Fungal Symbionts of Endangered Crocanthemum Canadense (Cistaceae) in Nova Scotia

Botany

Fungal symbionts of endangered Crocanthemum canadense (Cistaceae) in Nova Scotia

Journal: Botany

Manuscript ID cjb-2020-0187.R2

Manuscript Type: Article

Date Submitted by the 14-Jan-2021 Author:

Complete List of Authors: Byers, Philicity; Acadia University Faculty of Pure and Applied Sciences, Department of Earth and Environmental Science Evans, Rodger; Acadia University Faculty of Pure and Applied Sciences, Department of Biology Migicovsky,Draft Zoë; Dalhousie University Faculty of Agriculture, Department of Plant, Food and Environmental Sciences Walker, Allison; Acadia University Faculty of Pure and Applied Sciences, Department of Biology

Keyword: AMF, Cistaceae, endophyte, sand barren, Crocanthemum

Is the invited manuscript for consideration in a Special Not applicable (regular submission) Issue? :

Fungal symbionts of endangered Crocanthemum canadense (Cistaceae) in Nova Scotia

Philicity R. M. Byers*1,4, Rodger C. Evans2,5, Zoë Migicovsky3,6, and Allison K. Walker2,7

1Department of Earth and Environmental Science, Faculty of Pure and Applied Science, Acadia

University, Wolfville, Nova Scotia, B4P 2R6, Canada.

2Department of Biology, Faculty of Pure and Applied Science, Acadia University, Wolfville,

Nova Scotia, B4P 2R6, Canada.

3Department of Plant, Food and Environmental Sciences, Faculty of Agriculture, Dalhousie

University, Truro, NS, B2N 5E3, CanadaDraft

*Corresponding author. Telephone: 1 (902) 523-3425

4email: [email protected]

5email: [email protected]

6email: [email protected]

7email: [email protected]

Abstract

Crocanthemum canadense (L.) Britt. (Cistaceae) is critically imperiled in Nova Scotia. The

decline of Nova Scotian C. canadense is largely due to the loss of the Annapolis Valley

sand barrens habitat. Fungal symbionts may aid in nutrient and water acquisition as well as

plant defenses. The role of fungal associations with C. canadense is unknown; our goal

was to identify fungal symbionts to inform ongoing conservation research. We isolated

fungi from eighteen C. canadense plants collected from Greenwood, Nova Scotia. Using

ITS rDNA barcoding of fungal cultures, we identified 58 fungal taxa. ITS2 meta-amplicon

barcoding of roots and rhizosphere soil revealed 241 fungi with basidiomycetes accounting

for 53.8% of reads. Chaetothyriales sp., Mycetinis scorodonius, Acidomelania panicicola,

and Scleroderma citrinum were the mostDraft abundant root associates based on meta-amplicon

data. We quantified percent root colonization of arbuscular mycorrhizal fungi (AMF) using

root staining and microscopy. The average AMF colonization rate of the roots was 29.6%

(n=18). We provide a foundation for understanding the fungal community in this declining

habitat and the first account of fungal symbionts in the above- and below-ground tissues

and rhizosphere of C. canadense. Identifying fungi influencing endangered Nova Scotian

C. canadense is valuable for developing conservation strategies.

Keywords: AMF, Cistaceae, Crocanthemum, endangered species, endophyte, mycorrhiza, sand barren

Introduction

Crocanthemum canadense (L.) Britt. (Cistaceae) is a small, herbaceous perennial

classified as critically imperiled in Nova Scotia by the Nova Scotia Endangered Species Act

(Figure 1A–B) (Newell 2007). The global distribution of C. canadense is confined to Eastern

North America with Canadian populations in Nova Scotia, Quebec, and Ontario (Figure 2)

(Newell 2007). Surveys in 2006 estimated that roughly 5 000 C. canadense plants remained in

Nova Scotia (Newell 2007). More recently, Newell (2018) estimated that 7 100 – 7 800 C.

canadense plants remained in Nova Scotia. Two of the three remaining Nova Scotian C.

canadense populations inhabit the remnants of the Corema conradii-dominated sand barrens that

are unique to Eastern North America (Carbyn et al. 2006). The soil profile is acidic, rapidly

drained, and composed of well sorted sandDraft from glaciofluvial parent material (Neily et al. 2017).

The sand barren habitat has been reduced to less than 3% of its original span in the Annapolis

Valley, Nova Scotia, leaving little habitat intact for native plants favouring this environment

(Carbyn et al. 2006).

Habitat loss is largely due to land conversion for anthropogenic purposes and the invasion

of the introduced species Pinus sylvestris L. (Scots Pine) (Carbyn et al. 2006; Newell 2007).

Human-caused suppression of natural disturbances, including wildfires and grazing, further

reduced the availability of open habitat for native vascular plants like C. canadense. Without

these ecosystem processes reducing the growth of larger vegetation, C. canadense has been

unable to compete and establish as extensively. It is important to protect this habitat along with

the native flora and their concurrent fungal symbionts to maintain a healthy ecosystem.

Fungi perform vital ecosystem services by decomposing and recycling organic matter,

making otherwise scarce or inaccessible nutrients available for uptake by plants; some fungi can

improve these services by forming direct relationships with plant roots, called mycorrhizae

(Bonfante 2001). Arbuscular mycorrhizal fungi (AMF), from the phylum Glomeromycota, can benefit plant hosts in many ways and are especially beneficial when water and nutrients are scarce (Smith and Read 1997). Fungal endophytes asymptomatically inhabit living plant tissue and can reduce abiotic and biotic stresses on plant hosts by providing resistance to pathogenic fungi, herbivorous insects, and variations in environmental conditions, including temperature and moisture (Owen and Hundley 2004; Rodriguez et al. 2004; Porras-Alfaro and Bayman 2011).

Endophytic and mycorrhizal symbionts may provide a significant ecological advantage for the establishment, stress tolerance, and survival of the three remaining populations of the critically imperilled plant, C. canadense, in Nova Scotia.

Malloch and Thorn (1985) first investigatedDraft mycorrhizal associations with various species of Cistaceae in central and eastern Canada, including the species of interest in this study.

All species examined were found to have some evidence of ectomycorrhizae. A later study conducted in Nova Scotia investigated the ectomycorrhizal associations with two Cistaceae species: Hudsonia ericoides L. and Hudsonia tomentosa Nutt. using microscopic analysis

(Massicotte et al. 2010). Samples were collected west of Kingston near the current sites of interest (Massicotte et al. 2010). Studies investigating the fungal symbionts of Cistaceae plants have largely focused on ectomycorrhizal symbionts (Malloch and Thorn 1985; Dickie et al.

