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. 2022 Mar 25;3:860777. doi: 10.3389/ffunb.2022.860777

Molecular-Based Diversity Studies and Field Surveys Are Not Mutually Exclusive: On the Importance of Integrated Methodologies in Mycological Research

Jonathan Cazabonne 1,*,†, Lachlan Bartrop 2, Glen Dierickx 3,4,†, Yusufjon Gafforov 5,6,7,†, Tina A Hofmann 8,†, Thomas E Martin 9,†, Meike Piepenbring 10,†, Mauro Rivas-Ferreiro 11,†, Danny Haelewaters 3,8,9,12,*,†
PMCID: PMC10512293  PMID: 37746218

Abstract

Understanding and describing the diversity of living organisms is a great challenge. Fungi have for a long time been, and unfortunately still are, underestimated when it comes to taxonomic research. The foundations were laid by the first mycologists through field observations. These important fundamental works have been and remain vital reference works. Nevertheless, a non-negligible part of the studied funga escaped their attention. Thanks to modern developments in molecular techniques, the study of fungal diversity has been revolutionized in terms of tools and knowledge. Despite a number of disadvantages inherent to these techniques, traditional field-based inventory work has been increasingly superseded and neglected. This perspective aims to demonstrate the central importance of field-based research in fungal diversity studies, and encourages researchers not to be blinded by the sole use of molecular methods.

Keywords: fieldwork, taxonomy, monitoring, fungal diversity, fungal conservation, genetics

Introduction

  We must collect, collect, and collect.

—Richard P. Korf (Reinventing taxonomy, 2005)

When we want to understand ecosystems and how they work, fungi are a keystone group. They contribute to many important ecological processes, as nutrient recyclers, dominant decomposers, regulators of the natural environment, and mutualists (Dighton, 2003; Bardgett and van der Putten, 2014; Powell and Rillig, 2018), and are indispensable elements in the composition and functioning of ecosystems (Christensen, 1993).

Fungi are found in practically all ecosystems and their predominance facilitates interconnectivity with other groups of organisms, constituting a considerable diversity of associations (Hock, 2012). Most of these are poorly studied, such as certain endophytic associations with plants (Dastogeer and Wylie, 2017), and many mysteries remain to be discovered regarding particular associations, including insect–fungal (Vega and Blackwell, 2005; Haelewaters et al., 2021a), algal–fungal (Hawksworth, 1988), bacterial–fungal (Deveau et al., 2018), and bat–fungal associations (Cunha et al., 2020). Some of them result in tripartite (e.g., Cardoza et al., 2006; Afkhami and Stinchcombe, 2016; Haelewaters et al., 2018) or even quadripartite (e.g., Currie, 2000; Piepenbring, 2015) interactions. Many associations are yet to be discovered, since only 2–6% of the estimated 2.2–6 million species are formally described (Taylor et al., 2014; Hibbett et al., 2016; Hawksworth and Lücking, 2017).

Due to “the magnitude of fungal diversity” (fide Hawksworth, 2001) and the large number of existing threats facing them, fungi should be an important conservation priority (Moore et al., 2001; Heilmann-Clausen et al., 2014). Paradoxically, the fungal kingdom has only very recently received a slight interest in international or national programs for the protection and conservation of biodiversity (Velázquez et al., 2010; Sadiković and Kuštera, 2013), especially when it comes to little-studied morphological groups such as microfungi (Arnolds, 2001; Gonçalves et al., 2021). To date, only 0.0091 to 0.025% of the estimated fungal diversity have an assessed conservation status (IUCN, 2022). Growing awareness of the lack of consideration and equivalence toward the fungal kingdom has prompted the launch of a global initiative to make fungi one of the priorities within conservation and agricultural policy frameworks: the FF&F proposal, Fauna, Flora, and Funga (Kuhar et al., 2018).

These initiatives, combined with a renewed interest in fungi transcending the boundaries of the scientific community (Irga et al., 2018), have raised a broader awareness of the knowledge gap in worldwide fungal biodiversity. Studying fungal biodiversity introduces considerable challenges in terms of ecological, taxonomic, systematic, and phylogenetic knowledge, and of fungal protection, conservation, and enhancement (Hibbett et al., 2016). Indeed, knowing what one is protecting allows one to protect it better—an important challenge considering the complexity of fungal identification and ongoing conflicts regarding species concepts (Lücking et al., 2020).

Methods for studying fungal biodiversity have evolved dramatically since mycology emerged as a scientific discipline. From an initial focus on field inventories and macroscopic observations, mycologists progressed to examinations of cellular structures, made possible by the advent of light microscopy. Electron microscopy later allowed for the analysis of ultrastructural characteristics, and, in recent decades, DNA characteristics can be evaluated due to the development of molecular techniques such as high-throughput sequencing (Schmit and Lodge, 2005; Oberwinkler, 2018). With increasingly complex methods and ever-growing quantities of data, there has been a trend for mycologists to become more specialized in dealing with particular subfields such as cytology, physiology, species delimitation, phylogenetics, phylogenomics, comparative genomics, ecology, et cetera. As a result, traditional field inventories, morphological investigation for identification, and alpha taxonomy are frequently neglected in favor of molecular methods and sequence data (Drew, 2011; Ríos-Saldaña et al., 2018).

The Molecular Era: New Insights in Fungal Diversity

Taxonomic investigations of fungi based on morphological features of sexual and/or asexual states (teleomorph, anamorph) were challenging up until the early 2000s, because anamorph–teleomorph connections were often difficult to assess and many morphological characteristics are prone to convergency or are indistinguishable (e.g., yeasts, conidiogenesis). Taxa were classified within frameworks that were useful for that time but not necessarily phylogenetically informative—as was later discovered (e.g., Taylor et al., 2000). For example, the red-pigmented yeasts were traditionally grouped into Rhodotorula and Sporobolomyces, two asexual genera that are highly polyphyletic in their original sense, with many species having been combined in other genera (Fell et al., 2000; Aime et al., 2006; Bauer et al., 2006; Haelewaters et al., 2021b). Cryptic (i.e., morphologically identical) species can only be differentiated by a molecular (DNA) approach (Lücking et al., 2014).

Important developments of molecular methods have allowed fundamental advances in many fields of mycology (Morange et al., 1998). DNA barcoding (Kress and Erickson, 2012) has emerged as a tool to help differentiate and identify fungal collections, and may reveal morphologically and ecologically unpredictable systematic relationships (e.g., Gómez-Zapata et al., 2021). It has become the norm in taxonomy and phylogenetic studies, and is also being used in other subfields such as ethnomycology (Teke et al., 2018) and paleomycology (Bernicchia et al., 2006). Open-access sequence databases such as NCBI GenBank (https://www.ncbi.nlm.nih.gov/genbank/) have allowed for quick and easy identifications of sequenced specimens based on multiple molecular markers. By using base-pair sequences of DNA as characteristics to discriminate species and individuals, aided by high throughput sequencing techniques (e.g., Edwards et al., 2006), molecular approaches have become a vital component of studies in fungal diversity (e.g., Anderson and Cairney, 2004; Dentinger et al., 2016; Gafforov et al., 2017, 2020; Haelewaters et al., 2018; Dirks and Russell, 2020), biology (Oliver and Schweizer, 1999; Osiewacz, 2002), evolution (Bruns et al., 1991; Berbee and Taylor, 2001, 2010; Fraser and Heitman, 2004; Spatafora et al., 2018; Butler et al., 2021), biogeography (Peay and Matheny, 2016; Wang et al., 2021), and ecology (Weiss et al., 2004; Peay et al., 2008; Peay, 2014; van der Linde et al., 2018).

Metagenomics is the analysis of the collective genomes of organisms in their natural environment independent of specimens (Nilsson et al., 2019). The use of environmental DNA (eDNA) has revealed new branches of the Fungal Tree of Life (Tedersoo et al., 2014; Khan et al., 2020) and presented evidence of cryptic fungi in many environments (Jones et al., 2011; Baeza et al., 2017; Léveillé-Bourret et al., 2021; Runnel et al., 2021). The use of eDNA in biodiversity studies has made it possible to obtain new understandings of hitherto hidden fungal diversity within any given ecosystem. Without new mass sequencing technologies, the discovery of fungal communities present in air (Rosa et al., 2020), snow (Penton et al., 2013; Rosa et al., 2020), aquatic ecosystems (Grossart et al., 2019), biotic substrates (e.g., plants, insects, animals), and even abiotic substrates (e.g., rocks Bjelland and Ekman, 2005; Egidi et al., 2014; Selbmann et al., 2014; Liu et al., 2021) would not have been possible.

In addition to being crucial to basic fungal research, molecular tools also contribute to applied fungal research, as in forest management (Glaeser and Lindner, 2011; Stewart et al., 2018; Hagge et al., 2019; Purahong et al., 2019; Tomao et al., 2020) or ecosystem restoration programs (Nuñez et al., 2021). Other questions have recently been re-evaluated thanks to molecular techniques, such as the impact of natural (Vargas-Gastélum et al., 2015; Pérez-Izquierdo et al., 2020) and anthropogenic (Borriello et al., 2012; Lienhard et al., 2014; Tomao et al., 2020; Kim et al., 2021) disturbances on fungal diversity. Through the detection of endangered species (Gordon and van Norman, 2015), molecular methods have also enabled a new appreciation of fungal conservation priorities (Geml et al., 2014).

