Diversity and host preference of fungi co-inhabiting Cenococcum mycorrhizae

f u n g a l e c o l o g y 1 7 ( 2 0 1 5 ) 8 4 e9 5

available at www.sciencedirect.com

ScienceDirect journal homepage: www.elsevier.com/locate/funeco

Diversity and host preference of fungi co-inhabiting Cenococcum mycorrhizae Gavin KERNAGHAN*, Glenn PATRIQUIN Biology Department, Mount St. Vincent University, 166 Bedford Hwy., Halifax, NS B3M 2J6, Canada

article info

abstract

Article history:

Diverse fungal assemblages colonize the fine feeder roots of woody plants, including

Received 1 December 2014

mycorrhizal fungi, fungal root endophytes and soil saprotrophs. The fungi co-inhabiting

Revision received 30 March 2015

Cenococcum geophilum ectomycorrhizae (ECM) of Abies balsamea, Betula papyrifera and Picea

Accepted 3 April 2015

glauca were studied at two boreal forest sites in Eastern Canada by direct PCR of ITS rDNA.

Available online xxx

50 non-Cenococcum fungal sequence types were detected, including several potentially

Corresponding editor:

mycorrhizal species as well as fungal root endophytes. Non-melanized ascomycetes

Kabir G Peay

dominated, in contrast to the dark septate endophytes (DSE) reported in most culture dependent studies. The results demonstrate significant differences in root associated

Keywords:

fungal assemblages among the host species studied. Fungal diversity was also host

Boreal forest

dependent, with P. glauca roots supporting a more diverse community than A. balsamea.

Dark septate endophytes

Differences in root associated fungal communities may well influence ecological inter-

Ectomycorrhizae

actions among host plant species.

Fungal communities

ª 2015 Elsevier Ltd and The British Mycological Society. All rights reserved.

Fungal diversity Helotiales Root endophytes

Introduction Ectomycorrhizae (ECM) consist of both fine root and fungal tissues, resulting in a unique and metabolically active habitat within the soil ecosystem. A healthy ECM can support a wide variety of soil organisms, including a diverse array of fungi other than the dominant ectomycorrhizal symbiont (Bergero et al., 2000; Kernaghan et al., 2003; Menkis et al., 2005; Bergemann and Garbelotto, 2006; Morris et al., 2008; Urban et al., 2008; Wright et al., 2009). These secondary fungi may
zares and be other mycorrhizal species within the root (Ca Trappe, 1993; Olsson et al., 2000; Morris et al., 2008; Toju et al., 2014), or saprobes on the surface and associated mycorrhizosphere (Foster and Marks, 1967; Fogel, 1988).

Ectomycorrhizae also harbor a wide range of asymptomatic fungal endophytes, which colonize the root tissue internally and appear to be as common as mycorrhizal fungi (Mandyam and Jumpponen, 2005; Weishampel and Bedford, 2006; Toju et al., 2013a). Fungal endophytes can be found colonizing all types of plant tissue (Rodriguez et al., 2009), but the species assemblages associated with roots appear distinct from those colonizing shoots and leaves (Addy et al., 2005; Summerbell, 2005). Although the ecological role of these fungi is unclear (Mandyam and Jumpponen, 2005; Mayerhofer et al., 2013), some may be latent pathogens (Schulz et al., 1999) or saprotrophs (Kernaghan, 2013). Others may provide protection from soil pathogens (Narisawa et al., 2004), or improve plant growth

* Corresponding author. Tel.: þ1 902 457 6328; fax: þ1 902 457 6455. E-mail address: [email protected] (G. Kernaghan). http://dx.doi.org/10.1016/j.funeco.2015.05.001 1754-5048/ª 2015 Elsevier Ltd and The British Mycological Society. All rights reserved.

Diversity and host preference of fungi

through phytohormone production (Khan et al., 2012), drought tolerance (Barrow, 2003), or improved access to soil phosphorus (Barrow and Osuna, 2002) or nitrogen (Upson et al., 2009). Unlike mycorrhizal fungi, fungal root endophytes lack highly evolved absorptive structures involved in carbon and nutrient exchange (Brundrett, 2006), instead forming structures such as microsclerotia and hyphal coils within the host root (O’Dell et al., 1993; Vohn
ık et al., 2003). Also, ECM fungal hyphae explore the soil environment and generally colonize fine roots only, while fungal root endophytes are more likely to be restricted to the roots, although they may grow throughout the root system (Menkis et al., 2004; Rodriguez et al., 2009; Toju et al., 2013a). Much of the research into fungal root endophytes has focused on the dark septate endophytes, or DSE (Ahlich and € nig et al., 2002; Alberton et al., 2010; Sieber, 1996; Gru Newsham, 2011); a morphological classification encompassing fungal root endophytes with highly melanized hyphae. Although DSE are common, woody roots and ECM are colonized by a much more diverse array of fungi, including many non-melanized species, as demonstrated by several culture dependent (Schild et al., 1988; Fisher et al., 1991; Girlanda and Luppi-Mosca, 1995; Girlanda et al., 2002; Hoff et al., 2004; Summerbell, 2005; Kernaghan and Patriquin, 2011), and culture independent studies (Kernaghan et al., 2003; Kwa
sna et al., 2008; Urban et al., 2008; Tedersoo et al., 2009; Izumi and Finlay, 2011; Toju et al., 2013a). Recent evidence also indicates that some fungal root endophytes exhibit preference for particular host plants € nig et al., 2008; Tedersoo et al., 2009; Kernaghan and (Gru Patriquin, 2011; Quilliam and Jones, 2012; Tejesvi et al., 2013), although examples of specificity at levels seen in some ECM fungi are lacking (Molina et al., 1992; Toju et al., 2013a). There is also increasing evidence that some fungal root endophytes prefer ECM formed by particular species of ectomycorrhizal fungi (Urban et al., 2008; Tedersoo et al., 2009; Izumi and Finlay, 2011; Yamamoto et al., 2014), potentially representing a second level of root endophyte selection. The occurrence of host preference, in combination with the range of possible effects that root associated fungi may have on their host plants, has motivated our research into differences in root associated fungal communities among host trees. In a previous study (Kernaghan and Patriquin, 2011), we investigated species composition, diversity and host preference in fungal root endophyte communities of ECM of boreal trees using culture-based techniques. To focus on differences among host plant species, and to avoid the potential influence of preference for ectomycorrhizal symbionts, ECM formed by Cenococcum geophilum (Cg) were focused on. Cg forms distinctive ECM that often dominate forest soils. It is widely dispersed, with a very broad host range and a relatively even distribution across soil horizons and successional stages (LoBuglio, 1999; Dickie et al., 2002). Although genetic variability in Cenococcum appears to be quite high (Douhan and Rizzo, 2005), the Cg ECM of a given host plant species should still represent a relatively homogeneous set of habitats for the study of root associated fungi. Here, we revisit questions regarding the species composition, diversity and host preference of fungi associated with the

85

Cg ECM of boreal trees, using direct PCR and cloning to eliminate the biases inherent in culture-dependant studies. We expected that the diversity of fungi associated with Cg ECM was underestimated in our previous work (Kernaghan and Patriquin, 2011) due to fast growing fungi such as Phialocephala fortinii outcompeting slower growing species. We also expected that a culture-independent approach would reveal obligate root associates that are not readily amenable to culture.

