Seeking the needle in the haystack: absence of mycorrhizal fungi outside of the plant rhizosphere associated with an endangered Australian orchid.
Co-occurrence and abundance of suitable mycorrhizal fungi are expected to be important drivers for orchid seedling establishment and development, as well as mature plant distribution. However, limited information is available on the occurrence and spatial patterns of orchid mycorrhizal fungi in soil independent of the orchid host. In this study, we investigated the in situ distribution of Tulasnella spp. associated with the critically endangered Australian orchid Diuris fragrantissima. We implemented a meta-barcoding approach (fungal ITS1 region) targeting three soil sources: orchid rhizosphere, orchid-associated bulk soil and bulk soil from the orchid native site. The quality-filtered data set revealed that the occurrence of Tulasnella spp. in situ is restricted to the orchid rhizosphere, suggesting that a limited number of potential recruitment micro-sites with suitable mycorrhizal taxaexists in the Diuris fragrantissima natural habitat. The meta-barcoding approach also revealed a distinctive fungal community associated with the orchid rhizosphere. Overall, NGS technology has proven to be a suitable method for large-scale screening of fungal isolates in orchid-associated soil.
Seed germination, transition to seedling and early development of terrestrial orchids in natural conditions are reliant on colonization by mycorrhizal fungal symbionts (Leake, 1994, Harley and Smith 1983). Lack of an extensive root system makes nutrient acquisition by fungal symbionts critical throughout the life of the orchid (Harley and Smith 1983). Despite the diversity of fungi associated with terrestrial orchids varying across habitats, or even between seedlings and adult plants (Rasmussen 1995, Currah et al. 1997 and Brundrett 2006), many such orchids have been found to form mycorrhizal associations with a relatively narrow diversity of fungi in the ‘rhizoctonia’ alliance. Rhizoctonia fungi are a polyphyletic assemblage that encompasses pathogens, endophytes, saprotrophs and mycorrhizal fungi (Warcup 1981, Sivasithamparam 1993, Rasmussen 1995, Currah et al. 1997 and Roberts 1999), including important orchid mycorrhizal genera such as Ceratobasidium, Tulasnella, Thanatephorus, and Sebacina (Weiss et al. 2004; Dearnaley 2007).
Co-occurrence and abundance of suitable mycorrhizal fungi is expected to be an important driver not only for orchid seedling establishment and development, but also mature plant occurrence, and the significance of distribution/dispersal linkages between orchids and fungi has been documented in several studies (reviewed in McCormick and Jacquemyn 2014). While limited fungal presence may represent a major constraint especially for those orchids displaying narrow fungal specificity (Brundrett et al. 2003; Bonnardeaux et al. 2007; Dearnaley 2007; Ogura-Tsujita et al. 2009), dynamics and ecology of fungi independent from the orchids they associate with have not been clearly addressed yet (Bahram, Peay and Tedersoo 2015, McCormick et al. 2012, 2016), although the majority of orchid mycorrhizae are free-living fungi that can grow without the host plant (Dearnaley et al. 2012, McCormick and Jacquemyn 2014, Rasmussen et al. 2015). Several seed germination experiments have demonstrated that symbiont presence is usually limited to patches where the orchid also occurs (e.g. Batty et al. 2001; Phillips et al. 2011). Indeed, germination often declines with increasing distance from adult plants, suggesting that the abundance of suitable mycorrhizal fungi also decreases (McKendrick et al. 2000, 2002, Batty et al. 2001 Diez 2007 and Jacquemyn et al. 2012). However, some orchid mycorrhizae have been shown to occur independently from orchids (i.e. > 5 m from the nearest conspecific orchid; e.g. McKendrick et al. 2002; Tesitelova et al. 2012; Oja et al. 2015), indicating that suitable fungi can be present outside orchid patches. Given the importance of fungi as drivers of orchid dispersal (Batty et al. 2002, Jacquemyn et al. 2014, McCormick et al. 2016), being able to resolve the distribution of mycorrhizal fungi in both occupied and unoccupied orchid habitats is likely to be a crucial requirement to identify appropriate microsites for reintroduction activities.
‘Baiting’ techniques using orchid seed packets are commonly implemented to investigate the distribution of orchid mycorrhizal fungi (e.g. Perkins and McGee 1995; McKendrick et al. 2000; Batty et al. 2001; Brundrett et al. 2003; Otero et al. 2007, McCormick et al. 2016). However, this method can only detect locations where both fungi and environmental conditions are appropriate for seed germination, while actual fungal occurrence may be more widespread (McCormick et al. 2016). Recently, DNA metabarcoding using Next Generation Sequencing (NGS) has emerged as an effective tool for monitoring fungal biodiversity and orchid mycorrhizal fungi occurrence (Jacquemyn et al. 2014, Oja et al. 2015, Waud et al. 2016). A high-throughput DNA sequencing approach allows for retrieval and identification of microbial genetic material directly from environmental samples and could represent an efficient, non-invasive and easy-to-standardize sampling approach for soil fungi (Oja et al. 2015).
Diuris fragrantissima is critically endangered Australian native orchid (EPBC Act, 1998) which grows in association with a narrow taxonomic range of mycorrhizal fungi (Tulasnella spp.) within the cosmopolitan family Tulasnellaceae (Smith et al. 2010). The limited survival of cultivated plants of D. fragrantissima after reintroduction has been correlated with the loss of associated fungi during the transition from ex situ to in situ (Smith 2006). The presence of microsites containing suitable mycorrhizal fungi able to assist reintroduced orchids is predicted to be a necessary prerequisite for their long-term survival in the wild (Smith et al. 2010). Nevertheless, monitoring the presence of the fungus using traditional ‘baiting’ techniques has proven to be unreliable in this situation (Smith 2006). In this study, we used metabarcoding to investigate distribution of orchid mycorrhizal fungi in the orchid rhizosphere and adjacent soil, as well to characterize fungal diversity across the natural habitat of D. fragrantissima.
Material and Methods
Site Description and Soil Sampling
Diruis fragrantissima, Sunshine Diuris, is a perennial, terrestrial orchid endemic to the basalt plains immediately to the west of Melbourne (Australia) in the Victorian Volcanic Plain IBRA Bioregion (sensu DEH 2000). Following habitat degradation, weed invasion, predation by introduced herbivores, altered burning regimes and illicit collection, a catastrophic decline in range and abundance has impacted this native species (Murphy et al. 2008, Smith 2006). To date, only a single wild population, comprising about 30 plants, remains in a grassland reserve within a fenced area of approximately 1200 m2 located in Sunshine (Melbourne, Victoria, Australia) (Murphy et al. 2008). The site is dominated by several Australian native species, including Themeda triandra, Austrodanthonia sp., Dianella longifolia, D. revoluta, Tricoryne elatior, Pimelea humilis and Dichanthium sericeum subsp. sericeum. Orchids are located in the north-eastern side of the fenced area, in a patch measuring approximately 15×10 m. The soil is shallow heavy clay, with several exposed basalt boulders also present (Murphy et al. 2008).