There now is a common consensus that human activities and their predicted intensification in coming decades are severe enough to represent a sixth ‘Anthropocene' mass-extinction event (Barnosky et al., 2011; Ceballos et al., 2015; IPBES, 2019). Associated biodiversity declines will include the fungi specific to the affected habitats (Antonelli et al., 2020), hence the urgent need to identify these at-risk species swiftly, so actions can be taken to protect them.

Limitations of Modern Methods

Despite recent improvements in molecular biology, important limitations must be taken into account. The identification of fungi based solely on sequence analysis presents great risks and can lead to false identification (Haelewaters et al., 2018; Hofstetter et al., 2019) and erroneous ecological conclusions (Wutkowska et al., 2019) including under- or overrepresentation of fungal groups in biodiversity surveys (George et al., 2019). Defining species boundaries accurately is crucial for diversity estimations as it determines whether organisms are members of the same phylogenetic group. Notwithstanding, many fungi are only known from metabarcoding studies (Wu et al., 2019) and cannot be accommodated within the current framework of fungal nomenclature due to lack of physical specimens or cultures that can serve as types (Ryberg and Nilsson, 2018). The valid publication of new names (sensu Turland et al., 2018) recommends specimen-based molecular methods. However, the cultivation approach favors rapidly growing fungi (e.g., Penicillium) and excludes biotrophic species (Zhang et al., 2010; Cordier et al., 2012). Methods for isolation can also skew diversity estimates, e.g., the choice of growing medium.

Indeed, the success of identifying species based on sequences available in GenBank greatly depends on the amount of available reference sequences (Nilsson et al., 2006). No or very few reference sequences are available for many rare and threatened fungal species as well as poorly known fungi in unexplored geographic areas (Piepenbring et al., 2018; Haelewaters et al., 2021c). This has direct consequences on our understanding of fungal diversity and phylogenetic relationships, as topologies are strongly influenced by taxon sampling. An increasing number of sequences not identified to species level or identified as “uncultured fungus” are present in online sequence databases; this, too, makes molecular-based identification problematic (Hibbett et al., 2011; Nilsson et al., 2014). Nevertheless, efforts have been made to maximize the probability of accurate sequence-based identification, e.g., the UNITE database (Abarenkov et al., 2010) and the use of the term “species hypothesis”—a cluster of sequences that share 97–99% similarity (Kõljalg et al., 2013). However, despite these advances, the greatest challenge in fungal identification and barcoding will mostly, but not only, remain of a taxonomic nature (Nilsson et al., 2006).

The success and efficacy of molecular protocols, including DNA extraction, PCR amplification, and sequencing can be questionable, which may lead to partial or poor-quality sequences. DNA extraction remains difficult for certain fungal groups, requiring specialized protocols or modifications of standard kits (Haelewaters et al., 2015; Sundberg et al., 2018; Meswaet et al., 2021). PCR amplification may depend on taxon-specific primers to target universal or secondary (meta-) barcoding markers (Liu et al., 2020; Reynolds et al., 2021) that are selective for certain taxa. In fungal groups where the internal transcribed spacer (ITS) region of the ribosomal RNA gene does not provide sufficient resolution, secondary DNA barcodes need to be used, representing additional inputs of time and cost (Lücking et al., 2020).

In addition to the pitfalls in sequence-based identification and current public sequence databases, homopolymer regions are also problematic. In addition to practical and technical issues, the use of molecular methods may lead to statistical problems, such as the question of the normalization of fungal taxa abundance among samples (McMurdie and Holmes, 2014; Weiss et al., 2017; Lin and Peddada, 2020).

Lack of funding and training is also a major concern. Molecular work is expensive and highly specialized, and requires specific equipment not available for many mycologists (Hibbett et al., 2016). This is a problem especially in remote and understudied areas in the tropics (Haelewaters et al., 2021a,c) and some temperate zones, such as Central Asia (Gafforov, 2017; Cheek et al., 2020; Abdurazakov et al., 2021).

Field-Based Research: The Solution?

Field surveys are one of the oldest and most fundamental techniques for studying biodiversity, through the observation and description of biological organisms living on Earth. Unfortunately, their value as viewed by the scientific community has diminished and today species records are rarely a priority for researchers. A decline in the relative importance of field-based science has been observed in most taxa (Ríos-Saldaña et al., 2018). This may be particularly acute in mycology, perhaps due to the cryptic nature of fungi, resulting in tedious and costly species identifications that require expertise from mycologists, taxonomists, and evolutionary biologists. This is a worrying trend, given field surveys generate essential new information that cannot be obtained through the prism of sequencing, and are ideal instruments for training the next generation.

Taxonomy

Alpha-taxonomy (i.e., taxonomy in its historical sense) establishes taxonomic units based on phenotypic traits (Simpson, 1960). Traditionally, this was achieved mainly by the presentation of monographs and allowed for the study of the variability of informative characteristics. Diagnosable morphological features can be attained rapidly in the field as well as by light microscopy in the lab, and multiple specimens of species warrant insights into morphological variation, which forms the basis of species delimitation (Noordeloos and Antonín, 2008; Kerekes and Desjardin, 2009; Phengsintham et al., 2013). Some characteristics important for the morphological identification of species can only be observed on freshly collected specimens, including the original color, taste, smell, chemical reactions, e.g., in the case of fruiting bodies of Russula spp. (Adamčík et al., 2019), and the organization of oil drops in ascospores (Baral, 1992). Field survey methods (Mueller et al., 2004) allow access to macro- and micro-morphology of a given taxon that is combined with previously or newly generated sequences. This “fusion” between phenotypic and molecular genetic data, referred to as integrative taxonomy (Dayrat, 2005; Pante et al., 2015), is essential for species delimitation and, thus, formal description of taxa new to science (Simões et al., 2013).

Geographic Records of Known Species

Despite a generalized decline in the importance of traditional taxonomic research (Drew, 2011), new records of known species should be properly documented and published. To establish a species concept new to science is easy, but it is more valuable to apply an old name to a new collection, although this takes more time and effort. As a result, the new information is connected to existing data. With every new collection, the knowledge of any given species is improved; this includes infraspecific variation of morphological and molecular characteristics, interaction with other organisms (host range in the case of pathogens), and geographic distribution. A high number of geographic records are required to become aware of changes in distributional areas and general patterns of spatial distribution, which is critical in the light of climate change and for species that are disease agents of humans, animals, or cultivated plants. Good knowledge of geographic distributions is indispensable for phylogeographic studies and forms the basis of assessments for the International Union for Conservation of Nature (IUCN) Red List of Threatened Species and designation of Alliance for Zero Extinction (AZE) sites (https://zeroextinction.org/).

Ecology

Opportunistic field sampling, the most common approach used by citizen scientists (Mueller, 2017), can be of great value with regard to fungal studies, particularly when exploring poorly-studied and remote locations (Rivas-Ferreiro, 2021). Long-term field monitoring, however, has the potential to generate longitudinal datasets necessary to understand the role of fungi in ecosystems. Not only do data from such monitoring studies enable hypothesis-driven research (Korf, 2005), they inform on fungal reproduction (i.e., sexual states), phenology (e.g., Jumbam et al., 2019), life cycles (i.e., methods of spore dispersal), nourishment (i.e., saprophytic, endophytic, etc.), and functional diversity. Ecological monitoring also reveals key information about ecological adaptations, i.e., insects attracted by volatiles (Davis and Landolt, 2013), nematode-trapping structures (Zhang et al., 2013), secotioid basidiomata as adaptations to dry environments (Claridge et al., 2000a,b), the dominance of sporocarps in periods with high precipitation (e.g., López-Quintero et al., 2012; Piepenbring et al., 2015), et cetera.

Traits

In addition to capturing ecological information as outlined above, observations made in the field allow to integrate biotic and abiotic data, resulting in information critical for fungal conservation studies. This includes ecological observations concerning the relevance of, e.g., climate (temperate, tropical), seasonality, weather, environmental factors (pH, water, nutrient availability, substrate, soil characteristics), and habitat, as well as insights concerning dispersal strategies, interactions with other organisms (natural enemies, hosts), and responses to anthropogenic pressures. These data also facilitate the discovery of abiotic and biotic connections between fungal clades and biodiversity more broadly (Kendrick, 2011; Peay et al., 2016). Meta-analysis and ecological studies rely on key spatial data obtained from surveys (Shrestha and Bawa, 2014; Yan et al., 2017) and temporal variations of fungal diversity governed by natural and anthropogenic environmental change can advise conservation efforts (Haelewaters et al., 2021c; Kaishian, 2021).

Cultures

Field surveys allow the isolation of fungal collections from habitats such as plant leaves, fruit skins, and insect guts that are common sources of endophytes, yeasts, and yeast-like fungi, often representing untapped diversity preserved in a metabolically inactive state (Boundy-Mills et al., 2015; Sharma et al., 2019). From here, results obtained by molecular methods can be supplemented with morphological characteristics (including photographs of colonies on distinct culture media; Pitt et al., 1983), spectral analyses (Petisco et al., 2008; Rodrigues et al., 2011), recognition of anamorphic and teleomorphic states (Hawksworth, 2011), and screening for biochemical profiles (Suh et al., 2008).

