungi to be grouped into new modules in the most complete network (including all fungi) (Fig. 6, panels 1.1 or 1.2). On the other hand, if changes are induced by a split of the group of bacteria/archaea that were previously associated with the same highly abundant fungi, we expect that abundant fungi from the same module in the more restricted network would not be grouped together in the most complete network (Fig. 6, panels 1.1 or 2.1). These two processes can affect the shifts in the network either independently or simultaneously. If neither of those processes occur, shifts between the networks will be minimized. No changes between the networks structure will occur when rare fungi are evenly distributed across modules in the most complete network and highly abundant fungi keep sharing the same module (Fig. 6, panel 2.2).
Based on our network analysis we saw significant patterns in both outlined processes (Fig. 6, panel 1.1). Including rare members of the fungal community resulted in newly emerging modules, exclusively containing rare fungi. Furthermore, highly abundant fungi did not share the same module when rare fungi were considered (Fig. 5).
The generation of new modules harboring only rare fungi (Fig. 5a, c) suggests that these low abundant fungi could facilitate novel bacterial/archaeal assemblages. In addition, the uneven distribution of highly abundant fungi among the different modules of the most complete network suggests that the association patterns between bacteria and some highly abundant fungi are weak, and they might tend to similarly associate to other highly abundant fungi when more detailed information about their association patterns is included in the network by considering the presence of rare fungi.
In summary, our network analysis demonstrates that rare fungi can alter the potential for microbe-microbe interactions, leading to new fungal-bacterial community assemblies. Our study further emphasizes the importance of focusing on small-scale fungal-bacterial/archaeal co-occurrences among individual mycorrhizal root-tips, as it holds the potential to deepen our understanding of the complex interplay of the symbiotic partners in the widespread and vital tripartite ectomycorrhizal symbiosis.
Approximately three- to four-year-old Fagus sylvatica trees were sampled from a beech forest in Klausen-Leopoldsdorf (Austria). Based on stereoscopic examination of selected plant roots, all trees showed abundant ectomycorrhization at the time of replanting. Specifically, twenty-seven trees were transferred into individual split-root boxes (as this study was part of a bigger experiment, see detailed information on experimental setup in previous studies using the same plants35,61). The trees were planted in a mixture of soil (collected from the tree sampling site, A-horizon, 4 mm sieved) and perlite (soil:perlite (8:1, v/v)35,61 and grown in a greenhouse under ambient (outdoor) sunlight and temperature for a year. After 12 months of plant growth, 13 plants were sampled for this study (Supplementary Table 1). During the plant harvest, boxes were carefully taken apart. Plant root systems were gently shaken, soil falling off, together with the soil that remained in the box, was 2 mm sieved and declared bulk soil. Root pieces were cut off and shaken in 1× Phosphate-buffered saline (PBS, pH = 8) for 30 min. After root pieces were transferred into new tubes, the remaining soil slurry was centrifuged (8000× g, 4 min), the supernatant discarded, and the pellet considered rhizosphere soil.
To obtain mycorrhizal root-tips (about 1-3 mm long terminal ends of the individual roots), roots were washed in tap water, the root system split into at least 6 parts and screened for total mycorrhization using stereoscopes35,61 (see an example for one of the root parts in Supplementary Material Fig. S2 in Gorka et al.35). Mycorrhization (i. e. the percentage of plant root-tips colonized by ectomycorrhizal fungi, estimates on average 85%) was similar across all investigated root systems. Individual mycorrhizal root-tips (for an example, see Supporting Information Fig. S1 in Mayerhofer et al.61) were sampled from different root parts. All samples were frozen at −80 °C until further processing.
Total nucleic acid extraction
Total nucleic acids were extracted according to Angel et al.62, using a mechanical and chemical lysing approach (as proposed in63 by combining bead-beating with phenol-chloroform extraction. 0.4 g of bulk soil and rhizosphere soil were used. For mycorrhizal root-tips (directly weighed into bead-beating tubes), the extraction protocol was slightly adapted by performing four bead-beating steps and by subsequently transferring only the aqueous phase without the phenolic phase. Bulk and rhizosphere samples were resuspended in 200 µl Low TE Buffer and mycorrhizal root-tips in 30 µl. Immediately after extraction, humic substances were removed with the OneStepTMPCR Inhibitor Removal Kit (Zymo Research, Irvine, CA, USA). DNA was quantified by using the Quant-iTTM PicoGreen®dsDNA assay (ThermoFisher Scientific, Waltham, MA, USA).