We collected soil samples from three distinct sources: orchid rhizosphere soil (in direct contact with the orchid root), orchid bulk soil (adjacent to the orchid plants), and site bulk soil (Figure 1). Firstly, to explore the composition of the fungal community directly influenced by the orchid roots, two samples from the rhizosphere soil (~1 mm distance from the root surface and 3 cm away from root tip) from four native plants in situ (‘orchid rhizosphere soil’) were collected, for a total of eight samples. Secondly, to investigate the orchid ‘zone of influence’ — i.e. how far from the plant we can still find traces of its mycorrhizal symbiont — five plants at the external boundaries of the orchid patch were further examined by collecting adjacent to the plant (0 cm, but not in contact with the root) along with three points at incremental distance from each plant (10, 20, 40 cm). Samples were taken along one and four transects in the directions north, south, east and west, depending on the orchid location (some transects were not possible due to presence of rocks), for a total of 35 additional samples (‘orchid bulk soil’). Sampling was performed using a soil corer (2 cm diameter) to a depth of 10 cm, following Prober et al. (2015). Thirdly, to investigate the distribution of Tulasnella spp. and the occurrence of possible micro-sites suitable for orchid translocation, soil samples were collected from the site at which the orchid was present (‘site bulk soil’). The sample pool comprised soils from areas where orchids were and were not present. To achieve systematic sampling, a 5 m2 grid was designated within the fenced area. For each plot, the middle point was identified and four soil cores (2 cm diameter x 10 cm length) were collected within a small radius (25 cm) of the grid centre. The soil cores were combined as one sample for analysis. Given the patchy distribution of Tulasnella spp. in the Diuris natural habitat (Smith 2006), combining several cores increases the chance to detect the fungus of interest. A total of 44 samples were collected from the orchid reserve, including six samples (A1, A2, A3, B1, B2, and B3) from the orchid-occupied area. In addition, ten soil samples were collected outside the reserve to test the occurrence of Tulasnella spp. in adjacent zones that have potential for reintroduction.
Rhizosphere soil was collected in June 2015, while site and orchid bulk soil were collected in July 2015. All samples were stored in sterile falcon tubes at -20 °C before DNA extraction. The data-set included freshly isolated Tulasnella sp. pure cultures from three of the examined plants, all of which successfully germinated seedlings (Neil Anderton, personal communication). The experimental workflow is summarised in Figure 1.
DNA extraction, PCR amplification and Library Preparation
Soil samples were homogenized and DNA was extracted from 0.25 g soil using the PowerSoil™ Soil DNA Isolation Kit (Geneworks, Australia). DNA from three Tulasnella sp. cultures (~1 cm diameter on agar) was used as positive control. To improve the efficiency of the DNA extraction protocol, we employed a longer cell lysis step (15 min) with samples heated to 65 °C (instead of room temperature) in PowerBead, following Jiang et al. 2015. The DNA was quantitated with a ND1000 spectrophotometer (NanoDrop Technologies, Wilmington, Delaware) and standardized to 5.0 ng/µl.
As the 5.8S rDNA region is hyper variable in Tulasnella species and all known primers have multiple mismatches to many species (Tedersoo et al. 2014; Oja et al. 2015), fungi identification was based on the ITS1 region obtained from the full internal transcribed spacer (ITS) region. Fungal ITS region was amplified using primers ITS1F/ ITS4 (Gardes and Bruns 1993; White et al. 1990), whose ability to amplify ITS region of Diuris-associated Tulasnellaceae has been previously demonstrated by Smith et al. (2010). However, to further confirm the suitability of ITS1F-ITS4 primers for identifying the presence of Tulasnellaceae in soil, we conducted a pilot MiSeq run, where the performance of these primers was compared with the Tulasnella-specific primers ITS1F-ITS4Tul (Taylor and McCormick 2008). The pilot run included rhizosphere soil and the Tulasnella spp. pure cultures. The two primer pairs performed similarly in retrieving Tulasnella spp. from soil (see Supplementary Material 1) and therefore primers ITS1F/ITS4 were chosen over ITS1F-ITS4Tul because they are universal (Schoch et al. 2012), allowing for investigation of both Tulasnella spp. distribution and whole soil fungal community composition.
Library preparation required a first amplification carried out in a total volume of 20 μl using 10 ng of DNA, 12.5 μl of KAPA Taq ReadyMix (Kapa Biosystems), and 10 μM each of forward and reverse primers (ITS1F and ITS4 respectively). PCR conditions were 6 min at 95 °C, followed by 35 cycles of 30 s at 95 °C, 30 s at 55 °C, and 30 s at 72 °C. Final extension was done at 72 °C for 5 min. A second PCR was performed to add individual Illumina indexes to each sample, using 25 μl of the amplicon obtained from the first PCR, with the following conditions: 6 min at 95 °C, followed by 8 cycles of 30 s at 95 °C, 30 s at 55 °C, 30 s at 72 °C, and final at 72 °C for 5 min. Each PCR product was purified with Agencourt AMPure XP SPRI magnetic beads (Beckman and Coulter). Indexed PCR products were normalized and pooled after quantification with a Qubit 2.0 Fluorometer and the Qubit dsDNA HS Assay Kit (Invitrogen). Paired-end sequencing (MiSeq Reagent Kit v3; 600-cycles) was carried out in-house on an Illumina MiSeq sequencer.
Molecular identification and bioinformatics
Each single-end read was analysed separately following the strategy implemented by Harrison et al. (2016), and only forward reads, which include the ITS1 regions, were investigated. First, reads were truncated to 280 bp and then filtered using USEARCH. v8.1.1831 (Edgar 2010). After quality-control analyses, sequences were dereplicated and clustered into operational taxonomic units (OTUs) using USEARCH. OTU clustering was performed at 97% identity threshold and taxonomy was assigned with RDP Classifier against the UNITE fungal ITS reference dataset (Kõljalg et al. 2005), at 85% confidence threshold. The UNITE species hypothesis (Kõljalg et al. 2013) for OTU matches of 97% or better was recorded.