Fungarium Collections

Specimens collected through field surveys are often deposited in fungaria, after having been slowly dried, packaged, and labeled. Fungarium collections represent a treasure trove of data because they represent traces of past and current species diversity and occurrences (Paton et al., 2020). Even if unidentified, fungarium collections may be useful for future researchers who may investigate morphological or molecular characteristics (Brock et al., 2009; Osmundson et al., 2013; Dentinger et al., 2016). Specimens from field surveys, as a best practice, are accompanied by field notes with information about characteristics that are only present in situ (e.g., hygrophany, smell, substrate) or that are no longer observable after drying (e.g., chemical reactions, colors, texture).

Checklists

The value of checklists resulting from field surveys is increasingly appreciated by many researchers worldwide (Piepenbring et al., 2011, 2020; Haelewaters et al., 2021c). A checklist of fungi for a given area presents information about the history of mycological research in that area, generates data necessary to compare the area with other regions with respect to species diversity, helps to detect and monitor invasive species, and advances taxonomic research with new collections of “forgotten” species or “old names” (Piepenbring et al., 2020). Some researchers have made it a matter of principle to publish fungal species checklists (e.g., Marinho et al., 2018; Diederich and Ertz, 2020; Fryar et al., 2020). This helps fill the knowledge gaps on fungal diversity, particularly in under-sampled regions. Recent funded initiatives have highlighted the critical value of publishing checklists supported with molecular data (e.g., Haelewaters et al., 2018; Gafforov et al., 2020); we highly encourage our mycological colleagues to do the same. Fungal species occurrence data, whether from protected areas or public lands, through amateur or grant-funded, scientific surveys, are critical to science.

Training

Field-based projects can and should bring together researchers spanning junior to senior scientists as well as students and interns from diverse backgrounds. Teaching in the field is an effective way to educate young mycologists. Collaborators in the area may identify interested and talented students who will participate in every part of the research, in addition to local people to be trained as parataxonomists. Such education should go further than collecting fungi; it may include seminar talks by senior participants, presentations by more experienced students, interactions in the field, and laboratory work (Piepenbring and Yorou, 2017).

Unity Makes Strength: Onward Together!

Fungal diversity can only be unraveled and interpreted by combining molecular and organismic approaches that take into account the complexity and wholeness of members of the kingdom. In the field, only a small part of the existing fungal diversity can be seen through above- or belowground spore-bearing structures that are easily observed. A bias exists toward well-known species of “macrofungi” (Gonçalves et al., 2021). Indeed, “microfungi”—e.g., endophytes, epiphytes, arbuscular mycorrhizal fungi, coprophilous fungi—are neglected in fungal diversity and conservation studies. Most fungal diversity resides underground and in living or dead organisms, by means of hyphal networks, most of them unable to produce sporocarps (Halme et al., 2012). The vast majority of fungal taxa studies are only possible by combining molecular and field survey methods to an integrative double-sided focus.

In order to know the funga worldwide, we need field survey projects, particularly in poorly studied areas. Funding programs focusing on fieldwork, capacity building (to provide access to molecular equipment, especially in the tropics and the Global South), and taxonomy are key. An example is the U.S. National Science Foundation Poorly Sampled and Unknown Taxa (PurSUiT) category. In addition, grants should include funds to hire specialized taxonomists to accelerate species identifications. In remote areas lacking mycologists, such as in most of the tropics, Indigenous people may possess ancestral knowledge on fungi. To go further, we must encourage the establishment of field surveys by actively soliciting contributions of local communities. Such integrative parataxonomy can be of great value in terms of knowledge and scientific popularization. In the same vein, integrative taxonomy is directly linked to the use of this powerful combined method. Taxonomy should be based on a combination of molecular phylogenetic, morphological, and ecological data. Fungal biodiversity or ecological studies based on molecular data only, or equally without such molecular sequence data, can be considered as partly “blind.”

In every fungal diversity study, the collect-and-sequence approach (Truong et al., 2017) should be employed as standard. Where these two main methods are employed together, new perspectives can be explored. For example, eDNA and field surveys can be complementary tools when it comes to studying fungal communities and forest management (Yan et al., 2018). Cryptic fungi (e.g., soil fungi, epiphytes, endophytes) that cannot (or are very difficult to) collect or culture on axenic media can be molecularly identified and associated with ecological information (seasonality, spatial distribution) through employing an integrated methodology. We do note that for a few ecosystems, molecular approaches remain our best bet. For example, desert ecosystems, Antarctica, deep sea sediments, and other extreme ecosystems, where sporocarps are never seen and for which culturing techniques are very challenging (i.e., culturing of fungi under high pressure, or under very specific nutrient availability). Likewise, some taxa that we have failed to find in metabarcoding studies and that are unculturable but that can be readily observed and collected in the field, may continue to benefit from “traditional” taxonomic studies.

Integrative methodologies are crucial for improving our understanding and knowledge of global fungal diversity, both from a fundamental and applied research perspective. Indeed, integrative approaches are poised to become even more crucial for fungal conservation as new initiatives arise to protect fungi (Kuhar et al., 2018) and to consider all fungal species in biodiversity goals (Gonçalves et al., 2021). Only a tiny fraction of the total fungal biodiversity on the planet is known, and there is a huge need (i) to improve our understanding of the fundamental role of fungi in the functioning of ecosystems, (ii) to characterize the patterns of the distribution of biodiversity and the dynamics of fungal communities, whether within endangered habitats or not, (iii) to improve taxonomic knowledge, and (iv) to fully integrate the fungi into global conservation networks, such as the IUCN Red List, AZE sites, and Key Biodiversity Area (KBD) designations. Before it is too late.

Data Availability Statement

The original contributions presented in the study are included in the article/supplementary material, further inquiries can be directed to the corresponding author/s.

Author Contributions

JC and DH: conceptualization and writing—original draft and visualization. JC, LB, GD, YG, TAH, TEM, MP, MR-F, and DH: writing—review and editing. All authors have read and agreed to the published version of the manuscript. All authors approved the submitted version.

Funding

This work was supported in part by the Research Foundation—Flanders (Fellowship Fundamental Research 1126221N to GD, financed by the Research Institute for Nature and Forest; Junior Postdoctoral Fellowship 1206620N to DH), Panama's Secretaría Nacional de Ciencia, Tecnología e Innovación (SENACYT, grant number 121-2017-4-EIE17-016 to TAH), the German Federal Ministry of Education and Research (BMBF) (grant number 01DG20015FunTrAf to MP), the Spanish Ministry of Education (Predoctoral Fellowship FPU20/02280 to MR-F), and the U.S. National Science Foundation (DEB-2018098 and DEB-2127290 to DH).

Conflict of Interest

TEM and DH were employed by Operation Wallacea Ltd. The remaining authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Publisher's Note

All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.

Acknowledgments

Claus Bässler (Goethe-Universität Frankfurt am Main) is thanked for critical help with references. We also thank Adriana L. Romero-Olivares (New Mexico State University) for numerous suggestions that improved the manuscript.