Sample preparation for identification of microbial and fungal communities via MiSeq Illumina sequencing
A multiplexed barcoded amplicon sequencing approach64 was used to amplify the 16S rRNA gene and the fungal ITS1 region from extracted DNA samples. Briefly, the 16S rRNA was amplified with the primer set H-515F- mod and H-806R-mod targeting general bacteria and archaea (515 F: 5’-H- GTGYCAGCMGCCGCGGTAA-3’; 806 R: 5’-H-GGACTACNVGGGTWTCTAAT- 3’). For details on amplification protocols see supplementary materials (Supplementary Methods 1). The fungal ITS1 region was amplified with primers ITS1F (5’-H- GCTATGCGCGAGCTGCCTTGGTCATTTAGAGGAAGTAA -3‘) and ITS2 (5‘-H-GCTATGCGCGAGCTGCGCTGCGTTCTTCATCGATGC-3‘) as described in detail in Gorka et al.35 and sequencing was performed using the Illumina MiSeq platform, v3 chemistry, and 2 × 300 bp sequencing mode at Microsynth AG (Balgach, Switzerland).
Visualization techniques of mycorrhizal root-tips and associated microorganisms
Mycorrhizal root-tips were fixed in 1% glutaraldehyd (GA) for 2 h and washed by exchanging GA with K2HPO4:KH2PO4 buffer three times. To preserve the native structure of the samples, they were treated with 1% osmiumtetroxide (OsO4) solution for 2 h and washed with MQ water thrice. Samples were dehydrated through an ethanol/acetone series (50%-100%) and subsequently critical point dried (25 cycles, EM CPD300, Leica Microsystem, Wetzlar, Germany) by replacing water with CO2 through the intermediate fluid acetone. Critical point dried samples were fixed on stubs with carbon stickers and sputter-coated with palladium-platinum (Pd/Pt) for 100 secs at 80 mA (High Resolution Fine Coater JFC-2300 HR, JEOL, Freising, Germany). Morphological structures of different mycorrhizal root-tips and their colonization with microorganisms were investigated by scanning electron microscopy (SEM Jeol IT 300, JEOL, Freising, Germany).
Paired sequence reads were merged using BBmerge v.37.6165 with “strict” setting (requiring exact match), a minimum overlap of 50 bp after clipping 3’-prime ends with quality scores below 20. In total, 884136 16S rRNA gene and 498541 ITS reads were generated. Exact amplicon sequence variants (ASVs) were determined with DADA266 based on the entire dataset (“pool=T” setting) with standard settings. The ASVs were further grouped into percentage-identity-independent “operational taxonomic units (OTUs)” with SWARM267 in fastidious mode with limit of a large swarm for grafting set at 20. Taxonomy was assigned to OTU-centroids by Last-common-ancestor (LCA) algorithm using rRNA-secondary-structure-aware SINA aligner v.1.2.1168 and the SILVA SSU138 database69. Unite database (version 8.2) was used for fungal taxonomic classification. Fungal lifestyles were defined based on genus level or on higher levels in case genus was not available, according to the FUNguild database implemented in R70.
Highly abundant OTUs from the extraction blanks were removed from the 16S rRNA dataset (11 OTUs) and from the ITS1 dataset (7 OTUs). Further, OTUs not classified at kingdom or domain level were excluded from both datasets. OTUs classified as mitochondria or chloroplasts (58 OTUs) were removed from the 16S rRNA dataset and OTUs not classified as fungi at kingdom level (44 OTUs) were removed from the fungal ITS1 dataset. The average number of reads per individual sample after removing reads that correspond to these aforementioned OTUs was ~6000 for ITS and ~9000 for 16S rRNA genes (detailed information on the final number of reads per sample can be found in the Supplementary Data 1). The final datasets contained 1323 fungal OTUs (encompassing 13 phyla, 41 classes) and 6784 bacterial OTUs/ 23 archaeal OTUs (encompassing 42 phyla, 107 classes, 309 families). Because archaeal OTUs represented only 0.3% (23 OTUs) of all 16S OTUs, bacterial and archaeal OTUs were analyzed together.