Materials and methods Sampling ECM samples were collected from two boreal forest sites; Mount Mackenzie, Cape Breton Highlands National Park, Nova Scotia and the Lac Duparquet Teaching and Research
miscamingue, Que
bec. The sites are Forest, Abitibi-Te 1 400 km apart and support similar mixtures of mature Abies balsamea, Betula papyrifera and Picea glauca with sparse understory vegetation dominated by herbaceous species including Cornus canadensis and Clintonia borealis, as well as ferns such as Osmundastrum cinnamomeum. Ericaceous shrubs including Kalmia and Vaccinium spp. are also present but not dominant. More detailed site descriptions are given in Kernaghan and Patriquin (2011). At each site, four 2 m2 sampling plots were established approximately 50 m apart. Each plot supported all three dominant tree species (A. balsamea, B. papyrifera and P. glauca; at least one of each per plot). During the snow free period between May and Sep., root systems were traced from the base of one individual of each of the three tree species on each plot (four root systems per tree species per site, for a total of 24 root systems) to the fine roots and associated ECM. Dominant ECM types were identified on the basis of morphology, and included C. g., Laccaria bicolor, Tomentella spp., Russula spp. Tylospora sp., “Piceirhiza bicolorata” and Cortinarius spp. Twenty healthy looking Cg ECM were collected from each root system, surface sterilized in 15 % hydrogen peroxide and frozen at 20 in AP1 extraction buffer (Qiagen) prior to DNA extraction.

DNA extraction and PCR Genomic DNA was extracted from each set of 20 ECM using the DNeasy Plant Mini kit (Qiagen). ECM frozen in AP1 buffer were ground in a ceramic mortar and incubated at 65  C for 30 min prior to following the manufacturer’s instructions. Fungal ITS rDNA was amplified from the extracted genomic DNA using the fungal specific primers ITS-1F (Gardes and Bruns, 1993) and ITS-4 (White et al., 1990) in 25 ml reactions containing 2.5 U taq polymerase (New England Biolabs), 2.5 ml 10X PCR buffer (Promega) (0.05 M KCl, 0.01 M TriseHCl, 0.01 % Triton X-100), 2.5 mM of each primer, 2.5 mM MgCl2 and 0.2 mM dNTPs. In cases of PCR failure, reactions were amplified under the same conditions, but using GoTaq master mix (Promega). Cycling parameters were as described in Kernaghan and Patriquin (2011).

86

Gel separation and cloning To separate Cg amplicons from those of other fungi, we took advantage of the w500 bp CgSSU intron (Shinohara et al., 1996, 1999) that results in the characteristically large PCR products (1 100 bp) from amplifications of Cg ITS using forward primers targeting the 30 end of the SSU and the 50 end of the LSU rDNA. Amplicons were separated by running 20 ml of PCR product on 1 % low melt agarose in TAE buffer for 60 min at 90 V. After ethidium bromide staining, gel slices containing amplicons ranging from 400 to 1 000 bp were excised, excluding the band of Cg DNA at 1 100 bp. The agarose gel slices were then purified using the Qiaex II gel purification system (Qiagen) according to the manufacturer’s instructions. Bands at 1 100 bp were also excised from select gels and purified to confirm that the 1 100 bp band represented Cg DNA. Excised 400e1 000 bp amplicons were then ligated into pGEM-T Easy Vector System II (Promega) and used to transform NEB 5-alpha competent Escherichia coli cells (New England Biolabs). Twenty white colonies from each transformation were then randomly selected for amplification by colony PCR, using the same protocol used for the ECM samples, except that the initial denaturation step was increased to 7 min and the annealing temperature increased to 60  C. Each amplicon generated by colony PCR was subsequently digested with TaqI (Bougoure et al., 2007) and run on 2 % agarose gels stained with ethidium bromide, resulting in 20 RFLP patterns from each of the ECM samples.

G. Kernaghan, G. Patriquin

In five samples, the sequence data were dominated by Cg, likely because of a lack of the CgSSU intron in some of the rDNA, resulting in Cg amplicons in the same size range as our target fungi. These samples, together with a sixth, which had a high proportion of chimeric sequences, were not further analyzed, leaving samples from three replicate trees per host species per site, instead of the original four.

Statistical analyses Diversity indices (Fisher’s alpha) were calculated on the basis of the number and proportion of RFLP patterns generated from each sample using PAST version 2.04 (http://folk.uio.no/ ohammer/past/). Fungal richness and diversity values were compared among hosts using the Kruskal-Wallis test. Species accumulation curves and associated bootstrap and Jackknife richness estimates were produced for each host tree species using Estimate 8.2.0 (http://viceroy.eeb.uconn.edu/estimates). Relationships among sequence types and host tree species were further explored using the following analyses in R (R Development Core Team, 2010): permutational multivariate analysis of variance (PERMANOVA) using the ADONIS function in Vegan (http://CRAN.R-project.org/package/vegan), non-metric multidimensional scaling (NMDS) with BrayCurtis distances using the metaNMDS function in Vegan, and indicator species analysis using the multipatt function of the indicspecies package (http://cran.r-project.org/web/packages/indicspecies).

Sequencing Classification of fungi detected The number of distinct patterns within each set of 20 RFLPs was determined, and 25 % of the amplicons representing each pattern were sequenced at the McGill University and Genome
bec Innovation Centre with an ABI PRISM 3730XL DNA Que Analyzer system using ITS1 (White et al., 1990) and ITS4 as forward and reverse primers, respectively. If the resulting sequences revealed that a single RFLP type represented more than one sequence type, then all of the amplicons showing that RFLP pattern were subsequently sequenced.

Sequence analysis Sequence quality was determined on the basis of KB basecaller (ABI) scores, and low quality sequences were excluded. Sequence contigs were then assembled and edited using Sequencher 4.9 (Gene Codes, Ann Arbor, MI) and clustered using the fungal ITS pipeline (Nilsson et al., 2009), which queries sequences against the International Sequence Data Base (INSD) using BLAST and groups sequences that share at least 50 % of their best 15 matches. Sequences within each group were subsequently blasted against one another using BLAST (bl2seq) to ensure that within group sequence variability did not exceed 3 %. Chimeric sequences were identified by blasting the ITS1 and ITS2 regions separately and removed. Examples of each remaining sequence group were then compared to those on the INSD using blastn. As much as possible, sequence identifications were based on BLAST matches to sequences derived from cultures or fruit bodies, rather than to “environmental” sequences. All high quality sequences were deposited in GenBank (KC876130eKC876356).

Sequence types were categorized on the basis of inferred melanization, mycorrhizal status and type and level of root association. Categorization into either heavily melanized (DSE) or not heavily melanized was made on the basis of both € nig et al., published descriptions (e.g. Domsch et al., 1993; Gru 2011) and live cultures obtained during previous work at the same field sites (Kernaghan and Patriquin, 2011). Fungi in culture were considered heavily melanized if their mycelia darkened within the first month of growth, and not heavily melanized if their mycelia were persistently un-pigmented. Categorization with respect to level of root association (strong, partial, weak or unknown) was based both on liter€ nig et al., 2011) and on ature (e.g. Domsch et al., 1993; Gru matching International Sequence Data Base accessions in which the isolation habitat is reported. Fungi were considered strongly root associated when more than 75 % of the habitats were reported as roots, partially associated when roots were between 50 % and 75 % of habitats, and weakly associated if roots accounted for fewer than 50 %. All mycorrhizal species were considered to be strong root associates. Grouping by mycorrhizal status (ECM, ericoid, nonmycorrhizal and unknown) was mainly based on literature reports (e.g. Chambers et al., 2008; Rinaldi et al., 2008) although the following approaches were taken for two common sequence types that lacked significant GenBank matches (other than ‘environmental’ sequences). Helotiaceae sp. III was classified as ECM on the basis of sequences obtained from ECM occurring on the Cape Breton site, which were morphologically similar to the P. bicolorata morphotype described by

Diversity and host preference of fungi

87

Vr alstad et al. (2000). Helotiaceae sp. VI was classified as potentially ericoid mycorrhizal (ERM) on the basis of in vitro mycorrhizal synthesis using the methods of Villarreal-Ruiz et al. (2004). Sterile V. macrocarpon seedlings were paired with pure cultures (ARSL 070907.33, ARSL 190907.14) isolated from the same sites (Kernaghan and Patriquin, 2011) and with ITS sequences identical to those of Helotiaceae sp. VI detected in the present study. Mycorrhizal syntheses were carried out in Petri dishes on low carbon media, and Oidiodendron maius and L. bicolor were used as positive controls for ERM and ECM formation, respectively. The ECM hosts Abies, Betula and Picea, failed to form ECM with the Helotiaceae sp. VI isolates.