References

  1. Abarenkov K., Nilsson R. H., Larsson K. H., Alexander I. J., Eberhardt U., Erland S., et al. (2010). The UNITE database for molecular identification of fungi–recent updates and future perspectives. New Phytol. 186, 281–285. 10.1111/j.1469-8137.2009.03160.x [DOI] [PubMed] [Google Scholar]
  2. Abdurazakov A. A., Bulgakov T. S., Kholmuradova T. N., Gafforov Y. S. (2021). Powdery mildew fungi (Erysiphaceae) of the Fergana Valley (within Uzbekistan): a first annotated checklist. Nov. Sist. Nizs. Rast. 55, 55–78. 10.31111/nsnr/2021.55.1.55 [DOI] [Google Scholar]
  3. Adamčík S., Looney B., Cabon M., Jančovičová S., Adamčíková K., Avis P. G., et al. (2019). The quest for a globally comprehensible Russula language. Fungal Divers. 99, 369–449. 10.1007/s13225-019-00437-2 [DOI] [Google Scholar]
  4. Afkhami M. E., Stinchcombe J. R. (2016). Multiple mutualist effects on genomewide expression in the tripartite association between Medicago truncatula, nitrogen-fixing bacteria and mycorrhizal fungi. Mol. Ecol. 25, 4946–4962. 10.1111/mec.13809 [DOI] [PubMed] [Google Scholar]
  5. Aime M. C., Matheny P. B., Henk D. A., Frieders E. M., Nilsson R. H., Piepenbring M., et al. (2006). An overview of the higher-level classification of Pucciniomycotina based on combined analyses of nuclear large and small subunit rDNA sequences. Mycologia 98, 895–905. 10.1080/15572536.2006.11832619 [DOI] [PubMed] [Google Scholar]
  6. Anderson I. C., Cairney J. W. (2004). Diversity and ecology of soil fungal communities: increased understanding through the application of molecular techniques. Environ. Microbiol. 6, 769–779. 10.1111/j.1462-2920.2004.00675.x [DOI] [PubMed] [Google Scholar]
  7. Antonelli A., Fry C., Smith R. J., Simmonds M. S. J., Kersey P. J., Pritchard H. W., et al. (2020). State of the World's Plants and Fungi 2020. London: Royal Botanic Gardens, Kew. [Google Scholar]
  8. Arnolds E. (2001). The future of fungi in Europe: threats, conservation and management. Brit. Mycol. Soc. Symp. Ser. 22, 64–80. 10.1017/CBO9780511565168.005 [DOI] [Google Scholar]
  9. Baeza M., Barahona S., Alcaíno J., Cifuentes V. (2017). Amplicon-metagenomic analysis of fungi from antarctic terrestrial habitats. Front. Microbiol. 8, 2235. 10.3389/fmicb.2017.02235 [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Baral H.-O. (1992). Vital versus herbarium taxonomy: morphological differences between living and dead cells of ascomycetes, and their taxonomic implications. Mycotaxon 44, 333–390. [Google Scholar]
  11. Bardgett R. D., van der Putten W. H. (2014). Belowground biodiversity and ecosystem functioning. Nature 515, 505–511. 10.1038/nature13855 [DOI] [PubMed] [Google Scholar]
  12. Barnosky A. D., Matzke N., Tomiya S., Wogan G. O. U., Swartz B., Quental T. B., et al. (2011). Has the Earth's sixth mass extinction already arrived? Nature 471, 51–57. 10.1038/nature09678 [DOI] [PubMed] [Google Scholar]
  13. Bauer R., Begerow D., Sampaio J. P., Wei,β M., Oberwinkler F. (2006). The simple-septate basidiomycetes: a synopsis. Mycol. Prog. 5, 41–66. 10.1007/s11557-006-0502-0 [DOI] [Google Scholar]
  14. Berbee M. L., Taylor J. W. (2001). “Fungal molecular evolution: gene trees and geologic time,” in Systematics and Evolution, eds. McLaughlin D. J., McLaughlin E. G., Lemke P. A. (New York, NY: Springer–Verlag; ), 229–245. 10.1007/978-3-662-10189-6_10 [DOI] [Google Scholar]
  15. Berbee M. L., Taylor J. W. (2010). Dating the molecular clock in fungi – how close are we? Fungal Biol. Rev. 24, 1–16. 10.1016/j.fbr.2010.03.001 [DOI] [Google Scholar]
  16. Bernicchia A., Fugazzola M. A., Gemelli V., Mantovani B., Lucchetti A., Cesari M., et al. (2006). DNA recovered and sequenced from an almost 7000 y-old Neolithic polypore, Daedaleopsis tricolor. Mycol. Res. 110, 14–17. 10.1016/j.mycres.2005.09.012 [DOI] [PubMed] [Google Scholar]
  17. Bjelland T., Ekman S. (2005). Fungal diversity in rock beneath a crustose lichen as revealed by molecular markers. Microb. Ecol. 49, 598–603. 10.1007/s00248-004-0101-z [DOI] [PubMed] [Google Scholar]
  18. Borriello R., Lumini E., Girlanda M., Bonfante P., Bianciotto V. (2012). Effects of different management practices on arbuscular mycorrhizal fungal diversity in maize fields by a molecular approach. Biol. Fertil. Soils 48, 911–922. 10.1007/s00374-012-0683-4 [DOI] [Google Scholar]
  19. Boundy-Mills K., Hess M., Bennett A. R., Ryan M., Kang S., Nobles D., et al. (2015). The United States Culture Collection Network (USCCN): enhancing microbial genomics research through living microbe culture collections. Appl. Environ. Microbiol. 81, 5671–5674. 10.1128/AEM.01176-15 [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Brock P. M., Döring H., Bidartondo M. I. (2009). How to know unknown fungi: the role of a herbarium. New Phytol. 181, 719–724. 10.1111/j.1469-8137.2008.02703.x [DOI] [PubMed] [Google Scholar]
  21. Bruns T. D., White T. J., Taylor J. W. (1991). Fungal molecular systematics. Annu. Rev. Ecol. Syst. 22, 525–564. 10.1146/annurev.es.22.110191.002521 [DOI] [Google Scholar]
  22. Butler G., Heitman J., Idnurm A., James T. Y. (2021). On a special collection in MMBR on sex in fungi: molecular mechanisms and evolutionary implications. Microbiol. Mol. Biol. Rev. 85, e00094–e00021. 10.1128/MMBR.00094-21 [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Cardoza Y. J., Paskewitz S., Raffa K. F. (2006). Travelling through time and space on wings of beetles: a tripartite insect-fungi-nematode association. Symbiosis 41, 71–79. [Google Scholar]
  24. Ceballos G., Ehrlich P. R., Barnosky A. D., García A., Pringle R. M., Palmer T. M. (2015). Accelerated modern human-induced species losses: entering the sixth mass extinction. Sci. Adv. 1, e1400253. 10.1126/sciadv.1400253 [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Cheek M., Nic Lughadha E., Kirk P., Lindon H., Carretero J., Looney B., et al. (2020). New discoveries: plants and fungi. Plants People Planet 2, 371–388. 10.1002/ppp3.10148 [DOI] [Google Scholar]
  26. Christensen M. (1993). The fungal community: its organization and role in the ecosystem. BioScience 43, 787–789. 10.2307/1312325 [DOI] [Google Scholar]
  27. Claridge A. W., Barry S. C., Cork S. J., Trappe J. M. (2000a). Diversity and habitat relationships of hypogeous fungi. II. factors influencing the occurrence and number of taxa. Biodivers. Conserv. 9, 175–199. 10.1023/A:1008962711138 [DOI] [Google Scholar]
  28. Claridge A. W., Cork S. J., Trappe J. M. (2000b). Diversity and habitat relationships of hypogeous fungi. I. study design, sampling techniques and general survey results. Biodivers. Conserv. 9, 151–173. 10.1023/A:1008941906441 [DOI] [Google Scholar]
  29. Cordier T., Robin C., Capdevielle X., Fabreguettes O., Desprez-Loustau M.-L., Vacher C. (2012). The composition of phyllosphere fungal assemblages of European beech (Fagus sylvatica) varies significantly along an elevation gradient. New Phytol. 196, 510–519. 10.1111/j.1469-8137.2012.04284.x [DOI] [PubMed] [Google Scholar]
  30. Cunha A. O., Bezerra J. D. P., Oliveira T. G. L., Barbier E., Bernard E., Machado A. R., et al. (2020). Living in the dark: bat caves as hotspots of fungal diversity. PLoS ONE 15, e0243494. 10.1371/journal.pone.0243494 [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Currie C. R. (2000). The ecology and evolution of a quadripartite symbiosis, examining the interactions among Attine ants, fungi, and actinomycetes. Ph.D. thesis. Toronto, ON: University of Toronto. [Google Scholar]
  32. Dastogeer K. M., Wylie S. J. (2017). “Plant–fungi association: role of fungal endophytes in improving plant tolerance to water stress,” in Plant–Microbe Interactions in Agro-Ecological Perspectives, D. P. eds. Singh, Singh H. B, Prabha R. (Singapore: Springer Nature; ), 143–159. 10.1007/978-981-10-5813-4_8 [DOI] [Google Scholar]
  33. Davis T. S., Landolt P. J. (2013). A survey of insect assemblages responding to volatiles from a ubiquitous fungus in an agricultural landscape. J. Chem. Ecol. 39, 860–868. 10.1007/s10886-013-0278-z [DOI] [PubMed] [Google Scholar]
  34. Dayrat B. (2005). Towards integrative taxonomy. Biol. J. Linn. Soc. 85, 407–415. 10.1111/j.1095-8312.2005.00503.x [DOI] [Google Scholar]