Statistics and reproducibility
For comparisons among bulk soil, rhizosphere and mycorrhizal root-tips, samples derived from seven trees were used (Supplementary Table 1), resulting in eight bulk samples, seven rhizosphere samples and 31 mycorrhizal root-tip samples. For all analyses comparing mycorrhizal root-tips amongst each other, 31 mycorrhizal root-tip samples of six additional trees (4-6 root-tips per tree) were included in the analysis, leading to a total of 62 mycorrhizal root-tip samples derived from a total of 13 trees (Supplementary Table 1).
Data analyses and statistics were performed in R (version 3.6.3). Packages “phyloseq”71 and “vegan”72 were used for sequence data analyses. Graphs were plotted in “ggplot2”73. Package “ggtern”74 was used for ternary plots and package “UpSetR”75 was used to create upset plots. See supplementary material (Supplementary Methods 2) for details on statistics.
Most network analyses of soil microbial communities rely on a statistical inference of association metrics based on pairwise correlations of taxa across a sufficiently large number of soil samples (i.e. co-occurrence networks)76. This is necessary because it is impossible to directly quantify small-scale microbial interactions, in the relatively large size of soil samples. In our approach we circumvented this need by performing a sampling design at an ecologically meaningful spatial scale (as described for instance in Fortuna et al.77 and Montesinos-Navarro et al.78), that allows to observe which fungi and bacteria tend to inhabit the same individual mycorrhizal root-tip. Thus, rather than using statistical inference to estimate coefficients of association between taxa across samples, our study uses direct counts of root-tips on which a pair of fungi and bacteria/archaea are found, thereby relying on a fine-tuned experimental design to characterize the observed networks.
We constructed weighted bipartite networks of fungal and bacterial/archaeal communities of 62 mycorrhizal root-tips. The networks were calculated with relative abundance data. In the bipartite networks, each element of the adjacency matrix corresponds to the number of root-tips on which a given fungal and bacterial taxa co-exist. We then estimated the modularity of this bipartite network to determine the assemblies of bacterial taxa that tend to inhabit the same root-tip as a given set of fungi. In more detail, we constructed five bipartite networks, each considering 6632 bacterial OTUs and different relative abundance cut-offs for fungal OTUs per individual root-tip. As such we created five different networks that either considered only dominant fungi or also less abundant ones. Network “25%” included all fungal OTUs with a relative abundance higher than 25 percent (26 OTUs), network “10%” consisted of all fungal OTUs >10% relative abundance (40 OTUs), network “5%” included all fungal OTUs >5% rel. abundance (54 OTUs), network “1%” consisted of all fungal OTUs >1% rel. abundance (126 OTUs), network “0.1%” included all fungal OTUs >0.1% rel. abundance (416 OTUs). We then computed modularity in all the networks and chose three networks (network “10%”, network “5%” and network “0.1%”) for further analysis. We compared the resulting structure between the networks considering only the highly abundant fungi (networks with >10% or >5% relative abundance) with the most complete network including also other less abundant fungi (>0.1% relative abundance: network “0.1%”). No substantial changes among the three networks would suggest that mainly the highly abundant fungal OTUs shape the assembly of co-existing bacteria, while contrasting network structures will suggest that the consideration of rare fungi generate fungal-bacterial/archaeal community assemblies.
We propose an ecological framework to assess two non-alternative processes through which rare fungi may contribute to changes in the fungal/bacterial network structure. We assess the presence of each process based on whether consideration of rare fungi results in new modules in the more complete network, or in changes in taxa composition in pre-existing modules.
