One stop shop IV: taxonomic update with molecular phylogeny for important phytopathogenic genera: 76–100 (2020)

Abstract

This is a continuation of a series focused on providing a stable platform for the taxonomy of phytopathogenic fungi and fungus-like organisms. This paper focuses on one family: Erysiphaceae and 24 phytopathogenic genera: Armillaria, Barriopsis, Cercospora, Cladosporium, Clinoconidium, Colletotrichum, Cylindrocladiella, Dothidotthia,, Fomitopsis, Ganoderma, Golovinomyces, Heterobasidium, Meliola, Mucor, Neoerysiphe, Nothophoma, Phellinus, Phytophthora, Pseudoseptoria, Pythium, Rhizopus, Stemphylium, Thyrostroma and Wojnowiciella. Each genus is provided with a taxonomic background, distribution, hosts, disease symptoms, and updated backbone trees. Species confirmed with pathogenicity studies are denoted when data are available. Six of the genera are updated from previous entries as many new species have been described.

Contents and contributors (main contributors underlined)

Newly discussed genera and family

• 76. Armillaria – B Chuankid, M Stadler

• 77. Barriopsis – IS Manawasinghe, RS Jayawardena

• 78. Cercospora – ID Goonasekara

• 79. Clinoconidium – AK Gautam, S Avasthi

• 80. Cylindrocladiella – D Harischandra, RS Jayawardena

• 81. Dothidotthia – C Senwanna

• 82. Erysiphaceae – KK Liyanage, RS Jayawardena, KD Hyde

• 83. Fomitopsis – V Papp, B Palla, D Papp

• 84. Ganoderma – KK Hapuarachchi, T Luangharn, O Raspe

• 85. Golovinomyces – RS Jayawardena

• 86. Heterobasidium – V Papp, B Palla, D Papp

• 87. Meliola – S Hongsanan, XY Zeng

• 88. Neoerysiphe – RS Jayawardena

• 89. Nothophoma – IS Manawasinghe, RS Jayawardena

• 90. Phellinus – V Papp, B Palla, D Papp

• 91. Pseudoseptoria – A Karunarathna, RS Jayawardena

• 92. Stemphylium – RS Jayawardena, KD Hyde

• 93. Thyrostroma – C Senwanna, KD Hyde

• 94. Wojnowiciella – D Harischandra, RS Jayawardena

Updated genera

• 95. Cladosporium – NG Liu, RS Jayawardena

• 96. Colletotrichum – RS Jayawardena, KD Hyde

• 97. Mucor – VG Hurdeal, HB Lee

• 98. Phytophthora – CS Bhunjun, RS Jayawardena

• 99. Pythium – CS Bhunjun, RS Jayawardena

• 100. Rhizopus – VG Hurdeal, HB Lee

Introduction

This is the fourth paper in the One Stop Shop series focusing on providing a stable platform for the taxonomy of plant pathogenic fungi and fungus-like organisms. Genera included in this series are associated with plant diseases, and when the data are available we discuss the species that have been established as pathogens using Koch’s postulates. Some genera, however, are not well-known plant pathogens and some may be emerging pathogens, and need further studies to confirm their pathogenicity. Hyde et al. (2014) launched this series and stated its specific aims.

Three issues of One Stop Shop (OSS) have been published treating 73 genera and two families of plant pathogenic fungi and fungus-like organisms (Hyde et al. 2014; Jayawardena et al. 2019a, b, Table 1). In this fourth contribution, a further 24 genera and one family are treated, providing clarification of their taxonomy and classification. Six of the entries are updates from previous entries as many changes have occurred in these genera. For each entry, the background of the genus, disease symptoms, host distribution, pathogen biology and epidemiology, morphological based identification, molecular-based identification, updated phylogeny and recommended genetic markers are provided and discussed. All contributed entries will be placed in the database, http://www.onestopshopfungi.org. The main outcome of this series is to enhance the current understanding of plant pathogens and gain better insights into the current classification, providing a stable taxonomy and phylogeny for plant pathogens. This will provide a definitive classification for mycologists and plant pathologists to accurately identify causal agents of disease and to implement accurate control strategies.

Table 1 All entries treated in One stop shop (OSS) series

Materials and methods

Photo plates of the symptoms of the disease and morphological characters are given, when available. Classification follows Wijayawardene et al. (2020).

For the treated taxa, all species that have been published until 30 March 2020 are included in the phylogenetic analyses. Sequence data from ex-type, ex-epitype or authentic or reference/voucher strains for each species were retrieved from GenBank. Sequence data from single gene regions were aligned using Clustal Xv.1.81 (Thompson et al. 1997) and further alignment of the sequences carried out using the default settings of MAFFT v.7 (Katoh and Toh 2008; http://mafft.cbrc.jp/alignment/server/), and manual adjustment was conducted using BioEdit where necessary. Gene regions were also combined using BioEdit v.7.0.9.0 (Hall 1999). Primers for each gene locus can be found in the bibliography related to the phylogeny presented in each genus. Phylogenetic analyses consisted of maximum likelihood (ML), maximum parsimony (MP) and Bayesian posterior probability (BYPP). Maximum parsimony analysis was performed using PAUP (Phylogenetic Analysis Using Parsimony) v. 4.0b10 (Swofford 2002) to obtain the most parsimonious trees. Maximum likelihood analyses were also performed in raxmlGUIv.0.9b2 (Silvestro and Michalak 2010) or RAxML-HPC2 on XSEDE (8.2.8) on the CIPRES science gateway platform (http://www.phylo.org; Miller et al. 2010). Bayesian inference was conducted using MrBayes v. 3.2.6 on the CIPRES science gateway platform (http://www.phylo.org; Miller et al. 2010) or stand-alone MrBayes v.3.1.2 (Ronquist and Huelsenbeck 2003). MrModeltest v. 2.3 (Nylander 2004) or jModeltest v. 2.1.4 (Darriba et al. 2012) was used for the statistical selection of the best-fit model of nucleotide substitution to parametrize the analyses.

Results

76. Armillaria (Fr.) Staude, Schwämme Mitteldeutschl. 28: xxviii, 130 (1857)

Background

Armillaria is a plant pathogenic genus in the phylum Basidiomycota, family Physalacriaceae (He et al. 2019), collectively referred to as shoestring root-rot fungi or honey mushrooms. Armillaria can cause root-rot disease in a wide variety of woody hosts worldwide. Armillaria has undergone significant revision in the past 20 years. The genus once accommodated any white-spored agaric with broadly attached gills and an annulus (Volk et al. 1996). Armillaria mellea is the type species. Most Armillaria species have the potential to infect healthy and stressed trees, they differ in their pathogenicity to their hosts and under certain circumstances, they behave as obligate saprobes. Most Armillaria species are facultative necrotrophs causing root and butt rot on a broad range of woody plants affecting a variety of forest, shade, ornamental and orchard trees and shrubs. Some Armillaria species cause significant economic losses to forest trees and in nursery plantations. Armillaria root disease is found in many temperate and tropical forests throughout the world. This fungus spreads mainly through the interaction of tree roots. As saprotrophs, Armillaria species are important wood decomposers that contribute to nutrient cycling in forest ecosystems. As pathogens, they infect and eventually kill susceptible trees, which impacts forest structure, composition and succession. Trees that are used for fibre or lumber production, as well as trees located in recreation sites, are affected by these diseases. Such Armillaria infections may cause yield reduction and tree mortality in silvicultural and agricultural tree plantations and provoke economic losses.