  35. Dentinger B. T. M., Gaya E., O'Brien H., Suz L. M., Lachlan R., Díaz-Valderrama J. R., et al. (2016). Tales from the crypt: genome mining from fungarium specimens improves resolution of the mushroom tree of life. Biol. J. Linn. Soc. 117, 11–32. 10.1111/bij.12553 [DOI] [Google Scholar]
  36. Deveau A., Bonito G., Uehling J., Paoletti M., Becker M., Bindschedler S., et al. (2018). Bacterial–fungal interactions: ecology, mechanisms and challenges. FEMS Microbiol. Rev. 42, 335–352. 10.1093/femsre/fuy008 [DOI] [PubMed] [Google Scholar]
  37. Diederich P., Ertz D. (2020). First checklist of lichens and lichenicolous fungi from Mauritius, with phylogenetic analyses and descriptions of new taxa. Plant Fungal Syst. 65, 13–75. 10.35535/pfsyst-2020-0003 [DOI] [Google Scholar]
  38. Dighton J. (2003). Fungi in Ecosystem Processes. New York, NY: Marcel Dekker, Inc. 10.1201/9780203911440 [DOI] [Google Scholar]
  39. Dirks A., Russell S. D. (2020). DNA barcoding of macrofungi from the 2018 Smith Foray: new fungal records for Wisconsin and the United States of America. Great Bot. 59, 191–201. [Google Scholar]
  40. Drew L. W. (2011). Are we losing the science of taxonomy? As need grows, numbers and training are failing to keep up. BioScience 61, 942–946. 10.1525/bio.2011.61.12.4 [DOI] [Google Scholar]
  41. Edwards R. A., Rodríguez-Brito B., Wegley L., Haynes M., Breitbart M., Peterson D. M., et al. (2006). Using pyrosequencing to shed light on deep mine microbial ecology. BMC Genom. 7, 57. 10.1186/1471-2164-7-57 [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Egidi E., de Hoog G. S., Isola D., Onofri S., Quaedvlieg W., de Vries M., et al. (2014). Phylogeny and taxonomy of meristematic rock-inhabiting black fungi in the Dothideomycetes based on multi-locus phylogenies. Fungal Divers. 65, 127–165. 10.1007/s13225-013-0277-y [DOI] [Google Scholar]
  43. Fell J. W., Boekhout T., Fonseca A., Scorzetti G., Statzell-Tallman A. (2000). Biodiversity and systematics of basidiomycetous yeasts as determined by large-subunit rDNA D1/D2 domain sequence analysis. Int. J. Syst. Evol. Microbiol. 50, 1351–1371. 10.1099/00207713-50-3-1351 [DOI] [PubMed] [Google Scholar]
  44. Fraser J. A., Heitman J. (2004). Evolution of fungal sex chromosomes. Mol. Microbiol. 51, 299–306. 10.1046/j.1365-2958.2003.03874.x [DOI] [PubMed] [Google Scholar]
  45. Fryar S. C., Hyde K. D., Catcheside D. E. (2020). A checklist of marine fungi from Australia. Mycotaxon 135, 465. 10.5248/135.465 [DOI] [Google Scholar]
  46. Gafforov Y. (2017). A preliminary checklist of ascomycetous microfungi from Southern Uzbekistan. Mycosphere 8, 660–696. 10.5943/mycosphere/8/4/12 [DOI] [Google Scholar]
  47. Gafforov Y., Ordynets A., Langer E., Yarasheva M., Gugliotta A. M., Schigel D., et al. (2020). Species diversity with comprehensive annotations of wood- inhabiting poroid and corticioid fungi in Uzbekistan. Front. Microbiol. 11, 598321. 10.3389/fmicb.2020.598321 [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Gafforov Y., Riebesehl J., Ordynets A., Langer E., Yarasheva M., Ghobad-Nejhad M., et al. (2017). Hyphodontia (Hymenochaetales, Basidiomycota) and similar taxa from Central Asia. Botany 95, 1041–1056. 10.1139/cjb-2017-0115 [DOI] [Google Scholar]
  49. Geml J., Gravendeel B., van der Gaag K. J., Neilen M., Lammers Y., Raes N., et al. (2014). The contribution of DNA metabarcoding to fungal conservation: diversity assessment, habitat partitioning and mapping red-listed fungi in protected coastal Salix repens communities in the Netherlands. PLoS ONE 9, e99852. 10.1371/journal.pone.0099852 [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. George P. B., Creer S., Griffiths R. I., Emmett B. A., Robinson D. A., Jones D. L. (2019). Primer and database choice affect fungal functional but not biological diversity findings in a national soil survey. Front. Environ. Sci. 7, 173. 10.3389/fenvs.2019.00173 [DOI] [Google Scholar]
  51. Glaeser J. A., Lindner D. L. (2011). Use of fungal biosystematics and molecular genetics in detection and identification of wood-decay fungi for improved forest management. For. Pathol. 41, 341–348. 10.1111/j.1439-0329.2010.00681.x [DOI] [Google Scholar]
  52. Gómez-Zapata P. A., Haelewaters D., Quijada L., Pfister D. H., Aime M. C. (2021). Notes on Trochila (Ascomycota, Leotiomycetes), with new species and combinations. MycoKeys 78, 21–47. 10.3897/mycokeys.78.62046 [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Gonçalves S. C., Haelewaters D., Furci G., Mueller G. M. (2021). Include all fungi in biodiversity goals. Science 373, 403. 10.1126/science.abk1312 [DOI] [PubMed] [Google Scholar]
  54. Gordon M., van Norman K. (2015). Bridgeoporus nobilissimus is much more abundant than indicated by the presence of basidiocarps in forest stands. North Am. Fungi 10, 1–28. [Google Scholar]
  55. Grossart H.-P., Van den Wyngaert S., Kagami M., Wurzbacher C., Cunliffe M., Rojas-Jimenez K. (2019). Fungi in aquatic ecosystems. Nat. Rev. Microbiol. 17, 339–354. 10.1038/s41579-019-0175-8 [DOI] [PubMed] [Google Scholar]
  56. Haelewaters D., Dirks A. C., Kappler L. A., Mitchell J. K., Quijada L., Vandegrift R., et al. (2018). A preliminary checklist of fungi at the Boston Harbor islands. Northeast. Nat. 25, 45–76. 10.1656/045.025.s904 [DOI] [Google Scholar]
  57. Haelewaters D., Gorczak M., Kaishian P., De Kesel A., Blackwell M. (2021a). “Laboulbeniomycetes, enigmatic fungi with a turbulent taxonomic history,” in Encyclopedia of Mycology, ed. Zaragoza Ó. (Oxford: Elsevier; ), 263–283. 10.1016/B978-0-12-819990-9.00049-4 [DOI] [Google Scholar]
  58. Haelewaters D., Gorczak M., Pfliegler W. P., Tartally A., Tischer M., Wrzosek M., et al. (2015). Bringing Laboulbeniales to the 21st century: enhanced techniques for extraction and PCR amplification of DNA from minute ectoparasitic fungi. IMA Fungus 6, 363–372. 10.5598/imafungus.2015.06.02.08 [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Haelewaters D., Peterson R. A., Nevalainen H., Aime M. C. (2021b). Inopinatum lactosum gen. & comb. nov., the first yeast-like fungus in Leotiomycetes. Int. J. Syst. Evol. Microbiol. 71, 4862. 10.1099/ijsem.0.004862 [DOI] [PubMed] [Google Scholar]
  60. Haelewaters D., Schoutteten N., Medina-van Berkum P., Martin T. E., Verbeken A., Aime M. C. (2021c). Pioneering a fungal inventory at Cusuco National Park, Honduras. J. Mesoam. Biol. 1, 111–131. [Google Scholar]
  61. Hagge J., Bässler C., Gruppe A., Hoppe B., Kellner H., Krah F.-S., et al. (2019). Bark coverage shifts assembly processes of microbial decomposer communities in dead wood. Proc. R. Soc. B Biol. Sci. 286, 20191744. 10.1098/rspb.2019.1744 [DOI] [PMC free article] [PubMed] [Google Scholar]
  62. Halme P., Heilmann-Clausen J., Räm,ä T., Kosonen T., Kunttu P. (2012). Monitoring fungal biodiversity – towards an integrated approach. Fungal Ecol. 5, 750–758. 10.1016/j.funeco.2012.05.00521040971 [DOI] [Google Scholar]
  63. Hawksworth D. L. (1988). The variety of fungal-algal symbioses, their evolutionary significance, and the nature of lichens. Bot. J. Linn. Soc. 96, 3–20. 10.1111/j.1095-8339.1988.tb00623.x [DOI] [Google Scholar]
  64. Hawksworth D. L. (2001). The magnitude of fungal diversity: the 1.5 million species estimate revisited. Mycol. Res. 105, 1422–1432. 10.1017/S095375620100472528752818 [DOI] [Google Scholar]
  65. Hawksworth D. L. (2011). Naming Aspergillus species: progress towards one name for each species. Med. Mycol. 49, S70–S76. 10.3109/13693786.2010.504753 [DOI] [PubMed] [Google Scholar]
  66. Hawksworth D. L., Lücking R. (2017). Fungal diversity revisited: 2.2 to 3.8 million species. Microbiol. Spectr. 5, 1–17. 10.1128/microbiolspec.FUNK-0052-2016 [DOI] [PMC free article] [PubMed] [Google Scholar]
  67. Heilmann-Clausen J., Barron E. S., Boddy L., Dahlberg A., Griffith G. W., Nordén J., et al. (2014). A fungal perspective on conservation biology. Conserv. Biol. 29, 61–68. 10.1111/cobi.12388 [DOI] [PubMed] [Google Scholar]
  68. Hibbett D., Abarenkov K., Kõljalg U., Öpik M., Chai B., Cole J., et al. (2016). Sequence-based classification and identification of Fungi. Mycologia 108, 1049–1068. 10.3852/16-130 [DOI] [PubMed] [Google Scholar]