Results Fungi detected Our PCR-cloning strategy detected 50 distinct non-Cg ITS sequence types that varied by at least 3 % (including 18 singletons) (Table 1, Table S1). Of these, ascomycetes accounted for 56 % of the sequence types (species) and 75 % of the amplicons and basidiomycetes represented 42 % of the sequence types and 24 % of the amplicons. The Mucoromycotina made up only 1 % of amplicons and 2 % of sequence

Table 1 e Matches between sequences obtained from Cg ECM and GenBank or UNITE databases GenBank accessions

Closest database matcha

Percent coverage

Ascomycetes Archaeorhizomycete sp. I

KC876130eKC876141

99

419/461

90.8

Archaeorhizomycete sp. II

KC876142eKC876143

99

440/465

94.6

Cladophialophora chaetospira Cladophialophora sp.

KC876147eKC876148

100

586/598

97.9

100

569/603

94.4

Dermateaceae sp. I

KC876152eKC876155

91

498/500

99.6

Dermateaceae sp. II

KC876156eKC876168

89

375/419

89.5

Dermateaceae sp. III

KC876169eKC876174

100

498/554

89.8

Exophiala sp.

KC876175

90

547/570

96

Helotiaceae sp. III

KC876176eKC876185

98

556/558

99.6

Helotiaceae sp. V

KC876186eKC876187

98

557/558

99.8

Helotiaceae sp. VI

KC876188eKC876217

98

552/561

98.4

Hyaloscyphaceae sp. I

KC876218

85

466/470

99.1

Hyaloscyphaceae sp. IV

KC876220

85

467/468

99.7

Hyaloscyphaceae sp. V

KC876221

100

508/553

91.8

Hypocrea pachybasioides

KC876222

94

564/568

99.2

Lachnum sp.

88

462/480

96.2

Meliniomyces bicolor

KC876223e KC876224 KC876227eKC876237

98

532/542

98.1

Meliniomyces variabilis

KC876238eKC876247

98

546/548

99.6

Meliniomyces vraolstadiae

KC876248eKC876251

98

538/538

Oidiodendron maius

KC876252eKC876254

82

433/435

99.5

Penicillium montanense

KC876255

Uncultured Pezizomycotina clone SLUBC35 (FJ152542) Ectomycorrhizal root tip 93-sepA_Ny1.EB23.5 (AF476985.1) Cladophialophora chaetospira strain CBS 491.70 (EU035405.1) Cladophialophora chaetospira strain CBS 514.63 (EU035406.1) Dermataceae sp. I GK-2010 strain ARSL 190907.53 (HQ157901.1) Cryptosporiopsis ericae voucher UAMH 10419 (AY853167.1) Phialocephala virens strain CBS 452.92 (AF486132.1) Exophiala moniliae strain CBS 520.76 (GU225948.1) Helotiaceae sp. III GK-2010 strain ARSL 070907.34 (HQ157918.1) Helotiaceae sp. V GK-2010 strain ARSL 190907.54 (HQ157878.1) Helotiaceae sp. VI GK-2010 strain ARSL 070907.9 (HQ157913.1) Hyaloscyphaceae sp. I GK-2010 strain ARSL 230507.52 (HQ157858.1) Hyaloscyphaceae sp. IV GK-2010 strain ARSL 180907.20 (HQ157957.1) Hyphodiscus hymeniophilus strain MUCL 40275 (DQ227258.1) Hypocrea pachybasioides strain ARSL 190907.26 (HQ157910.1) Lachnum pygmaeum isolate ARON3255.H (AJ430218.1) Meliniomyces bicolor strain ARSL 180907.22 (HQ157926.1) Meliniomyces variabilis strain ARSL 220507.4I (HQ157933.1) Meliniomyces vraolstadiae strain ARSL 250507.3 (HQ157841.1) Oidiodendron maius strain ARSL 230507.19 (HQ157849.1) Penicillium montanense strain ARSL 180507.8 (HQ157860.1)

100

555/563

98.6

Phylotype

KC876149eKC876151

Identity

Percent similarity

100

(continued on next page)

88

G. Kernaghan, G. Patriquin

Table 1 e (continued ) GenBank accessions

Closest database matcha

Percent coverage

Phialocephala fortinii s.l.

KC876256eKC876272

100

545/552

98.7

Phialocephala sphaeroides

KC876273eKC876274

93

839/849

98.8

Rhizoscyphus ericae aggregate sp. I Rhizoscyphus ericae aggregate sp. II Thysanophora penicillioides Venturia sp. I

KC876275eKC876282

87

443/475

93.2

100

510/545

93.5

KC876144 KC876284eKC876285

100 100

530/531 499/550

99.8 90.7

Venturia sp. II

KC876286eKC876287

Phialocephala fortinii voucher BB64_310_Of_Fa_ 011106 (DNA614) (HM190137.1) Phialocephala sphaeroides strain UAMH 10279 (AY524844.1) Hymenoscyphus ericae isolate 101 (AF069505.1) Meliniomyces bicolor isolate MBI-1 (EF093180.1) Thysanophora penicillioides (AB213271.1) Venturia tremulae var. tremulae strain CBS 257.38 (EU035475.1) Venturia hystrioides strain CBS 117727 (EU035459.1)

100

492/541

90.9

Basidiomycetes Amanita virosa

KC876288

96

627/629

99.6

Clavulina sp.

KC876289eKC876294

96

649/659

98.4

Clavulinaceae sp.

KC876295

100

590/720

81.9

Cortinarius sp. I Cortinarius sp. II Cortinarius sp. III Galerina sp.

KC876296 KC876297 KC876298eKC876299 KC876301eKC876305

100 100 94 89

420/426 566/577 590/597 588/606

98.6 98.1 98.8 97

Hydnodontaceae sp. Inocybe sp. Lactarius camphoratus Lactarius sp.

KC876306eKC876308 KC876309eKC876310 KC876311 KC876312eKC876313

100 88 100 93

568/661 577/587 701/702 667/673

85.9 98.2 99.8 99.1

Lactarius tabidus

KC876314

100

599/605

99

Mycena sp. I

KC876315eKC876321

100

698/700

99.7

Mycena sp. II Mycena sp. III

KC876322eKC876324 KC876325

99 93

671/676 642/657

99.2 97.7

Mycena sp. IV

KC876326eKC876330

100

685/686

99.0

Piloderma sp.

KC876331eKC876340

95

580/580

Pucciniomycete sp.

KC876343eKC876346

98

451/575

78.4

Russula sp. Tremellodendron sp.

KC876347eKC876348 KC876350

100 95

689/690 568/593

99.8 95.8

Mycena sp. IV

KC876326eKC876330

100

685/686

99.0

Tricholomataceae sp.

KC876351

Amanita virosa voucher RV96No.37 (EU909449.1) Clavulina cf. cristata BIO 10088 (EU862212) Clavulina cf. amethystina O 175524 (EU862208.1) Cortinarius flexipes var. inolens (UDB001228) Cortinarius diasemospermus (UDB002161) Cortinarius comptulus (UDB002198) Galerina lubrica specimen voucher O 154034 (AJ585471.1) Trechispora farinacea (EU909231.1) Inocybe leptophylla BJ920801 (UDB000620) Lactarius camphoratus KF01-23 (UDB000045) Lactarius chrysorrheus voucher TENN61713 (FJ596871.1) Lactarius tabidus voucher BB22_104_Aeh_Pi_ 230407 (DNA745) (HM189833.1) Mycena sp. GK-2010 strain ARSL 190907.23II (HQ157912.1) Mycena aff. murina F14062 (AF335444.1) Mycena sanguinolenta voucher TENN59879 (JF908390.1) Mycena silvae-nigrae voucher CC 13-12 (KF359604.1) Ectomycorrhizal root tip (Piloderma) 192_Ny3.B1-31.4 (AF476984) Helicobasidium purpureum II voucher BR 37479-37 (AY292431.1) Russula betularum AF-C04 (UDB002437) Tremellodendron pallidum voucher F:PRL4391 (GQ166897.1) Mycena silvae-nigrae voucher CC 13-12 (KF359604.1) Leucopaxillus tricolor voucher JMP0050 (EU819413.1)

99

598/673

88.8

Mucoromycotina Umbelopsis sp.