2004; Comandini et al. 2006; Massicotte et al. 2010; Leonardi et al. 2018). Insufficient exploration of other ecological fungal groups associated with Cistaceae limits our understanding of the fungal interactions occurring.

The fungal symbionts of the declining Nova Scotian C. canadense populations remain unexplored but must be studied to acquire a better understanding of their ecological relationships

and symbioses to preserve the vital elements of their deteriorating habitat. This study was

conducted to inform future conservation efforts and had three main objectives: (1) to isolate and

identify endophytic fungi within C. canadense plants to characterize the fungal assemblage

associated with this plant population; (2) to determine the taxonomic diversity of the fungal

assemblage within C. canadense roots and sandy rhizosphere soil using ITS2 metabarcoding; (3)

to measure arbuscular mycorrhizal colonization of C. canadense roots to verify and quantify

AMF presence in this endangered plant.

Materials and Methods

Sample collection We collected eighteen C. canadenseDraft plants across six sand barren study sites at 14 Wing Canadian Forces Base Greenwood, Kings County, Nova Scotia (hereafter CFB Greenwood) on 3

and 4 July 2018. These six plots were set up as part of ongoing research projects investigating C.

canadense and are home to the largest remaining C. canadense population in Nova Scotia. Five

of the six plots were 10 m x 10 m and less than 200 m from one another. The sixth plot was

approximately 500 m from the other plots and was 10 m x 25 m. All plots had full sun exposure

or partial shade with ground cover primarily consisting of grasses, ferns, asters, blackberry, and

broom crowberry (Figure 1C–D). Upon the emergence of C. canadense flowers, we harvested

three plants from each site including the corresponding root system. To select plants, we used an

opportunistic sampling approach based on visual assessments to select plants that were similar in

size, had at least one open flower, and were more than 1 m apart. Using a garden trowel, we

excavated a 10 cm x 10 cm section of rhizosphere soil surrounding each of the plants to a depth

of 10 cm to obtain the root system. We cleaned the trowel with 70% ethanol after collecting each

plant. Soil was then shaken from the roots of each sample, collected in Ziploc bags, and

transported on ice to Acadia University.

Soil chemistry and nutrient analysis

Approximately 250 mL of soil was measured from the root zone of each plant sampled

using a measuring cup. One pooled soil sample weighing approximately 450 g (500 mL) was

analyzed for each site. Each of the six soil samples contained equal parts of soil collected from

the root zone of the three plants from the corresponding site. The samples were transported in

large Ziploc bags for nutrient analysis at the Nova Scotia Department of Agriculture,

Laboratory Services in Truro, Nova Scotia. The laboratory used the Modified Mehlich III method to determine the nutrient contentDraft in the soil (Mehlich 1984).

Plant tissue preparation

All of the plants we collected were similar in size with some variation in the size of the root systems. We divided each plant into leaf, upper stem, lower stem, and root subsamples to investigate endophytic fungi using culture-based methods. Large leaves (>10 mm) were cut into thirds, medium leaves (5 to 10 mm) into halves, and smaller leaves (<5 mm) were left whole. We divided the upper and lower stem subsamples at the midpoint of the stem. We cut both the stem and root samples into 1 cm long sections. To surface sterilize the tissue, we used a series of immersions in separate plastic Petri dishes (100 mm x 15 mm) starting with 70% ethanol for 1 minute, 10% NaOCl for 3 minutes, 70% ethanol for 30 seconds, and a final 30 second rinse in autoclaved distilled water (Irizarry 2010). We surface sterilized additional root tissue using the method outlined by Danielson (1984). For this method, we quickly immersed the roots in 95% ethanol, followed by 30% H2O2 for 15 seconds, with a final rinse in autoclaved distilled water.

We used two sterilization methods for the roots to help recover a greater diversity of culturable

fungi. Following the two sterilization procedures, a subset of each tissue type was plated on the

appropriate media, left for 5 minutes, and then moved to a new plate to test for contamination.

The test plates were observed for contamination to indicate the efficacy of the sterilization

protocols.

Isolating endophytic fungi

We plated the sterilized leaf, stem, and root subsamples on separate Petri dishes (100 mm

x 15 mm) containing 2% MEA media. The root material that we subjected to the sterilization

method by Danielson (1984) was plated on two different media types, 2% MEA and a modified version of Melin-Norkran’s (MMN) agar,Draft in 100 mm x 15 mm Petri dishes to increase the likelihood of culturing a greater diversity of fungi (Marx 1969). We modified the MMN agar by

replacing the sucrose with dextrose and adding 100 ppm streptomycin, 50 ppm of

chlortetracycline, and 5 ppm benomyl (Danielson 1984). MMN agar favours the growth of

basidiomycetes as many of these fungi are tolerant to benomyl, an antifungal compound, and

streptomycin prevents the overgrowth of bacteria.

We sealed all plates with parafilm before placing them in the dark at room temperature to

grow. We made weekly observations to monitor fungal emergence. Following the appearance of

growth, we transferred emergent fungi to new 60 mm x 15 mm Petri plates with the

corresponding media to isolate individual fungal cultures or we held the plates at 4C to slow

fungal growth until we could complete the transfers. We used a dissecting microscope to observe

the hyphae for verification of pure cultures based on the uniformity of characters. This was

repeated until we obtained axenic cultures. Based on the colony morphology and apparent

growth rate, we grouped the isolates from each tissue type into morphotypes. To reduce the

number of downstream replicates, we selected a representative isolate for each morphotype and

collected separate portions of the axenic culture for cryopreservation and DNA extraction. For

morphotypes found in more than one tissue type, we selected one isolate from each tissue for

further analysis. For cryopreservation, we preserved mycelium in sterile 10% glycerol at -80C

and sterile distilled water at 4C.

DNA extractions and amplification from cultures

We used a Qiagen DNeasy® UltraClean® Microbial kit (Carlsbad, California) for DNA extraction from cultures using the protocol provided with the kit. We amplified the DNA extracts with the Biometra TGradient Thermocycler (Biometra, Jena, Germany) using the following thermocycling protocol: (1) 95°C for 3 minutes;Draft (2) 95°C for 1 minute; (3) 56°C for 45 seconds; (4) 72°C for 90 seconds; (5) 72°C for 10 mins; and (6) Hold at 4°C. We repeated steps (2) to (4) for 34 cycles. Each reaction contained 12.5 L Bio-Rad Master Mix for PCR (2x) (Bio-Rad

Laboratories Inc., Mississauga, Ontario), 9.5 L molecular biology grade water, 1 L ITS4

primer (10 M), 1 L ITS5 primer (10 M), and 1 L template DNA. We targeted and amplified

the ITS barcoding region of fungal rDNA using the primer pair ITS4 and ITS5 which were M13-

tagged (White et al. 1990).