Armillaria species are expected to become more aggressive during drought and thus enhance root rot (La Porta et al. 2008; Kolb et al. 2016; Kubiak et al. 2017). The incidence of Armillaria related root disease is likely to increase as temperatures increase and precipitation decreases due to climate change (Sturrock et al. 2011). Whilst the ability of the pathogen to sporulate, spread and infect is affected by temperature and moisture, factors that stress host trees directly may be just as critical to a successful invasion of host tissues. It seems likely that the disease will become more severe in the future, wherever Armillaria susceptible tree species are subjected to increased levels of climate stress (Klopfenstein et al. 2009). Currently, Armillaria root disease causes large growth/volume losses (e.g., 16–55%) in areas of western and North America (Filip and Goheen 1984; Cruickshank et al. 2011; Lockman and Kearns 2016). Armillaria root disease is typically more severe in trees that are maladapted to climate-induced stress (Ayres and Lombardero 2000; Kliejunas et al. 2009; Sturrock et al. 2011). Thus, it is likely that climate change will further exacerbate damage from Armillaria root disease, which can further predispose trees to beetle attack (e.g. Hertert et al. 1975; Tkacz and Schmitz 1986; Goheen and Hansen 1993).

Armillaria mellea is an edible species that has long been used as a Traditional Chinese Medicine. Some of Armillaria species are is believed to be able to improve health and prevent various diseases, such as insomnia, pain, and neurasthenia. Extracts of A. mellea exhibit anti-oxidative, anti-inflammatory and immune-modulatory activities. Armillaria mellea can also induce maturation of human dendritic cells. The chemical constituents isolated from A. mellea include sesquiterpenoids, steroids, triterpenoids, adenosine and resin acids. Armillariol C is a furan-based natural product isolated from Armillaria species. A xylosyl 1,3-galactofucan (AMPS-III) was isolated and identified as a novel anti-inflammatory agent from this species.

Classification—Basidiomycota, Agaricomycotina, Agaricomycetes, Agaricomycetidae, Agaricales, Physalacriaceae (He et al. 2019)

Type species—Armillaria mellea (Vahl) P. Kumm.

Distribution—Worldwide, mostly in temperate areas (northern and southern hemisphere) and some in tropical areas.

Disease symptoms—Armillaria root disease, shoestring root rot

Symptoms caused by this fungus can be categorized into two categories:

Crown symptoms—branch dieback, crown thinning, chlorosis, reddening of foliage or heavier than normal production of cones.

Basal symptoms—the fungus can grow up from the roots in the inner bark in some tree species and causes basal cankers above the infected roots. Resinosis (exudation of resin) can be observed in resinous conifers. In some plants, decayed roots or decay in the inner wood of stem bases can be observed. Species cause a white rot of wood. In white rot, wood often has a bleached, whitish appearance and are spongy or stringy, and maybe wet. Black lines called “zone lines” are usually seen in the decayed wood. These lines are curved planes in the wood, sometimes called “pseudosclerotial plates”, composed of thickened, dark fungal cells. They may play a role in the protection of Armillaria from unfavourable conditions or other fungi that attempt to invade its territory, including other individuals of the same species. Actively decaying wood may be luminescent, producing a faint glow in the dark (Baumgartner and Rizzo 2002; Worrall 2004; Klopfenstein 2009).

There are three major signs of Armillaria root disease in the field.

Mycelial fans can always be seen in infected and recently killed trees. These are white mats of fungal mycelium between the inner bark and wood that are generally substantial and have a mushroom odour.

Rhizomorphs are commonly associated with infection and are often attached to infected roots, but they may also be attached to the surface of uninfected roots. Depending on the species these may be few, small, fragile, hard to find or abundant and robust. Rhizomorphs can be cylindrical in soil or flattened under bark, reddish-brown to black branched and have a cream-coloured tip when actively growing (Guillaumin and Legrand 2013).

Mushrooms that have honey-brown caps can be seen in clusters near or on the base of trees.

Hosts—Many angiosperms and gymnosperms (especially conifers) in native, planted forests, orchards and vineyards (Farr and Rossman 2020).

Pathogen biology, disease cycle and epidemiology

Sexual reproduction results in the diploid mycelium. Such a mycelium is the dominant phase that is found growing in wood, growing through the soil as rhizomorphs, and killing trees. Armillaria species can be dispersed through airborne sexual basidiospores which will establish a new infection center. These taxa do not reproduce asexually but disperse by growing mycelium which is the most common source of infection, through root contacts or root grafts or by growing through the soil as rhizomorphs. Mycelium in colonized roots and the rhizomorphs produced serve as the most common mode of infection and may survive for up to 50 years or more in stumps, depending on the climate, size of the stump, and other factors (Baumgartner and Rizzo 2002; Worrall 2004; Klopfenstein 2009).

Morphology-based identification and diversity

Armillaria has included only white-spored wood-inhabiting agarics with broadly attached to decurrent gills and macroscopic black to reddish-brown rhizomorphs. Armillaria basidiomes are easily recognized by their caespitose habit, annulus and honey colour. It is, however, extremely difficult to identify some species due to the lack of morphological apomorphies (Watling et al. 1991; Pegler 2000). Besides, basidiomata are often not available to differentiate species, which further complicates the taxonomy of Armillaria (Harrington and Wingfield 1995). In this regard, Armillaria provides a clear example of where a phylogenetic approach can contribute significantly to its taxonomy. Until the late 1970s, Armillaria mellea was considered by most researchers to be a polymorphic species with a wide host range and distribution. Herink (1973), among others, suspected that this single species might be a species complex. However, since the morphology of basidiomata is difficult to study because of overlapping and inconsistent traditionally used morphological characters, other avenues of research were pursued. Hintikka (1973) developed a technique that allowed the determination of mating types in Armillaria. Using a modification of this method, Korhonen (1978a) was able to distinguish five European biological species. The cumbersome nature of the mating-type method of species identification prompted a search for other techniques for identifying collections. They were able to separate all North American species (NABS) of Armillaria except for A. calvescens and A. gallica, which are apparently very closely related (Anderson and Stasovski1992). Ten species of Armillaria in North America have been confirmed from multiple studies utilizing a combination of morphological, biological and phylogenetic species concepts (Anderson and Ullrich 1979; Anderson and Stasovski 1992; Burdsall and Volk 1993; Kim et al. 2006; Ross-Davis et al. 2012). Before, A. mellea shows great variability in morphology and hosts. These species were first separated using interfertility tests using cultures of Armillaria haploid tester strains and morphology. Now, A. mellea is considered as an independent species, with two North American biological species (Bérubé and Dessureault 1989; Volk et al. 1996) (Fig. 1).

Fig. 1

Disease cycle of Armillaria mellea (redrawn from Agrios 2005)

Molecular-based identification and diversity

Problems surrounding the identification of Armillaria have led to important advances in developing robust but rapid DNA techniques. Such techniques have initially included DNA-base composition (Jahnke et al. 1987) DNA-DNA hybridization (Miller et al. 1994), sequence analyses of the IGS-1(Anderson and Stasovski 1992) and ITS (Coetzee et al. 2001a, b), RFLPs without PCR (Smith and Anderson 1989) and RFLPs of IGS-1 amplicons (Harrington and Wingfield 1995). Although several of these techniques might pose some problems (Pérez‐Sierra et al. 2000), by their relative simplicity they have gradually replaced traditional, morphological methods.

The amount of DNA sequence data on Armillaria species has increased substantially since the first publication on the phylogeny of the genus in the northern hemisphere (Anderson and Stasovski 1992). As with many other fungal genera, the focus of such studies initially was set on species of Europe and North America (Chillali et al. 1998; Coetzee et al. 2000b). Later, substantial datasets for species in Africa, Australasia and southeast Asia have become available (Terashima et al. 1998; Coetzee et al Coetzee et al. 2000a, 2001a). At present, ITS, IGS-1 and tef1 sequences are available in GenBank for the best-known species of Armillaria. However, there are disjunctions in data sets and relatively little is known about species from Indo-Malaysia and South America. Armillaria fruiting bodies are produced seasonally and not every year; they are, therefore, often not available during fieldwork (Kile et al. 1991).

