Skip to main content

New fungal primers reveal the diversity of Mucoromycotinian arbuscular mycorrhizal fungi and their response to nitrogen application

Abstract

Background

Arbuscular mycorrhizas (AM) are the most widespread terrestrial symbiosis and are both a key determinant of plant health and a major contributor to ecosystem processes through their role in biogeochemical cycling. Until recently, it was assumed that the fungi which form AM comprise the subphylum Glomeromycotina (G-AMF), and our understanding of the diversity and ecosystem roles of AM is based almost exclusively on this group. However recent evidence shows that fungi which form the distinctive 'fine root endophyte’ (FRE) AM morphotype are members of the subphylum Mucoromycotina (M-AMF), so that AM symbioses are actually formed by two distinct groups of fungi.

Results

We investigated the influence of nitrogen (N) addition and wheat variety on the assembly of AM communities under field conditions. Visual assessment of roots showed co-occurrence of G-AMF and M-AMF, providing an opportunity to compare the responses of these two groups. Existing ‘AM’ 18S rRNA primers which co-amplify G-AMF and M-AMF were modified to reduce bias against Mucoromycotina, and compared against a new ‘FRE’ primer set which selectively amplifies Mucoromycotina. Using the AM-primers, no significant effect of either N-addition or wheat variety on G-AMF or M-AMF diversity or community composition was detected. In contrast, using the FRE-primers, N-addition was shown to reduce M-AMF diversity and altered community composition. The ASV which responded to N-addition were closely related, demonstrating a clear phylogenetic signal which was identified only by the new FRE-primers. The most abundant Mucoromycotina sequences we detected belonged to the same Endogonales clades as dominant sequences associated with FRE morphology in Australia, indicating that closely related M-AMF may be globally distributed.

Conclusions

The results demonstrate the need to consider both G-AMF and M-AMF when investigating AM communities, and highlight the importance of primer choice when investigating AMF community dynamics.

Background

Arbuscular mycorrhizal fungi (AMF) are considered to form the most common and important symbiosis in terrestrial ecosystems [65]. Fossilised spores and hyphae in plant roots from the Ordovician date the first association of fungi with land plants to 460 million years ago and provide evidence for the significance of mycorrhizas in the evolution of vascular plants [46, 57]. Arbuscular mycorrhizas (AM) are traditionally considered to be formed by fungi within the monophyletic Glomeromycotina subphylum [59]. However, more recently a further group of arbuscle-forming fungi has been recognised within the Mucoromycotina subphylum [41, 69]. The species complex of the distinctive ‘fine root endophyte’ (FRE) arbuscular fungi was formerly collected within the species Glomus tenue within Glomeromycotina [17, 62], but molecular analyses have shown that these fungi are phylogenetically different to Glomeromycotina AMF (G-AMF; [41]). Consequently, Glomus tenue (basionym Rhizophagus tenuis) was renamed Planticonsortium tenue [69] and the new genus resides within the Endogonales order within the subphylum Mucoromycotina [2, 3, 40]. To date this is the only described species of M-AMF. Co-occurrence of both G-AMF and Mucoromycotina AMF (M-AMF) has frequently been reported across plant species and environments [41, 52]. The detection of both M-AMF and G-AMF in fossils of early land plants from the Rhynie Chert (ca. 407 million years ago) indicates early co-occurrence of these two groups in plant-fungal associations [60].

Several factors have resulted in the occurrence of M-AMF in nature, and their contributions to ecosystem processes, being overlooked. DNA primers designed to profile AMF communities have been targeted at G-AMF and fail to amplify M-AMF [41]. Furthermore, spores of M-AMF are much smaller (≤ 20 µm) than those of G-AMF (30 µm–1 mm), and as a result M-AMF are not recorded during fungal community assessments using spore morphology [41]. The characteristic fine hyphae (0.1–4 µm) and fan-shaped arbuscules enable differentiation of M-AMF from G-AMF by microscopy [33, 40, 41]. But this requires high power microscopy, such that M-AMF are not detected in studies which assess root colonisation under low magnification. M-AMF have thinner cell walls than G-AMF and form hyphal ropes, but the two groups of fungi have similar colonisation behaviour and ultrastructural features [2, 14].

Analysis of the distribution of FRE indicates that like G-AMF, M-AMF are globally distributed and found in both natural and agricultural soils, although there are indications that M-AMF are absent from tropical and subtropical biomes [4, 40]. Regardless, their functional significance is far less understood than that of G-AMF [40]. Recently, nutritional mutualism involving exchange of fungal phosphorus (P) and nitrogen (N) in return for plant carbon (C) has been shown for M-AMF in axenic systems [20]. Furthermore, comparative studies with G-AMF in association with liverworts showed higher N-uptake by M-AMF, while the reverse was true for P, suggesting that these groups of fungal endophytes could have complementary roles in nutrient acquisition [13].

Agricultural practices such as N-addition have negative effects on biodiversity in plant [9] and soil microbial [26, 74] communities and cause wider environmental impacts such as eutrophication of surface and groundwater bodies [10]. In particular, N-addition can decrease abundance of G-AMF in soil and roots of agricultural plants [15] and result in proliferation of ‘weedy’ G-AMF in natural systems [5]. In agricultural soils, optimised N-management is important to promote G-AMF diversity [30], and can be decisive for the success of G-AMF inoculation [12].

In contrast to the wealth of knowledge on the impacts of agricultural management practices and environmental variables on the diversity and community composition of G-AMF [15, 48], understanding of the responses of M-AMF are largely limited to morphological root colonisation data with a dominance of data from Australia and New Zealand [41]. Recently, Albornoz et al. [3] showed that M-AMF and G-AMF communities had similar responses to key environmental variables, including soil P concentration and pH within regionally co-located pasture sites. However, a continent-wide analysis across multiple ecosystems provided clear evidence for different responses of the two groups of fungi to edaphic and climatic variables, indicating that these groups of fungi have distinct but overlapping ecological niches [4]. In wheat and field pea in southern Australia, Ryan and Kirkegaard [51] showed that P-addition decreased the colonisation of M-AMF in roots more so than that of G-AMF, while, in the glasshouse Jeffery et al. [21] showed that the impact of 5 rates of P addition on colonisation of pasture legumes differed between M-AMF and G-AMF. However, the role of soil P, let alone soil N, in shaping M-AMF communities has received little attention. Sigüenza et al. [56] indicated that M-AMF root colonisation may be favoured by high N environments, and furthermore at the continental scale, relative abundance of M-AMF was favoured by availability of soil mineral-N [4]. Given the potentially contrasting contributions of G-AMF and M-AMF to host N nutrition [13], there is a need for comparative understanding of the role of N in shaping the diversity and composition of these contrasting AM communities.

The importance of host variety for determining G-AMF colonisation and growth responses has been demonstrated in many crops such as clover [53], onion [61], potato [1], sorghum [29], maize [55] and durum wheat [12]. However, most evidence for variety selection of G-AMF comes from studies of common wheat (Triticum aestivum. Host variety-dependent variations in mycorrhizal colonisation of wheat have been widely reported (Azcón and Ocampo 1981) [19, 27] and have been linked to nutrient use efficiency [24, 63] and abiotic stress tolerance [28]. There has been a long-term debate about the breeding background which determines mycorrhizal responsiveness of wheat [19, 27, 35, 75], and the results are inconclusive. However, most plant variety-differences in AMF selection are based on root colonisation assessments, and less is known about impacts on mycorrhizal community composition [61]. Mao et al. [34] provided evidence that G-AMF community assembly dynamics in wheat can operate at a variety level, with 21 varieties showing significantly different G-AMF mycobiomes under field conditions, and that varietal composition was correlated with drought stress tolerance. However, these variations had no effects on wheat performance [34]

There are several studies which report morphological evidence for M-AMF symbioses in agricultural crops, including wheat [18, 51, 52, 58], but to our knowledge, relationships between M-AMF and plant variety has not been investigated. Most recent studies on M-AMF in agricultural systems come from southern Australia where wheat was shown to have an AM fungal community dominated by M-AMF (present in 80% of AM colonised root length) abundance of which was impacted by previous crop and P-addition [51]. However, at the community level, understanding of M-AMF in agricultural systems is limited, and is contributed primarily from a survey of subterranean clover in pasture systems across southern Australia [3], and additional data from single fields of a broad range of crops (from wheat to fruit orchards) across a range of biomes in Albornoz et al. [4]. In all cases root associated communities contained multiple putative M-AMF taxa, but the taxa richness was lower than G-AMF communities [3, 4, 40]. Interestingly, the survey across Australian biomes by Albornoz et al. [4] found putative M-AMF had greater distribution and abundance in the soils of agricultural fields than adjacent natural ecosystems, indicating that, in contrast to G-AMF, agricultural practices may favour M-AMF.