On the one hand, a clumped distribution of rare fungi in new modules (vs. an even distribution among pre-existing modules) will suggest the emergence (or not) of specific bacterial associations with rare fungi. On the other hand, the higher (or lower) the proportion of highly abundant fungi that remain in the same module in both networks, the lower (or higher) the effect of rare fungi-induced changes in the similarity patterns of bacterial associations between the highly abundant fungi. We used a Pearson’s Chi-squared test for count data to assess whether rare fungi are evenly distributed across modules in the most complete network, and a t-test to assess whether the proportion of highly abundant fungi that keep sharing the same module in the most complete networks is significantly different from 1.
Modularity identifies groups of nodes that tend to interact more with each other than with the network as a whole. Different algorithms can be used to evaluate multiple partitions of nodes and select the optimal one that maximizes modularity.
We determined modularity using the DIRTLPAwb algorithm79 implemented in the function “computeModules” in the R package “bipartite”80. This algorithm maximizes weighted modularity in bipartite networks at a higher speed than other algorithms such as QuanBiMo, making it attractive for detecting the modularity of larger networks79. A maximum of 1000 MCMC steps with a tolerance level of 10−10 was used in 100 iterations, retaining the iterations with the highest likelihood value as the optimal modular configuration. We tested whether our networks were significantly more modular than random networks by running the same algorithm in 100 random networks, with the same linkage density as the empirical one, which served as null model81. Modularity significance was tested for each iteration by comparing the empirical versus the random modularity indices using a z-score test82. After testing the modularity of our network, we determined the number of modules83 and identified the bacterial and fungal taxa ascribed to each module with the function “listModuleInformation” in the R package bipartite 2.080.
Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.
The raw sequence data were deposited and released in the National Center for Biotechnology Information (NCBI) Short Read Archive under BioProject ID PRJNA778470. OTU tables, Metadata and Taxonomy files used in the analysis are deposited on figshare and available at https://doi.org/10.6084/m9.figshare.21276675.v1.
Brundrett, M. Diversity and classification of mycorrhizal associations. Biol. Rev. Camb. Philos. Soc. 79, 473–495 (2004).
Bonfante, P. & Anca, I.-A. Plants, Mycorrhizal Fungi, and Bacteria: A Network of Interactions. Annu. Rev. Microbiol. 63, 363–383 (2009).
Bogar, L.. & Peay, K.. Processes Maintaining the Coexistence of Ectomycorrhizal Fungi at a Fine Spatial Scale. in Biogeography of Mycorrhizal Symbiosis. Ecological Studies (Analysis and Synthesis) https://doi.org/10.1007/978-3-319-56363-3_4 (2017).
Rambelli, A. The Rhizosphere of Mycorrhizae. Ectomycorrhizae Their Ecol. Physiol. 299–349 https://doi.org/10.1016/b978-0-12-472850-9.50014-1 (1973).
Karlsson, A. E., Johansson, T. & Bengtson, P. Archaeal abundance in relation to root and fungal exudation rates. FEMS Microbiol. Ecol. 80, 305–311 (2012).
Bahram, M., Põlme, S., Kõljalg, U. & Tedersoo, L. A single European aspen (Populus tremula) tree individual may potentially harbour dozens of Cenococcum geophilum ITS genotypes and hundreds of species of ectomycorrhizal fungi. FEMS Microbiol. Ecol. 75, 313–320 (2011).
Yao, F., Vik, U., Brysting, A. K., Carlsen, T. & Halvorsen, R. Substantial compositional turnover of fungal communities in an alpine ridge-to-snowbed gradient. Mol. Ecol. 22, 5040–5052 (2013).
Gundale, M., Gustafsson, H. & Nilsson, M.-C. The sensitivity of nitrogen fixation by a feathermoss–cyanobacteria association to litter and moisture variability in young and old boreal forests. Can. J. Res. 39, 2542–2549 (2009).
Yoshida, N., Son, J. A., Matsushita, N., Iwamoto, K. & Hogetsu, T. Fine-scale distribution of ectomycorrhizal fungi colonizing Tsuga diversifolia seedlings growing on rocks in a subalpine Abies veitchii forest. Mycorrhiza 24, 247–257 (2014).
Morris, M. H., Pérez-Pérez, M. A., Smith, M. E. & Bledsoe, C. S. Multiple species of ectomycorrhizal fungi are frequently detected on individual oak root tips in a tropical cloud forest. Mycorrhiza 18, 375–383 (2008).
