Skip to main content

Advertisement

You are viewing the new article page. Let us know what you think. Return to old version

Dynamics of fungal communities during Gastrodia elata growth

Abstract

Background

Gastrodia elata is a widely distributed achlorophyllous orchid and is highly valued as both medicine and food. Gastrodia elata produces dust-like seeds and relies on mycorrhizal fungi for its germination and growth. In its life cycle, G. elata is considered to switch from a specific single-fungus relationship (Mycena) to another single-fungus relationship (Armillaria). However, no studies have investigated the changes in the plant-fungus relationship during the growth of G. elata in the wild. In this study, high-throughput sequencing was used to characterize the fungal community of tubers in different growth phases as well as the soils surrounding G. elata.

Results

The predominant fungi were Basidiomycota (60.44%) and Ascomycota (26.40%), which exhibited changes in abundance and diversity with the growth phases of G. elata. Diverse basidiomycetes in protocorms (phase P) were Hyphodontia, Sistotrema, Tricholoma, Mingxiaea, Russula, and Mycena, but the community changed from a large proportion of Resinicium bicolor (40%) in rice-like tubers (phase M) to an unidentified Agaricales operational taxonomic unit 1(OTU1,98.45%) in propagation vegetation tubers (phase B). The soil fungi primarily included Simocybe, Psathyrella, Conocybe, and Subulicystidium. Three Mycena OTUs obtained in this study were differentially distributed among the growth phases of G. elata, accounting for less than 1.0% of the total reads, and were phylogenetically close to Mycena epipterygia and M. alexandri.

Conclusions

Our data indicated that G. elata interacts with a broad range of fungi beyond the Mycena genus. These fungi changed with the growth phases of G. elata. In addition, these data suggested that the development of the fungal community during the growth of G. elata was more complex than previously assumed and that at least two different fungi could be involved in development before the arrival of Armillaria.

Background

Orchidaceae has been considered to be the most diverse and widely distributed plant family on earth [1]. Because orchids produce tiny, dust-like seeds [2], which lack the necessary energy reserves, a variety of mycorrhizal fungi are required to stimulate their germination and seedling growth, a strategy termed mycoheterotrophy [3, 4]. All orchids form protocorms at the beginning of germination [4, 5]. In the basal region of the protocorm, most plant cells are observed to harbor viable, coiled intracellular fungal hyphae (the pelotons) that eventually collapse and undergo lysis. After this early phase of seedling, most orchids develop green leaves and photosynthesize and are known as partial mycoheterotrophs. However, there are also > 200 achlorophyllous orchid species, which obtain their complete nutrients and carbon supply from mycorrhizal fungi and are known as full mycoheterotrophs [1, 6].

Because orchids are partially or fully dependent on mycorrhizal fungi, the mycorrhizal communities associated with orchids could be expected to mediate the abundance, spatial distribution and coexistence of terrestrial orchids in natural communities [7]. Some species of orchids have a narrow taxonomic group of mycorrhizal fungi [8]. For example, the fully mycoheterotrophic orchid Corallorhiza maculata is colonized exclusively by the same highly specific ectomycorrhizal fungi throughout the year [9]. Shefferson et al. (2008) found that the mycorrhizal fungi associated with orchids that colonized mine tailing hills were the same as those in pristine habitats [10]. Jacquemyn et al. (2014) indicated that cooccurring orchid species have distinctive mycorrhizal communities and show strong spatial segregation [11]. However, the easy germination and protocorm formation of most orchids in unoccupied habitats strongly suggest the ubiquitous occurrence and low specialization of mycorrhizal fungi [12,13,14]. For example, species in the genus Dactylorhiza have been associated with a diverse suite of mycorrhizal fungi [15]. Tĕšitelová et al. (2012) found evidence for low fungal specificity in four Epipactis species with divergent ecological preferences [16]. In addition, Cynorkis purpurea was found to be a generalist with diverse mycorrhizal fungi spanning 12 operational taxonomic units (OTUs) in three genera (Tulasnella, Ceratobasidium, and Sebacina) [17].

Interestingly, throughout their life history, the fungal specificity patterns in orchid seedlings and adults vary among orchid species [18]. The same fungi facilitate both seed germination and nutrition of the adult plants of Drakaea spp. [19]. However, other species switch from a single-fungus relationship to relationships with a variety of fungi, e.g., Tipularia discolor [20] and Orchis purpurea [21]. In addition, seedlings of Cephalanthera or Epipactis associate with a selection of fungi and continue with a wider selection in adulthood [16, 22].

Gastrodia elata is an achlorophyllous orchid and is primarily distributed in East Asia, Southeast Asia and Oceania [23, 24]. Its rhizome (mature tuber) is a popular herbal medicine in Asian countries, such as Korea, China and Japan [25, 26]. The rhizome has also been used as a food and is commonly cooked in soups. Like many other orchids, G. elata produces thousands of dust-like seeds (10,000–50,000 seeds per capsule) [27]. These seeds do not possess an endosperm and thus have to germinate and develop into protocorms, forming vegetative propagation corms and immature tubers only when adequate nutrients are obtained through the digestion of the specific fungi, Mycena spp. G. elata subsequently undergoes vegetative growth through an established symbiotic association with the compatible mycorrhizal fungus Armillaria mellea to yield mature tubers [28, 29]. Therefore, G. elata is widely considered to switch from a specific single-fungus relationship (Mycena) to another single-fungus relationship (Armillaria) from germination to adult growth [30]. Some species of Gastrodia, for example, the adult G. confusa, also gain carbon through a narrow taxonomic group of Mycena species [31].