Identification using the biological species concept with species identification based on sexual compatibility tests (Korhonen 1978a) has been examined for its utility by some mycologists, but its application was soon abandoned. This was because of complications due to the absence of known tester strains, lack of haploid strains, ambiguous mating interactions and degeneracy of cultures. For these reasons, DNA-based molecular techniques have finally been preferred in Armillaria taxonomy, either complementing other methods or on their own. The techniques utilized for the taxonomy of Armillaria species include comparisons of RFLPs (Harrington and Wingfield 1995), AFLPs (Pérez-Sierra et al. 2004), and the use of sequences from the ITS, IGS-1 and tef1 gene in phylogenetic studies (Coetzee et al. 2000b, 2001a; Maphosa et al. 2006; Kim et al. 2006). Phylogenetic methods have made it possible to differentiate the lineages of the genus in southern Argentina (Pildain et al. 2009). Lineages I and II grouped with A. novae-zelandiae and A. luteobubalina, respectively, while Lineages III and IV represented unique taxa that were closely related to A. hinnulea, Armillaria 4th species from New Zealand (established by Coetzee et al. 2001a, b) and Armillaria Group III from Kenya (Mwenje et al. 2006). Modern approaches to identification of Armillaria species are mostly based on the analyses of DNA sequences. The present study reconstructs the phylogeny of Armillaria based on a combined ITS, IGS and tef1 sequence data (Fig. 2, Table 2). However, insufficient data are available for the LSU gene region in GenBank. Then, it is difficult to have comparative phylogenetic analyses but the single gene analysis of each gene was carried out to compare the topology of the tree and clade stability. This phylogenetic tree is largely in accordance with earlier studies from Coetzee et al. (2018) and provides the most conclusive phylogeny of the genera to date. Genealogical concordance phylogenetic species recognition (GCPSR) using the concordance among several gene trees (Taylor et al. 2000; Dettman et al. 2003) to delineate species has become standard in fungal taxonomy. However, except for a few studies (Guo et al. 2016; Tsykun et al. 2013), this taxonomic method has not been widely implemented in Armillaria taxonomy. Sequences of the genomes of key species are already providing prospects to study the evolution and systematics of Armillaria. They are certain to lead to important breakthroughs regarding not only the taxonomy but the biology and ecology of these fungi in the future (Sipos et al. 2017).

Fig. 2

Phylogenetic tree generated by maximum likelihood analysis of combined ITS-IGS-tef1 sequence data of Armillaria species. Related sequences were obtained from GenBank. One hundred and thirty-nine strains are included in the analyses, which comprise 4557 characters including gaps. The tree was rooted with Guyanagaster lucianii (G31.4) and Guyanagaster necrorhizus (MCA 3950). Single gene analyses were carried out to compare the topology of the tree and clade stability. Tree topology of the ML analysis was similar to the MP and BYPP. ML phylogenetic tree inference was performed using RAxML version 8.2.12 on the CIPRES web server, using a mixed-model analysis and the GTRCAT model of substitution. The four partitions were defined as ITS, IGS, tef1 exons and tef1 introns. The best scoring RAxML tree with a final likelihood value of − 25308.198187 is presented. The matrix had 1957 distinct alignment patterns, with 65.74% of undetermined characters or gaps. Estimated base frequencies of ITS were as follows: A =0.227071, C =0.203923, G =0.235701, T =0.333305; substitution rates AC =0.628852, AG=3.751709, AT =1.365607, CG =1.467905, CT =2.788595, GT = 1.000000. Estimated base frequencies of IGS were as follows: A =0.244624, C =0.196588, G =0.242370, T =0.316418; substitution rates AC =0.954911, AG=3.055115, AT =1.041498, CG =1.278095, CT = 3.421100, GT = 1.000000. Estimated base frequencies of tef1 exons were as follows: A =0.228587, C =0.301128, G =0.255865, T =0.214420; substitution rates AC =0.905728, AG=3.660986, AT =1.564184, CG =0.648739, CT = 28.048363, GT = 1.000000. Estimated base frequencies of tef1 introns were as follows: A =0.215042, C =0.222693, G =0.185633, T =0.376631; substitution rates AC =1.170263, AG=5.878084, AT =0.847943, CG =1.087990, CT = 5.095797, GT = 1.000000; gamma distribution shape parameter α =0.1000000000. The maximum parsimonious dataset consisted of 2908 constant, 1172 parsimony-informative and 477 parsimony-uninformative characters. The parsimony analysis: CI = 0.610, RI = 0.861, RC = 0.525, HI = 0.390 in the first tree. Bayesian posterior probability was performed using the Markov chain Monte Carlo (MCMC) method implemented in MrBayes 3.2.6 with a mixed-model partition identical to the ones defined in the ML analysis. The best-fit nucleotide substitution model was separately determined for each partition with jModeltest version 2.1.10 on CIPRES, using the Akaike Information Criterion. K80+I, K80+I, SYM+G and HKY+G were selected as best-fit models for ITS, IGS, tef1 exons and tef1 introns, respectively. At the end of the runs, the average deviation of split frequencies was 0.016675. MP and RAxML bootstrap support value ≥  50% and BYPP ≥ 0.95 are shown, respectively, near the nodes. Holotype or ex-type strains are in bold

Table 2 DNA barcodes available for Armillaria

Recommended genetic marker (genus level)—ITS

Recommended genetic markers (species level)—ITS, IGS1, tef1

Additional genetic markers (species level)—LSU, tub2

Accepted number of species—There are 278 epithets in Index Fungorum (2020) listed for this genus. However, sequence data are only available for 31 species including 16 groups of unnamed species (Table 2).

References—Watling et al. (1991), Pegler (2000), Harrington and Wingfield (1995) (morphology); Coetzee et al. (2000a, b, 2001a, b), Maphosa et al. (2006), Mwenje et al. (2006), Kim et al. (2006), Coetzee et al. (2018) (molecular phylogeny).

77. Barriopsis A.J.L. Phillips, A. Alves & Crous, in Phillips et al., Persoonia 21: 39 (2008)

Background

Stevens (1926) originally described the type species of Barriopsis in Physlospora as Physlospora fusca and Petrak and Deighton (1952) transferred it to Phaeobotryosphaeria. The fungus that was considered by Stevens (1926), and Petrak and Deighton (1952) did not have apiculi on its ascospores and was not similar to Phaeobotryosphaeria which had small, hyaline apiculi on the ascospores. von Arx and Müller (1954) considered Phaeobotryosphaeria as a synonym of Botryosphaeria. Based on morphological difference and molecular sequence data, Phillips et al. (2008) introduced Barriopsis. Species of Barriopsis are mostly saprobic and weak pathogens (Phillips et al. 2013).

Classification—Ascomycota, Dothideomycetes, Incertae sedis, Botryosphaeriales, Botryosphaeriaceae

Type species—Barriopsis stevensiana A.J.L. Phillips & Pennycook

Distribution—Species appear to be confined to regions with tropical or subtropical climates including Australia, Cuba, Iran and Thailand (Phillips et al. 2008; Abdollahzadeh et al. 2009; Liu et al. 2012; Phillips et al. 2013; Doilom et al. 2014; Konta et al. 2016; Dissanayake et al. 2016; Hyde et al. 2018b; Burgess et al. 2019).