Identification of endophytic isolates

We sent the amplified DNA products from PCR to the Genome Quebec Innovation

Centre (McGill University, Montreal, QC, Canada) for Sanger sequencing. We used Geneious

11.1.4 (Kearse et al. 2012) to trim the low-quality regions on the ends of the resultant

chromatograms. To identify trimmed DNA sequences, we used the Basic Local Alignment

Search Tool (BLAST) (Altschul et al. 1990) to compare against known sequences in the UNITE

and the National Center for Biotechnology Information (NCBI) GenBank sequence databases.

We considered differences of less than 3% in two ITS sequences to be conspecific (Begerow et

al. 2010) and we assigned the identification that best matched each sequence.

DNA extractions and meta-amplicon sequencing

In addition to culture methods, we analyzed DNA extracted directly from the roots and

soil using meta-amplicon sequencing to further categorize the below-ground fungi, providing a

thorough analysis of the fungal diversity in the soil microbiome. We extracted DNA directly

from the roots and soil using a Qiagen DNeasy® PowerSoil® Kit (Qiagen Inc., Valencia, CA)

following the kit protocol. For each of the 18 soil samples, we did DNA extractions in triplicate, resulting in nine DNA extracts from theDraft soil per site. We did a single DNA extraction from the roots of each plant, resulting in three DNA extracts from the roots per site. For both root and soil

DNA extracts, we sent one pooled sample per site (a total of 12 samples) on ice for ITS2 meta-

amplicon barcoding to the Integrated Microbiome Resource Centre (IMR), Dalhousie University.

At IMR, the samples were PCR-amplified using the ITS4 and ITS86F primers in duplicate using

a 1:1 and 1:10 dilution of template DNA (Beeck et al. 2014). The PCR products for each sample

were pooled and run on the Illumina MiSeq™ (Illumina Inc., San Diego, California) for ITS2

meta-amplicon barcoding.

Bioinformatic processing and community analysis

Bioinformatic analyses were based on a workflow and code provided by

MicrobiomeHelper (Comeau et al. 2017) which is publicly available on GitHub

(https://github.com/LangilleLab/microbiome_helper/wiki). We have outlined the specific steps

taken for our study. First, FastQC version 0.11.8 was run to inspect read quality (Andrews 2010).

We removed reads that did not begin with primers and trimmed primer sequences using cutadapt

version 1.1. (Martin 2011). All subsequent steps were performed using QIIME2 version 2018.8

(Bolyen et al. 2018). Briefly, we stitched together forward and reverse reads using DADA2 and output amplicon sequence variants (ASVs), removing ASVs with a frequency of less than 13, which was 0.1% of mean sample depth (13 440) (Callahan et al. 2016). We used MAFFT to make a de novo multiple-sequence alignment of the ASVs (Katoh and Standley 2013) and created a phylogenetic tree using FastTree (Price et al. 2010). Next, we used the UNITE ITS database and a Naive-Bayes classifier to perform taxonomic classification (Nilsson et al. 2018).

Any ASVs classified as mitochondria or chloroplast were removed and only ASVs classified as belonging to the kingdom fungi were retained. We set a minimum sampling depth of 4 000 to exclude samples with a lower read number and all samples satisfied the criteria. Lastly, we used the command “diversity core-metrics-phylogenetic”Draft to generate a variety of alpha and beta diversity metrics including observed ASVs and UniFrac unweighted and weighted PCoA to be used in subsequent data visualizations.

Evaluation of arbuscular mycorrhizal colonization of roots

To quantify AMF colonization, we stained C. canadense roots following a modified version of the protocol described by Vierheilig et al. (1998). We cleaned the fine roots before cutting them into 5 cm long segments. Next, we boiled the root segments in 10% KOH to clear the tissue of pigmentation. We then stained the tissue with a 5% Shaeffer® Skrip black ink- vinegar solution (Sered, Slovak Republic). Finally, we partially decoloured the root sections in a

5% acetic acid soak. This process selectively stained the chitinous cell walls of fungi leaving dark, fungal structures apparent in the cleared plant tissue.

We drew transects 5 mm apart on the reverse of glass microscope slides using black fine tip Sharpie permanent markers to indicate points of measurement. We then positioned the

stained roots across the lines on the marked microscope slides with 10% glycerol and a glass

cover slip for observation at 400x under a Nikon Alphaphot-2 YS2 compound microscope

(Tokyo, Japan). One researcher analyzed each intersection for the presence of AMF until a count

of 50 intersections was reached. AMF structures occurring at each intersection out of 50 were

counted to calculate the colonization rate of the root tissue. We preserved voucher microscope

slides showing fungal structures associated with C. canadense roots and deposited them in the

E.C. Smith Herbarium, Acadia University, Wolfville, NS, Canada under accession numbers

ACAD19927F to ACAD19932F.

Data and statistical analysis All of our plots were made usingDraft the ggplot2 R package (Wickham 2016) in R to graphically display our results (R Core Team 2020). We customized our colour scheme for

Figure 3 using RColorBrewer version 1.1-2 (Neuwirth 2014). From our QIIME2 “diversity core-

metrics-phylogenetic” output, we created alpha and beta diversity visualizations (Wickham

2016; R Core Team 2019). We plotted ASV richness within each site and substrate to visualize

alpha diversity (Figure 4). To visualize beta diversity, we created UniFrac unweighted and

weighted PCoA plots to illustrate relative differences in the fungal communities between

samples (Figure 5B). To compare AMF colonization rates of the roots between sites, we

conducted a Kruskal-Wallis test in R (α = 0.05) (R Core Team 2020).

Results

Soil characteristics

There was little variation in soil chemistry between sites with the pH ranging from 4.55 –

5.20, cation exchange capacity ranging from 1.9 – 4.0 meq/100 g, and organic matter ranging

from 1.9 – 2.5%. The soil nutrients were fairly uniform across sites with the exception of site 4

which had higher amounts of calcium, magnesium, and phosphorous when compared to other sites1. We did not find a significant correlation between these soil characteristics and AMF colonization in the roots of C. canadense plants. A larger sample size, more controlled variables, and long-term observations would be needed to better model the AMF community in C. canadense roots.

Identification of cultured fungi

We isolated a total of 397 fungal isolates from the tissues of 18 C. canadense plants collected across six sites at CFB Greenwood with 159 isolates from the roots, 113 from the leaves, 81 from lower stems, and 44 from upper stems. This resulted in 227 fungal morphotypes with 94 isolated from the roots, 56 fromDraft the leaves, 50 from lower stems, and 27 from the upper stems. Further analysis was conducted for all 94 morphotypes isolated from the roots along with a subset of 15 morphotypes isolated from leaves, 12 from lower stems, and 8 from upper stems.