However, other Gastrodia species have been shown to associate with diverse litter- or wood-decaying fungi, e.g., Resinicium in G. similis [32] and Campanella and Marasmius in G. sesamoides [33]. Although three closely related species—G. confusa, G. pubilabiata, and G. nipponica—have different fungal specificities, no shift of fungal partners in the course of the life history occurs [34]. There have been no detailed studies on the fungal community associated with G. elata in the wild since 1989, when the fungus Mycena osmundicola was isolated from G. elata protocorms for the first time [35]. Although in vitro studies have shown that several Mycena species, including M. orchidicola, M. dendrobii, and M. anoectochili, promote the germination of G. elata, these fungi were isolated from the orchids Cymbidium sinense, Dendrobium candidum, and Anoectochilus roxburghii, respectively [36]. Therefore, a final conclusion has not yet been reached on the fungal switch during the development phases of G. elata.

To accurately describe mycorrhizal associations in G. elata, fungal communities should be assessed rather than the presence of individual fungal species, as has been previously shown [29]. In addition, more advanced detection techniques are required rather than traditional isolation and culture methods. Novel high-throughput sequencing methods, such as Illumina sequencing and PacBio sequencing, have recently been used to explore the fungal communities associated with orchids, including species from five different genera (Anacamptis, Neotinea, Orchis, Ophrys and Serapias) in Gargano National Park in southern Italy [11] and G. flavilabela in Taiwan [24]. The present study is the first example of the use of high-throughput sequencing to characterize the fungal community associated with G. elata in Zhaotong, China. We aimed to understand (1) the dynamics of the fungal community associated with G. elata in different developmental phases, (2) whether the Mycena species comprise the majority of fungi associated with G. elata during early development phases, and (3) the overlap between the fungal communities associated with G. elata and those in the surrounding soils.

Results

Fungal samples and ITS sequences

From the genomic DNA of each sample, the internal transcribed spacer 1 (ITS1) rDNA segments were amplified, and the PCR products were subjected to Illumina MiSeq sequencing. A total of 402,117 high-quality reads were obtained from 12 samples (including nine plant tuber samples and three methodological replicates of soil, which were performed to increase the recovery of fungi from the pooled soil samples), and 1000 OTUs were observed at a 97% similarity. Each sample contained 4531 to 41,185 reads with different phylogenetic OTUs ranging from 30 to 597 (additional table files provide greater detail [see Additional file 1: Table S1 and Additional file 2: Table S2]). After subsampling, 779 OTUs were observed at 97% similarity, with different OTUs ranging from 14 to 373 (additional files show these findings in greater detail [see Additional file 1: Table S1 and Additional file 3: Table S3]). The phase B samples provided the fewest OTUs and the greatest number of unidentified fungi OTUs. The OTU table after subsampling offers greater detail (see Additional file 3: Table S3).

All the rarefaction curves tended to approach the saturation plateau, suggesting a reasonable number of sequenced reads in each sample. The species accumulation curves showed that these fungal diversity indices declined with the growth of G. elata (Additional file 4: Figure S1).

Fungal communities of the tubers and surrounding soils

At the phylum level, the predominant fungi in all the samples, including the tubers and soils, were from Basidiomycota and Ascomycota, with average abundances of 60.44 and 26.40%, respectively (Fig. 1a). The annotated fungi in the protocorm (phase P) belonged to five phyla, with the three most abundant phyla being Ascomycota, Basidiomycota and Zygomycota, accounting for 42.33, 39.76 and 15.21%, respectively. The fungi of rice-like G. elata (phase M) were assigned to three phyla. Among them, Basidiomycota and Ascomycota were dominant, accounting for 39.29 and 36.62%, respectively. The fungi of the propagation vegetation tubers (phase B) included three phyla, with 98.48% from Basidiomycota. Basidiomycota and Ascomycota accounted for 60.44 and 26.41%, respectively, in the surrounding soils.

Fig. 1
figure1

Distribution histogram of fungal community structure. Distribution histogram of fungal community structure at the phylum level (a) in Basidiomycota (b) and in Ascomycota (c). “Other” includes those nonfungal and unidentified sequences at the phylum level. P: protocorms; M: rice-like G. elata; B: propagation vegetation tubers; and S: surrounding soils

At the genus level, the distribution of Basidiomycota and Ascomycota varied among the growth phases of G. elata (Fig. 1b, c). The abundance of Basidiomycota typically increased with the growth phases of G. elata (Fig. 1b). Although relatively diverse fungi were found in phase P, including Hyphodontia (7.8%), Sistotrema (4.58%), Tricholoma (3.58%), Mingxiaea (2.66%), Russula (1.79%), and Mycena (1.47%), in total, these genera accounted for nearly 20% of fungi in this phase. Resinicium was numerically dominant in phase M, with a percentage of 39.30%. Much less diverse fungi were found in the propagation vegetation tubers (phase B), and more than 98.45% were in Agaricales. Soil fungi belonged to four genera with an average percentage > 2.0%, including Simocybe (40.12%), Psathyrella (6.35%), Conocybe (3.91%), and Subulicystidium (2.30%).

In contrast, the abundance of Ascomycota decreased with the growth phases of G. elata (Fig. 1c). The fungi in phase P were primarily Mariannaea (11.07%), Trichoderma (6.48%), Clonostachys (5.56%), Lecanicillium (2.20%) and Cladophialophora (2.06%). However, the majority of fungi in phase M were Trichoderma (13.29%) and Clonostachys (11.08%), and phase B rarely contained Ascomycota fungi. The soils primarily contained Trichoderma (1.96%), Lasiosphaeris (1.70%), Ascotaiwania (1.21%) and Bactrodesmiastrum (1.02%).

Consequently, the distribution of the top 10 OTUs, which accounted for 63.65% of the total fungi detected, including six basidiomycetes, three ascomycetes and one unidentified fungus, were used to compare the difference in the fungal community among groups (Fig. 2, also see Additional file 3: Table S3 for the OTU table after subsampling). The average abundances of OTU71 (Mariannaea samuelsii, Ascomycota), OTU138 (Hyphodontia barba-jovis, Basidiomycota) and OTU271 (Hydnodontaceae, Basidiomycota) declined with the growth of G. elata. OTU71 and OTU271 were primarily found in the protocorms. Two Simocybe spp. from Basidiomycota, including OTU313 and OTU305, were almost exclusively detected in soils. An Agaricales OTU1 was the most abundant in phase B. In addition; other OTUs, including OTU16 (Resinicium bicolor, Basidiomycota), OTU43 (unidentified fungus), OTU51 (Clonostachys sp., Ascomycota) and OTU61 (Trichoderma harzianum, Ascomycota), showed an increase in the first two phases of G. elata (P and M) and then a decrease in phase B. Totally, the numbers of unique OTUs from different groups were much higher than those of shared OTUs (Fig. 3, also see Additional file 5: Table S4).