The FunGuild application (Nguyen et al. 2015) was used to assign fungal OTUs from the rarefied data to functional guilds. For this study, we only included results where a confidence ranking of ‘highly probable’ was achieved.
We examined whether the sampling effort was adequate to capture the fungal community richness by generating species rarefaction curves and species accumulation plots using the ‘rarecurve’ and ‘specaccum’ functions in the R library vegan (v. 2.3–4; Oksanen et al., 2013) using R 3.2.0 (R-Development-Core-Team 2015).
Before all downstream analyses, the dataset was rarefied to an even depth of 500 sequences per sample and singletons removed. The fungal community data were ordinated in Primer-E software (version 6, PRIMER-E Ltd. Plymouth, UK) using non-metric multidimensional scaling (nMDS) on untransformed data and on data subject to a second-root transformation, with Bray-Curtis metric used as dissimilarity measure. As the outcomes of the ordination-based analyses were qualitatively the same irrespective of the transformation used, we report only the second-root transformation in the results. Similarities between samples displayed using 2D multidimensional scaling (nMDS) were tested using a non-parametric permutation-based test ANOSIM (Analysis of Similarity) to investigate potential differences between fungal communities in the three different soil sources (‘rhizosphere’, ‘orchid bulk soil’ and ‘site bulk soil’), and patterns in fungal composition were investigated by generating a dendrogram with the UPGMA algorithm. SIMPROF (Similarity Profile Analysis) was used to determine whether any structure existed in by group mean at the 95% confidence interval.
Fungi from soil and roots exhibited different patterns in OTU richness at the phylum level. In soil samples, Ascomycota (48.0%) was the most OTU-rich phylum followed by Basidiomycota (38.1%), whereas this pattern was reversed in root samples (Basidiomycota 53.3% and Ascomycota 38.5%; Figure 1).
Other phyla taken together comprised c. 5% of OTUs for both sample types. In soil and root samples, 8.5 and 3.0% of OTUs, respectively, remained uni dentified at the phylum level.
Fungi from soil and roots exhibited different patterns in OTU richness at the phylum level. In soil samples, Ascomycota (48.0%) was the most OTU-rich phylum followed by Basidiomycota (38.1%), whereas this pattern was reversed in root samples (Basidiomycota 53.3% and Ascomycota 38.5%; Figure 1).
Other phyla taken together comprised c. 5% of OTUs for both sample types. In soil and root samples, 8.5 and 3.0% of OTUs, respectively, remained uni dentified at the phylum level
Tulasnella detection and fungal community description
The quality-filtered data set comprised a total of 892,288 sequences, which were assigned to 2,707 OTUs at the final clustering step with 97% similarity threshold. Rarefaction curves of total fungal OTUs for the rhizosphere sample reached a plateau more often compared with the site bulk soil and orchid bulk soil (Supplementary Figure 1A), while curves on species accumulation plots were saturated for all of the three soil sources (Supplementary Figure 1B).
Prior to singleton removal and rarefaction, a total of 1,661 fungal OTUs were identified from the rhizosphere soil, 1,306 from the orchid bulk soil and 1,710 from the site bulk soil. A total of 642 OTUs were recorded from the rhizosphere exclusively, while 459 were unique to the orchid bulk soil and 117 to the site bulk soil. The OTUs unique to the rhizosphere are reported in Supplementary Table 1. One OTU (detected from 38 sequences) identified as Tulasnella sp. (SH234355.06FU) was retrieved from the rhizosphere soil collected from two of the four analysed plants (35 and 3 sequences respectively, Supplementary Table 1). The same OTU was also retrieved from the fresh cultures used as positive control (2,415 sequences), while no Tulasnella sp. OTUs were retrieved from the other soil sources (orchid bulk soil and site bulk soil).
After rarefaction and removal of singletons, a total of 1,562 OTUs and 64 samples were retained. Overall, fungi from orchid rhizosphere soil, site bulk soil and orchid bulk soil exhibited similar patterns in OTU richness at the phylum level. In site bulk soil and orchid bulk soil samples, Ascomycota (60% and 59% respectively) was the most OTU-rich phylum, followed by Basidiomycota (20% and 21% respectively) (Figure 2A). Other phyla taken together comprised approximately 20% of OTUs for both sample types. A similar distribution was observed in the rhizosphere samples. Ascomycota (59%) represented the most OTU-rich and abundant phylum, followed by Basidiomycota (21%). Of other fungal groups, Chytridiomycota, and Glomeromycota were the most OTU-rich. Less than 3% of OTUs belonged to Zygomycota, and the remaining 5% of OTUs were unidentified at the phylum level (Figure 1A).
For our dataset, 51% of OTUs (786 of 1,562) were able to be assigned to a functional group match to the FUNGuild database at the genus level. Of these, 89.6% were assigned with a confidence level of ‘highly probable’, and 10.4% were given a confidence level of ‘probable’ or ‘possible’. The OTUs were grouped and averaged according to the soil source, and the functional analyses included only the trophic modes and guilds with a ‘highly probable’ confidence level (705 OTUs) (Figure 2B and C). Nevertheless, because fungal functional groups are not well defined for Australian grassland soils (Egidi et al. 2016), the assignments here must be interpreted with caution.
In terms of trophic groups, symbiotrophic fungi were the highest in abundance (502 OTUs), followed by saprotrophs (114 OTUs). Overall, symbiotroph-containing groups (pathotroph-symbiotroph and symbiotroph groups) were relatively more abundant in the rhizosphere (65% of the total community) compared to the other soil sources (<55% of the orchid bulk soil community at each distance from the host plant, and 56% in site bulk soil).
In terms of fungal guilds, arbuscular mycorrhizal fungi (AM) were the most frequently detected taxa (447 OTUs), followed by ectomycorrhizal (ECM) fungi (44 OTUs) and undefined dung saprotrophs (38 OTUs). ECM fungi were the only group that showed a distinct pattern in relative abundance in relation to soil source (Figure 2C), being more abundant in the orchid rhizosphere (6% of the total community) compared to orchid bulk soil (<3% of the total community at each distance from the host plant) and site bulk soil (4% of the total community).
Fungal community structure
The plot in Supplementary Figure 2 shows nMDS ordinations of samples from the three soil sources (rhizosphere, orchid bulk soil and site bulk soil); the distance between any two sample points reflects their relative similarity. ANOSIM revealed that variation in soil fungal community composition was positively correlated to soil source. The strongest grouping occurred between orchid rhizosphere soil and orchid bulk soil (R= 0.85, p=0.001), and between orchid rhizosphere soil and site bulk soil (R=0.82, p=0.001), while grouping between orchid bulk soil and site bulk soil was weaker (R=0.35, p=0.001). In the ordination space, the rhizosphere samples were clearly separated from the other samples (albeit relatively widely dispersed in the ordination space), with samples from orchid bulk soil falling between the rhizosphere samples and the site bulk soil samples.