In this study we investigated the influence of N-addition on the assembly of AM communities in two contrasting wheat cultivars, under field conditions. Preliminary visual assessment of roots at the field site showed co-occurrence of G-AMF and M-AMF, which provided an opportunity to compare the responses of these two groups of fungi to both crop variety and N-addition. To date, work investigating M-AMF community composition has used 18S rRNA primers designed by Sato et al. [54] which co-amplify M-AMF and G-AMF, but provide poor coverage of putative M-AMF and short (220 bp) sequence lengths [3, 4, 33, 41]. In the current study we modified the Sato forward primer to reduce bias against M-AMF, and designed new 18S rRNA Mucoromycotina specific primers which amplified longer sequences (440 bp), thus providing improved resolution of community composition and phylogeny.

Methods

Field site and agronomic management

The field trial was conducted in 2018–2019 at Nafferton Experimental Farm in Northumberland (54° 59′ 27.26″ N, 1° 54′ 26.96″ W, Stocksfield, UK). The soil type at the experimental field site is a uniform sandy clay loam [47] with a pH of 6.8 and Olsen extractable contents of 8.1 ± 1.49 mg L−1 P, 72 ± 8.86 mg L−1 potassium and 157 ± 6.18 mg L−1 magnesium. The interaction of N application and wheat variety was assessed in a randomised-block split-plot factorial design. There were three 24 × 24 m blocks across a 0.3 ha area. Each block contained a 6 m × 4 m plot of each the 4 treatments (i.e. (1). Aszita with no added N (n = 3) (2). Aszita with added N (n = 3) (3). Skyfall with no added N (n = 3) and (4). Skyfall with added N (n = 3). The factorial design provided 6 plots for assessing the effect of both N treatment and variety on fungal communities, with 3 replicates for each variety/ fertiliser combination. The winter wheat varieties Skyfall (released 2014 by RAGT Seeds Ltd., UK) and Aszita (released in 2004 by Getreidezüchtung Peter Kunz, Switzerland) were drilled in September 2018.

Skyfall has a conventional breeding background, and is a modern, high yielding, semi-dwarf variety, and one of the most widely grown winter wheats in the UK. Aszita has an organic breeding background and is characterised by long straw growth of more than 1 m and achieves low grain yields, but with high quality. Both varieties have been investigated in other studies on variety-dependent AM interaction: Skyfall acquired more P through AMF in comparison to two other modern wheat varieties and showed an intermediate root length colonisation of 34% in a study by Elliott et al. [11]. Another study using the same varieties identified Skyfall as particularly dependent on AM-mediated nutrient uptake [63]. Aszita on the other hand was revealed as one of the least colonised by AMF among 94 wheat varieties in a greenhouse screening study by Lehnert et al. [28].

Half of each wheat variety population was treated with fertiliser in the form of mineral-N (ammonium nitrate; Nitram 34.5% N, CF fertilisers UK Ltd.), which was added at a rate of 170 kg N ha−1 in two applications: 70 kg ha−1 in mid-April 2019 and 100 kg 2 weeks later. The control plots did not receive any fertiliser input and are further referred to as zero-N plots. Fungicides, herbicides and plant growth regulators were applied throughout the cropping season (as detailed in Table S1).

Root colonisation assessment

Root samples were collected from the topsoil layer of the twelve plots just after the start of stem elongation (growth stage 32, [73]) in May 2019. Shoots across a 0.25 m2 sampling area were cut at the stem base, prior to digging plants out. Roots from 15 to 20 cm soil depth were removed from 5–6 plants and pooled to provide a composite sample from each plot. Root samples were washed in tap water and stored in 50% ethanol until further processing. Roots were stained using the ink-vinegar method described by Vierheilig et al. [68]. First, roots were rinsed in tap water to remove ethanol residues. Then, samples were incubated in 10% KOH solution at 80 °C for two hours in the oven. Roots were rinsed with tap water and subsequently incubated in an 8% acetate/5% China ink solution overnight. Microscopy for the visual assessment of root colonisation was conducted on a total of 30 cm root length from each sample. Percentages of overall mycorrhizal frequency intensity as well as intensity of arbuscules, vesicles and hyphae were estimated based on the method described by Trouvelot et al. [66] using the INOQ Calculator Advanced [37].

DNA extraction and sequencing

DNA was extracted from 0.5 g of the root samples using the DNeasy PowerSoilPro Kit (Qiagen, Germany) following the manufacturer’s protocol. Prior to DNA extraction, samples were homogenised using a FastPrep-24™ (MP Biomedicals, USA) at 6 m s−1 for 2 × 40 s periods and an incubation step for 5 min at 4 °C between homogenisations. DNA concentrations were measured by fluorometric quantification (Qubit™ Fluorometer 3.0) using the Qubit™ dsDNA high sensitivity assay kit (Invitrogen, USA) following the manufacturer’s protocol. Amplicons were produced using two different primer sets targeting the 18S rRNA gene.

The first primer set (further referred to as AM-primers) targeted both Glomeromycotina and Mucoromycotina and was adapted from the primers published by Sato et al. [54] which amplify a 220 bp region of the 18S rRNA gene (Fig. S1). Alignment of Mucoromycotina and Glomeromycotina 18S rRNA sequences showed that the Sato et al. AMV4.5NF forward primer (AAGCTCGTAGTTGAATTTCG) had 2 nucleotide differences to Mucoromycotina at the 3′ end which drastically reduced amplification of Mucoromycotina relative to Glomeromycotina (Fig. S2). The redesigned AM forward primer AM-Sal-F (AAGCTCGTAGTTGAATTT) was based on AMV4.5NF with the two end 3′ nucleotides removed. The TestPrime 1.0 program (https://www.arb-silva.de/search/testprime/) within the Silva database ([44], (> 3 K Mucoromycota reference sequences) was used to evaluate the coverage of Mucoromycota sub-phyla using the suggested settings of 1 mismatch with 5 bases 0-mismatch zone for a realistic simulation of PCR behaviour. This showed that the new primer AM-Sal-F improved coverage of both Glomeromycotina and Endogonales, whilst also allowing the inclusion of the Umbelopsidales group of the Mucoromycotina, and the Mortierellomycotina sub-phylum (Table S2).

Comparative analysis of the sequencing performance of AMV4.5NF and AM-Sal-F primers in combination with the Sato et al. [54] AMDGR reverse primer was performed (Fig. S3). Roots were collected from 3 independent locations separated by 10 m, within the 2 ha Boddington Meadow Nature Reserve, Northamptonshire (52° 10′ 24″ N, 1° 16′ 44″ W), UK in August 2019. Composite samples of root biomass (grasses and herbs) were collected from 0 to 10 cm depth at each location. The site has never been ploughed, is rich in wildflowers, is grazed periodically with sheep and cut for hay at the end of the growing season. DNA extraction, PCR amplification, sequence analysis and bioinformatic processing was performed as described above for the samples. Testing revealed that the AM-Sal-F primer enriched Endogonales sequences tenfold compared to AMV4.5NF, associated with a decrease in non-target sequences (e.g. Agaricomycetes), although this was associated with reduced % of Glomerales and an increase in the percentage of Mortierellales reads (Fig. S3).