Fig. 2
figure2

Heatmap of the distribution of the top 10 OTUs among the different groups. The relative abundances are expressed as the richness of fungi transformed by log2(x + 1) in a given sample. P: protocorms; M: rice-like G. elata; B: propagation vegetation tubers; and S: surrounding soils

Fig. 3
figure3

Venn diagram for the four fungal communities. The numbers are the OTU number in each part. P: protocorm; M: rice-like tubers; B: propagation vegetation tubers; S: surrounding soils

Phylogenetics and distribution of Mycena sp. during G. elata germination

Three Mycena OTUs, designated OTU117, OTU148 and OTU155, were obtained in this study. Phylogenetically, all were combined into one clade with the two species M. epipterygia and M. alexandri. The strain BZZ was positioned in a different clade (Fig. 4). These three OTUs were distributed differently among the growth phases of G. elata. Phase M contained all three OTUs, but P only contained OTU155, while B had none. The surrounding soils contained only OTU117. The percentage of OTU155 was relatively higher than that of OTU117 and OTU148. However, in general, Mycena occupied a very low percentage of the total reads (an average of less than 1.0%).

Fig. 4
figure4

Phylogenetic tree of Mycenaspp. Three Mycena detected in this study were designated OTU117, OTU155 and OTU117. The strain BZZ was previously isolated from a protocorm of G. elata. Bootstrap values of > 50% are indicated at the branch nodes. GenBank accession numbers and their geographic isolation sources of reference sequences are shown in parentheses. The scale bar represents 3% dissimilarities among different Mycena sp.

Discussion

Currently, our knowledge of the fungal biology of G. elata is very limited, although this species is widely distributed and has high medicinal value [23, 25, 26]. This study is the first to use culture-independent molecular techniques to characterize the fungal community associated with G. elata. As previously reported for most other orchids [18], the broader range of fungi than previously assumed, involving many fungal families from the phyla Basidiomycota and Ascomycota, may be related to the germination and vegetation growth of G. elata (Fig. 1).

The most important finding in this study is that the fungal communities associated with G. elata changed with the developmental phases (Figs. 1, 2). Generally, seedling-mycobiont specificity appears to be narrower than in adult plants [21, 22]. However, fungal diversities declined with development phases of G. elata in this case (also see Additional file 4: Figure S1). For Basidiomycota, no OTUs with known names overlapped between the soils and G. elata tubers, with the exception of several rare unidentified OTUs (Fig. 1b). With the developmental phases of G. elata, the fungal species richness declined but the relative abundance of predominate OTUs increased, from diverse basidiomycetes (11 genera, with a percentage of 20%) in phase P to a single Agaricales OTU1 in phase B (with a percentage of 98%). In phase P, the shared genera in three replicates included Hyphodontia (Trechisporales), Sistotrema (Cantharellales), Russula (Russulales), and Mingxiaea (Tremellales). Hyphodontia, Sistotrema and Russula are common in the fungal communities associated with orchids, e.g., Erythrorchis altissima [37], Corallorhiza [38], and Limodorum [18, 39]. There is no report of Mingxiaea as a mycorrhizal fungus of orchids. The genus Resinicium is common in Gastrodia species, e.g., G. similis [32] and G. sesamoides [33]. In the present study, one species, Resinicium bicolor, was numerically dominant in phase M, but it was rarely observed in phase P and the surrounding soils and was completely absent in phase B. These OTUs were rare in soils. Interestingly, fungal species had limited overlap between soil and the tubers of G. elata (Figs. 1, 2). In soils, the dominant fungi, including Simocybe, Psathyrella and Conocybe, were absent in the G. elata tubers, regardless of the growth phase (Fig. 1). Among these genera, only Psathyrella was previously observed in the orchid Neottia ovata [40]. Similarly, the dominant fungi in the G. elata tubers, e.g., Resinicium, Mariannaea, Trichoderma and Clonostachys, were rare fungi in the soil samples. Hyphodontia and Sistotrema, which were the dominant fungi in group P, were absent in the soil samples (Fig. 1). However, it was possible to detect these fungi with increased sequencing depth. Our data indicated that the fungal colonization in G. elata was highly dynamic, and G. elata was likely to select different basidiomycete fungi throughout the growth process.

Diverse ascomycetes, including Clonostachys, Trichoderma, Mariannaea, Cladophialophora, Lecanicillium, Chaetosphaeria, Leptodontidium, and Fusarium, were observed in G. elata as well as in the soils (Fig. 1c). Most of these genera are common nonmycorrhizal endophytes associated with orchids [37, 41]. Clonostachys rosea and Trichoderma chlorosporum acted as potential biocontrol agents and promoted the growth of the orchid Dendrobium nobile [41]. Trichoderma harzianum was proven to significantly promote the seedling survival and growth of the orchid Guarianthe skinneri [42]. Fusarium, as an endophytic fungus, has been found to play an important role in the growth and survival of D. friedericksianum [43]. However, there is controversy about the role of these nonmycorrhizal endophytes in orchids. It is possible that surface contaminants could be mistakenly identified as orchid nonmycorrhizal endophytes [44]. Because no certain role of ascomycetes associated with G. elata is reported, one should be cautious about their role in the growth of G.elata.