In the cluster analysis (Fig. 3), there were four main clusters, one cluster was composed exclusively of samples from the rhizosphere which clustered together with relatively low overall similarity (20%), while samples from the orchid and bulk soil were spread throughout the other three clusters. Two of these three other clusters only had samples from site bulk soil (three samples and eight samples respectively) while a third major cluster comprised all samples from orchid bulk soil along with the other 23 samples from site bulk soil. Within this third major cluster, all the orchid bulk soil samples were within one major sub-cluster (along with three bulk soil samples all from the orchid-occupied areas). Consistently, composition differences visualised by cluster and ordination showed that the rhizosphere soil samples are separate cluster.
In terms of the ‘core’ community (OTUs present in at least 75% of the samples, and contributing to more than 5% of the community, Supplementary Table 2), that of the orchid rhizosphere was composed of 90 OTUs, dominated by Ascomycota (65 OTUs), followed by Chytridiomycota (6 OTU), Basidiomycota (4 OTUs), Zygomycota (3 OTU), and Glomeromycota (1 OTU), while 10 OTUs were unidentified at phylum level (Supplementary Table 3 A). Among the rhizosphere core community, ten OTUs were assigned with high probability to the ‘Symbiotroph’ trophic mode, being mainly AM (5 OTUs) and ECM (3 OTUs), while 13 OTUs were considered ‘Saprotrophs’, mainly ‘Dung-Saprotrophs’ (11 OTUs). The core communities for orchid bulk soil and site bulk soil had less taxa (51 and 9 OTUs respectively, Supplementary Table 2 B and C). Similar to rhizosphere core community, Ascomycota were significantly present in the core community of both orchid bulk soil (38 OTUs) and site bulk soil (7 OTUs). Five and one OTUs identified as Symbiotroph were present in both orchid bulk soil and site bulk soil respectively, with orchid bulk soil being dominated by ‘Saproptrophs’ (21 OTUs) and ‘Pathotrophs’ (10 OTUs).
Tulasnella spp. detection
One OTU identified as the mycorrhizal fungus Tulasnella sp. (SH199347.06FU) was successfully retrieved from the rhizosphere of three of the four analysed wild orchids. However, no Tulasnella OTUs were obtained from the bulk soils collected from the Diuris native site, nor the orchid bulk soil, regardless of the distance from the host plant. Despite having a crucial role for the germination and persistence of D. fragrantissima, Tulasnellaceae represent ‘rare’ members of the soil fungal community, accounting for less than 1% of the total rhizosphere fungi. This suggests that the fungus of interest is narrowly distributed and restricted solely to the soil directly associated with D. fragrantissima. The limited occurrence of Tulasnella spp. may be a consequence of the small volumes of soil that can be sampled in molecular methods, insufficient sequencing depth or high heterogeneity of fungal distribution at fine scales (McCormick et al. 2016). Nevertheless, we were able to cultivate the mycorrhizal symbiont from the roots of all the orchids that gave positive results for the presence of Tulasnella spp. in the metabarcoding sampling, indicating that the retrieved Tulasnella OTUs were all associated with the presence of a living, active fungus.
Consistent with our results, rare occurrence of orchid mycorrhizal fungi in situ has been reported in previous molecular surveys, which showed that mycorrhizae outside the orchid roots represent a small fraction of the soil community (Liu et al. 2015, Oja et al. 2015, Voyron et al. 2016) and are usually restricted to patches where the host plant also occurs (Oja et al. 2015; Jacquemyn et al. 2014, McCormick et al. 2016, Waud et al. 2016, Oja et al. 2016). The sporadic occurrence of orchid-associated Tulasnella spp. in the orchid native site may be related to the limited soil-exploration ability typical of mycorrhizal fungi in high root-density areas, such as orchid patches, where short-range exploration would represent an effective strategy in terms of carbon costs (Peay et al. 2011; Voyron et al. 2016). However, limited information is available on the dispersal mechanisms of orchid mycorrhizal fungi in soil (Voyron et al. 2016). Alternatively, Diuris mycorrhizae abundance may be limited by site-specific environmental conditions. Multiple edaphic features can correlate with the restricted supply of orchid mycorrhizae within habitat patches, including soil properties such as moisture, organic content, pH and nutrient levels (Batty et al. 2001; Diez 2007; McCormick and Jacquemyn 2014). The amount and quality of soil organic material are determining for the occurrence in soil of saprotrophic fungi, such as Tulasnella spp., and the co-occurrence of particular plant species providing suitable organic material could represent an important aspect in defining the fungal distribution (Rasmussen et al. 2015). The D. fragrantissima native site has undergone habitat degradation, altered fire regime, and weed invasion during the last 30 years, likely resulting in alteration of the original orchid habitat. Therefore, vegetation compositional changes in the orchid native site may have made the habitat unable to support Tulasnella spp. persistence independently from the host plant.
Fungal diversity in the orchid rhizosphere and native site
In addition to the in situ distribution of Tulasnella spp., our study aimed to characterize the total fungal community in the orchid rhizosphere, as well as the orchid native site, in both occupied and non-occupied areas. A clear compositional shift was identified in the fungal communities from the rhizosphere compared to the bulk soil, irrespective of the distance from the host plant. This change in composition was mainly attributable to the presence of several taxa enriched in or unique to the rhizosphere. In addition to Tulasnella sp., two OTUs ascribed to the rhizoctonia-forming order Sebacinales have been recovered from the orchid rhizosphere exclusively, together with a heterogeneous group of non-mycorrhizae forming taxa, mainly belonging to the orders Agaricales (3 OTUs), Chaetotyriales (9 OTUs), Helotiales (5 OTUs), Hypocreales (13 OTUs), Pezizales (2 OTUs), and Pleosporales (3 OTUs) (Supplementary Table 1). Fungi from these orders have been previously isolated from orchid roots in temperate and tropical regions (Bayman and Otero 2006, Roy et al. 2009; Herrera et al. 2010, Tesitelova et al. 2015, Ma et al. 2015) and they are often regarded as root pathogens or saprobes in dead cells (Oja et al. 2015, Rasmussen et al. 2015, Ma et al. 2015). Although their role cannot be fully addressed in this study, association with saprotrophic fungi in the rhizosphere could represent a beneficial strategy for the plant to get access to growth-limiting resources in natural conditions. Indeed, the presence of saprotrophs is associated with an increased availability of organic matter from senescing root structures and exudates, thus increasing the probability of orchid survival under nutrient-poor environments (McCormick et al. 2006, Jacquemyn et al. 2012, Rasmussen et al. 2015, Ma et al. 2015, Lee et al. 2015).