  69. Hibbett D. S., Ohman A., Glotzer D., Nuhn M., Kirk P., Nilsson R. H. (2011). Progress in molecular and morphological taxon discovery in Fungi and options for formal classification of environmental sequences. Fungal Biol. Rev. 25, 38–47. 10.1016/j.fbr.2011.01.001 [DOI] [Google Scholar]
  70. Hock B. (2012). Fungal Associations. Berlin: Springer–Verlag. 10.1007/978-3-642-30826-0 [DOI] [Google Scholar]
  71. Hofstetter V., Buyck B., Eyssartier G., Schnee S., Gindro K. (2019). The unbearable lightness of sequenced-based identification. Fungal Divers. 96, 243–284. 10.1007/s13225-019-00428-3 [DOI] [Google Scholar]
  72. IPBES (2019). Summary for policymakers of the global assessment report on biodiversity and ecosystem services of the Intergovernmental Science-Policy Platform on Biodiversity and Ecosystem Services. Bonn: Secretariat of the Intergovernmental Science-Policy Platform on Biodiversity and Ecosystem Services. [Google Scholar]
  73. Irga P. J., Barker K., Torpy F. R. (2018). Conservation mycology in Australia and the potential role of citizen science. Conserv. Biol. 32, 1031–1037. 10.1111/cobi.13121 [DOI] [PubMed] [Google Scholar]
  74. IUCN (2022). The IUCN Red List of Threatened Species. Version 2021-3. Available online at: https://www.iucnredlist.org (accessed January 20, 2022).
  75. Jones M. D., Forn I., Gadelha C., Egan M. J., Bass D., Massana R., et al. (2011). Discovery of novel intermediate forms redefines the fungal tree of life. Nature 474, 200–203. 10.1038/nature09984 [DOI] [PubMed] [Google Scholar]
  76. Jumbam B., Haelewaters D., Koch R. A., Dentinger B. T., Henkel T. W., Aime M. C. (2019). A new and unusual species of Hericium (Basidiomycota: Russulales, Hericiaceae) from the Dja Biosphere Reserve, Cameroon. Mycol. Prog. 18, 1253–1262. 10.1007/s11557-019-01530-1 [DOI] [Google Scholar]
  77. Kaishian P. J. (2021). Insects and their Laboulbeniales (Ascomycota, Fungi) of Lake Eustis and Emeralda Marsh Conservation Area: a case study on urbanization and diversity. Ecol. Evol. 11, 16618–16633. 10.1002/ece3.8246 [DOI] [PMC free article] [PubMed] [Google Scholar]
  78. Kendrick B. (2011). “Fungi: ecological importance and impact on humans,” in eLS (Hoboken, NJ: Wiley; ). 10.1002/9780470015902.a0000369.pub2 [DOI] [Google Scholar]
  79. Kerekes J., Desjardin D. E. (2009). A monograph of the genera Crinipellis and Moniliophthora from Southeast Asia including a molecular phylogeny of the nrITS region. Fungal Divers. 37, 101–152. [Google Scholar]
  80. Khan F., Kluting K., Tångrot J., Urbina H., Ammunet T., Eshghi Sahraei S., et al. (2020). Naming the untouchable – environmental sequences and niche partitioning as taxonomical evidence in fungi. IMA Fungus 11, 23. 10.1186/s43008-020-00045-9 [DOI] [PMC free article] [PubMed] [Google Scholar]
  81. Kim S., Axelsson E. P., Girona M. M., Senior J. K. (2021). Continuous-cover forestry maintains soil fungal communities in Norway spruce dominated boreal forests. For. Ecol. Manag. 480, 118659. 10.1016/j.foreco.2020.118659 [DOI] [Google Scholar]
  82. Kõljalg U., Nilsson R. H., Abarenkov K., Tedersoo L., Taylor A. F., Bahram M., et al. (2013). Towards a unified paradigm for sequence-based identification of fungi. Mol. Ecol. 22, 5271–5277. 10.1111/mec.12481 [DOI] [PubMed] [Google Scholar]
  83. Korf R. P. (2005). Reinventing taxonomy: a curmudgeon's view of 250 years of fungal taxonomy, the crisis in biodiversity, and the pitfalls of the phylogenetic age. Mycotaxon 93, 407–415. [Google Scholar]
  84. Kress W. J., Erickson D. L. (2012). DNA barcodes: methods and protocols. Methods Mol. Biol. 858, 3–8. 10.1007/978-1-61779-591-6_1 [DOI] [PubMed] [Google Scholar]
  85. Kuhar F., Furci G., Drechsler-Santos E. R., Pfister D. H. (2018). Delimitation of Funga as a valid term for the diversity of fungal communities: the Fauna, Flora & Funga proposal (FF&F). IMA Fungus 9, A71–A74. 10.1007/BF03449441 [DOI] [Google Scholar]
  86. Léveillé-Bourret É., Eggertson Q., Hambleton S., Starr J. R. (2021). Cryptic diversity and significant cophylogenetic signal detected by DNA barcoding the rust fungi (Pucciniaceae) of Cyperaceae–Juncaceae. J. Syst. Evol. 59, 833–851. 10.1111/jse.12740 [DOI] [Google Scholar]
  87. Lienhard P., Terrat S., Prévost-Bour,é N. C., Nowak V., Régnier T., Sayphoummie S., et al. (2014). Pyrosequencing evidences the impact of cropping on soil bacterial and fungal diversity in Laos tropical grassland. Agron. Sustain. Dev. 34, 525–533. 10.1007/s13593-013-0162-9 [DOI] [Google Scholar]
  88. Lin H., Peddada S. D. (2020). Analysis of compositions of microbiomes with bias correction. Nat. Commun. 11, 3514. 10.1038/s41467-020-17041-7 [DOI] [PMC free article] [PubMed] [Google Scholar]
  89. Liu B., Fu R., Wu B., Liu X. Z., Xiang M. (2021). Rock-inhabiting fungi: terminology, diversity, evolution and adaptation mechanisms. Mycology 13, 1–31. 10.1080/21501203.2021.2002452 [DOI] [PMC free article] [PubMed] [Google Scholar]
  90. Liu J., Haelewaters D., Pfliegler W. P., Page R. A., Dick C. W., Aime M. C. (2020). A new species of Gloeandromyces from Ecuador and Panama revealed by morphology and phylogenetic reconstruction, with a discussion of secondary barcodes in Laboulbeniomycetes taxonomy. Mycologia 112, 1192–1202. 10.1080/00275514.2020.1781496 [DOI] [PubMed] [Google Scholar]
  91. López-Quintero C. A., Straatsma G., Franco-Molano A. E., Boekhout T. (2012). Macrofungal diversity in Colombian Amazon forests varies with regions and regimes of disturbance. Biodivers. Conserv. 21, 2221–2243. 10.1007/s10531-012-0280-8 [DOI] [Google Scholar]
  92. Lücking R., Aime M. C., Robbertse B., Miller A. N., Ariyawansa H. A., Aoki T., et al. (2020). Unambiguous identification of fungi: where do we stand and how accurate and precise is fungal DNA barcoding? IMA Fungus 11, 14. 10.1186/s43008-020-00033-z [DOI] [PMC free article] [PubMed] [Google Scholar]
  93. Lücking R., Dal-Forno M., Sikaroodi M., Gillevet P. M., Bungartz F., Moncada B., et al. (2014). A single macrolichen constitutes hundreds of unrecognized species. Proc. Natl. Acad. Sci. U.S.A. 111, 11091–11096. 10.1073/pnas.1403517111 [DOI] [PMC free article] [PubMed] [Google Scholar]
  94. Marinho F., da Silva I. R., Oehl F., Maia L. C. (2018). Checklist of arbuscular mycorrhizal fungi in tropical forests. Sydowia 70, 107–127. 10.12905/0380.sydowia70-2018-0107 [DOI] [Google Scholar]
  95. McMurdie P. J., Holmes S. (2014). Waste not, want not: why rarefying microbiome data is inadmissible. PLoS Comp. Biol. 10, e1003531. 10.1371/journal.pcbi.1003531 [DOI] [PMC free article] [PubMed] [Google Scholar]
  96. Meswaet Y., Mangelsdorff R., Yorou N. S., Piepenbring M. (2021). Unravelling unexplored diversity of cercosporoid fungi (Mycosphaerellaceae, Mycosphaerellales, Ascomycota) in tropical Africa. MycoKeys 81, 69–138. 10.3897/mycokeys.81.67850 [DOI] [PMC free article] [PubMed] [Google Scholar]
  97. Moore D., Nauta M. M., Evans S. E., Rotheroe M. (2001). Fungal Conservation: Issues and Solutions. Cambridge: Cambridge University Press. 10.1017/CBO9780511565168 [DOI] [Google Scholar]
  98. Morange M., Lagerkvist C., Helix D. (1998). A history of molecular biology. Endeavour 22, 165–166. 10.1016/S0160-9327(99)80026-X [DOI] [Google Scholar]
  99. Mueller G. M. (2017). Progress in conserving fungi: engagement and red listing. BGjournal 14, 30–33. [Google Scholar]
  100. Mueller G. M., Bills G. F., Foster M. S. (2004). Biodiversity of Fungi: Inventory and Monitoring Methods. Burlington, MA: Elsevier Academic Press. [Google Scholar]
  101. Nilsson R. H., Anslan S., Bahram M., Wurzbacher C., Baldrian P., Tedersoo L. (2019). Mycobiome diversity: high-throughput sequencing and identification of fungi. Nat. Rev. Microbiol. 17, 95–109. 10.1038/s41579-018-0116-y [DOI] [PubMed] [Google Scholar]
  102. Nilsson R. H., Hyde K. D., Pawłowska J., Ryberg M., Tedersoo L., Aas A. B., et al. (2014). Improving ITS sequence data for identification of plant pathogenic fungi. Fungal Divers. 67, 11–19. 10.1007/s13225-014-0291-8 [DOI] [Google Scholar]
  103. Nilsson R. H., Ryberg M., Kristiansson E., Abarenkov K., Larsson K.-H., Kõljalg U. (2006). Taxonomic reliability of DNA sequences in public sequence databases: a fungal perspective. PLoS ONE 1, e59. 10.1371/journal.pone.0000059 [DOI] [PMC free article] [PubMed] [Google Scholar]