There is conflict regarding the fungal specificity of Gastrodia species. Some authors think that Gastrodia species have a strong preference to associate with Mycena fungi, e.g., Mycena cf. quiniaultensis and Mycena chlorophos in G. flavilabela [24] and several Mycena species in G. confusa [31]. Since the fungus M. osmundicola has been isolated and identified [28, 30], no novel fungi beyond the Mycena species have been reported to associate with G. elata in the wild until now, which misread that G. elata likely has a strong specificity for the fungal family Mycenaceae at the early stage of growth. However, in our study, the proportion of Mycena was very low across the different growth phases of G. elata as well as in the surrounding soils (also see Additional file 6: Table S5 for the distribution of Mycena spp. in each sample). Diverse Mycena species have been previously proven in vitro to promote the seed germination of G. elata; however, these fungi were isolated from other orchids [36, 45]. In addition, several studies have shown that another fungal species in six genera isolated from different species of orchids are also capable of promoting the seed germination of G. elata [46]. Therefore, our data indicated that the fungi involved in the germination of G. elata could come from outside the family Mycenaceae. Interestingly, our data indicated that three OTUs previously reported in Italy were grouped with M. epipterygia [47] and M. alexandri in Sweden (Fig. 4). In addition, no OTUs from Mycena in this study were closely related to M. osmundicola, a fungus widely applied in the artificial cultivation of G. elata [28, 30]. Mycena strain BZZ, which is currently used in the cultivation of G. elata in Zhaotong, was closely related to a strain Mycena sp. (Fig. 4), which was isolated from G. nipponica in Japan [34]. These data suggest that Mycena strains being able to promote the germination of G. elata may be distributed globally.

Traditionally, G. elata is considered to switch from a specific single-fungus relationship (Mycena) to another single-fungus relationship (Armellaria) during its life cycle [30, 48]. Mycena must be replaced by A. mellea late in protocorm development [28], assuming that this replacement provides the G. elata with access to the greater reserves of organic carbon contained in the massive woody substrates exploited by A. mellea [49, 50]. Our molecular data suggested that the switch was more complex than previously assumed. Because diverse fungi are associated with different growth phases of G. elata (Figs. 1, 2; also see the discussion above), multiple fungal switches may occur at different development phases after germination, e.g., from Resinicium in phase M to unknown Agaricales OTU1 in phase B. There is evidence that diverse litter- or wood-decaying fungi, such as Resinicium, Campanella and Marasmius species, can provide carbon and nutrients to Gastrodia species [32, 33]. Therefore, our data indicated that at least two different fungi other than Mycena species might be involved in development before the arrival of Armillaria to sustain the growth of G. elata into mature tubers. To achieve this, these two fungi need to be cultured and reinoculated successfully, as previously indicated [18]. Unfortunately, our multiple efforts to culture these two fungi failed.

Wild orchids have a broad range of germination stages characterized by a high degree of variability in either the timing of seed germination or the rate of seed development. This variability could have primarily resulted from the absence of mycelia of appropriate fungi at the seed microsite [3]. Therefore, the presence of specific mycorrhizal fungi contributes, at least partly, to the spatial distribution and coexistence of orchid species [51]. Recent studies suggest that mycorrhizal fungi have an aggregated distribution within the habitats of orchids [52]. Higher abundances of fungal symbionts are typically found close to adult plants [3, 53]. In a study of the fungi Tulasnella spp. associated with G. pubescens, McCormick et al. (2016) found that established plants of G. pubescens that are < 50 cm apart associate with a single abundant fungal genet, while those > 50 cm apart associate with multiple fungal genets [54]. In addition to the fungal community associated with the development of G. elata, we originally tried to design our experiments to assess the spatial variation of the fungi. However, it is difficult to contain all the growth phases of G. elata in a single hole (microsite). After nearly one year of cultivation, some holes had no germination (empty holes) and some halted at protocorms with no further development into immature tubers (e.g., phases M and B). Therefore, we had to mix tubers at the same growth phase from different holes (also see Additional file 7: Table S6 The tubers at different growth phases in five holes). These data indirectly indicated that the distribution of fungi associated with G. elata in this site was highly heterogeneous; therefore, it was worth detailing the fungal species inhabiting soils and plant litters in these microsites. Finally, no A. mellae were detected in any samples in this study, and the possible reason was the absence or very low occurrence of this fungus in this site. Although our report revealed the novelty in fungal communities associated with G. elata during the developmental phases, only few holes were successfully germinated in one geographic site, which undoubtedly caused a degree of fungal community bias. In the future, more research is needed to determine whether Resinicium and OTU1 are common fungal switches during different development phases and in different habitats of G. elata and at what time point following these switches Armillaria finally arrives to complete the life cycle of G. elata.

Conclusions

Gastrodia elata is an important herbal medicine in Asian countries. However, there has been no detailed study on the fungal community associated with the growth phases of G. elata in the wild since 1989. In the present study, G. elata in Zhaotong, a core production area in China, was selected, and high-throughput sequencing was used to characterize its fungal community. Tubers with different growth phases, along with their surrounding soils, were collected and sequenced. The data revealed that G. elata was associated with a broad range of fungi beyond Mycena species. These fungi changed with the growth phases of G. elata. In addition, this study suggested a complex switch of the fungal community with the growth of G. elata. The Mycena spp. were clearly not the majority of fungi associated with the germination and early growth of G. elata. This study is the first report on the fungal dynamics of G. elata and will provide key insight into the G. elata –mycorrhizal fungal interaction in the future.