Fungi may take part in a variety of roles in plant roots, including endophytic, ectomycorrhizal, or ectendomycorrhizal partnerships; the diversity of relationships and potential for beneficial symbioses led us to prioritize the DNA barcoding of root-associated fungi. We targeted the selection of fungi isolated from the above-ground tissue toward morphotypes occurring frequently and with a wide distribution throughout sites. These factors were considered valuable for future research on the ability of these fungi to form symbioses with this host and potentially aid in its conservation.

We extracted DNA from each of the 129 chosen morphotypes. We successfully obtained

DNA barcode sequences for 109 of the selected morphotypes. DNA extracted from the

1Results of the soil analyses can be found in the supplementary Table S1.

remaining 20 morphotypes, all of which were from the roots, was not successfully amplified or

did not yield a successful DNA barcode following Sanger sequencing. Our BLAST results

returned 58 unique fungi with 50 being matched to species with less than 3% sequence

divergence2. Successfully identified fungi belonged to four phyla: Ascomycota, Basidiomycota,

Zygomycota, and Mucoromycota.

Although we obtained positive species identifications, several of the identified species are

part of a species complex or are poorly resolved from closely related species when exclusively

comparing the ITS barcode region of fungal rDNA. This includes members of the genera

Fusarium (van Diepeningen et al. 2015), Phialocephala (Tanney et al. 2016), Trichoderma

(Lübeck et al. 2004; Samuels et al. 2006), and Tubakia (Braun et al. 2018). Identification of

isolates from these genera therefore haveDraft a significant resemblance to more than one closely

related species. Additionally, two of our DNA barcodes returned matches to sequences with less

than 97% identity between sequences. In both cases of uncertainty, isolates were assigned the

species identification designated by the best match in NCBI BLAST or UNITE with ‘cf.’ to

indicate uncertainty in our identifications. DNA barcodes that returned less than 97% identity

between sequences were closely related to and will be discussed as Apiognomonia cf. hystrix

(96% sequence similarity) and Lophodermium cf. pini-excelsae (95%).

Of the fungal isolates we sequenced, 10 classes were represented with the majority of

species belonging to Sordariomycetes. We isolated and identified a total of 42 species from the

roots, 14 from leaves, 6 from upper stems, and 11 from lower stems (Figure 3). Only two species

were common to all 6 sites, A. cf. hystrix and Pyrenophora tetrarrhenae. We found A. cf. hystrix

at least once in every tissue (Figure 3) and we isolated it from 7 of the 18 plants collected.

2 Barcode rDNA sequences and identifications can be found in the supplementary Table S2.

Pyrenophora tetrarrhenae was common to all above-ground tissue types and we isolated it from

16 of the 18 plants collected making it the most commonly isolated fungal associate of C. canadense. Most species were only isolated from one tissue type, with the exception of Arcopilus aureus, A. cf. hystrix, Cryptodiaporthe aubertii, Fusarium cf. sporotrichioides, Nemania serpens, Pilidium concavum, Phialocephala cf. fortinii, and P. tetrarrhenae. The fungi most commonly isolated from C. canadense plants included A. aureus colonizing 56% of the plants, A. cf. hystrix colonizing 39% of the plants, P. concavum colonizing 50% of the plants, P. cf. fortinii colonizing 44% of the plants, P. tetrarrhenae colonizing 89% of the plants, and

Sphaeronaemella fragariae colonizing 39% of the plants. These six fungi were the most broadly distributed, occurring in more than half of the sites, and the most common, colonizing more than one third of the plants sampled (Figure 3).Draft S. fragariae was the only one that we found in only one tissue type, the leaves.

Fungal community analysis using meta-amplicon barcoding

We determined the taxonomic diversity comprising the fungal community in the roots and soil at each of the six sites using ITS2 meta-amplicon barcoding. This is the first account of the fungal community composition in the roots and sandy rhizosphere of C. canadense. After filtering the meta-amplicon sequence results to remove low-quality reads and rare ASVs, we obtained a total of 158 095 reads with an average of 13 175 reads per sample. Overall, we found a diverse fungal assemblage associated with C. canadense in its native sand barren habitat with

241 fungi identified from the roots and rhizosphere. The taxonomic richness was consistently higher in the sand compared to the roots. Site 3 had the greatest richness in the sand and roots with 201 and 145 ASVs, respectively (Figure 4).

Our meta-amplicon sequencing of the roots and rhizosphere of C. canadense revealed

seven phyla comprising the fungal assemblage. Most of the reads belonged to Basidiomycota

(53.8%), Ascomycota (33.2%), and Mortierellomycota (5.3%), with the remaining reads

belonging to Glomeromycota, Mucoromycota, Zoopagomycota, and an unidentified phylum.

Overall, we identified 25 classes in the roots and sand of C. canadense. In the roots, the most

common fungal classes were Agaricomycetes (53.6%), Leotiomycetes (12.9%), and

Eurotiomycetes (12.8%). In the sand, the most common fungal classes were Agaricomycetes

(48.5%), Ascomycota sp. (9.5%), Eurotiomycetes (9.0%), Umbelopsidomycetes (7.4%),

Mortierellomycetes (7.2%), and Leotiomycetes (5.3%). In both the roots and sand,

Agaricomycetes was the dominant class encompassing 10 orders in the roots and 12 in the sand.

We illustrated a subset of the fungalDraft community structure associated with C. canadense

using the ITS2 meta-amplicon barcoding results (Figure 5). After we classified the ASVs to the

lowest taxonomic level identifiable from the UNITE ITS database, we visualized the top 15 most

abundant taxa, representing 62.1% of the total fungal assemblage (Figure 5). Of these abundant

taxa, 10 were from Basidiomycota, three from Ascomycota, one from Mucoromycota, and one

from an unclassified phylum. The taxon with the highest average relative abundance (RA) was

Chaetothyriales sp. with 12.1% in the roots and 7.4% in the sand. Followed by Chaetothyriales

sp., the most abundant root-associates were Mycetinis scorodonius (11.9%), Acidomelania

panicicola (9.6%), and Scleroderma citrinum (9.1%).

In some cases, the most abundant species in the roots were less dominant in the sand and

species that were abundant in the sand were less dominant in the roots (Figure 5). Clavaria

argillacea was one of the most abundant species identified with an average RA of 9.5% in the

sand and only 0.6% in the roots. The average RA of Scleroderma citrinum was 9.1% in the roots

and only 0.2% in the sand. We observed similar trends for many other prolific taxa.