Saari, S. K., Campbell, C. D., Russell, J., Alexander, I. J. & Anderson, I. C. Pine microsatellite markers allow roots and ectomycorrhizas to be linked to individual trees. N. Phytol. 165, 295–304 (2005).
Koide, R. T., Xu, B., Sharda, J., Lekberg, Y. & Ostiguy, N. Evidence of species interactions within an ectomycorrhizal fungal community. N. Phytol. 165, 305–316 (2005).
Kaldorf, M., Renker, C., Fladung, M. & Buscot, F. Characterization and spatial distribution of ectomycorrhizas colonizing aspen clones released in an experimental field. Mycorrhiza 14, 295–306 (2004).
Smith, M. E., Douhan, G. W. & Rizzo, D. M. Ectomycorrhizal community structure in a xeric Quercus woodland based on rDNA sequence analysis of sporocarps and pooled roots. N. Phytol. 174, 847–863 (2007).
Pena, R. et al. Girdling Affects Ectomycorrhizal Fungal (EMF) Diversity and Reveals Functional Differences in EMF Community Composition in a Beech Forest †. Appl. Environ. Microbiol. 76, 1831–1841 (2010).
Agerer, R. Colour Atlas of Ectomycorrhizae. Einhorn-Verlag Eduard Dietenberger GmbH, Schwäbisch-Gmünd, Ger. (1997).
Lang, C., Seven, J. & Polle, A. Host preferences and differential contributions of deciduous tree species shape mycorrhizal species richness in a mixed Central European forest. Mycorrhiza 21, 297–308 (2011).
Pellitier, P. T. & Zak, D. R. Ectomycorrhizal root tips harbor distinctive fungal associates along a soil nitrogen gradient. Fungal Ecol. 54, 101111 (2021).
Tedersoo, L. Biogeography of Mycorrhizal Symbiosis. 230, (2017).
Tedersoo, L. et al. 454 Pyrosequencing and Sanger sequencing of tropical mycorrhizal fungi provide similar results but reveal substantial methodological biases. N. Phytol. 188, 291–301 (2010).
Leberecht, M. et al. Ectomycorrhizal Communities on the Roots of Two Beech (Fagus sylvatica) Populations from Contrasting Climates Differ in Nitrogen Acquisition in a Common Environment. Appl. Environ. Microbiol. 81, 5957–5967 (2015).
Bomberg, M. & Timonen, S. Distribution of cren- and euryarchaeota in scots pine mycorrhizospheres and boreal forest humus. Microb. Ecol. 54, 406–416 (2007).
Bomberg, M., Jurgens, G., Saano, A., Sen, R. & Timonen, S. Nested PCR detection of Archaea in defined compartments of pine mycorrhizospheres developed in boreal forest humus microcosms. FEMS Microbiol. Ecol. 43, 163–171 (2003).
Tateishi, T., Yokoyama, K., Kohno, N., Okabe, H. & Marumoto, T. Estimation of mycorrhizal colonization of the roots of oak seedlings inoculated with an ectomycorrhizal fungus, Laccaria amethystea. Soil Sci. Plant Nutr. 49, 641–645 (2003).
Mogge, B., Loferer, C., Agerer, R., Hutzler, P. & Hartmann, A. Bacterial community structure and colonization patterns of Fagus sylvatica L. ectomycorrhizospheres as determined by fluorescence in situ hybridization and confocal laser scanning microscopy. Mycorrhiza 9, 271–278 (2000).
Haq, I. U., Zhang, M., Yang, P. & van Elsas, J. D. The Interactions of Bacteria with Fungi in Soil: Emerging Concepts. Adv. Appl. Microbiol 89, 185–215 (2014).
Timonen, S., Jørgensen, K. S., Haahtela, K. & Sen, R. Bacterial community structure at defined locations of Pinus sylvestris - Suillus bovinus and Pinus sylvestris - Paxillus involutus mycorrhizospheres in dry pine forest humus and nursery peat. Can. J. Microbiol. 44, 499–513 (1998).