A new primer set was devised to selectively amplify ~ 440 base pairs (bp) of the V4-6 region of the 18S rRNA gene of the Endogonales and related Mucoromycotina (further referred to as FRE-primers, Fig. S1). All Endogonales sequences (49) from the Silva database (July 2019) were aligned in MegAlign (DNASTAR Inc). Representative Glomeromycotina and other Mucoromycotina sequences were included. A forward primer was designed to a region downstream, but overlapping with the AM primer AMV4.5NF, which was distinct between the Glomeromycotina and the Mucoromycotina (6 nt mismatches within the 3′ region). The reverse primer had only 4 nt mismatches to the Glomeromycotina but in combination with the forward primer resulted in more specific amplification of the Endogonales/Mucoromycotina group. The FRE-F (GTTGAATTTTAGCCYTGGC) and FRE-R (CCCAAAAACTTTGATTTCTCW) primers amplify at 18S rRNA positions 619 and 1114 generating a 440 bp fragment.

Primer pairs were applied in separate PCRs using 15 ng of DNA in a master mix containing 12.5 µl 2 × Q5® High Fidelity Hot Start Master Mix (New England BioLabs® Inc., USA), 5 µl ddH2O and 1.25 µl of AM- (10 µM each) or FRE-primers (0.5 µM each) respectively. The PCR programme for both primer pairs started with denaturation at 98 °C for 30 s, followed by 35 cycles starting with polymerase activation at 98 °C for 10 s. Annealing of AM-primers was set at 60 °C for 15 s and for FRE-primers at 55 °C for 15 s. Both reactions were followed by elongation at 72 °C for 20 s. Final extension was conducted at 72 °C for 5 min. Size of amplicons was checked by gel electrophoresis. Following PCR, the DNA amplicons were purified using Agencourt AMPure XP beads (Beckman Coulter, Brea, CA, USA) according to the manufacturer’s instructions. The adapted amplicons were then modified by attaching indices and Illumina sequencing adapters using the Nextera XT Index Kit v.2 (Illumina, San Diego, CA, USA) by PCR as described in the manufacturer’s protocol. The amplicons were then purified and normalized using the SequalPrepTM Normalization Plate (96) Kit (Invitrogen) and quantitatively assessed using a Qubit 2.0 Fluorometer (Life Technologies). The final concentrations of the libraries was 4 nM. The libraries were sequenced using the MiSeq Reagent Kit v.3 600-cycle (Illumina) at The University of Warwick, UK.

Demultiplexed sequences with primer sequences removed were processed using the DADA2 pipeline [7] in Quantitative Insights into Microbial Ecology (QIIME 1.8., [8]). This pipeline removed low-quality reads (QC < 30), chimeras (using the consensus method) and singletons from the library. Taxonomy was assigned to amplicon sequencing variants (ASV) based on the SILVA132 2019 database [44]. ASV have been shown to outperform OTU based clustering methods when estimating the correct number of fungal species present in samples [22], and provide better reproducibility and finer granularity of compositional differences between samples, including improved detection of rare fungi [64]. Furthermore, ASV provide a better resource for comparisons across studies, which is particularly important for biogeographic analyses.

Sequencing of mycorrhizal communities in wheat roots with the AM primers (AM-Sal-F and AMDGR primers) which amplify both Mucoromycotina and Glomeromycotina yielded 300,371 sequences, and Mucoromycotina and Glomeromycotina accounted for 44% and 47.5% of all reads respectively. Only the sequences assigned to Mucoromycotina and Glomeromycotina were used for subsequent analyses. After removing reads from outside these groups, there were 241,089 sequences representing Mucoromycotina and Glomeromycotina, with between 9,381 and 33,529 reads per sample, assigned to 253 ASV (145 identified as Mucoromycotina and 108 as Glomeromycotina). BLAST using the nucleotide database Altschul et al. [6] was used to confirm correct taxonomic assignment of sequences. At the Order level, there were very few Diversisporales or Archaeosporales sequences, and no Paraglomerales sequences. All Mucoromycotina sequences were classified as Endogolales. There were approximately equal proportions of Endogonales and Glomerales sequences (Fig. S4).

The specific amplification of Mucoromycotina using FRE-primers produced 88,202 reads with 4230 to 11,895 reads per sample. After removing contaminants, the remaining 86,922 reads were grouped into 121 ASV which were solely assigned to the family Endogonaceae within the order Endogonales. Read numbers after filtering ranged from 3756 to 11,895 reads per sample. Only 5% of these sequences could be assigned to the genus Endogone, the remaining 95% were assigned to Endogonaceae (i.e. further taxonomic resolution was not possible).

Statistical analysis

RStudio [36, 45] was used to calculate alpha diversity (species evenness, observed richness and Shannon’s diversity index) which were then compared among experimental treatments using Kruskal–Wallis tests. Reads for these analyses were rarefied to the lowest sample count (3756 and 9381 for the FRE and AM primers respectively). For analysis of beta diversity unrarified read counts were normalised using DESeq2 [31], and used to calculate a Bray Curtis dissimilarity matrix which was visualised using non-metric multidimensional scaling (NMDS) to identify groups of samples based on similar ASV compositions.

The effect of variety and N-addition on the ratios of Glomeromycotina to Mucoromycotina sequences was compared by analysis of variance (ANOVA) in linear mixed-effect models using the nlme-package [43]. For both datasets, the dispersion of homogeneity within treatment groups was assessed using the betadisper-function from the vegan-package [39]. Groups showing a sufficient homogeneity of variance (p ≥ 0.05) were run in permutational multivariate analysis of variance (PERMANOVA) using the adonis-function of the vegan-package. Blocks were included as random effects. Treatment groups showing significant differences at P < 0.05 were run through similarity percentage (SIMPER) analyses to identify ASV that contributed to these differences. The results of this method were validated by Kruskal–Wallis testing. All plots were generated with ggplot2 [70] in R, which was also used to correlate root colonisation data with ASV relative abundances of Glomeromycotina and Mucoromycotina in the samples.

Phylogenetic analysis

The Mucoromycotina sequences amplified with the AM and FRE primers were aligned with a broad range of reference sequences, including environmental sequences which have been associated with FRE morphology in Australian samples. This included operational taxonomic units (OTUs) associated with FRE in Trifolium subterraneum pastures collected from across Australia [3] i.e. TS OTUs 7, 18, 43, 49, 110, 152, 289, 350, 432 and 980) and OTUs associated with FRE in T. subterraneum grown in pasture soil from Western Australia (i.e. KX434777, KX434773, KX434782, KX434776, KX434780 and KX434781) [40]. In particular, KX434777 is identical to the only described culture of an FRE forming fungus, Planticonsortium tenue [4]. Mucoromycotina sequences which were abundant in agricultural soils across Australia were also included [4], i.e. FREOZ OTUs 2, 43, 170. Additionally, we included sequences of Mucoromycotina associated with lower land plants (hornworts, and liverworts) (KC708405, KC708440, KC708398, KJ952232, KJ952217, KC708443, KC708419, KR779282, JF414224, JF414221, JF414209, LC429236). We included sequences from cultures of the saprotrophic fungi Vinositunica radiata (LC431088), Sphaerocreas pubescens (AB752291) and Endogone pisiformis (DQ322628) and a sporocarp of the saprophyte Endogone lactiflua (JF414204). A sequence of the putative ectomycorrhizal fungus Endogone oregonensis (JF414208) was also included. Glomeromycotan sequences including Acaulospora laevis (Y17633), Archaeospora (MH629023 and HF954887), Diversispora aurantia (AM713432), and a selection of ASV generated from our samples using AM primers (ASVs 451, 464, 415,270, 335, 323 and 254) were used as an outgroup.