Methods

Description of the sampling site

Zhaotong is located in the northeast of Yunnan Province, China, and is well known for its high-quality G. elata production [55]. Our study site was thus selected in the core producing area of G. elata, Xiaocaoba, Zhaotong (27°48′7.99 N, 104°15′57.96E). This site grows sporadic trees and shrubs, including Sorbus pohuashanensis, Castanopsis delavayi, Dipentodon sinicus, Corylus chinensis, Phoebe zhennan, Cerasus serrula, Acer oliverianum, Alnus cremastogyne, and Cunninghamia sp. Traditionally, wild G. elatawas distributed in this area. To bait mycorrhizal fungi, 10 planting holes (80 cm (L) × 50 cm (W) × 20 cm (H)) were randomly dug in nearly 100 m2 of the area in July 2016. The litter of the trees (primarily Fagaceae tree leaves, which are traditionally thought to be good sources for mycorrhizal fungi growth on G. elata [48]) were collected and laid on the bottom of holes. Seeds of G. elata (from 8 to 10 capsules per hole) were well distributed onto plant materials and covered with a 5–10 cm layer of soil. In August 2017, to obtain G. elata tubers at different development phases, these holes were uncovered and G. elata tubers were collected (Fig. 5). Unfortunately, a half of holes failed to germinate. Among the holes with tubers, most of them failed to contain all the growth phases of G. elata in a single hole (see Additional file 7: Table S6 The tubers at different growth phases in five holes). We collected all tubers and categorized these tubers according to their size into three groups representing different growth phases of G. elata, including protocorms (phase P, size 0.5–1 mm), rice-like tubers (phase M, size 3-5 mm), and propagation vegetation tubers (phase B, size > 8 mm). The different phases of G. elata tubers from different holes were evenly mixed in equal proportions and divided into three replicates, which contained dozens of tuber individuals without damage in order to avoid the cross contamination of endophytic fungi. The G. elata tubers were washed with a large amount of sterile water after two minutes of surface disinfection with 0.5% sodium hypochlorite and crushed, then immediately stored at − 80 °C for subsequent analysis. To verify whether there is an overlap of fungal community between G. elata tubers and the surrounding soils, we collected 500 g of soil from each hole and a total of 2500 g of soil was obtained from 5 holes. Then, we repeatedly mixed the soils using method of coning and quartering until they were well mixed, after which we randomly took three subsamples of soils as methodological replicates to increase the recovery of fungi from the pooled samples. Soils were immediately stored at − 80 °C for subsequent analysis. In this study, a total of 12 samples, including three 250 mg samples of tubers from each growth phase and three 250 mg samples of soils, were used for DNA extraction.

Fig. 5
figure5

Sampling sites of G. elata. Planting holes of G. elata in the wild (a); the uncovered planting holes with G. elata tubers (b and c); the different growth phases of G. elata (d). P: protocorm; M: rice-like G. elata; B: propagation vegetation tubers of G. elata

DNA extraction and library construction

The genomic DNA of the plant tubers was extracted using CTAB [56]. Genomic DNA from the surrounding soils was extracted using a commercial DNA extraction kit (PowerSoil® DNA Isolation Kit, USA). The DNA quality was monitored using 0.8% agarose gel electrophoresis, and the extracted DNA was diluted to a concentration of 1 ng/μL and stored at − 20 °C until further processing. The diluted DNA was used as a template for the PCR amplification of fungal ITS genes with the barcoded primers and a HiFi Hot Start Ready Mix (KAPA, USA).

The ITS region of the fungi was amplified using the primers 1743F (5′-CTTGGTCATTTAGAGGAAGTAA-3′) and 2043R (5′-GCTGCGTTCTTCATCGATGC-3′) [57]. The 5′-barcoded amplicons were generated by PCR under conditions of 2 min at 95 °C, followed by 35 cycles at 95 °C for 30 s, annealing at 55 °C for 1 min, extension at 72 °C for 1 min, and a final extension at 72 °C for 10 min. Amplicon quality was visualized using gel electrophoresis, purified with AMPure XP beads (Agencourt) and amplified for another round of PCR. After purification with the AMPure XP beads again, the final amplicon was quantified using a Qubit dsDNA assay kit. Equal amounts of purified amplicon were pooled for subsequent sequencing at the Shanghai OE Biotech. Co., Ltd. (Shanghai,China).

Data analysis

Raw sequencing data were in the FASTQ format. Paired-end reads were preprocessed using Trimmomatic software [58] to detect and remove ambiguous bases (N). We also removed low-quality sequences with average quality score below 20 using the sliding window trimming approach. After trimming, paired-end reads were assembled using FLASH software [59]. The parameters of assembly were 10 bp of minimal overlapping, 200 bp of maximum overlapping and a 20% maximum mismatch rate. The sequences were further denoised as follows: reads with ambiguous, homologous sequences or below 200 bp were abandoned. Reads with 75% of bases above Q20 were retained. Reads with chimeras were detected and removed. These steps were performed using QIIME software (version 1.8.0) [60].

Clean reads were subjected to primer sequence removal and clustering to generate OTUs using UPARSE software with a 97% similarity cutoff [61]. The representative read of each OTU was selected using the QIIME package. All representative reads were annotated and blasted against the Unite database using BLAST [62]. At the phylum level, nonfungal reads and unidentified reads were merged as “Other” for further analysis.

Species accumulation curves [63] were analyzed using QIIME. All the resulting curves tended to approach the saturation plateau, indicating that the number of sequenced reads in each sample was reasonable (see Additional file 4: Figure S1 for species accumulation curves). The probability of new species detected declined significantly at ~ 5000 reads per sample with the exception of the soil. To compare the fungal diversity in tubers across all growth phases, we had to sacrifice some species to ensure that all the samples had the same sequencing depth because one sample of G. elata (P1) had the lowest reads of 4531 (an additional table of the description of sample tags also shows this in detail [see Additional file 1: Table S1]). Fortunately, the numbers of OTUs for G. elata had not significantly declined at the depth of 4530 tags. The sequences were randomly subsampled to avoid differences based on sequencing effort, leaving 4530 reads for further analysis as previously described [64, 65]. Three subsamples of soils as methodological replicates were thus pooled into one sample, and only mean values were compared in further analyses.

Venn diagram was used to measure the overlap between four fungal communities and was conducted using the “VennDiagram” package in R software. Heatmapping of the top 10 OTUs used for fungi distribution in different groups was performed using the R package “pheatmap.” Other histogram and line charts were plotted using Excel 2007 and GraphPad Prism 7 (GraphPad Software, Inc., CA, USA).