KC876352eKC876356

99

585/606

96.5

Phylotype

KC876283

Umbelopsis isabellina PP82a (AJ876493.1)

Identity

Percent similarity

100

a Sequence accession numbers beginning with UDB are from the Unite Data Base, all others are from GenBank.

types. At the family level, the Helotiaceae and Dermateaceae were most common, followed by the Tricholomataceae (mainly on B. papyrifera at the Quebec site) and the Archaeorhizomycetaceae, although not detected on P. glauca. Members of 16 other families were detected at relatively low abundances (Fig 1A and B).

Fungal richness and diversity Species accumulation curves indicate that only 16 non-Cg species were detected on A. balsamea, while 26 and 25 species were found on B. papyrifera and P. glauca respectively, even though similar numbers of amplicons were analyzed for each host

Diversity and host preference of fungi

Betula

40

Q

Betula CB

Q

Picea CB

Q

0

20

40

60

80

100

Abies CB

Fig 1 e Proportions of fungal families detected in association with Cenococcum ectomycorrhizae of Abies balsamea, Betula papyrifera and Picea glauca. (A) by host tree species; (B) by host tree species and site.

Richness

Sebacinaceae Myxotrichaceae Hypocreaceae Amanitaceae Trichocomaceae Venturiaceae Umbelopsidaceae Pucciniomycete Hydnodontaceae Hyaloscyphaceae Herpotrichiellaceae Cortinariaceae Russulaceae Strophariaceae Clavulinaceae Atheliaceae Archaeorhizomycetaceae Tricholomataceae Dermataceae Helotiaceae

J

20

B Obs

10

0 0

20

40

60

80

100

120

Amplicons

B

40

J

B

30

Richness

20

40

60

30

B % of amplicons

A

Picea

80

Abies

0

% of amplicons

100

A

89

Obs 20

10

0 0

20

40

60

80

100

120

Amplicons

Characterization of detected fungi Based on literature descriptions and comparison with cultures isolated from the same field sites (Kernaghan and Patriquin, 2011), approximately 70 % of the fungi for which melanization could be inferred were not strongly melanized (i.e. not considered DSE) (Fig 3A and B). Although melanization levels could not be inferred for all fungal species, weakly or nonmelanized species would still dominate even if all of the undetermined fungi were melanized. This trend is evident in all three host species, although B. papyrifera appears to support the highest proportion of melanized fungi. For the fungi for which the strength of root association could be inferred, the majority (88 % of amplicons and 61.5 % of sequence types) appeared to represent fungi considered to be strongly associated with plant roots. Only a small number (4 % of amplicons and 15 % of sequence types) appear to be generalists with little preference for plant roots. All three

C

J

40

B

30

Richness

(Fig 2AeC). Richness estimators (Bootstrap and Jackknife 2) predict that actual species richness may be from 18 to 27 for A. balsamea, 30e37 for B. papyrifera and 30e44 for P. glauca. The average non-Cg fungal species richness per sample of 20 ECM was 4.17  0.54 (SE) for A. balsamea, 6.0  0.89 for B. papyrifera and 6.2  0.70 for P. glauca, and was not significantly different among hosts (p ¼ 0.13). The average non-Cg fungal species richness per individual ECM (across all host species) was estimated at 0.27. With respect to fungal species diversity, A. balsamea samples showed the lowest average Fisher’s a value at 1.79  0.42 (SE), while B. papyrifera had an average a of 3.25  0.63 and P. glauca averaged 6.04  2.59. The diversity of the A. balsamea samples was significantly lower than that of the P. glauca samples (p ¼ 0.03).

Obs 20

10

0 0

20

40

60

80

100

120

Amplicons

Fig 2 e Species accumulation curves for root associated fungi on (A) Abies balsamea, (B) Betula papyrifera and (C) Picea glauca. Obs; observed species, J; jackknife 2 estimator, B; bootstrap estimator.

hosts were dominated by strong root associates, but the proportion was largest in A. balsamea (Fig 3C and D). The mycorrhizal status and type was inferred for 65 % of the fungi detected. Of these, 24 % of the amplicons and 45 % of the sequence types were deemed to represent nonmycorrhizal fungi, 37 % and 45 % represented ECM fungi and 39 % and 10 % represented fungi with the potential to form ERM. P. glauca supported the largest proportion of potentially ECM fungi, while A. balsamea supported the largest proportion of potentially ERM fungi (Fig 3E and F). However, these categories may not necessarily be exclusive; for example, some species may form either ECM or ERM depending on the host (Grelet et al., 2010).

50

50 100 150 200 250 300 350

G. Kernaghan, G. Patriquin

30 20 10

Species

0 Betula

Total

Picea

Abies

Betula

Picea

50

50 100 150 200 250 300 350

Abies

D

30 20 10

Species

40

C

0

0

Abies

Betula

Picea

Total

Abies

Betula

Picea

50

50 100 150 200 250 300 350

Total

F

30 0

0

10

Species

40

E

20

Amplicons

Total

Amplicons

B

40

A

0

Amplicons

90

Total

Abies

Betula

Picea

Total

Abies

Betula

Picea

Fig 3 e Categorization of fungi associated with Cenococcum ectomycorrhizae on the basis of number of amplicons and number of sequence types. (A, B) melanization: grey; undetermined, white; not heavily melanized, black; heavily melanized (DSE). (C, D) level of root association: grey; undetermined, white; strongly root associated, hatched; partial preference for roots, black; no clear association with roots. (E, F) mycorrhizal status: grey; undetermined, white; non-mycorrhizal, hatched; potentially ectomycorrhizal, black; potentially ericoid mycorrhizal.

Distributions of fungal species Permutational multivariate analysis of variance (PERMANOVA) indicated that there were no significant differences in co-inhabiting fungal communities between sites or among plots, as well as no significant site  plot interaction. However, fungal communities differed significantly among hosts (p ¼ 0.0034) with no significant host  site interaction (Table 2).

NMDS (stress ¼ 0.152) also indicated that relatively distinct communities of fungi co-inhabited the Cg ECM of the three hosts sampled (Fig 4). Fungal species to the lower right of the diagram, such as Helotiaceae sp. VI and Archaeorhizomycete spp., were mainly detected on A. balsamea, while those to the left of the diagram, such as Mycena sp. I and Piloderma sp., mainly colonized P. glauca. Fungal species near the top center of the diagram, such as Rhizoscyphus ericae agg. I and Mycena sp. IV, were characteristic of B. papyrifera. Those placed

Diversity and host preference of fungi

91

Table 2 e Results of permutational multivariate analyses of variance comparing fungal communities across plots and hosts

Plot by site analysis

Host by site analysis

Degrees of freedom

Sum of squares

Mean of sum of squares

FModel

R2

p

1 3 2 11 17 1 2 2 12 17

0.4952 1.4508 0.7934 4.3714 7.1109 0.4952 1.5063 0.7077 4.4016 7.1109

0.49523 0.48361 0.3967 0.3974 1 0.49523 0.75317 0.35386 0.3668 1

1.24618 1.21695 0.99824

0.06964 0.20403 0.11158 0.61475

0.2262 0.1796 0.4744

1.35014 2.05336 0.96473

0.06964 0.21184 0.09953 0.61899

0.1654 0.0034 0.5128

Site Plot Site*Plot Residuals Total Site Host Site*Host Residuals Total

Significant differences (p < 0.05) are in bold.