  104. Noordeloos M. E., Antonín V. (2008). Contribution to a monograph of marasmioid and collybioid fungi in Europe. Czech Mycol. 60, 21–27. 10.33585/cmy.60103 [DOI] [Google Scholar]
  105. Nuñez N. F., Maggia L., Stenger P. L., Lelièvre M., Letellier K., Gigante S., et al. (2021). Potential of high-throughput eDNA sequencing of soil fungi and bacteria for monitoring ecological restoration in ultramafic substrates: the case study of the New Caledonian biodiversity hotspot. Ecol. Eng. 173, 106416. 10.1016/j.ecoleng.2021.106416 [DOI] [Google Scholar]
  106. Oberwinkler F. (2018). “How to understand cryptogams? The development of research methods and their impact on the knowledge of cryptogams,” in Biodiversity and Ecology of Fungi, Lichens, and Mosses. Kerner von Marilaun Workshop 2015 in memory of Josef Poelt, Vol. 34, ed. Blanz P. (Budapest: Austrian Academy of Sciences; ), 106–186. [Google Scholar]
  107. Oliver R. P., Schweizer M. (1999). Molecular Fungal Biology. Cambridge: Cambridge University Press Oliver. 10.1017/CBO9781139163972 [DOI] [Google Scholar]
  108. Osiewacz H. D. (2002). Genes, mitochondria and aging in filamentous fungi. Ageing Res. Rev. 1, 425–442. 10.1016/S1568-1637(02)00010-7 [DOI] [PubMed] [Google Scholar]
  109. Osmundson T. W., Robert V. A., Schoch C. L., Baker L. J., Smith A., Robich G., et al. (2013). Filling gaps in biodiversity knowledge for macrofungi: contributions and assessment of an herbarium collection DNA barcode sequencing project. PLoS ONE 8, e62419. 10.1371/journal.pone.0062419 [DOI] [PMC free article] [PubMed] [Google Scholar]
  110. Pante E., Schoelinck C., Puillandre N. (2015). From integrative taxonomy to species description: one step beyond. Syst. Biol. 64, 152–160. 10.1093/sysbio/syu083 [DOI] [PubMed] [Google Scholar]
  111. Paton A., Antonelli A., Carine M., Forzza R. C., Davies N., Demissew S., et al. (2020). Plant and fungal collections: current status, future perspectives. Plants People Planet 2, 499–514. 10.1002/ppp3.1014132076703 [DOI] [Google Scholar]
  112. Peay K. G. (2014). Back to the future: natural history and the way forward in modern fungal ecology. Fungal Ecol. 12, 4–9. 10.1016/j.funeco.2014.06.001 [DOI] [Google Scholar]
  113. Peay K. G., Kennedy P. G., Bruns T. D. (2008). Fungal community ecology: a hybrid beast with a molecular master. BioScience 58, 799–810. 10.1641/B580907 [DOI] [Google Scholar]
  114. Peay K. G., Kennedy P. G., Talbot J. M. (2016). Dimensions of biodiversity in the Earth mycobiome. Nat. Rev. Microbiol. 14, 434–447. 10.1038/nrmicro.2016.59 [DOI] [PubMed] [Google Scholar]
  115. Peay K. G., Matheny P. B. (2016). “The biogeography of ectomycorrhizal fungi – a history of life in the subterranean,” in Molecular Mycorrhizal Symbiosis, ed. Martin F. (Hoboken, NJ: John Wiley & Sons, Ltd.), 341–361. 10.1002/9781118951446.ch19 [DOI] [Google Scholar]
  116. Penton C. R., StLouis D., Cole J. R., Luo Y., Wu L., Schuur E. G., et al. (2013). Fungal diversity in permafrost and tallgrass prairie soils under experimental warming conditions. Appl. Environ. Microbiol. 79, 7063–7072. 10.1128/AEM.01702-13 [DOI] [PMC free article] [PubMed] [Google Scholar]
  117. Pérez-Izquierdo L., Zabal-Aguirre M., Verdú M., Buée M., Rincón A. (2020). Ectomycorrhizal fungal diversity decreases in Mediterranean pine forests adapted to recurrent fires. Mol. Ecol. 29, 2463–2476. 10.1111/mec.15493 [DOI] [PubMed] [Google Scholar]
  118. Petisco C., Downey G., Murray I., Zabalgogeazcoa I., García-Criado B., García-Ciudad A. (2008). Direct classification of related species of fungal endophytes (Epichloë spp.) using visible and near-infrared spectroscopy and multivariate analysis. FEMS Microbiol. Lett. 284, 135–141. 10.1111/j.1574-6968.2008.01186.x [DOI] [PubMed] [Google Scholar]
  119. Phengsintham P., Braun U., McKenzie E. H. C., Chukeatirote E., Hyde K. D. (2013). Monograph of cercosporoid fungi from Thailand. Plant Pathol. Quar. 3, 67–138. 10.5943/ppq/3/2/2 [DOI] [Google Scholar]
  120. Piepenbring M. (2015). Introduction to Mycology in the Tropics. St. Paul, MN: American Phytopathological Society. 10.1094/9780890546130 [DOI] [Google Scholar]
  121. Piepenbring M., Caballero E., Fournier J., Guzmán G., Hou C.-L., Kirschner R., et al. (2011). Pioneer forays for fungi in the Darién Province in eastern Panama: quintuplicating the knowledge on fungi in this area by five days of fieldwork. Biodivers. Conserv. 20, 2511–2526. 10.1007/s10531-011-0085-1 [DOI] [Google Scholar]
  122. Piepenbring M., Hofmann T. A., Miranda E., Cáceres O., Unterseher M. (2015). Leaf shedding and weather in tropical dry-seasonal forest shape the phenology of fungi–lessons from two years of monthly surveys in southwestern Panama. Fungal Ecol. 18, 83–92. 10.1016/j.funeco.2015.08.004 [DOI] [Google Scholar]
  123. Piepenbring M., Lotz-Winter H., Hofmann T. A. (2018). Incentives and challenges for mycologists in the tropics. Biosyst. Ecol. Ser. 34, 481–514. [Google Scholar]
  124. Piepenbring M., Maciá-Vicente J. G., Codjia J. E. I., Glatthorn C., Kirk P., Meswaet Y., et al. (2020). Mapping mycological ignorance – checklists and diversity patterns of fungi known for West Africa. IMA Fungus 11, 13. 10.1186/s43008-020-00034-y [DOI] [PMC free article] [PubMed] [Google Scholar]
  125. Piepenbring M., Yorou N. S. (2017). Promoting teaching and research on African fungi by field schools on tropical mycology in Benin. IMA Fungus 8, 74–77. 10.1007/BF03449467 [DOI] [Google Scholar]
  126. Pitt J. I., Hocking A. D., Glenn D. R. (1983). An improved medium for the detection of Aspergillus flavus and A. parasiticus. J. Appl. Bacteriol. 54, 109–114. 10.1111/j.1365-2672.1983.tb01307.x [DOI] [PubMed] [Google Scholar]
  127. Powell J. R., Rillig M. C. (2018). Biodiversity of arbuscular mycorrhizal fungi and ecosystem function. New Phytol. 220, 1059–1075. 10.1111/nph.15119 [DOI] [PubMed] [Google Scholar]
  128. Purahong W., Wubet T., Krüger D., Buscot F. (2019). Application of next-generation sequencing technologies to conservation of wood-inhabiting fungi. Conserv. Biol. 33, 716–724. 10.1111/cobi.13240 [DOI] [PubMed] [Google Scholar]
  129. Reynolds N. K., Jusino M. A., Stajich J. E., Smith M. E. (2021). Understudied, underrepresented, and unknown: Methodological biases that limit detection of early diverging fungi from environmental samples. Mol. Ecol. Resour. 10.1111/1755-0998.13540 [DOI] [PubMed] [Google Scholar]
  130. Ríos-Saldaña C. A., Delibes-Mateos M., Ferreira C. C. (2018). Are fieldwork studies being relegated to second place in conservation science? Glob. Ecol. Conserv. 14, e00389. 10.1016/j.gecco.2018.e00389 [DOI] [Google Scholar]
  131. Rivas-Ferreiro M. (2021). Exploring the potential of opportunistic surveys and sequence databases: revealing the fungal diversity of Madagascar and the arrival of ectomycorrhizal fungi in the island. M.Sc. thesis. London: Royal Botanic Gardens, Kew. [Google Scholar]
  132. Rodrigues P., Santos C., Venâncio A., Lima N. (2011). Species identification of Aspergillus section Flavi isolates from Portuguese almonds using phenotypic, including MALDI-TOF ICMS, and molecular approaches. J. Appl. Microbiol. 111, 877–892. 10.1111/j.1365-2672.2011.05116.x [DOI] [PubMed] [Google Scholar]
  133. Rosa L. H., Pinto O. H. B., Šantl-Temkiv T., Convey P., Carvalho-Silva M., Rosa C. A., et al. (2020). DNA metabarcoding of fungal diversity in air and snow of Livingston Island, South Shetland Islands, Antarctica. Sci. Rep. 10, 21793. 10.1038/s41598-020-78630-6 [DOI] [PMC free article] [PubMed] [Google Scholar]
  134. Runnel K., Drenkhan R., Adamson K., Lõhmus P., Rosenvald K., Rosenvald R., et al. (2021). The factors and scales shaping fungal assemblages in fallen spruce trunks: a DNA metabarcoding study. For. Ecol. Manag. 495, 119381. 10.1016/j.foreco.2021.119381 [DOI] [Google Scholar]
  135. Ryberg M., Nilsson R. H. (2018). New light on names and naming of dark taxa. MycoKeys 30, 31–39. 10.3897/mycokeys.30.24376 [DOI] [PMC free article] [PubMed] [Google Scholar]
  136. Sadiković D., Kuštera M. (2013). Fungal conservation: protected species of fungi in South Serbia region. Biol. Nyssana 4, 35–40. [Google Scholar]