Phylogenetic analyses

Because Mycena has been widely proven to germinate G. elata [35, 36, 48], the ITS sequences of 20 Mycena references were downloaded from the NCBI database (sequence information of Mycena spp. is shown in Additional file 8: Table S7). Combined with our three Mycena sequences obtained from high-throughput sequencing and a strain of Mycena sp. (BZZ) previously isolated from G. elata protocorms in the Zhaotong area, a total of 24 partial ITS sequences were used for phylogenetic analyses. Bayesian (BI) analyses were performed using MrBayes v. 3.2.1 [66] based on the models selected by jModeltest v.2.1.4.The analyses lasted until the average standard deviation of split frequencies was below 0.01 with trees saved every 1000 generations. The first 25% of the trees were removed as the burn-in phase, and the remaining trees were used to calculate posterior probabilities. Posterior probability values of the BI analyses (BPP) over 95.0% were considered significant.

Availability of data and materials

All data analyzed during this study are included in this published article and its supplementary information files. Additional raw data from sequencing and quality control datasets are available from the corresponding author upon reasonable request.

Abbreviations

BLAST:

Basic local alignment search tool

ITS:

Internal transcribed spacer

OTU:

Operational taxonomic units

References

  1. 1.

    Merckx V. Mycoheterotrophy: the biology of plants living on fungi. Berlin: Springer; 2013.

  2. 2.

    Arditti J, Ghani AKA. Tansley review no. 110. Numerical and physical properties of orchid seeds and their biological implications. New Phytol. 2000;145(3):367–421.

  3. 3.

    Batty AL, Dixon KW, Brundrett M, Sivasithamparam K. Constraints to symbiotic germination of terrestrial orchid seed in a mediterranean bushland. New Phytol. 2001;152(3):511–20.

  4. 4.

    Leake JR. Myco-heterotroph/epiparasitic plant interactions with ectomycorrhizal and arbuscular mycorrhizal fungi. Curr Opin Plant Biol. 2004;7(4):422–8.

  5. 5.

    Peterson RL, Farquhar ML. Mycorrhizas: integrated development between roots and fungi. Mycologia. 1994;86(3):311–26.

  6. 6.

    Leake JR. The biology of myco-heterotrophic (‘saprophytic’) plants. New Phytol. 2006;127(2):171–216.

  7. 7.

    Oja J, Kohout P, Tedersoo L, Kull T, Kãμljalg U. Temporal patterns of orchid mycorrhizal fungi in meadows and forests as revealed by 454 pyrosequencing. New Phytol. 2015;205(4):1608–18.

  8. 8.

    Barrett CF, Freudenstein JV, Taylor DL, Kõljalg U. Rangewide analysis of fungal associations in the fully mycoheterotrophic Corallorhiza striata complex (Orchidaceae) reveals extreme specificity on ectomycorrhizal Tomentella (Thelephoraceae) across North America. Am J Bot. 2010;97(4):628–43.

  9. 9.

    Taylor DL, Bruns TD. Population, habitat and genetic correlates of mycorrhizal specialization in the ‘cheating’ orchids Corallorhiza maculata and C. mertensiana. Mol Ecol. 1999;8(10):1719–32.

  10. 10.

    Shefferson R, Kull T, Tali K. Mycorrhizal interactions of orchids colonizing Estonian mine tailings hills. Am J Bot. 2008;95(2):156–64.

  11. 11.

    Jacquemyn H, Brys R, Merckx VS, Waud M, Lievens B, Wiegand T. Coexisting orchid species have distinct mycorrhizal communities and display strong spatial segregation. New Phytol. 2014;202(2):616–27.

  12. 12.

    Roy M, Watthana S, Stier A, Richard F, Vessabutr S, Selosse M. Two mycoheterotrophic orchids from Thailand tropical dipterocarpacean forests associate with a broad diversity of ectomycorrhizal fungi. BMC Biol. 2009;7(1):1–17.

  13. 13.

    Jacquemyn H, Honnay O, Cammue BP, Brys R, Lievens B. Low specificity and nested subset structure characterize mycorrhizal associations in five closely related species of the genus Orchis. Mol Ecol. 2010;19(18):4086–95.

  14. 14.

    Hert KD, Jacquemyn H, Provoost S, Honnay O. Absence of recruitment limitation in restored dune slacks suggests that manual seed introduction can be a successful practice for restoring orchid populations. Restor Ecol. 2013;21(2):159–62.

  15. 15.

    Jacquemyn H, Deja A, Hert KD, Bailarote BC, Lievens B. Variation in Mycorrhizal associations with Tulasnelloid Fungi among populations of five Dactylorhiza species. PLoS One. 2012;7(8):e42212.

  16. 16.

    Tĕšitelová T, Tĕšitel J, Jersáková J, RÍhová G, Selosse M. Symbiotic germination capability of four Epipactis species (Orchidaceae) is broader than expected from adult ecology. Am J Bot. 2012;99(6):1020–32.

  17. 17.

    Rafter M, Yokoya K, Schofield EJ, Zettler LW, Sarasan V. Non-specific symbiotic germination of Cynorkispurpurea (Thouars) Kraezl., a habitat-specific terrestrial orchid from the central highlands of Madagascar. Mycorrhiza. 2016;26(6):541–52.

  18. 18.

    Rasmussen H, Dixon K, Jersáková J, Těšitelová T. Germination and seedling establishment in orchids: a complex of requirements. Ann Bot. 2015;116(3):391.

  19. 19.

    Phillips RD, Barrett MD, Dixon KW, Hopper SD. Do mycorrhizal symbioses cause rarity in orchids? J Ecol. 2011;99(3):858–69.

  20. 20.

    McCormick MK, Whigham DF, O'Neill J. Mycorrhizal diversity in photosynthetic terrestrial orchids. New Phytol. 2004;163(2):425–38.

  21. 21.