Some of the taxa unique in the rhizosphere, such as Purpureocillium lavendulum and Cordyceps bassiana, are commonly found in plant rhizospheric soil (e.g. Lan et al. 2017, Ownley et al. 2008), and their presence implies both host protection against insects and plant growth enhancing functions. The enrichment of these fungi in the orchid rhizosphere community suggests that orchid roots and the immediate surroundings may stimulate growth and proliferation of selected beneficial fungi. In turn, the orchid rhizosphere may support the fungal community by offering a sheltered environment for survival and persistence (Selosse and Martos 2014, Oja et al. 2015).
Molecular traces of fungi from the family Geoglossaceae (e.g. Trichoglossum hirsutum) were observed to occur in the orchid rhizosphere (Supplementary Table 1). These taxa are usually associated with nutrient poor grasslands of the Northern hemisphere, being typically found in unfertilized or unimproved grasslands, lawns and pastures (Griffith et al. 2013). Although an association between Geoglossaceae and grassy plant roots has been hypothesized (Wang et al. 2011), the mechanisms and dynamics of this alliance are yet to be defined, preventing us to assess the role of these fungi in the Diuris rhizosphere.
In our survey, molecular evidence of the occurrence of both AM and ECM fungi in the D. fragrantissima ‘core’ rhizosphere community was observed (Supplementary Table 2A). ECM fungi have been previously isolated from both myco-heterotrophic (Martos et al. 2009; Ogura-Tsujita et al. 2009) and photosynthetic orchids roots (Bidartondo et al. 2004; Selosse et al. 2004; Dearnaley 2007), and their presence in the rhizosphere suggests a beneficial role in the acquisition of released nutrients, such as N and P (Nurfadilah et al. 2014). In contrast, arbuscular mycorrhizae have been found only occasionally in the Orchidaceae (Shefferson et al. 2005). Given the ability of AM to transport compounds between multiple plant species through common hyphal networks (Babikova et al. 2014), we hypothesize an active role for this fungal group in mediating the underground communication between D. fragrantissima and the surrounding plants. Further studies are required to corroborate this hypothesis on the role of arbuscular mycorrhizae in the orchid rhizosphere.
While the Diuris rhizosphere fungal community showed a sharp distinctiveness in composition, compositional changes between the orchid bulk soil and the site bulk soil were less pronounced (Figure 3). Consistent with previous studies (Porras Alfaro et al. 2011, Prober et al. 2015, Egidi et al. 2016), most of the fungi characterising the grassland belonged to the phyla Ascomycota, Glomeromycota and Basidiomycota, being mainly symbiotrophic (e.g. Glomus sp.), plant pathogens (e.g Ilyonectria mors-panacis, Drechslera sp.), dung saprotrophs (e.g. Podospora glutinans) and litter saprotrophs (e.g. Mortierella spp., Penicillium spp., Clavaria spp.) (Supplementary Table B and C). Several of these taxa have been previously reported from grassland soil also in association with biological soil crusts (States and Christensen 2001, Porras Alfaro et al. 2011). Penicillium and Mortierella in particular are commonly known as late-stage colonizers in decomposing litter (Osono and Takeda 2007), and their presence suggests an increase in the decomposition and organic material recycling rates, possibly contributing to preserve the soil nutrient status (Baath 1981). However, the retrieval of Clavaria spp. suggests a nutrient-poor ecosystem. Indeed, together with Geoglossaceae, members of the family Clavariaceae are traditionally considered indicator species for unmanaged grasslands (Griffith et al. 2013).
The diversity, taxonomy and ecology of fungal communities in Australian grassy ecosystems are still largely unexplored compared to their counterparts in the Northern hemisphere (McMullan-Fisher et al. 2011), hampering our ability to infer conclusions on the value of these taxa as nutrient status or grassland quality indicators in this situation. The low proportion of retrieved sequences matched at the species level highlights the importance of populating sequence databases with more sequences from material identified to species. Lack of species-level identifications precludes in depth analysis of the role of particular OTUs, which can be possible for individual known species, drawing on knowledge of their particular life history and distribution. However, even without species-level identifications, the community of molecular OTUs recovered in this study is available for future meta-analyses of the growing set of metabarcode data available for Australian environments (e.g. Tedersoo et al., 2014; Prober et al., 2015; Egidi et al. 2016) (GenBank accession number SUB2559717)
While the symbiont of the highly endangered Diuris fragrantissima was recovered from in situ plants, it was not detected elsewhere, among or near the extant orchid population. This suggests that a very limited number of potential recruitment sites with suitable Tulasnella spp. exist for D. fragrantissima. The absence of suitable fungi outside the orchid patch suggests that appropriate micro-sites are essentially saturated with orchids. Many factors or combinations of factors may affect the distribution of orchid fungi, which in turn may affect germination potential and distribution of orchid plants in the field. Given the dependence of terrestrial orchids on mycorrhizae for development and survival, correlating environmental factors to fungal distribution is critical for producing a self-perpetuating orchid population, ultimately the aim of any reintroduction activity. However, lack of detection of Tulasnella away from orchid roots means that it is not possible to make comparison against other variables. Further exploration of the presence of Tulasnella symbionts of common species of Diuris, where plant recruitment is occurring away from existing plants, will be instructive.
An unprecedented diversity of fungi has been retrieved from D. fragrantissima rhizosphere soil, suggesting interaction with multiple fungal partners in the naturally occurring Diuris population. More detailed investigations are required for characterization of fungi and fungal ecotypes associated with this endangered orchid, as well as their role in plant development. Overall, Next-Generation Sequencing technology has proved to be a powerful method for detection of the symbiont of a rare orchid in situ and for large-scale screening of the associated fungal community.
Commonwealth of Australia (1999). Environment Protection and Biodiversity Conservation Act 1999.
Bååth, E. (1981). Microfungi in a clear-cut pine forest soil in central Sweden. Canadian Journal of Botany, 59(7), 1331-1337.
Babikova, Z., Johnson, D., Bruce, T., Pickett, J., and Gilbert, L. (2014). Underground allies: How and why do mycelial networks help plants defend themselves? BioEssays, 36(1), 21-26.