0.3

Cortinarius sp. III Inocybe sp. Phialocephala sphaeroides

0.1

0.2

Mycena sp. IV

Rhizoscyphus ericae agg. I

Hydnodontaceae sp. I

Cladophialophora chaetospira Russula sp. Dermateaceae sp II Lactarius camphoratus

Helotiaceae sp. III

0.0

Meliniomyces bicolor Mycena sp. I Lachnum sp. Umbelopsis sp.

Dermateaceae sp. III Helotiaceae sp. VI

Venturia sp. II Cladophialophora sp. Hypocrea sp. Venturia sp. I

Archaeorhizomycete sp. II

Archaeorhizomycete sp. I

-0.1

Puccinomycete sp.

Piloderma sp. Galerina sp.

Lactarius sp. Dermateaceae sp. I

Clavulina sp.

Phialocephala fortinii

Helotiaceae sp.V

Meliniomyces variabilis

-0.2

Meliniomyces vraolstadiae

-0.3

-0.2

Mycena sp II

-0.1

0.0

0.1

0.2

0.3

Fig 4 e NMDS analysis showing the distribution of sequence types on host trees. Host tree species (site scores) are indicated by ellipses based on standard deviations around their centroids (white; Picea glauca; light gray; Betula papyrifera; dark gray; Abies balsamea). Fungal species are represented by open circles, or closed circles where 2 or 3 species scores overlap. Singleton fungal species are not shown.

towards the center bottom, such as Clavulina sp. and Meliniomyces bicolor, were common to all three hosts. Of the 50 species of co-inhabiting fungi detected, only six (Clavulina sp., Dermateaceae sp. II, Helotiaceae sp. III, M. bicolor, M. variabilis and P. fortinii s.l.) were found on all three host trees (Table S1). Other than these, two were common to A. balsamea and B. papyrifera (Archaeorhizomycete sp. I and Helotiaceae sp. VI), two common to B. papyrifera and P. glauca (Cladophialophora chaetospira and Hydnodontaceae sp.), and only Galerina lubrica was common to A. balsamea and P. glauca. The majority (78 %) of co-inhabiting fungi were detected from only one host, and although many species were rare (36 % singletons), others were relatively common on certain hosts.

Indicator species analysis determined that two fungi, Helotiaceae sp. VI and R. ericae aggregate sp. I, exhibited both specificity and fidelity for A. balsamea and B. papyrifera, respectively (Table 3). In the case of P. glauca, Piloderma sp. exhibited host preference, but lacked fidelity (found in 3/6 samples), resulting in a non-significant p value (0.073).

Discussion We detected 50 species of fungi associated with Cg ECM, and found that fungal diversity and species composition varied significantly among the three host tree species sampled. The

92

G. Kernaghan, G. Patriquin

Table 3 e Indicator species results for the two most significant fungi for each host species Host Abies

Fungi

Helotiaceae sp. VI Dermateaceae III Betula Rhizoscyphus ericae agg. sp. I Mycena sp. IV Picea Piloderma sp. Helotiaceae sp. III

Association index

p

0.861 0.707 0.816 0.577 0.707 0.698

0.0064 0.074 0.0146 0.2896 0.0726 0.0958

Significant indicator species (p < 0.05) are in bold.

majority of fungi detected were ascomycetes, with the Helotiaceae as the largest group overall. It is becoming clear that members of the Helotiaceae, such as Meliniomyces spp., are not only important components of fungal root endophyte communities, but are also potential ericoid or ectomycorrhizal symbionts (Vr alstad et al., 2000; Grelet et al., 2010; Vohn
ık et al., 2013). The Dermateaceae was the next most common family, including highly melanized species such as Phialocephala spp. that would be considered DSE (Jumpponen and Trappe, 1998). Members of the Archaeorhizomycetaceae were also relatively common, although not detected on P. glauca. These represent a newly discovered but ancient fungal lineage, several species of which are common root associates of boreal tree species (Rosling et al., 2011, 2013). Close to half of the species we detected were basidiomycetes (42 %), much higher than the 6 % detected from the same type of samples using culture-based methods (Kernaghan and Patriquin, 2011). This is likely due to the fact that many basidiomycetes grow slowly (or not at all) in culture, and are often outgrown by faster groups (Thorn et al., 1996). For example, Hoff et al. (2004) obtained only a small number of basidiomycete cultures from Douglas-fir roots, even though basidiomycete selective media were used. Kwa
sna et al. (2008) compared culture-based and direct PCR methods of assessing root associated fungi and found that the resulting data sets were dominated by ascomycetes and basidiomycetes, respectively. A direct comparison of our current PCR based data with our previous culture dependent data (Kernaghan and Patriquin, 2011) revealed somewhat higher richness (50 vs 31 taxa respectively) and a 23 % overlap in fungal species identity. Furthermore, fungi that grow slowly in culture are better represented by direct PCR, while faster fungi dominate the culture based data. For example, the very slow growing Helotiaceae VI represented 19 % of all amplicons but only 9 % of cultures, while the much faster P. fortinii (DSE) represented only 5 % of amplicons, but 33 % of cultures. The basidiomycete community in the present study was dominated by members of the Tricholomataceae, particularly species of Mycena. Although Mycena spp. are generally considered soil and wood saprotrophs, or mycorrhizal symbionts of the Orchidaceae (Selosse et al., 2010), they have recently been reported as endophytes within the leaves of bryophytes (Davey et al., 2013) and the roots of grasses (Yuan et al., 2010; Tejesvi et al., 2013), ericaceous plants (Bougoure et al., 2007) and several other woody hosts, including Abies, Acer, Betula, Larix, Picea, Quercus and Schisandra (Kwa
sna et al., 2008; Kernaghan and Patriquin, 2011; Toju et al., 2013b; Vohn
ık

et al., 2013). As a genus, Mycena appears to form root endophytic associations with a broad range of hosts and clearly warrants further investigation in this regard. Many of the other basidiomycetes we detected in association with Cg ECM are ectomycorrhizal, including Clavulina, Cortinarius, Piloderma and Russula. As sequences from these genera were relatively uncommon, they may simply arise from exploratory hyphae adhering to the surface of the Cg ECM. However, in the case of Piloderma sp., there appeared to be a preference for P. glauca (association index of 0.707; p ¼ 0.073), which may indicate an endophytic association or secondary ectomycorrhizal colonization (Wu et al., 1999; Olsson et al., 2000). Species accumulation curves indicate that our sampling intensity was sufficient for A. balsamea, which exhibited relatively low richness, but further sampling of B. papyrifera and P. glauca would likely reveal even more root associated fungi. Comparisons with studies by other researchers is complicated by their inclusion of multiple ECM types, but the fungal richness detected in the present study is similar to that found on the roots of several plant species by pyrosequencing (Toju et al., 2013a,b). Although the relatively low fungal richness associated with A. balsamea Cg ECM could be due to a direct influence of the plant species, with the roots of A. balsamea simply accepting fewer fungal associates, it may instead be due to competitive exclusion by certain fungal endophytes. A. balsamea roots exhibited the lowest overall diversity and the highest colonization by members of the Helotiaceae (most A. balsamea samples were dominated by Helotiaceae species VI), while the situation in P. glauca was reversed. Some Helotiaceae, such as Ascocoryne and Hymenoscyphus spp. produce antifungal compounds (Hosoya, 1998; Morath et al., 2012; Halecker et al., 2014), and preliminary co-culture experiments indicate that Helotiaceae sp. VI is inhibitory to the growth of other fungal root endophytes (Kernaghan, unpublished). With respect to general characteristics of the fungal communities, we found that fungi lacking strong melanization were more abundant than heavily pigmented species (e.g. DSE), in terms of both number of sequence types and number of amplicons. Although DSE fungi such as Philaocephala have traditionally been considered the dominant endophytes of tree roots (Rodriguez et al., 2009), this is likely due to their rapid growth in culture. Our results agree with those of Vohn
ık et al. (2013), who found that non-melanized endophytes, such as species of Helotiaceae, can be dominant components of the fungal root endophyte community. Also, the vast majority of fungi detected appear to be strongly root associated (either root endophytes or non-Cg ectomycorrhizal fungi), with only a few that would also be expected in the soil. This is indicative of the fact that plant roots support fungal assemblages distinct from the surrounding environment; a prerequisite for the observed variation in fungal communities among hosts. Slightly more sequence types were classified as representing potentially mycorrhizal (ericoid or ectomycorrhizal) fungi than non-mycorrhizal fungi, although the mycorrhizal status remains unknown for a significant proportion. In terms of the number of amplicons generated (e.g. abundance), putatively mycorrhizal species also dominated over nonmycorrhizal species (especially on A. balsamea), although