Khetmalas, M. B., Egger, K. N., Massicotte, H. B., Tackaberry, L. E. & Clapperton, M. J. Bacterial diversity associated with subalpine fir (Abies lasiocarpa) ectomycorrhizae following wildfire and salvage-logging in central British Columbia. Can. J. Microbiol. 48, 611–625 (2002).
Uroz, S., Oger, P., Morin, E. & Frey-klett, P. Distinct Ectomycorrhizospheres Share Similar Bacterial Communities as Revealed by Pyrosequencing-Based Analysis of 16S rRNA Genes. Appl. Environ. Microbiol. 78, 3020–3024 (2012).
Vik, U. et al. Different bacterial communities in ectomycorrhizae and surrounding soil. Sci. Rep. 3, (2013).
Aslani, F., Tedersoo, L., Põlme, S., Knox, O. & Bahram, M. Global patterns and determinants of bacterial communities associated with ectomycorrhizal root tips of Alnus species. Soil Biol. Biochem. 148, 107923 (2020).
Tedersoo, L., May, T. W. & Smith, M. E. Ectomycorrhizal lifestyle in fungi: Global diversity, distribution, and evolution of phylogenetic lineages. Mycorrhiza 20, 217–263 (2010).
Marupakula, S., Mahmood, S. & Finlay, R. D. Analysis of single root tip microbiomes suggests that distinctive bacterial communities are selected by Pinus sylvestris roots colonized by different ectomycorrhizal fungi. Environ. Microbiol. 18, 1470–1483 (2016).
Marupakula, S. et al. Bacterial microbiomes of individual ectomycorrhizal Pinus sylvestris roots are shaped by soil horizon and differentially sensitive to nitrogen addition. Environ. Microbiol. 19, 4736–4753 (2017).
Gorka, S. et al. Rapid transfer of plant photosynthates to soil bacteria via ectomycorrhizal hyphae and its interaction with nitrogen availability. Front. Microbiol. 10, 1–20 (2019).
Van Der Heijden, M. G. A. & Horton, T. R. Socialism in soil? The importance of mycorrhizal fungal networks for facilitation in natural ecosystems. J. Ecol. 97, 1139–1150 (2009).
Kennedy, P. G. & Bruns, T. D. Priority effects determine the outcome of ectomycorrhizal competition between two Rhizopogon species colonizing Pinus muricata seedlings. N. Phytol. 166, 631–638 (2005).
Kennedy, P. G., Peay, K. G. & Bruns, T. D. Root tip competition among ectomycorrhizal fungi: Are priority effects a rule or an exception? Ecology 90, 2098–2107 (2009).
Koide, R. T., Fernandez, C. & Petprakob, K. General principles in the community ecology of ectomycorrhizal fungi. Ann. Sci. 68, 45–55 (2011).
Rinta-Kanto, J. M. & Timonen, S. Spatial variations in bacterial and archaeal abundance and community composition in boreal forest pine mycorrhizospheres. Eur. J. Soil Biol. 97, 103168 (2020).
De Boer, W., Folman, L. B., Summerbell, R. C. & Boddy, L. Living in a fungal world: Impact of fungi on soil bacterial niche development. FEMS Microbiol. Rev. 29, 795–811 (2005).
Van Hees, P. A. W. et al. Oxalate and ferricrocin exudation by the extramatrical mycelium of an ectomycorrhizal fungus in symbiosis with Pinus sylvestris. N. Phytol. 169, 367–378 (2006).
Johansson, E. M., Fransson, P. M. A., Finlay, R. D. & van Hees, P. A. W. Quantitative analysis of soluble exudates produced by ectomycorrhizal roots as a response to ambient and elevated CO2. Soil Biol. Biochem. 41, 1111–1116 (2009).
Siddiqui, Z. A., Akhtar, M. S. & Futai, K. Mycorrhizae: Sustainable Agriculture and Forestry. (Springer Netherlands, 2008). https://doi.org/10.1007/978-1-4020-8770-7.
Frey-Klett, P., Garbaye, J. & Tarkka, M. The mycorrhiza helper bacteria revisited. N. Phytol. 176, 22–36 (2007).