Bahram, M., Peay, K. G., and Tedersoo, L. (2015). Local‐scale biogeography and spatiotemporal variability in communities of mycorrhizal fungi. New Phytologist, 205(4), 1454-1463.
Bálint, M., Schmidt, P. A., Sharma, R., Thines, M., & Schmitt, I. (2014). An Illumina metabarcoding pipeline for fungi. Ecology and evolution, 4(13), 2642-2653.
Batty, A. L., Dixon, K. W., Brundrett, M., and Sivasithamparam, K. (2001). Constraints to symbiotic germination of terrestrial orchid seed in a mediterranean bushland. New Phytologist, 152(3), 511-520.
Batty, A. L., Dixon, K. W., Brundrett, M. C., and Sivasithamparam, K. (2002). Orchid conservation and mycorrhizal associations. In Microorganisms in plant conservation and biodiversity (pp. 195-226). Springer Netherlands.
Bayman, P., and Otero, J. T. (2006). Microbial endophytes of orchid roots. In Microbial root endophytes (pp. 153-177). Springer Berlin Heidelberg.
Bidartondo, M. I., Burghardt, B., Gebauer, G., Bruns, T. D., and Read, D. J. (2004). Changing partners in the dark: isotopic and molecular evidence of ectomycorrhizal liaisons between forest orchids and trees. Proceedings of the Royal Society of London B: Biological Sciences, 271(1550), 1799-1806.
Brundrett, M. C., Scade, A., Batty, A. L., Dixon, K. W., and Sivasithamparam, K. (2003). Development of in situ and ex situ seed baiting techniques to detect mycorrhizal fungi from terrestrial orchid habitats. Mycological Research, 107(10), 1210-1220.
Brundrett, M. C. (2006). Understanding the roles of multifunctional mycorrhizal and endophytic fungi. In Microbial root endophytes (pp. 281-298). Springer Berlin Heidelberg.
Bonnardeaux, Y., Brundrett, M., Batty, A., Dixon, K., Koch, J., and Sivasithamparam, K. (2007). Diversity of mycorrhizal fungi of terrestrial orchids: compatibility webs, brief encounters, lasting relationships and alien invasions. Mycological Research, 111(1), 51-61.
Currah, R. S., Zelmer, C. D., Hambleton, S., and Richardson, K. A. (1997). Fungi from orchid mycorrhizas. In Orchid Biology (pp. 117-170). Springer Netherlands. Currah, R. S., Zelmer, C. D., Hambleton, S., and Richardson, K. A. (1997). Fungi from orchid mycorrhizas. In Orchid Biology (pp. 117-170). Springer Netherlands.
Dearnaley, J. D. (2007). Further advances in orchid mycorrhizal research. Mycorrhiza, 17(6), 475-486.
Dearnaley, J. D. W., Martos, F., and Selosse, M. A. (2012). 12 Orchid Mycorrhizas: Molecular Ecology, Physiology, Evolution and Conservation Aspects. In Fungal associations (pp. 207-230). Springer Berlin Heidelberg.
DEH 2000. Revision of the Interim Biogeographic Regionalisation of Australia (IBRA) and the Development of Version 5.1. – Summary Report. Department of the Environment and Heritage, Canberra.
Diez, J. M. (2007). Hierarchical patterns of symbiotic orchid germination linked to adult proximity and environmental gradients. Journal of Ecology, 95(1), 159-170.
Edgar, R. C. (2010). Search and clustering orders of magnitude faster than BLAST. Bioinformatics, 26(19), 2460-2461.
Egidi, E., McMullan-Fisher, S., Morgan, J. W., May, T., Zeeman, B., and Franks, A. E. (2016). Fire regime, not time-since-fire, affects soil fungal community diversity and composition in temperate grasslands. FEMS Microbiology Letters, 363(17), fnw196.
Gardes, M., and Bruns, T. D. (1993). ITS primers with enhanced specificity for basidiomycetes‐application to the identification of mycorrhizae and rusts. Molecular Ecology, 2(2), 113-118.
Griffith, G. W., Gamarra, J. G. P., Holden, E. M., Mitchel, D., Graham, A., Evans, D. A., … and Smith, S. L. (2013). The international conservation importance of Welsh ‘waxcap’ grasslands. Biological Conservation. #details#
Hadley, G. (1970). Non‐specificity of symbotic infection in orchid mycorrhiza. New Phytologist, 69(4), 1015-1023.
Harrison, J. G., Urruty, D. M., and Forister, M. L. (2016). An exploration of the fungal assemblage in each life history stage of the butterfly, Lycaeides melissa (Lycaenidae), as well as its host plant Astragalus canadensis (Fabaceae). Fungal Ecology, 22, 10-16.
Harley, J. L., and Smith, S. E. (1983). Mycorrhizal symbiosis. Academic Press Inc..
Herrera, P., Suárez, J. P., and Kottke, I. (2010). Orchids keep the ascomycetes outside: a highly diverse group of ascomycetes colonizing the velamen of epiphytic orchids from a tropical mountain rainforest in Southern Ecuador. Mycology, 1(4), 262-268.
Jacquemyn, H., Brys, R., Lievens, B., and Wiegand, T. (2012). Spatial variation in below‐ground seed germination and divergent mycorrhizal associations correlate with spatial segregation of three co‐occurring orchid species. Journal of Ecology, 100(6), 1328-1337.
Jacquemyn, H., Brys, R., Merckx, V. S., Waud, M., Lievens, B., and Wiegand, T. (2014). Coexisting orchid species have distinct mycorrhizal communities and display strong spatial segregation. New Phytologist, 202(2), 616-627.
Jiang, W., Liang, P., Wang, B., Fang, J., Lang, J., Tian, G., … and Zhu, T. F. (2015). Optimized DNA extraction and metagenomic sequencing of airborne microbial communities. Nature protocols, 10(5), 768-779.
Kõljalg, U., Larsson, K. H., Abarenkov, K., Nilsson, R. H., Alexander, I. J., Eberhardt, U., … and Pennanen, T. (2005). UNITE: a database providing web‐based methods for the molecular identification of ectomycorrhizal fungi. New Phytologist, 166(3), 1063-1068.
Kõljalg, U., Nilsson, R. H., Abarenkov, K., Tedersoo, L., Taylor, A. F., Bahram, M., … and Douglas, B. (2013). Towards a unified paradigm for sequence‐based identification of fungi. Molecular ecology, 22(21), 5271-5277.