Phylogenetic trees were built with MrBayes v.3.2.6 using a Bayesian phylogenetic inference approach [49] in which sequences of different lengths can be aligned by inserting gaps in the alignment to account for insertions or deletions in some sequences. This approach allowed us to align full 18S rRNA reference sequences together with the different sized AM and FRE sequences, and to identify sequences which were identical despite different sizes. Two separate MC3 runs with randomly generated starting trees were carried out for four million generations each with one cold and three heated chains. The evolutionary model applied a GTR substitution matrix, with a four-category autocorrelated gamma correction. All parameters were estimated from the data. The trees were sampled every 1000 generations and the first million generations discarded as burn-in. All phylogenetic analyses were carried out on the Cipres server [38]. The resulting phylogenetic tree was visualised in R using ggtree v3.6.2 [72], and posterior probability values > 0.8 are shown. The phylogenetic diversity accessed by the AM-primers and FRE-primers, independently, was estimated using Faith’s phylogenetic diversity metric.

Results

Mycorrhizal root colonisation

Root colonisation by AMF was high, with the frequency of colonisation (F%) around 60% and AM intensities (M%) between 33 and 37% (Table S3). There was no significant effect of N-addition or wheat variety on AMF colonisation frequency, intensity, or percentage of arbuscules, vesicles or hyphae. Wheat roots supported both coarse AMF and FRE morphologies indicating colonisation by both G-AMF and M-AMF (Fig. 1A–D). Visible FRE morphology included finely branched arbuscules and intercalary swellings on thin hyphae (Fig. 1A, B). Coarse and FRE colonisation was intermixed and it was not possible to produce an accurate assessment of the percentage root length colonised by each morphotype.

Fig. 1
figure 1

Microscopy images of fine root endophyte morphology linked to Mucormycotina arbuscular mycorrhizas (M-AMF) and coarse morphology associated with Glomeromycotina arbuscular mycorrhizas (G-AMF) in wheat roots. A Left arrow marks hyphae with intercalary swellings of G-AMF, right arrow marks arbuscules of G-AMF. B Arbuscules (a) and hyphae (h) of M-AMF, arrows point towards intercalary swellings. C Arbuscules and hyphae of G-AMF. D Vesicles (v), arbuscules and hyphae of G-AMF. Scale bars indicate 50 µm at ×10 (A) and ×40 (B–D) magnification

Composition of Glomeromycotina and Mucoromycotina communities based on AM-primers

ANOVA showed that the relative abundances of Mucoromycotina and Glomeromycotina sequences (Table S4), and the ratio of Mucoromycotina to Glomeromycotina sequences (Table S5) were not significantly affected by variety or N-addition and there was no significant interaction between variety and N-addition. Alpha diversity of Mucoromycotina and Glomeromycotina was not significantly different between the varieties or N-treatments when analysed together (Fig. S5) or separately (Figs. S6, S7). Similarly PERMANOVA (Table S6) analysis showed that beta-diversity of the Glomeromycotina and Mucoromycotina communities when analysed together, or separately, was not significantly affected by either variety or N-addition. There was no significant correlation between AMF root colonisation and the relative abundance of Glomeromycotina (R = − 0.15, p = 0.63) or Mucoromycotina ASV (R = − 0.21, p = 0.5).

Composition of Mucoromycotina communities using FRE-primers

Shannon diversity of Mucoromycotina was significantly higher when mineral-N was applied, relative to the zero-N treatment (Fig. 2), and this was driven by trends in greater number of observed ASV, and greater eveness in the mineral-N relative to the zero-N treatment. However there was no significant difference in Shannon diversity between the two wheat hosts. PERMANOVA (Table 1) and NMDS (Fig. 3) analysis showed that beta-diversity of the Mucoromycotina community profile was significantly affected by N application, which contributed to 22% of community variation (P < 0.028). Wheat variety had no significant effect on beta diversity.

Fig. 2
figure 2

Box plot showing ASV evenness, number of observed ASV and Shannon diversity index of Mucoromycotina communities. Sequencing was performed using Mucoromycotina specific FRE primers. A Comparison of mineral N application with zero-input treatments (n = 6). B Comparison of wheat varieties (n = 6). Numbers indicate p-values for pairwise comparison by Kruskal–Wallis test (ns = not significant, * significant p ≤ 0.05)

Table 1 Permutational multivariate analysis of variance of the effect of wheat variety (Aszita vs. Skyfall) and nitrogen treatment (mineral-N vs. zero-N treatments) on the composition of Mucoromycotina
Fig. 3
figure 3

Non-metric multidimensional scaling plot of Bray Curtis dissimilarity of Mucoromycotina communities in roots of two wheat varieties with and without mineral-N application. Sequencing was performed using Mucoromycotina specific FRE primers

SIMPER analysis identified 12 ASV which contributed > 1% to the dissimilarity between Mucoromycotina communities in the zero-N and mineral N-addition treatments (Table 2), and for five of these ASV the relative abundance was also significantly different between the N treatments. ASV3 contributed 22.14% of the dissimilarity between N treatments. The relative abundance of this ASV was significantly (P < 0.05) increased by 20% in roots with mineral-N addition. ASV 7 contributed 4.63% of the variation in community composition between N treatments, with relative abundance increasing significantly (P < 0.05) from 1.51 in the zero-N treatment to 7.77% in the mineral-N treatment. ASV17, 18 and 21 contributed to less than 2% of the variation between N treatments, with significant (P < 0.05) increases in relative abundance from 0.38 to 0.53% in the zero-N treatment to between 1.75 and 2.56% in the mineral-N treatment. A number of further ASV contributed to the dissimilarity between N treatments (ASV1, 2, 8, 5, 10, 13, 15) but none of these showed a significant difference in relative abundance between treatments.

Table 2 Similarity percentage (SIMPER) analysis of ASV contributing to dissimilarity of Mucoromycotina communities in wheat roots from zero-N and mineral-N treatments (n = 6).

Phylogenetic analysis of Mucoromycotina sequences

A phylogenetic tree containing 145 ASV identified using AM primers, 121 ASV identified using FRE-primers, and reference Mucoromycotinian and Glomeromycotinian OTU sequences was visualised (Fig. 4). While this indicated that the two primer sets accessed a similar breadth of Mucoromycotinian diversity, Faith’s phylogenetic diversity was estimated to be greater for the ASV accessed using AM-primers (35.68) than those using FRE-primers (30.27). Nonetheless, the tree contained four main clades (referred to as Clades 1–4), with sub-clades largely represented by the ASV which were unique to each of the primer sets.

Fig. 4
figure 4

Phylogenetic assessment of ASV accessed using AM and FRE primers. Outer ring fill colour denotes sequence source. Clade labels denote the four main Mucoromycotina clades formed. The five FRE primer amplified ASV found to have significant differences in their relative abundance between mineral-N treatments are highlighted in pink. Branches with a grey circle have a posterior probability value > 0.8. Reference sequences are described in the Methods, section Phylogenetic analysis. Environmental sequences associated with fine root endophyte colonisation of roots include TS OTUs 7, 18, 49, 110, 152, 289, 350 and 432, from Trifolium subterraneum pastures across Australia [3] and sequences KX434777, KX434773, KX434782, KX434776, KX434780 and KX434781 from T. subterraneum grown in pasture soil from Western Australia [40]. Environmental sequence OTU4 from Orchard et al. [40], which shows 100% similarity to Planticonsortium tenue [4], is shown in bold (Clade 1)

Clade 1 (Fig. 4, Fig. S8) included four of the five dominant sequences amplified with FRE primers (FREASV 1 (9.8% reads), 3 (22.3%), 5 (11.3%) and 8 (5.3%)) and three dominant ASV amplified by the AM-primers (ASV 17 (24.2%), 37 (9.1%) and 21 (6.0%)). This clade also contained sequence from the only described FRE culture, Planticonsortium tenue (KX434777), and other abundant sequences associated with FRE morphology from previous studies: KX434773 from glasshouse grown T. subterraneum [41], TS OTU43, OTU110 and TS OTU432 from pasture sites in Australia [3], and OTU2, from agricultural soils in Australia [4]. Further inspection of Clade 1 revealed the presence of three sub-clades: Clade 1a contained the five ASV which displayed significant differential abundance between mineral-N treatments. These sequences were found to have 97–98% similarity to one another.