    Jacquemyn H, Brys R, Cammue BPA, Honnay O, Lievens B. Mycorrhizal associations and reproductive isolation in three closely related Orchis species. Ann Bot. 2011;107(3):347–56.

  22. 22.

    Bidartondo MI, Read DJ. Fungal specificity bottlenecks during orchid germination and development. Mol Ecol. 2008;17(16):3707–16.

  23. 23.

    Chen SC, Tsi ZH, Lang KY, Zhu GH. Flora reipublicae popularis sinicae. Beijing: Science Press; 1999.

  24. 24.

    Liu T, Li CM, Han YL, Chiang TY, Chiang YC, Sung HM. Highly diversified fungi are associated with the achlorophyllous orchid Gastrodia flavilabella. BMC Genomics. 2015;16(1):185.

  25. 25.

    Ahn EK, Jeon HJ, Lim EJ, Jung HJ, Park EH. Anti-inflammatory and anti-angiogenic activities of Gastrodia elata Blume. J Ethnopharmacol. 2007;110(3):476–82.

  26. 26.

    Tang W, Eisenbrand G. Gastrodia elata Bl. Chinese drugs of plant origin. Berlin: Springer; 1992.

  27. 27.

    Chen YY, Bao ZX, Qu Y, Li W, Li ZZ. Genetic diversity and population structure of the medicinal orchid Gastrodia elata revealed by microsatellite analysis. Biochem Syst Ecol. 2014;54:182–9.

  28. 28.

    Xu JT, Guo SX. Retrospect on the research of the cultivation of Gastrodia elata Bl, a rare traditional Chinese medicine. Chin Med J. 2000;113(8):686–92.

  29. 29.

    Xu JT. Studies on the life cycle of Gastrodia elata. Acta Academiae Medicinae Sinicae. 1989;11(4):237–41.

  30. 30.

    Xu JT, Mu C. The relation between growth of Gastrodia elata protocorms and fungi. Acta Bot Sin. 1990;32(1):26–31.

  31. 31.

    Oguratsujita Y, Gebauer G, Hashimoto T, Umata H, Yukawa T. Evidence for novel and specialized mycorrhizal parasitism: the orchid Gastrodia confusa gains carbon from saprotrophic Mycena. Proc Biol Sci. 2009;276(1657):761–7.

  32. 32.

    Martos F, Dulormne M, Pailler T, Bonfante P, Faccio A, Fournel J, et al. Independent recruitment of saprotrophic fungi as mycorrhizal partners by tropical achlorophyllous orchids. New Phytol. 2009;184(3):668–81.

  33. 33.

    Dearnaley JDW, Bougoure JJ. Isotopic and molecular evidence for saprotrophic Marasmiaceae mycobionts in rhizomes of Gastrodia sesamoides. Fungal Ecol. 2010;3(4):288–94.

  34. 34.

    Kinoshita A, Ogura-Tsujita Y, Umata H, Sato H, Hashimoto T, Yukawa T. How do fungal partners affect the evolution and habitat preferences of mycoheterotrophic plants? A case study in Gastrodia. Am J Bot. 2016;103(2):207.

  35. 35.

    Xu J, Guo S. Fungus associated with nutrition of seed germination of Gastrodia elataMycena osmundicola Lange. Acta Mycol Sin. 1989;8(3):221–6.

  36. 36.

    Guo S, Wang Q. Character and action of good strain on stimulating seed germination of Gastrodia elata. Mycosystema. 2001;20(3):408–12.

  37. 37.

    Oguratsujita Y, Gebauer G, Xu H, Fukasawa Y, Umata H, Tetsuka K, et al. The giant mycoheterotrophic orchid Erythrorchis altissima is associated mainly with a divergent set of wood-decaying fungi. Mol Ecol. 2018;27(5):1.

  38. 38.

    Taylor DL, Bruns TD. Independent, specialized invasions of ectomycorrhizal mutualism by two nonphotosynthetic orchids. Proc Natl Acad Sci U S A. 1997;94(9):4510–5.

  39. 39.

    Girlanda M, Selosse MA, Cafasso D, Brilli F, Delfine S, Fabbian R, et al. Inefficient photosynthesis in the Mediterranean orchid Limodorum abortivum is mirrored by specific association to ectomycorrhizal Russulaceae. Mol Ecol. 2006;15(2):491–504.

  40. 40.

    Jacquemyn H, Waud M, Merckx VS, Lievens B, Brys R. Mycorrhizal diversity, seed germination and long-term changes in population size across nine populations of the terrestrial orchid Neottia ovata. Mol Ecol. 2015;24(13):3269–80.

  41. 41.

    Yuan ZL, Chen YC, Yang Y. Diverse non-mycorrhizal fungal endophytes inhabiting an epiphytic, medicinal orchid (Dendrobium nobile): estimation and characterization. World J Microbiol Biotechnol. 2009;25(2):295–303.

  42. 42.

    Gutiarrezmiceli FA, Ayoratalavera T, Abudarchila M, Salvadorfigueroa M. Acclimatization of micropropagated orchid Guarianthe skinnerii inoculated with Trichoderma harzianum. Asian J Plant Sci. 2008;7(3):327–30.

  43. 43.

    Khamchatra NM, Dixon K, Chayamarit K, Apisitwanich S, Tantiwiwat S. Using in situ seed baiting technique to isolate and identify endophytic and mycorrhizal fungi from seeds of a threatened epiphytic orchid, Dendrobium friedericksianum Rchb.F. (Orchidaceae). Agric Nat Resour. 2016;50(1):8–13.

  44. 44.

    Ma X, Kang J, Nontachaiyapoom S, Wen T, Hyde KD. Non-mycorrhizal endophytic fungi from orchids. Curr Sci. 2015;109(1):72–87.

  45. 45.

    Park EJ, Lee WY. In vitro symbiotic germination of myco-heterotrophic Gastrodia elata by Mycena species. Plant Biotechnology Reports. 2013;7(2):185–91.