Nguyen, N. H. & Bruns, T. D. The Microbiome of Pinus muricata Ectomycorrhizae: Community Assemblages, Fungal Species Effects, and Burkholderia as Important Bacteria in Multipartnered Symbioses. Microb. Ecol. 914–921 https://doi.org/10.1007/s00248-015-0574-y (2015).
Offre, P. et al. Microdiversity of Burkholderiales associated with mycorrhizal and nonmycorrhizal roots of Medicago truncatula. FEMS Microbiol. Ecol. 65, 180–192 (2008).
Antony-Babu, S. et al. Black truffle-associated bacterial communities during the development and maturation of Tuber melanosporum ascocarps and putative functional roles. Environ. Microbiol. 16, 2831–2847 (2014).
Barbieri, E. et al. New evidence for bacterial diversity in the ascoma of the ectomycorrhizal fungus Tuber borchii Vittad. FEMS Microbiol. Lett. 247, 23–35 (2005).
Barbieri, E. et al. Occurrence and diversity of bacterial communities in Tuber magnatum during truffle maturation. Environ. Microbiol. 9, 2234–2246 (2007).
Garbaye, J. & Duponnois, R. Specificity and function of Mycorrhization helper bacteria (MHB) associated with the Pseudotsuga menziesii―Laccaria laccata symbiosis. Symbiosis (Phila., PA) 14, 335–344 (1992).
Labbé, J. L., Weston, D. J., Dunkirk, N., Pelletier, D. A. & Tuskan, G. A. Newly identified helper bacteria stimulate ectomycorrhizal formation in Populus. Front. Plant Sci. 5, 1–10 (2014).
Schrey, S. D. & Tarkka, Æ. M. T. Friends and foes: streptomycetes as modulators of plant disease and symbiosis. Antonie van Leeuwenhoek, Int. J. Gen. Mol. Microbiol. 1, 11–19 (2008).
Igiehon, N. O., Babalola, O. O. & Aremu, B. R. Genomic insights into plant growth promoting rhizobia capable of enhancing soybean germination under drought stress. BMC Microbiol 19, 1–22 (2019).
Liu, Q. et al. Response of Sugarcane Rhizosphere Bacterial Community to Drought Stress. Front. Microbiol. 12, 1–14 (2021).
Santos-Medellín, C., Edwards, J., Liechty, Z., Nguyen, B. & Sundaresan, V. Drought Stress Results in a Compartment-Specific Restructuring of. MBio 8, 1–15 (2017).
Boersma, F. G. H., Warmink, J. A., Andreote, F. A. & Van Elsas, J. D. Selection of Sphingomonadaceae at the base of Laccaria proxima and Russula exalbicans fruiting bodies. Appl. Environ. Microbiol. 75, 1979–1989 (2009).
Uroz, S. et al. Effect of the mycorrhizosphere on the genotypic and metabolic diversity of the bacterial communities involved in mineral weathering in a forest soil. Appl. Environ. Microbiol. 73, 3019–3027 (2007).
Fitter, A. H. & Garbaye, J. Interaction between mycorrhizal fungi and other soil organisms. Plant Soil 159, 115–121 (1994).
Bonfante, P. & Genre, A. Mechanisms underlying beneficial plant - fungus interactions in mycorrhizal symbiosis. Nat. Commun. 1, 1–11 (2010).
Mayerhofer, W. et al. Recently photoassimilated carbon and fungus-delivered nitrogen are spatially correlated in the ectomycorrhizal tissue of Fagus sylvatica. N. Phytol. 232, 2457–2474 (2021).
Angel, R., Claus, P. & Conrad, R. Methanogenic archaea are globally ubiquitous in aerated soils and become active under wet anoxic conditions. ISME J. 6, 847–862 (2012).
Janowski, D., Wilgan, R., Leski, T., Karlinski, L. & Rudawska, M. Effective molecular identification of ectomycorrhizal fungi: Revisiting DNA isolation methods. Forests 10, 1–10 (2019).
Herbold, C. W. et al. A flexible and economical barcoding approach for highly multiplexed amplicon sequencing of diverse target genes. Front. Microbiol. 6, 1–8 (2015).
Bushnell, B., Rood, J. & Singer, E. BBMerge – Accurate paired shotgun read merging via overlap. PLoS One 12, 1–15 (2017).