Acidomelania panicicola and Mycetinis scorodonius were more abundant occupants of the roots while Ascomycota sp. and Umbelopsis sp. favoured the sand.

Using the ITS2 amplicon sequences, we performed UniFrac unweighted and weighted principal coordinate analysis (PCoA) to visualize the relative differences in the fungal communities (beta diversity). Distances between points on the PCoA plots represent relative differences in the fungal community found at each site and substrate. The unweighted UniFrac

PCoA provides a measure of community dissimilarity based on presence/absence, without regard for the frequency of observed ASVs. The weighted UniFrac PCoA considers the frequency of observed ASVs. Differences in the root and soil communities are more pronounced in the unweighted PCoA indicating that a highDraft diversity of low-frequency or rare species were influencing the composition of the fungal communities (Figure 5B). Differences between the substrates become less apparent in the weighted PCoA which instead shows a significant difference in site 2 compared to the other sites. This quantity-dependent trend is largely due to the high-frequency of two species, Clavaria argillacea and Scleroderma citrinum, at site 2 when compared to the other sites (Figure 5A).

AMF identification and quantification using meta-amplicon barcoding

Our meta-amplicon barcoding of the fungal community in the roots and rhizosphere provided relative abundances and taxonomic identifications of AMF which helped us better describe the relationship between AMF and C. canadense. We found the highest RA at site 5 with AMF accounting for 1.9% of the fungal assemblage in the sand and 0.2% in the roots

(Figure 6). The greatest RA of AMF in roots was found at site 4 with 0.7% of the assemblage identified as AMF taxa. It is important to note that this approach measured RAs of AMF rather

than total abundances; therefore, low RAs do not necessarily mean there is insignificant AMF

present, but rather there is little AMF compared to the all the other fungal taxa in the assemblage.

We identified two genera within the phylum Glomeromycota, Rhizophagus and Archaeospora.

Archaeospora sp. was only identified in the sand while Archaeosporales sp., Glomeraceae sp.,

and Rhizophagus sp. were found in the roots. We did not identify Glomeromycota in the roots at

sites 1, 2, or 6 (Figure 6). Our identifications represent novel reports of AMF associating with C.

canadense.

Measuring AMF colonization of roots using microscopic analysis

We used compound microscopy to examine the ink-stained C. canadense roots to measure root AMF colonization rates. WeDraft observed both AMF and dark septate endophytes in the roots of C. canadense. We designated AMF based on the criteria of being ink stained, non-

septate hyphal structures that penetrated within cortical cells of roots and in some cases formed

storage vesicles or arbuscules (Figure 7).

We observed the highest AMF colonization rates at site 3 with an average colonization

rate of 53% (Figure 7C). The highest colonization rate in the roots of an individual plant was

found at site 3 with 80% colonization of C. canadense roots by AMF. Site 3 also had the largest

variability in colonization rates (18-80%). Site 2 had the lowest colonization rates (average

3.3%). The lowest observed colonization rate in the roots of an individual plant was at site 2

(2%). Overall, the 18 plants had an average AMF colonization rate of 29.6%. Following a

Kruskal-Wallis test to analyze spatial trends in the AMF colonization rates, we did not find

evidence that there were statistically significant differences in AMF colonization rates between

sites (P = 0.06486, H(5) = 10.392) at α = 0.05.

Discussion

We found C. canadense to be colonized by a diverse community of fungal symbionts. We isolated and identified 58 species of fungi from above- and below-ground tissues, discovered 241 fungi in the roots and rhizosphere using metabarcoding, and established the presence of AMF associated with this critically imperiled plant.

Common fungal isolates and their distribution in plant tissues

By investigating plant parts independently, our research revealed information about the limitations and selectivity of the endophytic fungi colonizing this underexplored plant and habitat. Pyrenophora tetrarrhenae was the most abundant cultured species among the fungal isolates from C. canadense. We isolatedDraft this fungus from 16 of the 18 C. canadense plants collected from CFB Greenwood. In our study, P. tetrarrhenae was exclusively isolated from the above-ground tissue (leaves, lower stems, and upper stems). Furthermore, we did not find P. tetrarrhenae in the roots using metabarcoding. We isolated Arcopilus aureus from five of the six sites and from all of the tissue types, indicating low specificity with respect to plant tissue and almost ubiquitous colonization of C. canadense. Phialocephala cf. fortinii demonstrated a preference for the roots and lower stems but remained absent from the leaves and upper stems.

Six fungi (Pyrenophora tetrarrhenae, Arcopilus aureus, Pilidium concavum,

Phialocephala cf. fortinii, Apiognomonia cf. hystrix, and Sphaeronaemella fragariae) were frequently isolated from C. canadense. All six of these fungi were ascomycetes. Given their widespread colonization and the abundance of isolates recovered from the plant population at

CFB Greenwood, these fungi were thought to have significant relationships with C. canadense.

Although common within the plant population, the ecological roles performed by these fungi remain unclear and may be more generalist as some of these species are typical isolates of

endophytic screenings from various plants in many substrates and environments (Table 1) (Paul

1971; Grünig et al. 2008; Ariyawansa et al. 2014; González 2015; Dwibedi and Saxena 2018;

Egidi et al. 2019).

Fungal community structure in the roots and soil

Our metabarcoding results showed that the fungal assemblages associated with the roots

were less diverse, as assessed using total ASVs, than the soil (Figure 4). Additionally, the

compositions of these two communities were somewhat distinct. When not considering the

frequency of the species, as in the unweighted UniFrac PCoA, the pattern suggests that there are

distinct fungal assemblages in the roots and sand associated with C. canadense (Figure 5B).

The occurrence of different fungal community structures and fewer taxa in the roots

compared to the soil is common among Draftstudies analyzing fungal associates of plants with high-

throughput sequencing (Xu et al. 2012; Nallanchakravarthula et al. 2014). The richer fungal

community in the soil compared to the roots is predictable due to the selective recruitment of

root-associated fungi from the soil environment. Furthermore, the limited taxa capable of

associating with roots combined with the selectivity of the host plant itself results in fewer and

distinct taxa inhabiting the roots (Goldmann et al. 2016). Our results suggested a preference

toward C. canadense forming root-associated relationships with Agaricomycetes

(Basidiomycota), Leotiomycetes (Ascomycota), and Eurotiomycetes (Ascomycota).

Our use of 70% ethanol to sterilize sampling tools during soil collection may not have

entirely removed residual free DNA that could result in cross-contamination among our soil

samples. In the future, the use of a stronger sterilant like bleach or DNA AWAY would be

preferred to eliminate any possibility of contamination. Given our focus was on the most

abundant fungi that we discovered, this would likely not affect our results significantly; however,

there may be minimal effects on our beta diversity analyses causing increased similarity between the fungal assemblages in the soil.