  137. Schmit J. P., Lodge D. J. (2005). “Classical methods and modern analysis for studying fungal diversity,” in The Fungal Community, ed. Dighton J. (Boca Raton, FL: Marcel Dekker, Inc.), 193–214. 10.1201/9781420027891.ch10 [DOI] [Google Scholar]
  138. Selbmann L., Isola D., Egidi E., Zucconi L., Gueidan C., de Hoog G. S., et al. (2014). Mountain tips as reservoirs for new rock-fungal entities: Saxomyces gen. nov. and four new species from the Alps. Fungal Divers. 65, 167–182. 10.1007/s13225-013-0234-9 [DOI] [Google Scholar]
  139. Sharma S. K., Saini S., Verma A., Sharma P. K., Lal R., Roy M., et al. (2019). National agriculturally important microbial culture collection in the global context of microbial culture collection centres. Proc. Natl. Acad. Sci. India Sect. B Biol. Sci. 89, 405–418. 10.1007/s40011-017-0882-8 [DOI] [Google Scholar]
  140. Shrestha U. B., Bawa K. S. (2014). Impact of climate change on potential distribution of Chinese caterpillar fungus (Ophiocordyceps sinensis) in Nepal Himalaya. PLoS ONE 9, e106405. 10.1371/journal.pone.0106405 [DOI] [PMC free article] [PubMed] [Google Scholar]
  141. Simões M. F., Pereira L., Santos C., Lima N. (2013). “Polyphasic identification and preservation of fungal diversity: Concepts and applications,” in Management of Microbial Resources in the Environment, eds. Malik A., Grohmann E., Alves M. (Dordrecht: Springer; ), 91–117. 10.1007/978-94-007-5931-2_5 [DOI] [Google Scholar]
  142. Simpson G. (1960). Principles of Animal Taxonomy. New York, NY: Columbia University Press. 10.7312/simp92414 [DOI] [Google Scholar]
  143. Spatafora J. W., Aime M. C., Grigoriev I. V., Martin F., Stajich J. E., Blackwell M. (2018). “The fungal tree of life: from molecular systematics to genome-scale phylogenies,” in The Fungal Kingdom, eds. Heitman J., Howlett B. J., Crous P. W., Stukenbrock E. H., James T. Y., Gow N. A. R. (Washington, DC: ASM Press; ), 1–34. 10.1128/9781555819583.ch1 [DOI] [PMC free article] [PubMed] [Google Scholar]
  144. Stewart J. E., Kim M. S., Klopfenstein N. B. (2018). Molecular genetic approaches toward understanding forest-associated fungi and their interactive roles within forest ecosystems. Curr. For. Rep. 4, 72–84. 10.1007/s40725-018-0076-5 [DOI] [Google Scholar]
  145. Suh S.-O., Zhang N., Nguyen N., Gross S., Blackwell M. (2008). Lab Manual for Yeast Study. Baton Rouge, LA: Louisiana State University. [Google Scholar]
  146. Sundberg H., Ekman S., Kruys Å. (2018). A crush on small fungi: an efficient and quick method for obtaining DNA from minute ascomycetes. Methods Ecol. Evol. 9, 148–158. 10.1111/2041-210X.12850 [DOI] [Google Scholar]
  147. Taylor D. L., Hollingsworth T. N., McFarland J. W., Lennon N. J., Nusbaum C., Ruess R. W. (2014). A first comprehensive census of fungi in soil reveals both hyperdiversity and fine-scale niche partitioning. Ecol. Monogr. 84, 3–20. 10.1890/12-1693.1 [DOI] [Google Scholar]
  148. Taylor J. W., Jacobson D. J., Kroken S., Kasuga T., Geiser D. M., Hibbett D. S., et al. (2000). Phylogenetic species recognition and species concepts in fungi. Fungal Genet. Biol. 31, 21–32. 10.1006/fgbi.2000.1228 [DOI] [PubMed] [Google Scholar]
  149. Tedersoo L., Bahram M., Põlme S., Kõljalg U., Yorou N. S., Wijesundera R., et al. (2014). Global diversity and geography of soil fungi. Science 346, 1256688. 10.1126/science.1256688 [DOI] [PubMed] [Google Scholar]
  150. Teke N. A., Kinge T. R., Bechem E., Nji T. M., Ndam L. M., Mih A. M. (2018). Ethnomycological study in the Kilum-Ijim mountain forest, Northwest Region, Cameroon. J. Ethnobiol. Ethnomed. 14, 25. 10.1186/s13002-018-0225-8 [DOI] [PMC free article] [PubMed] [Google Scholar]
  151. Tomao A., Bonet J. A., Castaño C., de-Miguel S. (2020). How does forest management affect fungal diversity and community composition? Current knowledge and future perspectives for the conservation of forest fungi. For. Ecol. Manag. 457, 117678. 10.1016/j.foreco.2019.117678 [DOI] [Google Scholar]
  152. Truong C., Mujic A. B., Healy R., Kuhar F., Furci G., Torres D., et al. (2017). How to know the fungi: combining field inventories and DNA-barcoding to document fungal diversity. New Phytol. 214, 913–919. 10.1111/nph.14509 [DOI] [PubMed] [Google Scholar]
  153. Turland N. J., Wiersema J. H., Barrie F. R., Greuter W., Hawksworth D. L., Herendeen P. S., et al. (2018). “International Code of Nomenclature for algae, fungi, and plants (Shenzhen Code),” in The Nineteenth International Botanical Congress Shenzhen, China, July 2017. Regnum Vegetabile 159 (Glashütten: Koeltz Botanical Books; ). 10.12705/Code.2018 [DOI] [Google Scholar]
  154. van der Linde S., Suz L. M., Orme C. D. L., Cox F., Andreae H., Asi E., et al. (2018). Environment and host as large-scale controls of ectomycorrhizal fungi. Nature 558, 243–248. 10.1038/s41586-018-0189-9 [DOI] [PubMed] [Google Scholar]
  155. Vargas-Gastélum L., Romero-Olivares A. L., Escalante A. E., Rocha-Olivares A., Brizuela C., Riquelme M. (2015). Impact of seasonal changes on fungal diversity of a semi-arid ecosystem revealed by 454 pyrosequencing. FEMS Microbiol. Ecol. 91, fiv044. 10.1093/femsec/fiv044 [DOI] [PubMed] [Google Scholar]
  156. Vega F. E., Blackwell M. (2005). Insect-Fungal Associations: Ecology and Evolution. New York, NY: Oxford University Press. [Google Scholar]
  157. Velázquez M. S., Biganzoli F., Cabello M. N. (2010). Arbuscular mycorrhizal fungi in El Palmar National Park (Entre Rios Province, Argentina) – a protected reserve. Sydowia 62, 149–163. [Google Scholar]
  158. Wang X. W., Jiang J. H., Liu S. L., Gafforov Y., Zhou L. W. (2021). Species diversification of the coniferous pathogenic fungal genus Coniferiporia (Hymenochaetales, Basidiomycota) in association with its biogeography and host plants. Phytopathology. 112, 404–413. 10.1094/PHYTO-05-21-0181-R [DOI] [PubMed] [Google Scholar]
  159. Weiss M., Selosse M. A., Rexer K. H., Urban A., Oberwinkler F. (2004). Sebacinales: a hitherto overlooked cosm of heterobasidiomycetes with a broad mycorrhizal potential. Mycol. Res. 108, 1003–1010. 10.1017/S0953756204000772 [DOI] [PubMed] [Google Scholar]
  160. Weiss S., Xu Z. Z., Peddada S., Amir A., Bittinger K., Gonzalez A., et al. (2017). Normalization and microbial differential abundance strategies depend upon data characteristics. Microbiome 5, 104. 10.1186/s40168-017-0237-y [DOI] [PMC free article] [PubMed] [Google Scholar]
  161. Wu B., Hussain M., Zhang W., Stadler M., Liu X., Xiang M. (2019). Current insights into fungal species diversity and perspective on naming the environmental DNA sequences of fungi. Mycology 10, 127–140. 10.1080/21501203.2019.1614106 [DOI] [PMC free article] [PubMed] [Google Scholar]
  162. Wutkowska M., Vader A., Mundra S., Cooper E. J., Eidesen P. B. (2019). Dead or alive; or does it really matter? Level of congruency between trophic modes in total and active fungal communities in high arctic soil. Front. Microbiol. 9, 3243. 10.3389/fmicb.2018.03243 [DOI] [PMC free article] [PubMed] [Google Scholar]
  163. Yan D., Mills J. G., Gellie N. J., Bissett A., Lowe A. J., Breed M. F. (2018). High-throughput eDNA monitoring of fungi to track functional recovery in ecological restoration. Biol. Conserv. 217, 113–120. 10.1016/j.biocon.2017.10.035 [DOI] [Google Scholar]
  164. Yan Y., Li Y., Wang W. J., He J. S., Yang R. H., Wu H. J., et al. (2017). Range shifts in response to climate change of Ophiocordyceps sinensis, a fungus endemic to the Tibetan Plateau. Biol. Conserv. 206, 143–150. 10.1016/j.biocon.2016.12.023 [DOI] [Google Scholar]
  165. Zhang Y., Qiao M., Xu J., Cao Y., Zhang K.-Q., Yu Z.-F. (2013). Genetic diversity and recombination in natural populations of the nematode-trapping fungus Arthrobotrys oligospora from China. Ecol. Evol. 3, 312–325. 10.1002/ece3.450 [DOI] [PMC free article] [PubMed] [Google Scholar]
  166. Zhang Y., Zhang S., Wang M., Bai F., Liu X. (2010). High diversity of the fungal community structure in naturally-occurring Ophiocordyceps sinensis. PLoS ONE 5, e15570. 10.1371/journal.pone.0015570 [DOI] [PMC free article] [PubMed] [Google Scholar]

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