Callahan, B. J. et al. DADA2: High resolution sample inference from Illumina amplicon data. Nat. Methods 13, 4–5 (2016).
Mahé, F., Rognes, T., Quince, C., de Vargas, C. & Dunthorn, M. Swarmv2: Highly-scalable and high-resolution amplicon clustering. PeerJ 1–12 (2015) https://doi.org/10.7717/peerj.1420.
Pruesse, E., Peplies, J. & Glöckner, F. O. SINA: Accurate high-throughput multiple sequence alignment of ribosomal RNA genes. Bioinformatics 28, 1823–1829 (2012).
Quast, C. et al. The SILVA ribosomal RNA gene database project: Improved data processing and web-based tools. Nucleic Acids Res 41, 590–596 (2013).
Nguyen, N. H. et al. FUNGuild: An open annotation tool for parsing fungal community datasets by ecological guild. Fungal Ecol. 20, 241–248 (2016).
McMurdie, P. J. & Holmes, S. Phyloseq: An R Package for Reproducible Interactive Analysis and Graphics of Microbiome Census Data. PLoS One 8, e61217 (2013).
Okansen, J. et al. vegan: Community Ecology Package. (2019).
Wickham H. ggplot2: Elegant Graphics for Data Analysis. Springer-Verlag New York (2016).
Hamilton, N. E. & Ferry, M. Ggtern: Ternary diagrams using ggplot2. J. Stat. Softw. 87, 1–17 (2018).
Conway, J. R., Lex, A. & Gehlenborg, N. UpSetR: An R package for the visualization of intersecting sets and their properties. Bioinformatics 33, 2938–2940 (2017).
Guseva, K. et al. From diversity to complexity: Microbial networks in soils. Soil Biol. Biochem. 169, 108604 (2022).
Fortuna, M. A. et al. Nestedness versus modularity in ecological networks: Two sides of the same coin? J. Anim. Ecol. 79, 811–817 (2010).
Montesinos-Navarro, A., Segarra-Moragues, J. G., Valiente-Banuet, A. & Verdú, M. The network structure of plant-arbuscular mycorrhizal fungi. N. Phytol. 194, 536–547 (2012).
Beckett, S. J. Improved community detection in weighted bipartite networks. R. Soc. Open Sci. 3, 140536 (2016).
Dormann, C. F., Gruber, B. & Fründ, J. Introducing the bipartite Package: Analysing Ecological. Netw. R. N. 8/2, 8–11 (2008).
Guimera, R. & Amaral, L. A. N. Functional cartography of complex metabolic networks. Nature 433, 895–900 (2005).
Dormann, C. F. & Strauss, R. A method for detecting modules in quantitative bipartite networks. Methods Ecol. Evol. 5, 90–98 (2014).
Newman, M. E. J. Analysis of weighted networks. Phys. Rev. E - Stat. Phys., Plasmas, Fluids, Relat. Interdiscip. Top. 70, 9 (2004).
This work was financially supported by the Austrian Science Fund (FWF) project DK plus [W1257-B20]. AMN was supported by Spanish Ministry of Science, Innovation and Universities (RTI2018-099672-J-I00). CK received funding from the European Research Council (ERC) under the Europeam Union’s Horizon 2020 research and innovation program (grant agreement No 819446). SEM work was carried out at Core Facility Cell Imaging and Ultrastructure Research at the university of Vienna. The authors would like to thank Daniela Gruber for help in SEM preparation.
The authors declare no competing interests.
Peer review information
Communications Biology thanks Raffaella Balestrini, Julieta Alvarez-Manjarrez and the other, anonymous, reviewers for their contribution to the peer review of this work. Primary Handling Editors: Meritxell Riquelme and Luke R. Grinham. Peer reviewer reports are available.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
About this article
Cite this article
Dietrich, M., Montesinos-Navarro, A., Gabriel, R. et al. Both abundant and rare fungi colonizing Fagus sylvatica ectomycorrhizal root-tips shape associated bacterial communities. Commun Biol 5, 1261 (2022). https://doi.org/10.1038/s42003-022-04178-y