Clade 2 included most of the reference sequences from Australian pastures, including TS OTU18, 152, 289, 350 and 980, and a sequence obtained from Australian agricultural soils (FREOZ OTU170). Clade 3 represented novel diversity. Lastly, Clade 4 represented the remaining low abundance sequences amplified with both primer sets, and was associated with ectomycorrhizal and saprotrophic Endogonales. Notably reference sequences obtained from hornworts and liverworts were also found in clades 1, 2 and 4.

Discussion

In the current study we report findings using two new sets of primers. The AM-primer set is a modification of those published by Sato et al. [54], which co-amplify Mucoromycotina and Glomeromycotina, and have been used to profile putative M-AMF communities in previous studies [2,3,4, 33, 41]. The second, FRE-primers, specifically amplify Mucoromycotinian fungi, and provide a longer 440 bp sequence length than the Sato et al. [54] primers (220 bp). The AM-primers were used to investigate whether co-occurring G-AMF and M-AMF responded in a similar manner to different wheat varieties and N-addition, but neither factor had a significant effect on the diversity or composition of either Glomeromycotina or Mucoromycotina communities. Data generated with the FRE-primers also indicated no effect of wheat variety on Mucoromycotina community composition, but in contrast to the AM-primers, the FRE-primers indicated that N-addition induced changes to Mucoromycotina alpha and beta diversity, with marked effects on the relative abundance of a number of ASV. The dominant Mucoromycotina sequences amplified with both primer sets formed phylogenetic clades within the order Endogonales, along with sequences associated with FRE morphology from previous studies in Australia [3, 33, 41], and also the only described species of M-AMF to date, Planticonsortium tenue, isolated from New Zealand [4, 69].

The AM and FRE primer sets amplified similar richness of Mucoromycotina associated with wheat roots, although the AM-primers provided a greater phylogenetic range than the FRE-primers. Importantly, there were differences in the relative amplification of different phylogenetic clades between the primers. The FRE-primers detected shifts in community composition associated with N-addition that the AM-primers did not. This was shown to be the result of preferential amplification by the FRE-primers of a clade associated with FRE morphology in previous studies in Australia [3, 4, 33]. This highlights the importance of primer choice for characterising AMF communities and interpreting responses to environmental parameters. Similarly, a range of primer sets have been used to characterise G-AMF communities, and these have well known biases which can affect the evaluation of taxa richness and the relative abundance of families [23, 25].

Notably the AM-primers amplified a broad range of Mucoromycota in addition to Mucoromycotina, and furthermore this and earlier studies [3] show they provide very limited coverage of Diversisporales, Archaeosporales and Paraglomerales. Co-amplification of Mucoromycotina with other Mucoromycota sub phyla may be an advantage in some circumstances, such as to allow comparative understanding of ecological distribution patterns within the Mucoromycota. However, the limited coverage of Glomeromycota is problematic since Diversisporales, Archaeosporales and Paraglomerales are widely distributed and abundant, particularly within agricultural soils [16, 25]. Despite the longer fragment size amplified by the FRE primers, they provided less phylogenetic diversity than the AM primers, and in contrast to the AM primers, the FRE primers identified community responses to N application. To provide the most comprehensive analysis of AMF communities for metabarcoding analysis separate analyses of Glomeromycotina and Mucoromycotina is preferable. The use of widely used Glomeromycotina primers [25], together with our new Mucoromycotina specific FRE primers, provides the optimal approach.

The relatedness of Mucoromycotinian sequences detected in our study with the FRE forming P. tenue and Mucoromycotinian sequences associated with FRE morphology in Australia [3, 4, 33, 41], suggests a global distribution of Endogonales clades associated with FRE morphology. Despite being short amplicon sequences, several ASV were very close matches to sequences described from Australia, indicating that some M-AMF taxa could have a global distribution. Furthermore, several of the abundant Mucoromycotinian sequences we detected in roots showed close similarity to sequences associated with rhizoids of hornworts and liverworts, suggesting that these fungi could form both AM symbioses with higher plants and mycorrhiza-like associations with both early diverging vascular plants and non-vascular plants, akin to G-AMF. However, M-AMF sequences associated with non-vascular plants showed a broader phylogenetic distribution than the Mucormycotina sequences associated with FRE morphology from this study, and the earlier studies in Australia [3, 4, 33, 41].

In the current study, and in previous studies in which Mucoromycotina associated with FRE morphology have been characterised [3, 4, 33, 41], a diverse assemblage of Mucoromycotinian fungi has been detected. This could therefore suggest that there is a wide diversity of Mucoromycotina which can form AM associations, similar to G-AMF. Additionally, at some sites, such as the conventional high input agricultural location studied here, the richness of root associated Mucoromycotina ASV may even be higher than Glomeromycotina. To date only a single FRE species, P. tenue, has been described based on morphological evidence [69]. However, Thippayarugs et al. [62] identified a range of morphological characteristics which varied across published studies of FRE. They used 11 characteristics including hyphal surface smoothness, hyphal diameter and branching, vesicle diameter and shape, and staining intensity with trypan blue to differentiate five distinct FRE morphological groups colonising Trifolium subterraneum in Western Australia. There is a clear need to link molecular and morphological evidence to develop a taxonomic classification of M-AMF.

G-AMF may confer a range of benefits to their host, particularly P uptake, although they may also promote supply of other nutrients including N, calcium, magnesium and micronutrients. G-AMF may also provide the host with resistance against pests and disease, particularly soil-borne pathogens [15]. Understanding of functional diversity within G-AMF communities is limited, although there is evidence for trait-based variation between Glomeromycotina families. In particular, Gigasporaceae may form abundant extraradical mycelium which facilitates P uptake from soil and its subsequent translocation to the host. In contrast, Glomeraceae mycelium may mostly grow within the root, providing protection against soil-borne pathogens but lower benefits to host-P nutrition, while Acaulosporaceae may produce low hyphal biomass compared to the other groups, but equivalent P uptake to Glomeraceae, providing cost effective trade in C for P [32].

The functional significance of M-AMF is less clear, although there is evidence that M-AMF may provide N [20] and P to their host plant [41]. Comparative studies of Glomeromycotina and Mucoromycotina fungi associated with liverworts and hornworts showed that Mucoromycotina facilitated higher N uptake to the host, while the reverse was true for P. This could suggest complementary roles in nutrient acquisition by these groups [13]. The significance of diverse Mucoromycotina communities within roots, and their contribution towards the functional diversity of AM symbioses remains to be determined.

The tendency of G-AMF and M-AMF to intermingle in plant roots has frequently been described [21, 51, 71], and all samples analysed in the current study, using microscopy and molecular analysis, contained both AM groups. Albornoz et al. [3, 4] suggested that the two groups had overlapping ecological niches at the landscape scale, although there were different responses to temperature, pH and plant richness. In contrast to Glomeromycotina, nutrient availability was an important determinant of Mucoromycotina abundance, which increased with fertility.

In the current study, characterisation of Mucoromycotina with FRE-primers indicated shifts in community composition associated with N-addition, including changes to the dominant ASV. We have revealed that the response to N-addition was clearly linked to phylogeny, such that the five ASV with differential abundance between N treatments (ASV 3, 7, 17, 18 and 21) belonged to the same clade. Moreover, these ASV were closely related to one another within a sub-clade (Clade 1a) which was mostly accessed by the FRE-primers. The biological significance of these abundance shifts are unclear, however, since they could reflect direct impacts of N on Mucoromycotina, or indirect effects operating through the plant or other components of the root microbiome [42]. In contrast to G-AMF [67], agricultural management may favour M-AMF [4, 41, 56] and this could indicate a response to varied management practices including tillage and fertilisation.