Lan, X., Zhang, J., Zong, Z., Ma, Q., and Wang, Y. (2017). Evaluation of the Biocontrol Potential of Purpureocillium lilacinum QLP12 against Verticillium dahliae in Eggplant. BioMed Research International, 2017.
Leake, J. R. (1994). The biology of myco‐heterotrophic (‘saprophytic’) plants. New Phytologist, 127(2), 171-216.
Lee, Y. I., Yang, C. K., and Gebauer, G. (2015). The importance of associations with saprotrophic non-Rhizoctonia fungi among fully mycoheterotrophic orchids is currently under-estimated: novel evidence from sub-tropical Asia. Annals of botany, mcv085.
Liu, Q., Chen, J., Corlett, R. T., Fan, X., Yu, D., Yang, H., and Gao, J. (2015). Orchid conservation in the biodiversity hotspot of southwestern China. Conservation Biology, 29(6), 1563-1572.
Martos, F., Dulormne, M., Pailler, T., Bonfante, P., Faccio, A., Fournel, J., … and Selosse, M. A. (2009). Independent recruitment of saprotrophic fungi as mycorrhizal partners by tropical achlorophyllous orchids. New Phytologist, 184(3), 668-681.
Ma, X., Kang, J., Nontachaiyapoom, S., Wen, T., and Hyde, K. D. (2015). Non-mycorrhizal endophytic fungi from orchids. Current Science, 108, 1.
McCormick, M. K., Whigham, D. F., Sloan, D., O’Malley, K., and Hodkinson, B. (2006). Orchid–fungus fidelity: a marriage meant to last? Ecology, 87(4), 903-911.
McCormick, M. K., Lee Taylor, D., Juhaszova, K., Burnett, R. K., Whigham, D. F., and O’Neil, J. P. (2012). Limitations on orchid recruitment: not a simple picture. Molecular Ecology, 21(6), 1511-1523.
McCormick, M. K., and Jacquemyn, H. (2014). What constrains the distribution of orchid populations? New Phytologist, 202(2), 392-400.
McCormick, M. K., Taylor, D. L., Whigham, D. F., and Burnett, R. K. (2016). Germination patterns in three terrestrial orchids relate to abundance of mycorrhizal fungi. Journal of Ecology.
McKendrick, S. L., Leake, J. R., Taylor, D. L., and Read, D. J. (2000). Symbiotic germination and development of myco‐heterotrophic plants in nature: ontogeny of Corallorhiza trifida and characterization of its mycorrhizal fungi. New Phytologist, 145(3), 523-537.
McKendrick, S. L., Leake, J. R., Taylor, D. L., and Read, D. J. (2002). Symbiotic germination and development of the myco‐heterotrophic orchid Neottia nidus‐avis in nature and its requirement for locally distributed Sebacina spp. New Phytologist, 154(1), 233-247.
McMullan-Fisher Sapphire J. M., May Tom W., Robinson Richard M., Bell Tina L., Lebel Teresa, Catcheside Pam, York Alan (2011) Fungi and fire in Australian ecosystems: a review of current knowledge, management implications and future directions. Australian Journal of Botany 59, 70-90.
Murphy, A. H., Webster, A., Knight, C., and Lester, K. (2008). National Recovery Plan for the Sunshine Diuris Diuris fragrantissima. Department of Sustainability, East Melbourne.
Nurfadilah, S., Swarts, N. D., Dixon, K. W., Lambers, H., and Merritt, D. J. (2013). Variation in nutrient-acquisition patterns by mycorrhizal fungi of rare and common orchids explains diversification in a global biodiversity hotspot. Annals of botany, 111(6), 1233-1241.
Nguyen, N. H., Smith, D., Peay, K., and Kennedy, P. (2015). Parsing ecological signal from noise in next generation amplicon sequencing. New Phytologist, 205(4), 1389-1393.
Nguyen, N. H., Song, Z., Bates, S. T., Branco, S., Tedersoo, L., Menke, J., … and Kennedy, P. G. (2016). FUNGuild: an open annotation tool for parsing fungal community datasets by ecological guild. Fungal Ecology, 20, 241-248.
Ogura-Tsujita, Y., Gebauer, G., Hashimoto, T., Umata, H., and Yukawa, T. (2009). Evidence for novel and specialized mycorrhizal parasitism: the orchid Gastrodia confusa gains carbon from saprotrophic Mycena. Proceedings of the Royal Society of London B: Biological Sciences, 276(1657), 761-767.
Oja, J., Kohout, P., Tedersoo, L., Kull, T., and Kõljalg, U. (2015). Temporal patterns of orchid mycorrhizal fungi in meadows and forests as revealed by 454 pyrosequencing. New Phytologist, 205(4), 1608-1618.
Oja, J., Vahtra, J., Bahram, M., Kohout, P., Kull, T., Rannap, R., … and Tedersoo, L. (2016). Local-scale spatial structure and community composition of orchid mycorrhizal fungi in semi-natural grasslands. Mycorrhiza, #vol# 1-13.
Oksanen, J., Blanchet, F. G., Kindt, R., Legendre, P., Minchin, P. R., O’hara, R. B., … and Oksanen, M. J. (2013). Package ‘vegan’. Community ecology package, version, 2(9).
Osono, T., and Takeda, H. (2007). Microfungi associated with Abies needles and Betula leaf litter in a subalpine coniferous forest. Canadian journal of microbiology, 53(1), 1-7.
Otero, J. T., Flanagan, N. S., Herre, E. A., Ackerman, J. D., and Bayman, P. (2007). Widespread mycorrhizal specificity correlates to mycorrhizal function in the neotropical, epiphytic orchid Ionopsis utricularioides (Orchidaceae). American Journal of Botany, 94(12), 1944-1950.
Ownley, B. H., Griffin, M. R., Klingeman, W. E., Gwinn, K. D., Moulton, J. K., and Pereira, R. M. (2008). Beauveria bassiana: endophytic colonization and plant disease control. Journal of invertebrate pathology, 98(3), 267-270.
Peay, K. G., Kennedy, P. G., and Bruns, T. D. (2011). Rethinking ectomycorrhizal succession: are root density and hyphal exploration types drivers of spatial and temporal zonation? Fungal Ecology, 4(3), 233-240.
Perkins, A. J., and McGee, P. A. (1995). Distribution of the orchid mycorrhizal fungus, Rhizoctonia solani, in relation to its host, Pterostylis acuminata, in the field. Australian Journal of Botany, 43(6), 565-575.
Phillips, R. D., Barrett, M. D., Dixon, K. W., & Hopper, S. D. (2011). Do mycorrhizal symbioses cause rarity in orchids?. Journal of Ecology, 99(3), 858-869.