Diversity and host preference of fungi

many of these are root endophytes with the potential to form ericoid mycorrhizae on ericaceous hosts (Vohn
ık et al., 2013), rather than ectomycorrhizal symbionts of the hosts studied. Some fungi, such as Dermateaceae sp. III, Mycena sp. IV and Piloderma sp., were detected on only one host, but lacked fidelity in that they were not found in every sample of that host. However, Helotiaceae sp. VI exhibited sufficient levels of both preference and fidelity to be considered an indicator species for A. balsamea. Helotiaceae sp. VI also preferentially colonized A. balsamea on the basis of our culture dependent methods (Kernaghan and Patriquin, 2011). Although less common, R. ericae aggregate sp. I was also determined to be an indicator species (for B. papyrifera) in the present study. Both of these fungi are members of the Helotiales, as are several of the root endophytic fungi reported as exhibiting host preference € nig et al., 2008; Tedersoo et al., 2009; in other studies (Gru Quilliam and Jones, 2012; Tejesvi et al., 2013; Toju et al., 2013a). Although some helotialian species exhibit very broad host ranges, e.g. P. fortinii (Jumpponen and Trappe, 1998) and Meliniomyces spp. (Hambleton and Sigler, 2005), many others are specific to certain plant species, or even particular tissues € nzlin, 1984; Verkley, of those species (Breitenbach and Kra 1999; Wang et al., 2006). We made use of the fact that Cg rDNA often contains the CgSSU intron (Shinohara et al., 1996, 1999), giving it a relatively large ITS amplicon which can be separated from those of other fungi. Other approaches to the detection of ECM associated fungi include the use of ascomycete specific primers on basidiomycetous ECM (Tedersoo et al., 2009; Izumi and Finlay, 2011), but this does not detect ECM associated basidiomycetes, and cannot be used for ascomycetous ECM such as Cg (often a dominant ECM type). Another approach involves amplification with more general fungal primers followed by pyrosequencing (Toju et al., 2013a,b). Although a powerful approach, pyrosequencing of ECM can result in datasets composed of mainly ECM forming taxa (Jumpponen et al., 2010), rather than the ECM associated species targeted here. Also, as pyrosequencing is biased toward short amplicons, only the ITS1 or the ITS2 region of the rDNA is generally analyzed (Schmidt et al., 2013), whereas sequences spanning both regions are more taxonomically informative; an important consideration given that many fungal root endophytes are not easily assignable to genera. One drawback of our approach however is that not all samples contained the CgSSU, resulting in the loss of some replicate samples. We have used culture independent methods to add to the growing evidence for host preference in ECM associated fungi. We also detected significant among-host differences in fungal diversity. Given the wide range of potential interactions between root associated fungi and their host plants, differences in fungal species composition and diversity among host species may represent an important but unseen factor in community ecology.

Acknowledgments This work was made possible by a grant from the Natural Sciences and Engineering Research Council of Canada

93

(341671-2007). We thank Cape Breton Highlands National Park and the Lac Duparquet Teaching and Research Forest for field Logistics. We also thank Michael Mayerhofer for assistance with statistics and graphics.

Appendix A. Supplementary data Supplementary data related to this article can be found at http://dx.doi.org/10.1016/j.funeco.2015.05.001.

references

Addy, H.D., Piercey, M.M., Currah, R.S., 2005. Microfungal endophytes in roots. Canadian Journal of Botany 83, 1e13. Ahlich, K., Sieber, T.N., 1996. The profusion of dark septate endophytic fungi in non-ectomycorrhizal fine roots of forest trees and shrubs. New Phytologist 132, 259e270. Alberton, O., Kuyper, T.W., Summerbell, R.C., 2010. Dark septate root endophytic fungi increase growth of Scots pine seedlings under elevated CO2 through enhanced nitrogen use efficiency. Plant and Soil 328, 459e470. Barrow, J.R., Osuna, P., 2002. Phosphorus solubilization and uptake by dark septate fungi in fourwing saltbush, Atriplex canescens (Pursh) Nutt. Journal of Arid Environments 51, 449e459. Barrow, J.R., 2003. Atypical morphology of dark septate fungal root endophytes of Bouteloua in arid southwestern USA rangelands. Mycorrhiza 13, 239e247. Bergemann, S.E., Garbelotto, M., 2006. High diversity of fungi recovered from the roots of mature tanoak (Lithocarpus densiorus) in northern California. Canadian Journal of Botany 84, 1380e1394. Bergero, R., Perotto, S., Girlanda, M., Vidano, G., Luppi, A.M., 2000. Ericoid mycorrhizal fungi are common root associates of a Mediterranean ectomycorrhizal plant (Quercus ilex). Molecular Ecology 9, 1639e1649. Bougoure, D.S., Parkin, P.I., Cairney, J.W., Alexander, I.J., Anderson, I.C., 2007. Diversity of fungi in hair roots of Ericaceae varies along a vegetation gradient. Molecular Ecology 16, 4624e4636. € nzlin, F., 1984. Fungi of Switzerland. In: Breitenbach, J., Kra Ascomycetes, vol. 1. Verlag Mykologia, Luzern, Switzerland. Brundrett, M.C., 2006. Understanding the roles of multifunctional mycorrhizal and endophytic fungi. In: Schulz, B., Boyle, C., Sieber, T.N. (Eds.), Microbial Root Endophytes. Springer, Berlin, pp. 281e293.
zares, E., Trappe, J.M., 1993. Vesicular endophytes in roots of Ca the Pinaceae. Mycorrhiza 2, 153e156. Chambers, S.M., Curlevski, N.J.A., Cairney, J.W.G., 2008. Ericoid mycorrhizal fungi are common root inhabitants of nonEricaceae plants in a south-eastern Australian sclerophyll forest. FEMS Microbiology Ecology 65, 263e270. Davey, M.L., Heimdal, R., Ohlson, M., Kauserud, H., 2013. Hostand tissue-specificity of moss-associated Galerina and Mycena determined from amplicon pyrosequencing data. Fungal Ecology 6, 179e186. Dickie, I.A., Xu, B., Koide, R.T., 2002. Vertical niche differentiation of ectomycorrhizal hyphae in soil as shown by T-RFLP analysis. New Phytologist 156, 527e535. Domsch, K.H., Gams, W., Anderson, T.H., 1993. Compendium of Soil Fungi, vol. 1. IHW Verlag, Eching, Germany. Douhan, G.W., Rizzo, D.M., 2005. Phylogenetic divergence in a local population of the ectomycorrhizal fungus Cenococcum geophilum. New Phytologist 166, 263e271.