Ecological roles of frequently isolated and abundant fungi

We investigated the potential ecological roles of the six most frequently isolated fungi from our culture-based methods and the most abundant fungi detected using meta-amplicon sequencing to help understand the potential fungal relationships occurring with the Nova Scotian

C. canadense population (Table 1). The fungal assemblage at site 2 was unique as it was heavily dominated by two fungi, Clavaria argillacea in the sand (55.0%) and Scleroderma citrinum in the roots (53.9%) (Figure 5). Clavaria argillacea is frequently cited as an associate of Ericaceae plants and is commonly found on acidicDraft and sandy substrates (Olariaga et al. 2015); both features are found at CFB Greenwood. This fungus was also recovered from the roots of C. canadense

(3.2% RA at site 2), possibly occupying an endophytic role in this non-ericaceous plant host.

Unlike the other sites, the ground cover near the plants sampled at site 2 was heavily colonized by broom crowberry (Corema conradii, Ericaceae). The presence of this plant near C. canadense could contribute to the high RA of C. argillacea in the soil. The ground cover at other sites was more diverse with grasses, ferns, asters, blackberry, and trees in proximity to the sampled plants, which could increase the diversity of the belowground fungal assemblage.

Although sites 1 and 2 were closely situated, the two had distinct fungal assemblages; this may be due to vegetation differences between the two sites.

Scleroderma citrinum was the dominant fungus in the roots at site 2. Members of the genus Scleroderma form ectomycorrhizal relationships with a broad range of plants (Jeffries

1999; Comandini et al. 2006; Leonardi et al. 2018), including several members of Cistaceae

(Cistus monspeliensis (Demoulin 1983); Halimium halimifolium (Leonardi et al. 2018);

Hudsonia ericoides; and Hudsonia tomentosa (Malloch and Thorn 1985)). It is unclear how the

overwhelming abundance of C. argillacea and S. citrinum may affect C. canadense at site 2.

More information is needed to determine if these two fungi are aiding or harming the plants

either directly or by outcompeting other fungal partners such as AMF.

Acidomelania panicicola was another abundant root associate of C. canadense. It was

most abundant at sites 4 and 5, accounting for 28.3% and 17.8% of the fungal assemblage in the

roots at these sites respectively (Figure 5A). This globally distributed fungus colonizes plant

roots from acidic and infertile soils, including the comparable New Jersey Pine Barrens, and was

shown to promote root hair growth in switchgrass (Walsh et al. 2014, 2015). Mycetinis

scorodonius was also abundant in the roots of C. canadense, but the nature of this relationship is

likely saprotrophic rather than mycorrhizalDraft or endophytic (Petersen and Hughes 2017).

Fungal relationships with Cistaceae and barren habitats

There was substantial overlap in the fungal assemblage we isolated from C. canadense

and those isolated from plants in a similar pine barren habitat in New Jersey, United States (Luo

et al. 2017). Shared fungal symbionts included members of Arcopilus, Curvularia, Fusarium,

Lachnum, Mortierella, Mycena, Penicillium, Phialocephala, Trichoderma, and Umbelopsis.

With our metabarcoding results, eight additional genera were found in common between the two

habitats, including Acidomelania, Ceratobasidium, Chaetomium, Cenococcum, Metarhizium,

Mollisia, Pseudophialophora, and Talaromyces. Crocanthemum canadense is also an inhabitant

of the New Jersey pine barrens where the habitat closely resembles that of the sand barrens at

CFB Greenwood (Wittenberg 2015). Acidic, sandy, and nutrient poor soils with an understory

dominated by grasses, blueberries, and other Ericaceae plants are key features of both habitats.

These members of the fungal community could be common to barren habitats and may be important for maintaining the health of these ecosystems and their native plant populations.

The investigations of the mycorrhizal fungi associated with Cistaceae in North America by Malloch and Thorn (1985) revealed 13 species from the genera Amanita, Astraeus,

Cenococcum, Collybia, Inocybe, Laccaria, Leccinum, Marasmiellus, Russula, Scleroderma, and

Thelephora. According to these authors, these genera were likely forming ectomycorrhizae with

Cistaceae. Crocanthemum canadense, previously referred to by the name Helianthemum canadense, was sampled from Ontario and had an unspecified ectomycorrhizal association

(Malloch and Thorn 1985). In the roots, we found six of the genera previously described by

Malloch and Thorn (1985), including Amanita, Cenococcum, Inocybe, Leccinum, Russula, and

Scleroderma. Our findings of these potentialDraft symbionts in the roots of C. canadense further support their conclusion that these genera form symbiotic relationships with members of

Cistaceae. None of these potential symbionts were isolated in culture, which emphasizes the need for combined methodologies to better understand plant-fungal relationships.

Given our molecular findings of known ectomycorrhizal fungi in the root tissue and our microscopic findings of AMF colonizing C. canadense roots, it is possible that C. canadense is a dual-mycorrhizal host capable of forming partnerships with both ectomycorrhizal fungi and

AMF. Further investigations would be needed to confirm the presence of true ectomycorrhizal structures within the roots of C. canadense. Teste et al. (2020) confirmed dual-mycorrhizal status for three other species in the Cistaceae, including Fumana procumbens, H. chamaecistus, and H. ovatum. Smith and Read (1997) suggested Cistus were also dual-mycorrhizal hosts with mycorrhizal plasticity likely aiding in the re-establishment of plants following fire disturbance.

Flexibility with mycorrhizal partners may be an important adaptation for members of the

Cistaceae to survive in disturbed habitats.

AMF colonization of roots

We documented AMF colonization of C. canadense roots for the first time with average

colonization rates reaching 53% at site 3. High rates of observed AMF colonization in sites 3-6

indicate an important relationship that could be influencing the C. canadense population at CFB

Greenwood. The results of our AMF quantification using microscopic and metabarcoding

analyses were in agreement with the exception of site 6. We determined site 6 to have a

relatively high AMF colonization rate based on microscopic observations, but our meta-amplicon

barcoding revealed an absence of AMF in the roots at site 6. Sites 1 and 2 also lacked AMF

barcodes in the roots which was consistentDraft with our observations of low AMF colonization rates

at these sites. The limited or absent AMF at sites 1, 2, and 6 compared to substantial colonization

at other sites demonstrates that there are variable interactions with AMF throughout the

population. We presume that reduced AMF colonization of the roots could have implications for

the success of C. canadense plants since AMF are known to improve the growth and

reestablishment of plants (Sylvia et al. 1993; Morton et al. 2004).