Disease symptoms—Barriopsis species can be weak pathogens and their pathogenicities are uncertain (Phillips et al. 2008; Dissanayake et al. 2016). Barriopsis stevensiana and B. iraniana were isolated from infected branches, fruits and leaves with various disease symptoms, including dieback, canker, rot and necrosis, from Cupressus sempervirens, Mangifera indica, Citrus sp. and Olea sp. in northern and southern provinces of Iran (Abdollahzadeh et al. 2009). Species of this genus may be future emerging pathogens.

Hosts—Archontophoenix alexandrae, Cassia sp., Citrus sp., Mangifera indica, Olea sp. Tectona grandis (Phillips et al. 2008, 2013; Abdollahzadeh et al. 2009; Liu et al. 2012; Doilom et al. 2014; Konta et al. 2016; Dissanayake et al. 2016; Hyde et al. 2018b, 2020b).

Pathogen biology, disease cycle and epidemiology

Barriopisis in this article is considered as an emerging pathogen. Further studies to identify the biology, disease cycle and epidemiology are needed.

Morphological based identification and diversity

The sexual morph is characterized by brown aseptate ascospores that are widest in the center and lack terminal apiculi (Phillips et al. 2008, 2013; Doilom et al. 2014; Dissanayake et al. 2016; (Fig. 3)). Barriopsis archontophoenicis forms the sexual morph in culture medium after long periods of incubation (up to 6 months, Konta et al. 2016). The asexual morph is lasiodiplodia-like with hyaline conidia that become dark-brown and septate with irregular longitudinal striations (Stevens 1926). Abdollahzadeh et al. (2009) observed the asexual morphs of B. fusca and B. iraniana and confirmed that the morphology is similar to the description given by Stevens (1926). In their study, they revealed that this genus can be distinguished from other genera of Botryosphaeriaceae by the presence of visible striations on conidia at an early stage of development.

Fig. 3

Barriopsis stevensiana MFLU 19–1560. a Ascomata on dead twigs of Cassia sp. b Ascomata cut through horizontally showing the white contents with dark spots. c, d Sections through ascomata. e, f Ascospores. g Germinated ascospore. Scale bars: c, d = 200 µm, e, f = 20 µm, g = 100 µm

However, using morphology alone in identifying these species is not wise due to the overlapping of morphological characters within the genus. Therefore, the use of multi loci phylogeny along with morphology is recommended for this genus. Very little is known about the diversity and pathogenicity of this botryosphaeriaceous genus and future studies are needed to confirm its pathogenic nature.

Molecular based identification and diversity

Phillips et al. (2008) using SSU, ITS, LSU, tef1 and tub2 sequence data established Barriopsis which is sister to Phaeobotryon. Based on ITS and tef1 sequence data, Abdollahzadeh et al. (2009) introduced B. iraniana. Doilom et al. (2014) introduced B. tectonae based on ITS, tub2 and tef1 sequence data. In this study, it was mentioned that ITS and tub2 sequence data have lesser variation, while tef1 sequence data have considerable variation. Konta et al. (2016) added a new species, B. archontophoenicis with the use of ITS, LSU, SSU and tef1 sequence data. In this study, we construct the phylogenetic tree for the accepted species based on ITS and tef1 sequence data (Fig. 4).

Fig. 4

Phylogram generated from maximum likelihood analysis based on combined ITS, and tef1 sequence data of Barriopsis species and closely related taxa. Fifteen strains are in the combined sequence analyses, which comprise 865 characters including gaps. Diplodia mutila (CBS 112553 and CBS 230.30) was used as the outgroup taxa. Tree topology of the ML analysis was similar to the one generated from BI. The best scoring RAxML tree with a final likelihood value of − 2372.487246 is presented. The matrix had 201 distinct alignment patterns, with 12.30% of undetermined characters or gaps. Estimated base frequencies were as follows: A = 0.207721, C = 0.288041, G = 0.271092, T = 0.233145; substitution rates AC = 1.068561, AG = 2.489613, AT = 0.682766, CG = 1.417925, CT = 4.236517, GT = 1.000000; gamma distribution shape parameter α = 1.343820. RAxML bootstrap support value ≥ 50% and BYPP ≥ 0.95 are shown respectively, near the nodes. Ex-type strains are in bold

Recommended genetic marker (genus level)—ITS

Recommended genetic marker (species level) —tef1

Accepted number of species—There are six species epithets in Index Fungorum (2020), however only five species have DNA sequence data (Table 3).

Table 3 DNA barcodes available for Barriopsis

References—Phillips et al. (2008), Abdollahzadeh et al. (2009) (morphology and phylogeny); Dissanayake et al. (2016) (accepted number of species, phylogeny); Doilom et al. (2014), Konta et al. (2016) (new species).

78. Cercospora Fresen. ex Fuckel, Hedwigia 2(15): 133 (1863)

Background

Cercospora includes pathogens, saprobes and endophytes. Species are widely distributed, occurring on numerous flowering and ornamental plants, ferns, other fungi (as parasites), gymnosperms, grasses and other monocotyledons such as lilies, magnoliids and palms, mostly causing leaf spots. The well-known asexual morph, which is hyphomycetous, are among the largest groups of plant pathogenic fungi causing leaf spots, leading to diseases on many economically important crops (Agrios 2005; To-Anun et al. 2011; Groenewald et al. 2013; Guatimosim et al 2016; Park et al. 2017). Comparatively only a few sexual morphs have been studied (Hyde et al. 2013). A photosensitizing toxic compound named ‘cercosporin’ is responsible for Cercospora species inhabiting such a wide host range (Daub et al. 2005; Thomas et al. 2020).

Classification—Ascomycota, Dothideomycetes, Dothideomycetidae, Capnodiales, Mycosphaerellaceae

Type species—Cercospora apii Fresen., Beitr. Mykol. 3: 91 (1863)

Distribution—Worldwide

Disease symptoms—Leaf blights and spots

This disease affects the leaves, petioles, stems and peduncles of the tree. Infection and lesion formation initially occur on older leaves before progressing to newer ones. Small, brown flecks develop with a reddish border, expanding to circular spots with an ashy-grey centre. Concentric rings may be observed as individual lesions expand. This tissue becomes thin and brittle, and often drops out, leaving a ragged hole. These lesions often resemble frogeyes, giving this disease its common name. Severely affected leaves wither and die from coalescing lesions (Shane and Teng 1992; Steddom et al. 2005).

Species of Cercospora cause blights and spots on the leaves, petioles, stems and peduncles of trees. Often infection and lesion formation occurs on older leaves before progressing to newer ones. Common symptoms include small, brown lesions that develop with a reddish border, eventually expanding to larger circular or angular spots. Concentric rings may be observed as individual lesions expand. The tissue becomes thin and brittle, and often drops out, leaving a ragged hole. Severely affected leaves wither and die from coalescing lesions (Shane and Teng 1992; Steddom et al. 2005).

Hosts—Wide host range including plant genera in Amaranthaceae, Apiaceae, Asteraceae, Arecaceae, Chenopodiaceae, Convolvulaceae, Cryptogammaceae, Cucurbitaceae, Cyatheaceae, Dennstaedtiaceae, Dioscoreaceae, Euphorbiaceae, Fabaceae, Gunneraceae, Hydrangeaceae, Lamiaceae, Lygodiaceae, Musaceae, Myrtaceae, Onagraceae, Plumbaginaceae, Poaceae, Pteridaceae, Scrophulariaceae, Solanaceae, Thelypteridaceae and Urticaceae (Farr and Rossman 2020).