Porras-Alfaro, A., Herrera, J., Natvig, D. O., Lipinski, K., and Sinsabaugh, R. L. (2011). Diversity and distribution of soil fungal communities in a semiarid grassland. Mycologia, 103(1), 10-21.
Prober, S. M., Leff, J. W., Bates, S. T., Borer, E. T., Firn, J., Harpole, W. S., … and Cleland, E. E. (2015). Plant diversity predicts beta but not alpha diversity of soil microbes across grasslands worldwide. Ecology Letters, 18(1), 85-95.
Rasmussen, H. N., Dixon, K. W., Jersáková, J., and Těšitelová, T. (2015). Germination and seedling establishment in orchids: a complex of requirements. Annals of botany, mcv087.
Roberts, P. (1999). Rhizoctonia-forming fungi. Herbarium, Royal Botanic Gardens.
Roy, M., Watthana, S., Stier, A., Richard, F., Vessabutr, S., and Selosse, M. A. (2009). Two mycoheterotrophic orchids from Thailand tropical dipterocarpacean forests associate with a broad diversity of ectomycorrhizal fungi. BMC biology, 7(1), 51.
Schoch, C. L., Seifert, K. A., Huhndorf, S., Robert, V., Spouge, J. L., Levesque, C. A., … and Miller, A. N. (2012). Nuclear ribosomal internal transcribed spacer (ITS) region as a universal DNA barcode marker for Fungi. Proceedings of the National Academy of Sciences, 109(16), 6241-6246.
Selosse, M. A., Faccio, A., Scappaticci, G., and Bonfante, P. (2004). Chlorophyllous and achlorophyllous specimens of Epipactis microphylla (Neottieae, Orchidaceae) are associated with ectomycorrhizal septomycetes, including truffles. Microbial Ecology, 47(4), 416-426.
Selosse, M. A., and Martos, F. (2014). Do chlorophyllous orchids heterotrophically use mycorrhizal fungal carbon?. Trends in plant science, 19(11), 683-685.
Shefferson, R. P., WEIß, M. I. C. H. A. E. L., Kull, T. I. I. U., and Taylor, D. (2005). High specificity generally characterizes mycorrhizal association in rare lady’s slipper orchids, genus Cypripedium. Molecular Ecology, 14(2), 613-626.
Sivasithamparam, K. (1993). Ecology of root infecting pathogenic fungi in Mediterranean environments. Advances in Plant Pathology.
Smith, Z. F., James, E. A., and McLean, C. B. (2010). Mycorrhizal specificity of Diuris fragrantissima (Orchidaceae) and persistence in a reintroduced population. Australian Journal of Botany, 58(2), 97-106.
Smith, Z. (2006). Biology and Reintroduction of the Threatened Terrestrial Orchid Diuris fragrantissima (Sunshine Diuris). PhD dissertation.
States, J. S., Christensen, M., and Kinter, C. L. (2001). Soil fungi as components of biological soil crusts. Biological Soil Crusts: Structure, Function, and Management, 155-166.
Team, R. C. R: A language and environment for statistical computing. R Foundation for Statistical Computing; Vienna, Austria: 2015. version 3.2. 0.
Tedersoo, L., Bahram, M., Põlme, S., Kõljalg, U., Yorou, N. S., Wijesundera, R., … and Smith, M. E. (2014). Global diversity and geography of soil fungi. Science, 346(6213), 1256688.
Teˇšitelová, T., Teˇšitel, J., Jersáková, J., Rˇíhová, G., and Selosse, M. A. (2012). Symbiotic germination capability of four Epipactis species (Orchidaceae) is broader than expected from adult ecology. American Journal of Botany, 99(6), 1020-1032.
Taylor, D. L., and McCormick, M. K. (2008). Internal transcribed spacer primers and sequences for improved characterization of basidiomycetous orchid mycorrhizas. New Phytologist, 177(4), 1020-1033.
Voyron, S., Ercole, E., Ghignone, S., Perotto, S., and Girlanda, M. (2016). Fine‐scale spatial distribution of orchid mycorrhizal fungi in the soil of host‐rich grasslands. New Phytologist.
Wang, Z., Nilsson, R. H., Lopez-Giraldez, F., Zhuang, W. Y., Dai, Y. C., Johnston, P. R., and Townsend, J. P. (2011). Tasting soil fungal diversity with earth tongues: phylogenetic test of SATe alignments for environmental ITS data. PLoS One, 6(4), e19039.
Warcup, J. H. (1981). The mycorrhizal relationships of Australian orchids. New Phytologist, 87(2), 371-381.
Waud, M., Busschaert, P., Lievens, B., and Jacquemyn, H. (2016). Specificity and localised distribution of mycorrhizal fungi in the soil may contribute to co-existence of orchid species. Fungal Ecology, 20, 155-165.
Weiss, M., Selosse, M. A., Rexer, K. H., Urban, A., and Oberwinkler, F. (2004). Sebacinales: a hitherto overlooked cosm of heterobasidiomycetes with a broad mycorrhizal potential. Mycological Research, 108(9), 1003-1010.
White TJ, Bruns T, Lee SJ et al. (1990) Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. PCR protocols: a guide to methods and applications, 18:315-322.
Figure 1 Experimental workflow
Figure 2 Relative abundance of (A) fungal phyla, (B)fungal trophic modes, and (C) fungal guilds, compared across the orchid rhizosphere, the orchid bulk soil (at increasing distance from the host plant) and the site bulk soil. The functional analyses included only the trophic modes and guilds with a ‘highly probable’ confidence level (N=705 OTUs).
Figure 3 Cluster analysis illustrating the similarity of fungal communities in samples retrieved from the orchid rhizosphere, the orchid bulk soil and the site bulk soil. Clustering was generated by the UPGMA algorithm, using a Bray-Curtis similarity matrix generated from the relative abundances of the rarefied OTU table. The four main clusters are highlighted in grey. Broken lines mark the non-significantly different branches, according to SIMPROF analysis (p > 0.05). Circles = orchid rhizosphere soil; Quadrats = orchid bulk soil; Triangles = site bulk soil; Inverted triangles = site bulk soil from the orchid-occupied area.
Supplementary Figure 1 Rarefaction (A) and species accumulation curves (B) for samples retrieved from the orchid rhizosphere, the orchid bulk soil and the site bulk soil, respectively.
Supplementary Figure 2 Non-metric MDS ordination based on OTU composition of the soil fungal community in the three soil sources (999 permutations). Total R= 0.488, p= 0.001.
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