Given our microscopic observations of AMF colonizing C. canadense roots at sites 1, 2,

and 6, it is possible that a poor root sample was used for the DNA extraction. In most cases, fine

roots were chosen for both the microscopic and molecular examination of AMF; however, in

some cases there was limited fine root material available and the microscopic analysis was

prioritized. The use of thicker root samples could have influenced the molecular identification of

AMF in the roots at some sites. It is also important to note that given our limited root tissue, we

only counted 50 root-line intersections for each sample while it is suggested in the literature that

a minimum of 100 counts are necessary to improve the accuracy of colonization rates (Sun and

Tang 2012). Although we may not have employed the most rigorous methodology cited in the literature, our results still represent valuable information regarding the fungal symbionts of this endangered plant that has scarcely been researched prior to our work.

Both of the identified Glomeromycota genera, Rhizophagus and Archaeospora, are known to form arbuscular mycorrhizae with vascular plants (Li et al. 2014; Le Pioufle et al.

2019; Baltruschat et al. 2019). Species of Rhizophagus are found to increase drought tolerance and nutrient uptake in plants (Li et al. 2014; Le Pioufle et al. 2019). Given the typical growing conditions of the Cistaceae, i.e. dry, acidic, and low-nutrient soil, there is significant potential for

AMF to aid in the survival and success of these plants in seemingly unfavourable habitats. Future investigations are necessary to gain a betterDraft understanding of the function and ecological importance of AMF associated with C. canadense.

Method comparison

Root endophyte surveys using culture-based methods are common and often result in a fungal assemblage dominated by ascomycetes, including our current study (Hoff et al. 2004; Luo et al. 2017). Our metabarcoding results were in opposition to this trend where basidiomycetes were the predominant phylum represented. Mundra et al. (2015) found similar results when using metabarcoding to investigate the root endophytes of the herbaceous, perennial, and ectomycorrhizal plant, Bistorta vivipara (Polygonaceae). Basidiomycota accounted for more than two thirds of the reads (69%) identified in the roots of B. vivipara (Mundra et al. 2015).

This is likely because culture-dependent methods often favour fast-growing, dominant fungi while slow-growing or unculturable fungi are overlooked (Jeewon and Hyde 2007).

Culture-independent methods can capture a broader array of fungi, and thus reveal a

different composition of the fungal community (Dissanayake et al. 2018). From the roots, 29

fungal genera were identified using culture methods and 138 genera using meta-amplicon

barcoding, with 18 genera in common between the two methodologies. In culture, we isolated

only two fungi (Mycetinis scorodonius and Acidomelania panicicola) that were also highly

abundant in our meta-amplicon barcoding of the roots and rhizosphere soil.

DNA barcoding of fungal isolates allowed us to identify almost all fungi to species and

allowed for preservation of cultures for future research but did not provide a full representation

of the fungal community. Meta-amplicon barcoding from environmental samples is often unable

to reach the same level of specificity in taxonomic identifications and living specimens are not

available for future inoculation studies; Drafthowever, it does expand the coverage of the fungal

assemblage. As demonstrated here, a combination of methods is necessary to document

endophytic fungal diversity, especially when future study or use of ecologically significant fungi

is desirable, such as in native plant propagation and restoration.

Conclusion

To our knowledge, this is the first documentation of fungal symbionts associated with C.

canadense. Our results illustrate a baseline composition of the fungi associated with C.

canadense and the sand barren heathlands located at CFB Greenwood, Nova Scotia. These data

offer a starting point to help understand plant-fungus partnerships with this threatened plant in its

declining habitat.

From our meta-amplicon data, we discovered a fungal assemblage (mainly A. panicicola,

C. argillacea, Rhizophagus sp., and S. citrinum) typically associated with plants from dry,

infertile, and acidic soils. These fungi may aid in the adaptation of plants to these poor habitats.

Future studies are necessary to realize the ecological function and importance of these fungal symbionts with C. canadense. Our research also provided the first documentation of AMF colonizing the roots of C. canadense with average root colonization rates as high as 53%. AMF can significantly improve plant growth and reestablishment. Meta-amplicon barcoding indicated the presence of at least two genera of AMF, Rhizophagus and Archaeospora. Inoculating C. canadense with indigenous species of these beneficial fungi could enhance the propagation of this critically imperiled plant.

Our results helped improve our understanding of the fungal symbioses with

Crocanthemum canadense by identifying 58 species of culturable fungi, 241 fungi using metabarcoding, and documenting root colonization by AMF. Additionally, we present new fungal records from habitats outside of theirDraft documented distribution, hosted by a critically imperiled plant in a unique sand barren heathland. Our findings are currently integrated into research investigating endangered native plant-fungal partnerships, including improved propagation strategies to conserve this critically imperiled plant in Nova Scotia.

Acknowledgements

The authors thank S. Adams, A. Bunbury-Blanchette, R. Browne, T. d'Entremont, D.

Divanli, H. Machat, B. Robicheau, and V. Taylor for assistance with this project. We acknowledge the K.C. Irving Environmental Science research facilities at Acadia University, a

NS Department of Natural Resources scientific collection permit to RCE, a clearance to dig permit from 14 Wing CFB Greenwood, the Integrated Microbiome Resource Centre (IMR) at

Dalhousie University, and Genome Québec Innovation Centre (McGill University). The culture- based and microscopic analyses included in this research were completed as part of the lead author’s undergraduate thesis submitted to Acadia University. This research was funded by an

NSERC Undergraduate Student Research Award to PRMB (No. USRA—526403-2018), the

Acadia Co-op Program, the Acadia University FPAS Colville Award in Science to PRMB, the

Nova Scotia Cooperative Education Incentive subsidy, and the Arthur Irving Academy for the

environment research grant awarded to RCE, Kirk Hillier, and AKW. ZM was supported by the

National Science Foundation Plant Genome Research Program 1546869. AKW gratefully

acknowledges funding from an NSERC Discovery Grant (No. NSERC—2017-04325).