  46. 46.

    Liu H, Luo Y, Liu H. Studies of mycorrhizal fungi of Chinese Orchids and their role in Orchid conservation in China—A Review. Bot Rev. 2010;76(2):241–62.

  47. 47.

    Osmundson TW, Robert VA, Schoch CL, Baker LJ, Smith A, Robich G, et al. Filling gaps in biodiversity knowledge for macrofungi: contributions and assessment of an herbarium collection DNA barcode sequencing project. PLoS One. 2013;8(4):e62419.

  48. 48.

    Park EJ, Lee WY, Jin KA. In vitro propagation of myco-heterotrophic Gastrodia elata. Hortic Environ Biotechnol. 2012;53(5):415–20.

  49. 49.

    Cha JY, Igarashi T. Armillaria species associated with Gastrodia elata in Japan. For Pathol. 1995;25(6–7):319–26.

  50. 50.

    Sekizaki H, Kuninaga S, Yamamoto M, Asazu SN, Sawa S, Kojoma M, et al. Identification of Armillaria nabsnona in gastrodia tubers. Biol Pharm Bull. 2008;31(7):1410–4.

  51. 51.

    Jacquemyn H, Wiegand T. Spatial variation in below-ground seed germination and divergent mycorrhizal associations correlate with spatial segregation of three co-occurring orchid species. J Ecol. 2012;100(6):1328–37.

  52. 52.

    Malinova JT. Spatial aspects of seed dispersal and seedling recruitment in orchids. New Phytol. 2010;176(2):237–41.

  53. 53.

    Diez J. Hierarchical patterns of symbiotic orchid germination linked to adult proximity and environmental gradients. J Ecol. 2007;95(1):159–70.

  54. 54.

    McCormick MK, Taylor DL, Whigham DF, Burnett RK. Germination patterns in three terrestrial orchids relate to abundance of mycorrhizal fungi. J Ecol. 2016;104(3):744–54.

  55. 55.

    Sun ZG, Chen Z, Zhang M, Cheng DL, Zhong XB. Current situation of protection of Chaotong Gastrodia elata, a national geographical symbol product. Guizhou Agricultural Sciences; 2010.

  56. 56.

    Murray MG, Thompson WF. Rapid isolation of high molecular weight plant DNA. Nucleic Acids Res. 1980;8(19):4321–5.

  57. 57.

    Sun X, Lyu G, Luan Y, Zhao Z, Yang H, Su D. Analyses of microbial community of naturally homemade soybean pastes in Liaoning Province of China by Illumina Miseq sequencing. Food Res Int. 2018;111:50–7.

  58. 58.

    Bolger AM, Lohse M, Usadel B. Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics. 2014;30(15):2114–20.

  59. 59.

    Reyon D, Tsai SQ, Khayter C, Foden JA, Sander JD, Joung JK. FLASH assembly of TALENs for high-throughput genome editing. Nat Biotechnol. 2012;30(5):460.

  60. 60.

    Caporaso J, Kuczynski J, Stombaugh J, Bittinger K, Bushman F. QIIME allows integration and analysis of high-throughput community sequencing data. Nat Methods. 2010;7(5):335–6.

  61. 61.

    Edgar RC. UPARSE: highly accurate OTU sequences from microbial amplicon reads. Nat Methods. 2013;10(10):996–8.

  62. 62.

    Lobo I. Basic local alignment search tool (BLAST). J Mol Biol. 2008;215(3):403–10.

  63. 63.

    Heck KL, Belle GV, Simberloff D. Explicit calculation of the rarefaction diversity measurement and the determination of sufficient sample size. Ecology. 1975;56(6):1459–61.

  64. 64.

    Barfod KK, Roggenbuck M, Hansen LH, Schjørring S, Larsen ST, Sørensen SJ, et al. The murine lung microbiome in relation to the intestinal and vaginal bacterial communities. BMC Microbiol. 2013;13(1):303.

  65. 65.

    Logares R, Audic S, Bass D, Bittner L, Boutte C, Christen R, et al. Patterns of rare and abundant marine microbial eukaryotes. Curr Biol. 2014;24(8):813–21.

  66. 66.

    Ronquist F, Teslenko M, Mark PVD, Ayres DL, Darling A, Höhna S, et al. MrBayes 3.2: efficient Bayesian phylogenetic inference and model choice across a large model space. Syst Biol. 2012;61(3):539–42.

Download references

Acknowledgements

The authors thank Tian Zeng and Wen-Ti Zheng at Yunnan University for sample collection in the wild.

Funding

This work was funded by the National Natural Science Foundation of China (grant No. 31770585).

Author information

LC: experimental design, data analysis and manuscript writing; YCW:project design, performed experiments in the wild; LYQ: sequencing and data analysis; HYH:performed experiments in the wild; XLY: project design; MZY: experimental design and manuscript review; and HBZ:project design, data analysis and manuscript writing. All authors have read and approved this manuscript.

Correspondence to Ming-Zhi Yang or Han-Bo Zhang.

Ethics declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

Additional information

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Additional files

Additional file 1:

Table S1. Description of sample Tags. (DOCX 12 kb)

Additional file 2:

Table S2. OTUs table. (XLSX 74 kb)

Additional file 3:

Table S3. OTUs table after subsampled. (XLSX 61 kb)

Additional file 4:

Figure S1. Species accumulation curves. (TIF 2414 kb)

Additional file 5:

Table S4. Venn diagram for OTUs distribution. (XLSX 18 kb)

Additional file 6:

Table S5. Distribution of Mycena sp. in each sample. (DOCX 13 kb)

Additional file 7:

Table S6. The tubers at different growth phases in five holes. (DOCX 13 kb)

Additional file 8:

Table S7. Sequence information of Mycena spp. (DOCX 15 kb)

Rights and permissions

Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Keywords

  • Gastrodia elata
  • Growth phases
  • Mycena
  • High-throughput sequencing
  • Fungal dynamics