Interestingly no effect of wheat variety was detected on either Glomeromycotina or Mucoromycotina communities. Previous studies indicated that Aszita and Skyfall supported high and low abundances of AM, respectively [11, 28, 63]. However, our data suggest that variety preferences for AM may vary across sites, determined by local climate, soil properties and management practices. This highlights the problems of extrapolating the outcomes of plant-AM interactions across locations, and for managing AM communities to support crop nutrition and system sustainability [50].

Availability of data and materials

The raw sequence datasets and metadata reported in this study are available in the NCBI Sequence Read Archive under BioProject ID PRJNA1026851. The ASV sequences are deposited in the NCBI GenBank database under SUB13873553 (AM primers) and SUB13873261 (FRE primers).

References

  1. Alaux PL, César V, Naveau F, Cranenbrouck S, Declerck S. Impact of Rhizophagus irregluaris MUCL 41833 on disease symptoms caused by Phytophtera infestans in potato grown under field conditions. Crop Prot. 2018;107:26–33.

    Article  Google Scholar 

  2. Albornoz FE, Hayes PE, Orchard S, Clode PL, Nazeri NK, Standish RJ, Bending GD, Hilton S, Ryan MH. First cryo-scanning electron microscopy images and X-ray microanalyses of mucoromycotinian fine root endophytes in vascular plants. Front Microbiol. 2020;11:1–13.

    Article  Google Scholar 

  3. Albornoz FE, Orchard S, Standish RJ, Dickie IA, Bending GD, Hilton S, Lardner T, Foster KJ, Gleeson DB, Bougoure J, et al. Evidence for niche differentiation in the environmental responses of co-occurring mucoromycotinian fine root endophytes and glomeromycotinian arbuscular mycorrhizal fungi. Microb Ecol. 2021;81:864–73.

    Article  CAS  PubMed  Google Scholar 

  4. Albornoz FE, Ryan MH, Bending GD, Hilton S, Dickie IA, Gleeson DB, Standish RJ. Agricultural land-use favours Mucoromycotinian, but not Glomeromycotinian, arbuscular mycorrhizal fungi across ten biomes. New Phytol. 2022;233:1369–82.

    Article  PubMed  Google Scholar 

  5. Albornoz FE, Standish RJ, Bissett A, Prober SM. Richness of arbuscular mycorrhizal fungi increases with ecosystem degradation of temperate eucalypt woodlands. Plant Soil. 2023;488:255–71.

    Article  CAS  Google Scholar 

  6. Altschul S, Madden TL, Schaffer AA, Zhang J, Zhang Z, Miller W, Lipman D. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 1997;25:3389–402.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  7. Callahan BJ, McMurdie PJ, Rosen MJ, Han AW, Johnson AJA, Holmes SP. DADA2: high-resolution sample inference from Illumina amplicon data. Nat Methods. 2016;13:581–3.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  8. Caporaso JG, Kuczynski J, Stombaugh J, Bittinger K, Bushman FD, Costello EK, Fierer N, Pẽa AG, Goodrich JK, Gordon JI, et al. QIIME allows analysis of high-throughput community sequencing data. Nat Methods. 2010;7:335–6.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  9. Clark CM, Tilman D. Loss of plant species after chronic low-level nitrogen deposition to prairie grasslands. Nature. 2008;451:712–5.

    Article  CAS  PubMed  Google Scholar 

  10. Di HJ, Cameron KC. Nitrate leaching in temperate agroecosystems: sources, factors and mitigating strategies. Nutr Cycl Agroecosyst. 2002;64:237–56.

    Article  CAS  Google Scholar 

  11. Elliott AJ, Daniell TJ, Cameron DD, Field KJ. A commercial arbuscular mycorrhizal inoculum increases root colonization across wheat cultivars but does not increase assimilation of mycorrhiza-acquired nutrients. Plants People Planet. 2019;00:1–12.

    Google Scholar 

  12. Ercoli L, Schüßler A, Arduini I, Pellegrino E. Strong increase of durum wheat iron and zinc content by field-inoculation with arbuscular mycorrhizal fungi at different soil nitrogen availabilities. Plant Soil. 2017;419:153–67.

    Article  CAS  Google Scholar 

  13. Field KJ, Bidartondo MI, Rimington WR, Hoysted GA, Beerling DJ, Cameron DD, Duckett JG, Leake JR, Pressel S. Functional complementarity of ancient plant–fungal mutualisms: contrasting nitrogen, phosphorus and carbon exchanges between Mucoromycotina and Glomeromycotina fungal symbionts of liverworts. New Phytol. 2019;223:908–21.

    Article  CAS  PubMed  Google Scholar 

  14. Gianinazzi-Pearson VDM, Dexheimer J, Gianinazzi S. Ultrastructural and ultracytochemical features of a Glomus tenuis mycorrhiza. New Phytol. 1981;88:633–9.

    Article  Google Scholar 

  15. Gosling P, Hodge A, Goodlass G, Bending GD. Arbuscular mycorrhizal fungi and organic farming. Agric Ecosyst Environ. 2006;113:17–35.

    Article  Google Scholar 

  16. Gosling P, Proctor M, Jones J, Bending GD. Distribution and diversity of Paraglomus spp. in tilled agricultural soils. Mycorrhiza. 2014;24:1–11.

    Article  PubMed  Google Scholar 

  17. Hall IR. Species and mycorrhizal infections of New Zealand Endogonaceae. Trans Br Mycol Soc. 1977;68:341–56.

    Article  Google Scholar 

  18. Hetrick BAD, Bockus WW, Bloom J. The role of vesicular arbuscular mycorrhizal fungi in the growth of Kansas USA winter wheat triticum-aestivum. Can J Bot. 1984;62:735–40.

    Article  Google Scholar 

  19. Hetrick BAD, Wilson GWT, Cox TS. Mycorrhizal dependence of modern wheat cultivars and ancestors: a synthesis. Can J Bot. 1993;71:512–8.

    Article  Google Scholar 

  20. Hoysted GA, Field KJ, Sinanaj B, Bell CA, Bidartondo MI, Pressel S. Direct nitrogen, phosphorus and carbon exchanges between mucoromycotina ‘fine root endophyte’ fungi and a flowering plant in novel monoxenic cultures. New Phytol. 2023;238:70–9.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  21. Jeffery RP, Simpson RJ, Lambers H, Orchard S, Kidd DR, Haling RE, Ryan MH. Contrasting communities of arbuscule-forming root symbionts change external critical phosphorus requirements of some annual pasture legumes. Appl Soil Ecol. 2018;126:88–97.

    Article  Google Scholar 

  22. Joo L, Beirinckx S, Haegeman A, Debode J, Vandecasteele B, Baeyen S, Goormachtig S, Clement L, De Tender C. Daring to be differential: metabarcoding analysis of soil and plant-related microbial communities using amplicon sequence variants and operational taxonomical units. BMC Genomics. 2020;21:733.

    Article  Google Scholar 

  23. Kirkman ER, Hilton S, Sethuraman G, Elias DMO, Taylor A, Clarkson J, Soh AC, Bass D, Ooi GT, McNamara NP, et al. Diversity and ecological guild analysis of the oil palm fungal microbiome across root, rhizosphere, and soil compartments. Front Microbiol. 2022;13:209.