Cercospora apii causes leaf spot disease on celery and C. beticola on sugar beet (Braun et al. 2013; Guatimosim et al. 2016). The pathogen Cercospora cf. sigesbeckiae infects various plant families, including economically valuable crops such as soybean, causing ‘Cercospora leaf blight’, a disease characterized by leaf bronzing (Albu et al. 2016, 2017). Some other species identified as causative organisms of the leaf blight are C. kikuchii and C. cf. flagellaris (Soares et al. 2015; Rezende et al. 2020). The yield losses related to Cercospora disease have been reported from Canada, China, India and other regions in the USA and South America (Almeida et al. 2005; Cai et al. 2009; Hershman 2009; Wrather et al. 2010; Geisler 2013; Albu et al. 2017; Bandara et al. 2020). Cercospora is among the leading fungal pathogens that cause a severe threat to soybean, which is an important grain legume crop, by reducing seed production and quality (Arantes et al. 2020). Two notable pathogens on soybean are C. kikuchii (leaf blight and purple seed stain) and C. sojina (frogeye leaf spot) (Soares et al. 2015)

Other notable reports include Cercospora leaf spots, which are the most common and destructive of the Hibiscus diseases, often resulting in complete crop loss (Park et al. 2017) and more than 200 fungal species in association with various diseases of ‘kenaf’ (Hibiscus cannabinus) worldwide (Park et al. 2017). Key proteins and expression of genes that could inhibit the pathogen C. kikuchii in soybean (Arantes et al. 2020) have been investigated. However, based on previous reports, morphological characters, phylogeny and pathogenicity of Cercospora cf. nicotianae was identified as one of several cryptic species causing Cercospora leaf blight (Sautua et al. 2019, 2020). Thomas et al. (2020) proposed the expression of fungal cercosporin auto resistance genes and silencing of the cercosporin pathway as effective strategies to combat Cercospora diseases.

Pathogen biology, disease cycle and epidemiology

The taxa survive on undecomposed residues in soil, on weed hosts and seeds. Leaf spot disease is favoured by warm, wet weather. Severe outbreaks generally require a period of showery weather. Infection from germinating fungal spores occurs via penetration of leaf stomata by fungal hyphae. Spores spread in wind, rain, irrigation or via mechanical tools (Vereijssen 2004; Lin and Kelly 2018).

Morphological based identification and diversity

Cercospora has been widely applied to all kinds of dematiaceous hyphomycetous asexual morphs characterized by holoblastic conidiogenesis and some associated with “Mycosphaerella”-like sexual morphs (Hyde et al. 2013; Groenewald et al. 2013). Species resembling the genus type, C. penicillata, characterized by pigmented conidiophores, thickened and darkened conidiogenous loci and singly formed colourless conidia are identified as Cercospora sensu stricto (Ellis 1971, 1976). Chupp (1954) published a worldwide monograph of this group which listed 1,419 species. A vast number of studies related to Cercospora are based on morphology or confined to specific regions or hosts (Phengsintham et al. 2013a, b). Hence, more than 3000 species of Cercospora have been described (Pollack 1987), often as a result of taxa being considered as host-specific at a genus or family level (Crous and Braun 2003; Groenewald et al. 2005). However, based on morphological features of the structure of conidiogenous loci and hila, absence or presence of pigmentation in conidiophores and conidia, Crous and Braun (2003) revised the generic circumscription of Cercospora, resulting in the reduction of the number of species to 659. A series of publications related to Cercospora and its allied genera in Mycosphaerellaceae, along with illustrations and descriptions of sexual morphs was published by Braun et al. (2013, 2014, 2015a, b, 2016).

Molecular based identification and diversity

Cercospora is monophyletic (Stewart et al. 1999; Hyde et al. 2013). Groenewald et al. (2013) provided a comprehensive phylogenetic analysis of 360 isolates which included ITS, and protein-coding genes; translation elongation factor 1-alpha (tef1), actin (act), calmodulin (cal) and histone 3 (his). This provided a basis for the identification of Cercospora species, indicating most to be host-specific (Park et al. 2017). Bakhshi et al. (2018) subjected 170 Cercospora isolates to an eight-gene analysis (tef1, act, cal, his, tub2, rpb2, gapdh) which resulted in several new clades within the C. apii, C. armoraciae, C. beticola, C. cf. flagellaris and Cercospora sp. G. complexes. The combination of tef1, cal, tub2, rpb2 and gapdh provided high phylogenetic resolution for distinguishing Cercospora species with gapdh being the gene effective in distinguishing the species complexes (Bakhshi et al. 2018). The genomes for several species—Cercospora arachidicola, C. aff. canescens, C. cf. sigesbeckiae, C. kikuchii, C. sojina and C. zeae-maydis have been published, of which C. cf. sigesbeckiae and C. sojina are important soybean pathogens (Albu et al. 2017; Sautua et al. 2019). The mating-type genes of some asexual Cercospora species have been characterised (Groenewald et al. 2013), of which C. beticola, C. zeae-maydis and C. zeina are heterothallic, while only one mating type was discovered in populations of C. apii and C. apiicola (Groenewald et al. 2006, 2010).

In soybean cultivation regions such as China, Latin America or the USA, C. sojina occurs as several pathotypes named as races, and their existence differs from soybean cultivar-to-cultivar (Athow et al. 1962; Yorinori and Henechin 1978; Mian et al. 2008; Gu et al. 2020). Apart from being differentiated physiologically, several molecular genetic tools such as AFLPs (Amplified Fragment Length Polymorphisms), SSR markers and SNP markers have been utilized to characterize their population diversity (Gu et al. 2020). The combination of DNA sequence data with ecology, morphological and cultural characteristics named as the Consolidated Species Concept (Quaedvlieg et al. 2014) is an effective method for delimiting Cercospora species (Groenewald et al. 2013; Bakhshi et al. 2015, 2018). Here we provide an updated phylogenetic tree of combined ITS, tef1, act, cal, his, tub2, rpb2 and gapdh (Fig. 5).

Fig. 5

The most parsimonious tree generated by MP analysis of combined ITS, tef1, act, cal, his, tub2, rpb2 and gapdh sequence data of Cercospora species is presented. Related sequences were obtained from previous publications and GenBank. One hundred and fourteen strains are included in the analysis comprising 4222 characters including gaps, of which 2942 characters are constant, 514 characters are parsimony-uninformative and 766 are parsimony-informative. The parsimony analysis of the data matrix resulted in the maximum of 84 equally most parsimonious trees with a length of 3092 steps (CI = 0.557, RI=0.678, RC = 0.382, HI = 0.443) in the first tree. The tree was rooted with Septoria provencialis (CBS 118910). Tree topology of the MP analysis was similar to the ML and BYPP analyses. ML and MP bootstrap support values ≥70% and BYPP ≥0.95 (ML/ MP/ BYPP) are shown respectively near the nodes. Ex-type strains are in bold.

Recommended genetic markers (genus level)—LSU, ITS

Recommended genetic markers (species level)—ITS, tef1, act, cal, his, tub2, rpb2, gapdh

Accepted number of species—There are over 3100 epithets listed in Index Fungorum (2020), however, only 93 have DNA sequence data (Table 4).

Table 4 DNA barcodes available for Cercospora

References—Braun et al. (2013, 2014, 2015a, b, 2016) (morphology), Groenewald et al. (2013) (morphology, phylogeny), Albu et al. (2017) (morphology, phylogeny), Guatimosim et al. (2016) (morphology, phylogeny), Bakhshi et al. (2015, 2018) (morphology, phylogeny).