94

Fisher, P.J., Petrini, O., Petrini, L.E., 1991. Endophytic ascomycetes and deuteromycetes in roots of Pinus sylvestris. Nova Hedwigia 52, 11e15. Fogel, R., 1988. Interactions among soil biota in coniferous ecosystems. Agriculture, Ecosystems and Environment 24, 69e85. Foster, R.C., Marks, G.C., 1967. Observations on the mycorrhizas of forest trees. II. The rhizosphere of Pinus radiata D. Don. Australian Journal of Biological Science 20, 915e926. Gardes, M., Bruns, T.D., 1993. ITS primers with enhanced specificity for basidiomycetes: application to the identification of mycorrhizae and rusts. Molecular Ecology 2, 113e118. Girlanda, M., Ghignone, S., Luppi, A.M., 2002. Diversity of sterile root-associated fungi of two Mediterranean plants. New Phytologist 155, 481e498. Girlanda, M., Luppi-Mosca, A.M., 1995. Microfungi associated with ectomycorrhizae of Pinus halepensis Mill. Allionia 33, 93e98. Grelet, G.A., Johnson, D., Vr alstad, T., Alexander, I.J., Anderson, I.C., 2010. New insights into the mycorrhizal Rhizoscyphus ericae aggregate: spatial structure and cocolonization of ectomycorrhizal and ericoid roots. New Phytologist 188, 210e222. € nig, C.R., Queloz, V., Sieber, T.N., 2011. Structure of diversity Gru in dark septate endophytes: from species to genes. In: € , A.M., Frank, C. (Eds.), Endophytes of Forest Trees. Pirttila Springer, Netherlands, pp. 3e30. € nig, C.R., Queloz, V., Sieber, T.N., Holdenrieder, O., 2008. Dark Gru septate endophytes (DSE) of the Phialocephala fortinii slAcephala applanata species complex in tree roots: classification, population biology, and ecology. Botany 86, 1355e1369. € nig, C., Sieber, T., Rogers, S., Holdenrieder, O., 2002. Spatial Gru distribution of dark septate endophytes in a confined forest plot. Mycological Research 106, 832e840. Halecker, S., Surup, F., Kuhnert, E., Mohr, K.I., Brock, N.L., Dickschat, J.S., Junker, C., Schulz, B., Stadler, M., 2014. Hymenosetin, a 3-decalinoyltetramic acid antibiotic from cultures of the ash dieback pathogen, Hymenoscyphus pseudoalbidus. Phytochemistry 100, 86e91. Hambleton, S., Sigler, L., 2005. Meliniomyces, a new anamorph genus for root-associated fungi with phylogenetic affinities to Rhizoscyphus ericae (hHymenoscyphus ericae), Leotiomycetes. Studies in Mycology 53, 1e27. Hoff, J.A., Klopfenstein, N.B., McDonald, G.I., Tonn, J.R., Kim, M.S., Zambino, P.J., Hessburg, P.F., Rogers, J.D., Peever, T.L., Carris, L.M., 2004. Fungal endophytes in woody roots of Douglas-fir (Pseudotsuga menziesii) and ponderosa pine (Pinus ponderosa). Forest Pathology 34, 255e271. Hosoya, T., 1998. Application of underutilized microbial resources in drug discovery: a review focused on discomycetes. Annual Report of the Sankyo Research Laboratory 50, 15e40. Izumi, H., Finlay, R., 2011. Ectomycorrhizal roots select distinctive bacterial and ascomycete communities in Swedish subarctic forests. Environmental Microbiology 13, 819e830. Jumpponen, A., Jones, K.L., Mattox, J.D., Yaege, C., 2010. Massively parallel 454 sequencing of fungal communities in Quercus spp. ectomycorrhizas indicates seasonal dynamics in urban and rural sites. Molecular Ecology 19, 41e53. Jumpponen, A., Trappe, J.M., 1998. Dark-septate root endophytes: a review with special reference to facultative biotrophic symbiosis. New Phytologist 140, 295e310. Kernaghan, G., 2013. Functional diversity and resource partitioning in fungi associated with the fine feeder roots of forest trees. Symbiosis 61, 113e123. Kernaghan, G., Patriquin, G., 2011. Host associations between fungal root endophytes and boreal trees. Microbial Ecology 62, 460e473. Kernaghan, G., Sigler, L., Khasa, D., 2003. Mycorrhizal and root endophytic fungi of containerized Picea glauca seedlings

G. Kernaghan, G. Patriquin

assessed by rDNA sequence analysis. Microbial Ecology 45, 128e136. Khan, A.L., Hamayun, M., Kang, S.M., Kim, Y.H., Jung, H.Y., Lee, J.H., Lee, I.J., 2012. Endophytic fungal association via gibberellins and indole acetic acid can improve plant growth under abiotic stress: an example of Paecilomyces formosus LHL10. BMC Microbiology 12, 3. Kwa
sna, H., Bateman, G.L., Ward, E., 2008. Determining species diversity of microfungal communities in forest tree roots by pure-culture isolation and DNA sequencing. Applied Soil Ecology 40, 44e56. LoBuglio, K.F., 1999. Cenococcum. In: Cairney, J.W.G., Chambers, S.M. (Eds.), Ectomycorrhizal Fungi: Key Genera in Profile. Springer-Verlag, Berlin, pp. 287e309. Mandyam, K., Jumpponen, A., 2005. Seeking the elusive function of the root-colonising dark septate endophytic fungi. Studies in Mycology 53, 173e189. Mayerhofer, M., Kernaghan, G., Harper, K.A., 2013. The effects of fungal root endophytes on plant growth: a meta-analysis. Mycorrhiza 23, 119e128. Menkis, A., Allmer, J., Vasiliauskas, R., Lygis, V., Stenlid, J., Finlay, R., 2004. Ecology and molecular characterization of dark septate fungi from roots, living stems, coarse and fine woody debris. Mycological Research 108, 965e973. Menkis, A., Vasiliauskas, R., Taylor, A., Stenlid, J., Finlay, R., 2005. Fungal communities in mycorrhizal roots of conifer seedlings in forest nurseries under different cultivation systems, assessed by morphotyping, direct sequencing and mycelial isolation. Mycorrhiza 16, 33e41. Molina, R., Massicotte, H., Trappe, J., 1992. Specificity phenomena in mycorrhizal symbioses: community-ecological consequences and practical implications. In: Allen, M.F. (Ed.), Mycorrhizal Functioning: an Integrative Plant-fungal Process. Chapman and Hall, NY, pp. 357e423. Morath, S.U., Hung, R., Bennett, J.W., 2012. Fungal volatile organic compounds: a review with emphasis on their biotechnological potential. Fungal Biology Reviews 26, 73e83.
rez-Pe
rez, M.A., Smith, M.E., Bledsoe, C.S., 2008. Morris, M.H., Pe Multiple species of ectomycorrhizal fungi are frequently detected on individual oak root tips in a tropical cloud forest. Mycorrhiza 18, 375e383. Narisawa, K., Usuki, F., Hashiba, T., 2004. Control of Verticillium yellows in Chinese cabbage by the dark septate endophytic fungus LtVB3. Phytopathology 94, 412e418. Newsham, K.K., 2011. A meta-analysis of plant responses to dark septate root endophytes. New Phytologist 190, 783e793. Nilsson, R.H., Bok, G., Ryberg, M., Kristiansson, E., Hallenberg, N., 2009. A software pipeline for processing and identification of fungal ITS sequences. Source Code in Biology and Medicine 4, 1. O’Dell, T., Massicotte, H., Trappe, J.M., 1993. Root colonization of Lupinus latifolius Agardh. and Pinus contorta Dougl. by Phialocephala fortinii wang & Wilcox. New Phytologist 124, 93e100. € nzenberger, B., Mahmood, S., Erland, S., 2000. Olsson, P.A., Mu Molecular and anatomical evidence for a three-way association between Pinus sylvestris and the ectomycorrhizal fungi Suillus bovinus and Gomphidius roseus. Mycological Research 104, 1372e1378. Quilliam, R.S., Jones, D.L., 2012. Evidence for host-specificity of culturable fungal root endophytes from the carnivorous plant Pinguicula vulgaris (Common Butterwort). Mycological Progress 11, 583e585. R Development Core Team, 2010. R: a Language and Environment for Statistical Computing, Reference Index Version 2.10.1. R Foundation for Statistical Computing, Vienna, Austria. http:// www.R-project.org/. Rinaldi, A.C., Comandini, O., Kuyper, T.W., 2008. Ectomycorrhizal fungal diversity: separating the wheat from the chaff. Fungal Diversity 33, 1e45.