    Article  Google Scholar 

  24. Koide RT. Nutrient supply, nutrient demand and plant response to mycorrhizal infection. New Phytol. 1991;117:365–86.

    Article  CAS  PubMed  Google Scholar 

  25. Kohout P, Sudová R, Janoušková ČM, Hejda M, Pánková H, Slavíková R, Štajerová K, Vosátka M, Sýkorová Z. Comparison of commonly used primer sets for evaluating arbuscular mycorrhizal fungal communities: is there a univsersal solution? Soil Biol Biochem. 2014;68:482–93.

    Article  CAS  Google Scholar 

  26. Leff JW, Jones SE, Prober SM, Barberán A, Borer ET, Firn JL, Harpole WS, Hobbie SE, Hofmockel KS, Knops JMH, et al. Consistent responses of soil microbial communities to elevated nutrient inputs in grasslands across the globe. Proc Natl Acad Sci USA. 2015;112:10967–72.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  27. Lehnert H, Serfling A, Enders M, Friedt W, Ordon F. Genetics of mycorrhizal symbiosis in winter wheat (Triticum aestivum). New Phytol. 2017;215:779–91.

    Article  CAS  PubMed  Google Scholar 

  28. Lehnert H, Serfling A, Friedt W, Ordon F. Genome-wide association studies reveal genomic regions associated with the response of wheat (Triticum aestivum l.) to mycorrhizae under drought stress conditions. Front Plant Sci. 2018;871:1728.

    Article  Google Scholar 

  29. Leiser WL, Olatoye MO, Rattunde HFW, Neumann G, Weltzien E, Haussmann BIG. No need to breed for enhanced colonization by arbuscular mycorrhizal fungi to improve low-P adaptation of West Africa sorghums. Plant Soil. 2016;401:51–64.

    Article  CAS  Google Scholar 

  30. Liu W, Jiang S, Zhang Y, Yue S, Christie P, Murray PJ, Li X, Zhang J. Spatiotemporal changes in arbuscular mycorrhizal fungal communities under different nitrogen inputs over a 5-year period in intensive agricultural ecosystems on the North China Plain. FEMS Microbiol Ecol. 2014;90:436–53.

    CAS  PubMed  Google Scholar 

  31. Love MI, Huber W, Anders S. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol. 2014;15:1–21.

    Article  Google Scholar 

  32. Maherali M, Klironomos JN. Influence of phylogeny on fungal community assembly and ecosystem functioning. Science. 2007;316:1746–8.

    Article  CAS  PubMed  Google Scholar 

  33. Mansfield TM, Albornoz FE, Ryan MH, Bending GD, Standish RJ. Niche differentiation of Mucoromycotinian and Glomeromycotinian arbuscular mycorrhizal fungi along a 2-million-year soil chronosequence. Mycorrhiza. 2023;33:139–52.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  34. Mao L, Liu Y, Shi G, Jiang S, Cheng G, Li X, An L, Feng H. Wheat cultivars form distinctive communities of root-associated arbuscular mycorrhiza in a conventional agroecosystem. Plant Soil. 2014;374:949–61.

    Article  CAS  Google Scholar 

  35. Martín-Robles N, Lehmann A, Seco E, Aroca R, Rillig MC, Milla R. Impacts of domestication on the arbuscular mycorrhizal symbiosis of 27 crop species. New Phytol. 2018;218:322–34.

    Article  PubMed  Google Scholar 

  36. McMurdie PJ, Holmes S. phyloseq: an R package for reproducible interactive analysis and graphics of microbiome census data. PLoS ONE. 2013;8:1–11.

    Article  Google Scholar 

  37. Mercy L, 2017. INOQ calculator advanced. Evaluate the mycorrhizal rate according to a modified Trouvelot method. https://doi.org/10.13140/RG.2.2.13641.03684

  38. Miller MA, Pfeiffer W, Schwartz T. Creating the CIPRES science gateway for inference of large phylo- genetic trees. In: Proceedings of the gateway computing environments workshop (GCE). Institute of Electrical and Electronic Engineers;2010; pp. 1–8.

  39. Oksanen J, Blanchet FG, Friendly M, Kindt R, Legendre P, McGlinn D, Minchin PR, O'Hara RB, Simpson GL, Solymos P, et al. vegan: community ecology package. In. R package version 2.5-3. 2018. https://CRAN.R-project.org/package=vegan.

  40. Orchard S, Hilton S, Bending GD, Dickie IA, Standish RJ, Gleeson DB, Jeffery RP, Powell JR, Walker C, Bass D, et al. Fine endophytes (Glomus tenue) are related to Mucoromycotina, not Glomeromycota. New Phytol. 2017;213:481–6.

    Article  PubMed  Google Scholar 

  41. Orchard S, Standish RJ, Dickie IA, Renton M, Walker C, Moot D, Ryan MH. Fine root endophytes under scrutiny: a review of the literature on arbuscule-producing fungi recently suggested to belong to the Mucoromycotina. Mycorrhiza. 2017;27:619–38.

    Article  PubMed  Google Scholar 

  42. Picot E, Hale CC, Hilton S, Teakle GR, Schäfer H, Huang Y, Perryman M, West J, Bending GD. Contrasting responses of rhizosphere bacterial, fungal, protist, and nematode communities to nitrogen fertilization and crop genotype in field grown oilseed rape (Brassica napus). Front Sustain Food Syst. 2021;5:613269.

    Article  Google Scholar 

  43. Pinheiro J, Bates D, DebRoy S, Sarkar D, R Core Team. nlme: linear and nonlinear mixed effects models (2021). https://CRAN.R-project.org/package=nlme

  44. Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P, Peplies J, Glöckner FO. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res. 2014;41:590–6.

    Article  Google Scholar 

  45. RStudio Team. 2015. RStudio: integrated development environment for R, Boston, MA. http://www.rstudio.com/.

  46. Redecker D, Kodner R, Graham L. Glomalean fungi from the Ordovician. Science. 2000;289:1920–1.

    Article  CAS  PubMed  Google Scholar 

  47. Rempelos L, Almuayrifi MSB, Branski M, Tetrad-Jones C, Barkla B, Cakmak I, Ozturk L, Cooper J, Volakakis N, Hall G, et al. The effect of agronomic factors on crop health and performance of winter wheat varieties bred for the conventional and the low input farming sector. Field Crop Res. 2020;254: 107822.

    Article  Google Scholar 

  48. Rillig M, Aguilar-Trigueros CA, Camenzind T, Cavagnaro TR, Degrune F, Hohmann P, Lammel DR, In M, Roy J, Van der Heijden MGA, Yang G. Why farmers should manage the arbuscular mycorrhizal symbiosis. New Phytol. 2018;222:1171–5.

    Article  Google Scholar 

  49. Ronquist F, Teslenko M, van der Mark P, Ayres DL, Darling A, Höhna S, Larget B, Liu L, Suchard MA, Huelsenbeck JP. MrBayes 3.2: efficient Bayesian phylogenetic inference and model choice across a large model space. Syst Biol. 2012;61:539–42.

    Article  PubMed  PubMed Central  Google Scholar 

  50. Ryan MH, Graham JR. Little evidence that farmers should consider abundance or diversity of arbuscular mycorrhizal fungi when managing crops. New Phytol. 2018;220:1092–107.

    Article  PubMed  Google Scholar 

  51. Ryan MH, Kirkegaard J. The agronomic relevance of arbuscular mycorrhizas in the fertility of Australian extensive cropping systems. Agric Ecosyst Environ. 2012;163:37–53.

    Article  Google Scholar 

  52. Ryan MH, Van Herwaarden AF, Angus JF, Kirkegaard JA. Reduced growth of autumn-sown wheat in a low-P soil is associated with high colonisation by arbuscular mycorrhizal fungi. Plant Soil. 2005;270:275–86.

    Article  CAS  Google Scholar 

  53. Ryan MH, Kidd DR, Sandral GA, Yang Z, Lambers H, Culvenor RA, Stefanski A, Nichols PGH, Haling RE, Simpson RJ. High variation in the percentage of root length colonised by arbuscular mycorrhizal fungi among 139 lines representing the species subterranean clover (Trifolium subterraneum). Appl Soil Ecol. 2016;98:221–32.