79. Clinoconidium Pat., Bulletin de la Société Mycologique de France 14: 156 (1898)

Background

Clinoconidium is an important genus that causes smut disease on plants in the family Lauraceae. This genus was established by Patouillard (1898) and typified with Clinoconidium farinosum. Taxonomically, Clinoconidium is placed in Cryptobasidiaceae (Exobasidiales, Exobasidiomycetes, Basidiomycota) and characterized by aseptate, colourless, and globose to ovoid basidiospores which are dispersed individually. The name Clinoconidium was considered illegitimate because of the designation of an illegitimate type species name; however, it was later validated by Saccardo (1902).

Clinoconidium is a gall producing genus which was once named as Ustilago by Ito (1935, 1936) due to the presence of a powdery spore mass on the surface of the galls. This genus was also transferred to another gall producing genus Melanopsichium by Kakishima (1982). However, it was renamed as Clinoconidium as its sorus structure and spore features are quite different from those of Ustilago (Saccardo 1902). The spores of Ustilago species are formed from sporogenous hyphae, whereas this fungus produces spores from hymenial layers in the galls. Spore walls are comparatively thinner than those of Ustilago. The differentiation from Melanopsichium, a gall producing taxon on plants in Polygonaceae (Vánky 2013) includes variation in gall structures and sporulation. Melanopsichium produces spores in chambers formed inside of gall tissues, while this genus produces spores in peripheral lacunae on the surface of gall tissues. The morphological characters of these taxa showed its close similarity to Clinoconidium.

Classification—Basidiomycota, Ustilaginomycotina, Exobasidiomycetes, Exobasidiomycetidae, Exobasidiales, Cryptobasidiaceae

Type species—Clinoconidium farinosum Pat. ex Sacc. & P. Syd

Distribution—Brazil, China, Costa Rica, India, Japan, Panama, Spain, Taiwan and Venezuela

Disease symptoms—mainly observed as powdery pappus gall in fruits. Infection initiates on very young fruits, converted into round, wrinkled galls. The fruit galls are then covered with a powdery mass of spores during early days of infection, withering in the rainy season, leaving behind hard, earthy, brown galls. On Cinnamon, entire young fruits are molded with buff and spongy smut like taxa in the full bloom of disease. Interestingly this infection is restricted to fruits only (Fig. 6).

Fig. 6

Clinoconidium sp. on Cinnamomum sp. a host plant with infected and healthy fruits, b healthy fruits, c, d infected fruits at various stages of infection

Hosts—different plants of Lauraceae including, Apollonias barbujana, Cinnamomum burmannii, C. camphora, C. daphnoides, C. tamala, C. tenuifolium, Nectandra sp., Octea sp., Oreodaphne sp. and Phoebe neurophylla (Farr and Rossman 2020).

Morphological based identification and diversity

This is an important pathogenic genus; producing galls on shoot buds of host plants belonging to the family Lauraceae. Fruits of the host are completely or partially transformed into reddish-brown to dark brown, irregularly malformed, enlarged, globose to subglobose galls; larger than normal fruits. Hymenia formed in peripheral lacunae of the galls are pale yellow to whitish and covered by the host epidermis. Inner tissues of galls consist of hyphae and deformed plant cells. Hyphae are intercellular, hyaline, compact, septate, smooth-walled and lack clamp connections, while haustoria are intercellular, slightly lobed to irregular and observed in deformed host cells. Upon maturation, galls rupture, exposing orange to dark brown or creamish white spore masses which cover the entire infected young fruits. Sterile hyphae can be found intermingled between the basidia in some species and are indistinguishable from young basidia or absent in some species of Clinoconidium. Basidia are clavate, hyaline, depressed, difficult to observe and gastroid, densely aggregated in masses, formed in irregular fascicles from basally agglutinated hyphae and the wall is densely foveolate when mature. Basidiospores are ellipsoid, clavate, pyriform, fusoid, globose, subglobose to oval, aggregated in a creamish white to brown coloured masses on the surface of the galls, hyaline or wall pale brown to brown, rugose when mature; producing long branched hyphae with septa when germinated on culture media and proliferating sympodially.

Molecular based identification and diversity

There are seven epithets of Clinoconidium recorded on various plant hosts. Sequence data for Clinoconidium bullatum, C. cinnamomi, C. onumae and C. sawadae are available in GenBank, including sequence data for LSU and ITS. Clinoconidium farinosum and C. globosum lack sequence data in GenBank. ITS and LSU are the most suitable loci for delineation of species within the genus (Fig. 7).

Fig. 7

Phylogram generated from MP analysis based on combined sequences of LSU and ITS sequences of all the species of Clinoconidium with molecular data. Related sequences were obtained from GenBank. Five taxa are included in the analyses, which comprise 1100 characters including gaps, of which 910 characters are constant, 182 characters are parsimony-uninformative, eight characters parsimony-informative. The parsimony analysis of the data matrix resulted in the maximum of two equally most parsimonious trees with a length of 202 steps (CI = 0.980, RI 0.500, RC = 0.490, HI = 0.020) in the first tree Single gene analyses were carried out and compared with each species, to compare the topology of the tree and clade stability. The tree was rooted with Microbotryum violaceum (AFTOL-ID1819). Maximum parsimony bootstrap support value ≥ 50% and BYPP ≥ 0.9 are shown respectively near the nodes

Recommended genetic markers (genus level)—ITS, LSU

Recommended genetic markers (species level)—ITS, LSU

Accepted number of species—There are seven species epithets in Index Fungorum (2020), however, only four species have DNA molecular data (Table 5).

Table 5 DNA barcodes available for Clinoconidium

References—Hendrichs et al. (2003), Jiang and Kirschner (2016), Kakishima et al. (2017a, b) (morphology, phylogeny)

80. Cylindrocladiella Boesew., Canadian Journal of Botany 60 (11): 2289 (1982)

= Nectricladiella Crous & C.L. Schoch, Studies in Mycology 45: 54 (2000)

Background

Boeswinkel (1982) established Cylindrocladiella to accommodate five Cylindrocladium-like species producing small, cylindrical conidia. Even though the generic status of Cylindrocladiella was initially opposed by Crous and Wingfield (1993), later studies on morphological comparisons by Crous et al. (1994) and molecular data (Victor et al. 1998; Schoch et al. 2000) supported the establishment of Cylindrocladiella as a genus. This genus is commonly confused with the asexual morph of Calonectria but can be distinguished by clear morphological differences, such as aseptate stipe extensions, different branching patterns of the conidiophores and comparatively small, aseptate conidia. Although species are generally not regarded as important plant pathogens, correct identification is essential for disease control and biosecurity implications.

Classification—Ascomycota, Sordariomycetes, Hypocreomycetidae, Hypocreales, Nectriaceae

Type species—Cylindrocladiella parva (P.J. Anderson) Boesew.

Distribution—as a soil-borne fungus, the species in Cylindrocladiella have a cosmopolitan distribution in various geographically and climatically distinct regions around the world (Farr and Rossman 2020).

Disease symptoms—black-foot disease, damping-off, leaf spot, root rot and shoot die-back

Many species belonging to Cylindrocladiella are opportunistic plant pathogens but they are not considered as primary pathogens. They can be isolated associated with disease symptoms such as leaf spot, damping off and shoot die-back (Scattolin and Montecchio 2007; Pham 2018). Chocolate brown lesions around the shoots spread primarily to be followed by wilting of the shoot tip, reddish discolouration, dropping of leaves, and finally plant death (Brielmaier-Liebetanz et al. 2013). Characteristic symptoms of the black-foot disease include a reduction in root biomass and root hairs with sunken and necrotic root lesions (Agustí-Brisach and Armengol 2013). Symptoms of Cylindrocladiella root rot are black lesions on the tap and lateral roots, wilting and foliar necrosis, and the outer bark of the seedlings will crack and become loose (Sinclair and Lyon 2005).