Diversity and host preference of fungi

Rodriguez, R.J., White, J.F., Arnold, A.E., Redman, R.S., 2009. Fungal endophytes: diversity and functional roles. New Phytologist 182, 314e330. Rosling, A., Cox, F., Cruz-Martinez, K., Ihrmark, K., Grelet, G.A., Lindahl, B.D., Menkis, A., James, T.Y., 2011. Archaeorhizomycetes: unearthing an ancient class of ubiquitous soil fungi. Science 12, 876e879. Rosling, A., Timling, I., Taylor, D.L., 2013. Archaeorhizomycetes: patterns of distribution and abundance in soil. In: Horwitz, B.A., Mukherjee, P.K., Mukherjee, M.P., Kubicek, C. (Eds.), Genomics of Soil-and Plant-associated Fungi. Springer, Berlin, Heidelberg, pp. 333e349. Schild, D.E., Kennedy, A., Stuart, M.R., 1988. Isolation of symbiont and associated fungi from ectomycorrhizas of Sitka spruce. European Journal of Forest Pathology 18, 51e61.
lint, M., Greshake, B., Bandow, C., Ro € mbke, J., Schmidt, P.A., Ba Schmitt, I., 2013. Illumina metabarcoding of a soil fungal community. Soil Biology and Biochemistry 65, 128e132. € mmert, A.K., Dammann, U., Aust, H.J., Strack, D., Schulz, B., Ro 1999. The endophyte-host interaction: a balanced antagonism? Mycological Research 103, 1275e1283. Selosse, M.A., Martos, F., Perry, B.A., Padamsee, M., Roy, M., Pailler, T., 2010. Saprotrophic fungal mycorrhizal symbionts in achlorophyllous orchids: finding treasures among the ‘molecular scraps’? Plant Signaling & Behavior 5, 349. Shinohara, M.L., LoBuglio, K.F., Rogers, S.O., 1996. Group-I intron family in the nuclear ribosomal RNA small subunit genes of Cenococcum geophilum isolates. Current Genetics 29, 377e387. Shinohara, M.L., LoBuglio, K.F., Rogers, S.O., 1999. Comparison of ribosomal DNA ITS regions among geographic isolates of Cenococcum geophilum. Current Genetics 35, 527e535. Summerbell, R.C., 2005. Root endophyte and mycorrhizosphere fungi of black spruce, Picea mariana, in a boreal forest habitat: influence of site factors on fungal distributions. Studies in Mycology 53, 121e145. € rtel, K., Jairus, T., Gates, G., Po ~ ldmaa, K., Tedersoo, L., Pa Tamm, H., 2009. Ascomycetes associated with ectomycorrhizas: molecular diversity and ecology with particular reference to the Helotiales. Environmental Microbiology 11, 3166e3178. € , A.M., Ruotsalainen, A.L., 2013. Tejesvi, M.V., Sauvola, T., Pirttila Neighboring Deschampsia flexuosa and Trientalis europaea harbor contrasting root fungal endophytic communities. Mycorrhiza 23, 1e10. Thorn, R.G., Reddy, C.A., Harris, D., Paul, E.A., 1996. Isolation of saprophytic basidiomycetes from soil. Applied and Environmental Microbiology 62, 4288e4292. Toju, H., Sato, H., Tanabe, A.S., 2014. Diversity and spatial structure of belowground plantefungal Symbiosis in a mixed subtropical sorest of ectomycorrhizal and arbuscular mycorrhizal plants. PLoS One 9, e86566. Toju, H., Yamamoto, S., Sato, H., Tanabe, A.S., Gilbert, G.S., Kadowaki, K., 2013a. Community composition of rootassociated fungi in a Quercus-dominated temperate forest: “codominance” of mycorrhizal and root-endophytic fungi. Ecology and Evolution 3, 1281e1293.

95

Toju, H., Yamamoto, S., Sato, H., Tanabe, A.S., 2013b. Sharing of diverse mycorrhizal and root-endophytic fungi among plant species in an oak-dominated cool-temperate forest. PLoS One 8, e78248. Upson, R., Read, D.J., Newsham, K.K., 2009. Nitrogen form influences the response of Deschampsia antarctica to dark septate root endophytes. Mycorrhiza 20, 1e11. Urban, A., Puschenreiter, M., Strauss, J., Gorfer, M., 2008. Diversity and structure of ectomycorrhizal and co-associated fungal communities in a serpentine soil. Mycorrhiza 18, 339e354. Verkley, G.J.M., 1999. A monograph of Pezicula and its anamorphs. Studies in Mycology 44, 1e176. Villarreal-Ruiz, L., Anderson, I.C., Alexander, I.J., 2004. Interaction between an isolate from the Hymenoscyphus ericae aggregate and roots of Pinus and Vaccinium. New Phytologist 164, 183e192. ic , S., Bahor, E., Regvar, M., Vosa
tka, M., Vohn
ık, M., Lukanc Vodnik, D., 2003. Inoculation of Rhododendron cv. Belle-Heller with two strains of Phialocephala fortinii in two different substrates. Folia Geobotanica 38, 191e200.
, T., Bruzone, M.C., Kohout, P., Vohn
ık, M., Mrnka, L., Luke sova Fehrer, J., 2013. The cultivable endophytic community of Norway spruce ectomycorrhizas from microhabitats lacking ericaceous hosts is dominated by ericoid mycorrhizal Meliniomyces variabilis. Fungal Ecology 6, 281e292. Vr alstad, T., Fossheim, T., Schumacher, T., 2000. Piceirhiza bicolorata e the ectomycorrhizal expression of the Hymenoscyphus ericae aggregate? New Phytologist 145, 549e563. Wang, Z., Johnston, P.R., Takamatsu, S., Spatafora, J.W., Hibbett, D.S., 2006. Toward a phylogenetic classification of the Leotiomycetes based on rDNA data. Mycologia 98, 1065e1075. Weishampel, P.A., Bedford, B.L., 2006. Wetland dicots and monocots differ in colonization by arbuscular mycorrhizal fungi and dark septate endophytes. Mycorrhiza 16, 495e502. White, T.J., Bruns, T., Lee, S., Taylor, J., 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: Innis, M.A., Gelfand, D.H., Sninsky, J.J., White, T.J. (Eds.), PCR Protocols: a Guide to Methods and Applications. Academic Press, New York, pp. 315e322. Wright, S.H.A., Berch, S.M., Berbee, M.L., 2009. The effect of fertilization on the below-ground diversity and community composition of ectomycorrhizal fungi associated with western hemlock (Tsuga heterophylla). Mycorrhiza 19, 267e276. Wu, B., Nara, K., Hogetsu, T., 1999. Competition between ectomycorrhizal fungi colonizing Pinus densiflora. Mycorrhiza 9, 151e159. Yamamoto, S., Sato, H., Tanabe, A.S., Hidaka, A., Kadowaki, K., Toju, H., 2014. Spatial segregation and aggregation of ectomycorrhizal and root-endophytic fungi in the seedlings of two Quercus species. Plos One 9, e96363. Yuan, Z.L., Zhang, C.L., Lin, F.C., Kubicek, C.P., 2010. Identity, diversity, and molecular phylogeny of the endophytic mycobiota in the roots of rare wild rice (Oryza granulate) from a nature reserve in Yunnan, China. Applied and Environmental Microbiology 76, 1642e1652.