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Table 1. Known ecological roles of abundant fungal taxa identified using culture-dependent and culture-independent methods. DSE = dark septate endophyte, ECM = ectomycorrhizal, En = endophyte, ERM = ericoid mycorrhizal, Para = parasite, Path = pathogen, Sap = saprotroph. Fungi Ecological role Reference Culture-dependent Arcopilus aureus En (Dwibedi and Saxena 2018) Apiognomonia cf. hystrix En/path/sap (Sieber et al. 1990; Sogonov et al. 2008) Pilidium concavum Path/sap (Lopes et al. 2010; González 2015) Phialocephala cf. fortinii DSE/path (Sheridan 1977; Grünig et al. 2008) Pyrenophora tetrarrhenae En/path/sap (Paul 1971; Ariyawansa et al. 2014) Sphaeronaemella fragariae Sap (Senanayake et al. 2017) Culture-independent Acidomelania panicicola DSE (Walsh et al. 2014) Clavaria argillacea ERM/sap (Olariaga et al. 2015) Mycena sp. En/para/sap (Tedersoo and Smith 2013) Mycetinis scorodonius Sap (Petersen and Hughes 2017) Russula crustosa ECM (Hou et al. 2012) Scleroderma citrinum ECM/sap (Jeffries 1999; Leonardi et al. 2018) Sistotrema sp. ECM (Nilsson et al. 2006) Tomentella lapida ECM Draft(Dickie et al. 2004) Umbelopsis sp. ECM/En/Sap (Hoff et al. 2004; Summerbell 2005; Kowalik 2013) Note: Only fungi classified to genus or lower taxonomic ranking were included.

Figure Captions

Figure 1. Habitat and morphology of Crocanthemum canadense. (A) Open flower (modified from Evans, 2017). (B) Dehiscing fruit with seeds. (C) Site 2 at CFB Greenwood. (D) Site 6 at CFB Greenwood. A = Anther; C = Petal; K = Sepal; S = Seeds. Scale bars = 1 cm.

Figure 2. Provincial and state conservation status for Crocanthemum canadense according to NatureServe (2019). Figure was created using MapChart (Minas 2020).

Figure 3. Species isolated from surface sterilized leaves, upper stems, lower stems, and roots of Crocanthemum canadense plants across six sites at CFB Greenwood. Colour indicates taxonomic class. Number of plant-hosts out of a possible three per site is indicated by circle size.

Figure 4. Observed fungal ITS2 ASV richness in the sand and root mycobiomes of Crocanthemum canadense collected from six sites at CFB Greenwood, Nova Scotia in July 2018.

Figure 5. Fungal community composition in the roots and sand from sites 1-6 at CFB Greenwood based on ITS2 rDNA meta-amplicon barcoding. (A) Relative abundance of the dominant taxa comprising the fungal communities. (B) Qualitative and quantitative comparisons of the fungal community compositions inDraft the roots and sand across six sites using UniFrac unweighted and weighted PCoA plots, respectively. Shape represents substrate and colour indicates the collection site.

Figure 6. Relative abundances of Glomeromycota (AMF) in the sand and root mycobiomes of Crocanthemum canadense from six sites at CFB Greenwood as revealed by ITS2 rDNA meta- amplicon barcoding.

Figure 7. Crocanthemum canadense roots and colonization by AMF. (A) Typical Crocanthemum canadense root system. Scale bar = 3.5 cm. (B) Crocanthemum canadense roots showing selectively stained fungal structures using Shaeffer Skrip black pen ink under 400x magnification. Ar = Arbuscule; H = Hyphae; V = Vesicle. (C) AMF colonization rates of Crocanthemum canadense roots from six sites at CFB Greenwood (N = 3).

Draft

© The Author(s) or their Institution(s) Leaf Lower StemBotany Root Upper Stem Page 42 of 46 Perenniporia medulla−panis Class Mycetinis scorodonius Mycena citrinomarginata Agaricomycetes Mycena abramsii Dothideomycetes Marasmiellus tricolor Ceratobasidiaceae sp. 1 Eurotiomycetes Pyrenophora tetrarrhenae Leotiomycetes Pyrenophora biseptata Microbotryomycetes Drechslera sp. 3 Drechslera sp. 2 Mortierellomycetes Drechslera sp. 1 Mucoromycetes Curvularia protuberata Alternaria alternata Sordariomycetes Pleotrichocladium opacum Tremellomycetes Melanommataceae sp. 1 Zygomycetes Stagonospora bicolor Epicoccum nigrum Pseudoseptoria obscura Number Penicillium soppii 1 Penicillium simplicissimum Penicillium melinii 2 Lophodermium cf. pini−excelsae 3 Halenospora varia Phialocephala sp. 1 Phialocephala cf. fortinii Botrytis fabiopsis Lachnum virgineum Draft Godronia cassandrae

Species Acidomelania panicicola Pilidium concavum Sporobolomyces ruberrimus Mortierella alpina Mucor moelleri Cunninghamella elegans Nemania serpens Chaetomium umbonatum Arcopilus aureus Sphaeronaemella fragariae Neonectria candida Ilyonectria leucospermi Fusarium cf. sporotrichioides Fusarium cf. poae Fusarium cf. oxysporum Fusarium cf. avenaceum Trichoderma cf. koningiopsis Trichoderma cf. koningii Trichoderma cf. hamatum Trichoderma cf. asperellum Clonostachys rosea Colletotrichum cereale Tubakia cf. suttoniana Tubakia cf. californica Cryptodiaporthe aubertii Apiognomonia cf. hystrix Apiotrichum porosum Umbelopsis ramanniana Umbelopsis nana Umbelopsis dimorpha 1 2 3 4 5 6 ©1 The2 Author(s)3 4 5 or6 their1 Institution(s)2 3 4 5 6 1 2 3 4 5 6 Site Page 43250 of 46 Botany

200 ● ● ● ● 150 ● Draft● ● ● ● 100 ● ● ● 50 Substrate ● Roots

Relative Abundance (%) 0 1 2 3 4 5 6 1 2 3 4 5 6

Roots Sand Site #

Species (listed to lowest taxonomic level available) Serendipitaceae (family) Draft Russula crustosa (species) Sistotrema (genus) Tomentella lapida (species) Agaricomycetes (class) Thelephoraceae (family) Unclassified Mycena (genus) Umbelopsis (genus) Scleroderma citrinum (species) Acidomelania panicicola (species) Clavaria argillacea (species)

Mycetinis scorodonius (species) Ascomycota (phylum) Chaetothyriales (order) Other B UniFrac unweighted PCoA UniFrac weighted PCoA ● ● ● ●

● ● ● ● PCoA 2 (15.8%) PCoA 2 (23.2%)

● ● ● ● PCoA 1 (22.3%) PCoA 1 (30.9%) © The Author(s) or their Institution(s) ● 1 ● 3 ● 5 Site # Substrate ● roots sand ● 2 ● 4 ● 6 250

200 ● ● ● ● 150 ● ● ● ● ● 100 ● ● Page 45 of 46 Botany ●2.0 50 Substrate ● Roots

Richness (Observed ASVs) ● Sand 0 1.5 1 2 3 4 5 6 Site # Draft substrate 1.0 ● roots ● sand

0.5 Glomeromycota Relative Abundance (%) Relative Abundance Glomeromycota

0.0

Draft