Hosts—Species are soil-borne, weak pathogens of forestry, agricultural and horticultural crops. There are 270 records of Cylindrocladiella associated with different plant species (Farr and Rossman 2020). Among them, different Vitis species and Eucalyptus species are common hosts associated with different species of Cylindrocladiella.

Morphological based identification and diversity

Cylindrocladiella can be distinguished from related species by penicillate and/or subverticillate symmetrically branched conidiophores which produce small, cylindrical, 1-septate conidia and aseptate stipe extensions (Lombard et al. 2012). The generic status of Cylindrocladiella was earlier strongly contested (Sharma and Mohanan 1991), however, based on morphological evaluation and comparisons by Crous and Wingfield (1993) and Crous et al. (2017) confirmed its generic status. Victor et al. (1998) and Schoch et al. (2000) provided molecular data to support generic status. Lombard et al. (2012) in his revision of Cylindrocladiella mentioned that only two species have been recognized with their respective Nectricladiella sexual morph. Rossman et al. (2013) proposed that the generic name Cylindrocladiella be used rather than Nectricladiella. Lombard et al. (2015) showed that Cylindrocladiella formed a monophyletic group in Nectriaceae (Wijayawardene et al. 2020).

Molecular based identification and diversity

Using RFLPs and AT-DNA data, Victor et al. (1998) recognised seven species in the genus. Schoch et al. (2000) added another species based on ITS and partial tub2. Van Coller et al. (2005) introduced the use of his3 sequence data for this group. A combined multilocus phylogeny of his, tef1, tub2 and ITS was used by Lombard et al. (2012) which resulted in 18 new Cylindrocladiella species and several unresolved species complexes. Lombard et al. (2017) introduced six new species based on a combined ITS, tef1 and tub2 dataset. Pham (2018) introduced five new species based on his, tef1, tub2 and ITS sequence data and Marin-Felix et al. (2019) introduced two new species based on ITS, tef1 and tub2 sequence data. Here we reconstruct the phylogenetic analyses of these species based on ITS, tef1 and tub2 sequence data (Fig. 8).

Fig. 8

Phylogram generated from MP analysis based on combined sequences of ITS, tef1 and tub2 sequences of all the accepted species of Cylindrocladiella. Related sequences were obtained from GenBank. Fourty-six taxa are included in the analyses, which comprise 2460 characters including gaps. Single gene analyses were carried out and compared with each species, to compare the topology of the tree and clade stability. The tree was rooted with Gliocladiopsis sagariensis (CBS 19955). The best scoring RAxML tree with a final likelihood value of − 6772.195394 is presented. The matrix had 261 distinct alignment patterns, with 0.96% of undetermined characters or gaps. Estimated base frequencies were as follows: A = 0.230657, C = 0.279364, G = 0.252128, T = 0.237852; substitution rates AC = 1.388608, AG = 2.845402, AT = 2.389715, CG = 0.838197, CT = 7.220493, GT = 1.000000; gamma distribution shape parameter a = 0.650385. Maximum likelihood and MP bootstrap support value > 50% are shown respectively near the nodes. Ex-type strains are in bold

Recommended genetic markers (genus level)—ITS, LSU

Recommended genetic markers (species level)—his, tef1, tub2

Accepted number of species—There are 47 species epithets in Index Fungorum (2020). However, only 46 species have DNA sequence data (Table 6).

Table 6 DNA barcodes available for Cylindrocladiella

References—Crous and Wingfield (1993), Lombard et al. (2012) (morphology); Victor et al. (1998), Schoch et al. (2000), Lombard et al. (2015) (morphology, phylogeny).

81. Dothidotthia Höhn., Berichte der Deutschen Botanischen Gesellschaft 36: 312 (1918)

Background

Dothidotthia was assigned to Botryosphaeriaceae, because of its coelomycetous asexual morph, and characteristic peridium, pseudoparaphyses and asci (Barr 1989). Ramaley (2005) reported that Thyrostroma is the asexual morph of Dothidotthia based on the production of hyphomycetes in culture. Phillips et al. (2008), introduced a new family Dothidotthiaceae to accommodate Dothidotthia and considered Thyrostroma as the asexual morph of Dothidotthia. However, the links between the sexual and asexual morphs are not supported by molecular evidence. Recent molecular and morphology studies (Marin-Felix et al. 2017; Crous et al. 2019; Senwanna et al. 2019), based on a taxon sampling of current species indicates that Dothidotthia does not cluster near Thyrostroma. Thus, Dothidotthia is a distinct genus.

Classification—Ascomycota, Pezizomycotina, Dothideomycetes, Pleosporomycetidae, Pleosporales, Dothidotthiaceae

Type species—Dothidotthia symphoricarpi (Rehm) Höhn.

Distribution—in both temperate and tropical countries (Italy, Russia, Thailand, Ukraine and the USA)

Disease symptoms—species cause canker, dieback and leaf spot diseases on twig, branch, bark and leaf

Hosts—Pathogens of Acer negundo, Diapensia lapponica, Fendlera rupicola, Euonymus alatus, Robinia pseudoacacia, Verbena asparagoides (Barr 1989; Farr and Rossman 2020; Index Fungorum 2020).

Morphological based identification and diversity

In previous studies, the asexual morphs of Dothidotthia have been reported as Thyrostroma (Ramaley 2005), however, phylogenetic analyses indicated that Dothidotthia can be separated from Thyrostroma (Marin-Felix et al. 2017; Crous et al. 2016; Senwanna et al. 2019). Dothidotthia is characterized by fusiform to obclavate or obpyriform, 0–3-transversely septate conidia and a sexual morph with clavate, short pedicellate asci, ellipsoid, 1-septate ascospores (Fig. 9). The sexual morphs of Dothidotthia and Thyrostroma have similar morphological characteristics in shape and overlapping dimensions of asci and ascospores (Barr 1989; Ramaley 2005; Phillips et al. 2008; Hyde et al. 2013; Senwanna et al. 2019). However, Dothidotthia can be differentiated from Thyrostroma by peridium structure and conidial morphology and molecular phylogeny (Senwanna et al. 2019). Crous et al. (2019) introduced Neodothidotthia to accommodate N. negundinicola and Dothidotthia aspera was synonymized under N. negundinis based on analysis of LSU sequence data. However, Senwanna et al. (2019) showed that Neodothidotthia negundinicola and N. negundinis group with D. robiniae and D. symphoricarpi (type species). Furthermore, the conidial morphology of Neodothidotthia is similar to Dothidotthia symphoricarpi (Pseudotthia symphoricarpi) and D. robiniae (Phillips et al. 2008; Zhang et al. 2012; Crous et al. 2019; Senwanna et al. 2019). Therefore, Neodothidotthia had been treated as a synonym of Dothidotthia.

Fig. 9

Dothidotthia robiniae (MFLU 16-1704). a, b Sporodochia on the host surface. c Vertical section of sporodochium. d Conidiogenesis. e, g Conidia attached with the conidiogenous cells. f, h Conidia. i Germinated conidium. Scale bars: b = 1000 µm, c = 200 µm, d–i = 30 µm

Molecular based identification and diversity

Dothidotthia species can be separated from Thyrostroma based on LSU sequence data (Marin-Felix et al. 2017; Crous et al. 2019). Multigene phylogenetic analyses of a combined LSU, SSU, ITS and tef1 dataset for Dothidotthia is presented in this study, which is similar to Senwanna et al. (2019) (Fig. 10).