    Article  Google Scholar 

  54. Sato K, Suyama Y, Saito M, Sugawara K. A new primer for discrimination of arbuscular mycorrhizal fungi with polymerase chain reaction-denature gradient gel electrophoresis. Grassland Sci. 2005;51:179–81.

    Article  CAS  Google Scholar 

  55. Sawers RJH, Svane SF, Quan C, Grønlund M, Wozniak B, Gebreselassie MN, Gonzáez-Muñoz E, Montes RAC, Baxter I, Goudet J, et al. Phosphorus acquisition efficiency in arbusuclar mycorrhizal maize is correlated with the abundance of root-external hyphae and the accumulation of transcripts encoding PHT1 phosphate transporters. New Phytol. 2017;214:632–43.

    Article  CAS  PubMed  Google Scholar 

  56. Sigüenza C, Crowley DE, Allen EB. Soil microorganisms of a native shrub and exotic grasses along a nitrogen deposition gradient in southern California. Appl Soil Ecol. 2006;32:13–26.

    Article  Google Scholar 

  57. Simon L, Bousquet J, Lévesque RC, Lalonde M. Origin and diversification of endomycorrhizal fungi and coincidence with vascular land plants. Nature. 1993;363:67–9.

    Article  Google Scholar 

  58. Smith SE, Manjarrez M, Stonor R, McNeill A, Smith FA. Indigenous arbuscular mycorrhizal (AM) fungi contribute to wheat phosphate uptake in a semi-arid field environment, shown by tracking with radioactive phosphorus. Appl Soil Ecol. 2015;96:68–74.

    Article  Google Scholar 

  59. Spatafora JW, Chang Y, Benny GL, Lazarus K, Smith ME, Berbee ML, Bonito G, Corradi N, Grigoriev I, Gryganskyi A, James TY, O’Donnell K, Roberson RW, Taylor TN, Uehling J, Vilgalys R, White MM, Stajich JE. A phylum-level phylogenetic classification of zygomycete fungi based on genome-scale data. Mycologia. 2016;108:1028–46.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  60. Strullu-Derrien C, Kenrick P, Pressel S, Duckett JG, Rioult JP, Strullu DG. Fungal associations in Horneophyton ligneri from the Rhynie Chert (c. 407 million year old) closely resemble those in extant lower land plants: Novel insights into ancestral plant-fungus symbioses. New Phytol. 2014;203:964–79.

    Article  PubMed  Google Scholar 

  61. Taylor A, Pereira N, Thomas B, Pink DAC, Jones JE, Bending GD. Growth and nutritional responses to arbuscular mycorrhizal fungi are dependent on onion genotype and fungal species. Biol Fertil Soils. 2015;51:801–13.

    Article  Google Scholar 

  62. Thippayarugs S, Bansal M, Abbott LK. Morphology and infectivity of fine endophyte in a mediterranean environment. Mycol Res. 1999;103:1369–79.

    Article  Google Scholar 

  63. Thirkell TJ, Grimmer M, James L, Pastok D, Allary T, Elliott A, Paveley N, Daniell T, Field KJ. Variation in mycorrhizal growth response among a spring wheat mapping population shows potential to breed for symbiotic benefit. Food Energy Secur. 2022;11:e370.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  64. Tipton L, Zahn GL, Darcy J, Amend AS, Hynson NA. Hawaiian fungal amplicon sequence variants reveal otherwise hidden biogeography. Microb Ecol. 2021;83:48–57.

    Article  PubMed  Google Scholar 

  65. Tisserant E, Malbreil M, Kuo A, Kohler A, Symeonidi A, Balestrini R, Charron P, Duensing N, Frei Dit Frey N, Gianinazzi-Pearson V, et al. Genome of an arbuscular mycorrhizal fungus provides insight into the oldest plant symbiosis. Proc Natl Acad Sci USA. 2013;110:20117–22.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  66. Trouvelot A, Kough JL, Giannazzi-Pearson V. 1986. Mesure du taux de mycorhization VA d’un système radiculaire. Recherche de méthode d’estimation ayant une signification fonctionnelle. In: Aspects physiologiques et génétiques des mycorhizes: actes du 1er symposium européen sur les mycorhizes. Dijon: INRA, Paris, pp 217–221.

  67. Verbruggen E, Röling WFM, Gamper HA, Kowalchuk GA, Verhoef HA, van der Heijden MGA. Positive effects of organic farming on below-ground mutualists: large-scale comparison of mycorrhizal fungal communities in agricultural soils. New Phytol. 2010;186:968–79.

    Article  CAS  PubMed  Google Scholar 

  68. Vierheilig H, Coughlan AP, Wyss U, Piché Y. Ink and vinegar, a simple staining technique for arbuscular-mycorrhizal fungi. Appl Environ Microbiol. 1998;64:5004–7.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  69. Walker C, Gollotte A, Redecker D. A new genus, Planticonsortium (Mucoromycotina), and new combination (P. tenue), for the fine root endophyte, Glomus tenue (basionym Rhizophagus tenuis). Mycorrhiza. 2018;28:213–9.

    Article  PubMed  Google Scholar 

  70. Wickham H. ggplot2: elegant graphics for data analysis. New York: Springer; 2016.

    Book  Google Scholar 

  71. Yamamoto K, Shimamura M, Degawa Y, Yamada A. Dual colonization of Mucoromycotina and Glomeromycotina fungi in the basal liverwort, Haplomitrium mnioides (Haplomitriopsida). J Plant Res. 2019;132:777–88.

    Article  CAS  PubMed  Google Scholar 

  72. Yu G. Data integration, manipulation and visualization of phylogenetic trees. 1st ed. Cambridge: Chapman and Hall/CRC; 2022.

    Book  Google Scholar 

  73. Zadoks J, Chang T, Konzak C. A decimal growth code for the growth stages of cereals. Weed Res. 1974;14:415–21.

    Article  Google Scholar 

  74. Zhang T, Chen HYH, Ruan H. Global negative effects of nitrogen deposition on soil microbes. ISME J. 2018;12:1817–25.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  75. Zhang S, Lehmann A, Zheng W, You Z, Rillig MC. Arbuscular mycorrhizal fungi increase grain yields: a meta-analysis. New Phytol. 2019;222:543–55.

    Article  CAS  PubMed  Google Scholar 

Download references

Acknowledgements

We thank the Wildlife Trust for Bedfordshire, Cambridgeshire and Northamptonshire for permission to collect samples from Boddington Meadow

Funding

This project was funded by the UK Natural Environment Research Council Grant NE/S010270/1 and the European Union Marie Skłodowska-Curie Grant Agreement No. 722642.

Author information

Authors and Affiliations

Authors

Contributions

GB, MR, RS and FA conceived the work on Mucoromycotan fungi and GB and MR obtained funding. SH designed the primers and performed the sequencing and bioinformatic processing. PB and CS conceived the field experiment and obtained funding. NG, PB CS and LR designed the field experiment. LM and CW supervised the mycorrhizal visual analysis. MS performed the field sampling and DNA extraction supervised by NG, PB, LR, CS and LM. MS performed the bioinformatic analysis of M-AMF and G-AMF supervised by GB and SH. DB performed the sequence alignments and GM drew the trees and performed the phylogenetic analysis. MS and GB wrote the manuscript. All authors edited the manuscript.

Corresponding authors

Correspondence to Mirjam Seeliger or Gary D. Bending.

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.

Supplementary Information

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Seeliger, M., Hilton, S., Muscatt, G. et al. New fungal primers reveal the diversity of Mucoromycotinian arbuscular mycorrhizal fungi and their response to nitrogen application. Environmental Microbiome 19, 71 (2024). https://doi.org/10.1186/s40793-024-00617-x

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: https://doi.org/10.1186/s40793-024-00617-x

Keywords