Fig. 10

Phylogenetic tree generated by ML analysis of LSU, SSU, ITS and tef1 sequence data of Dothidotthia species. Related sequences were obtained from GenBank. The tree was rooted with Thyrostroma compactum (CBS 335.37) and T. lycii (MFLUCC 16-1170). Tree topology of the ML analysis was similar to the Bayesian analysis. The best scoring RAxML tree with a final likelihood value of − 5116.933762 is presented. The matrix had 115 distinct alignment patterns, with 25.41% of undetermined characters or gaps. Estimated base frequencies were as follows: A = 0.245094, C = 0.237101, G = 0.269739, T = 0.248067; substitution rates AC = 3.925871, AG = 7.445430, AT = 2.745308, CG = 2.728664, CT = 20.049514, GT = 1.000000; gamma distribution shape parameter α = 0.790240. Maximum likelihood bootstrap support values greater than 60% and BYPP probabilities ≥ 0.95 are indicated above the nodes. Ex-type (ex-epitype) and voucher strains are in bold

Recommended genetic markers (genus level)—LSU, SSU

Recommended genetic markers (species level)—ITS, tef1, rpb2 and tub2

Accepted number of species—There are 14 epithets listed in Index Fungorum (2020), however only four species have DNA molecular data (Table 7).

Table 7 DNA barcodes available for Dothidotthia

References—Barr (1989), Ramaley (2005) (morphology); Phillips et al. (2008), Zhang et al. (2012), Hyde et al. (2013), Marin-Felix et al. (2017), Crous et al. (2019), Senwanna et al. (2019) (morphology and phylogeny)

82. Erysiphaceae Tul. & C. Tul. [as ‘Erysiphei’], Select. fung. carpol. (Paris) 1: [191] (1861)

Background

Powdery mildews belong to Erysiphales of Ascomycota (Mori et al. 2000). Powdery mildews are one of the most prevalent and easily recognizable of plant diseases (Glawe 2008). Mucor erysiphe, published by Linnaeus (1753), was the first binomial referring to powdery mildew (now known as Phyllactinia guttata) (Braun and Cook 2012). Infections are often conspicuous owing to the profuse production of conidia that give them their common name. Powdery mildews are also models for basic research on host-parasite interactions, developmental morphology, cytology, and molecular biology (Glawe 2008). Erysiphaceae is obligately parasitic and as such, their life cycle depends completely on living hosts, from which they obtain nutrients without killing host cells and without which they are unable to survive. As they are obligate plant pathogens, researchers have not had the advantage of routinely cultivating these taxa on artificial media. However, many powdery mildews have been grown on detached leaves of their hosts (Hirose et al. 2005). Powdery mildews seldom kill their host, but are responsible for water and nutrient loss and impaired growth and development. They can increase respiration and transpiration and interfere with photosynthesis and reduce yields.

Changes in host range directly cause the niche separation of powdery mildews and thus may become a trigger of speciation in their evolution. It is possible that studying the evolutionary history of powdery mildews will not only reveal facts on fungal evolution but may also lead us to consider the evolutionary history of angiosperm plants (Takamatsu 2004; Matsuda and Takamatsu 2003; Hirata et al. 2000; Mori et al. 2000).

The first systematic trial to identify the conidial states of powdery mildews at the species level was made by Ferraris (1910), who grouped species of Oidium according to the size and shape of their conidia and provided a key to its species. Foex (1913), Jaczewski (1927), and Brundza (1934) contributed to the classification of the conidiophore types. Jaczewski (1927) introduced the terms ‘Euoidium and Pseudoidium’ for Oidium states with catenate and solitary conidia, respectively. Yarwood (1957) provided a survey on the Erysiphaceae, including the asexual morphs. Boesewinkel (1980) provided the first comprehensive key based on a combination of more than 12 morphological characteristics observed on conidia, conidiophores, appressoria, haustoria, fibrosin bodies, and mycelium. Braun (1987) issued a second comprehensive monograph of the Erysiphales encompassing all powdery mildew taxa known at that time. Shin and La (1993) and Shin and Zheng (1998) introduced some new morphological features of taxonomic relevance. A progressive report was provided by the work of Cook et al. (1997), who examined the surface of conidia by scanning electron microscopy and separated Oidium into eight subgenera. Braun (1999) discussed the classification of Erysiphaceae as proposed by Cook et al. (1997) and introduced some corrections and alterations. Fundamental innovations in the generic taxonomy of the group based on molecular and SEM examination and a better insight into the phylogeny are results of comprehensive investigations over the last decade (Takamatsu et al. 1998, 1999, 2000, 2005a, b, 2008; Matsuda and Takamatsu 2003; Hirose et al. 2005; Liberato et al. 2006; Braun and Cook 2012).

Classification—Ascomycota, Pezizomycotina, Leotiomycetes, Leotiomycetidae, Erysiphales

Type genus—Erysiphe R. Hedw. ex DC.

Distribution—worldwide

Disease symptoms—powdery mildew

The initial signs of infection appear on young leaves in the form of small, raised blisters, which cause the leaves to curl and expose the under surfaces. As the disease progresses, round, pinpoint powdery white spots dusting the upper surfaces of leaves, as well as stems and occasionally fruiting occurs. As the disease becomes severe, the spots will become larger, and more interconnected and irregular in shape. Over time they progress from younger to older leaves and the undersides of leaves. However, mature leaves are usually much less severely infected than new or young leaves. If the white patches (which have a granular, powdery texture) are wiped away, the growths will return in a matter of days. Severely infected leaves will turn yellow, dry out and drop from the plant. Buds and growing tips of shoots can also become infected, eventually becoming distorted and stunted (Bushnell and Allen 1962; Davis et al. 2001; Romero et al. 2003; Oberti et al. 2014; Saharan et al. 2019).

Hosts- The host range of this fungal group is strictly confined to angiosperms and powdery mildews have never been reported to infect ferns or gymnosperms (Amano 1986; Hirata et al. 2000; Takamatsu et al. 2010). They affect a wide range of angiosperms such as cereals and grasses, vegetables, ornamentals, weeds, shrubs, fruit trees, and broad-leaved shade and forest trees. Powdery mildews are considered as host-specific.

Pathogen biology, disease cycle and epidemiology

Powdery mildews tend to grow superficially, or epiphytically, on plant surfaces. During the growing season, hyphae are produced on both the upper and lower leaf surfaces, although some species are restricted to one leaf surface. Infections can also occur on stems, flowers or fruit. Specialized absorption cells, termed haustoria, extend into the plant epidermal cells to obtain nutrition. While most powdery mildews produce epiphytic mycelium, a few genera produce hyphae that are within the leaf tissue; this is known as endophytic growth. Conidia are produced on plant surfaces during the growing season. They develop either singly or in chains on conidiophores. Conidiophores arise from the epiphytic hyphae, or in the case of endophytic hyphae, the conidiophores emerge through leaf stomata. At the end of the growing season, powdery mildews produce ascospores, in a sac-like ascus enclosed in a fruiting body called a chasmothecium. The chasmothecium is generally spherical with no natural opening; asci with ascospores are released when a crack develops in the wall of the fruiting body. A variety of appendages may occur on the surface of the chasmothecia. These appendages are thought to act as the hooks of a velcro fastener, attaching the fruiting bodies to the host, particularly to the bark of woody plants, where they overwinter. They can survive winter conditions as dormant mycelia within the buds and other plant tissue of the host. These infected parts of the host can be the source of primary inoculum that can initiate further infection when conditions are right (Misra 2001; Amsalem et al. 2006; Heffer et al. 2006; Te Beest et al. 2008; Saharan et al. 2019; Fig. 11).

Fig. 11

The life cycle of a powdery mildew fungus on roses. Redrawn from Agrios (2005) and Mulbrhan et al. (2016)

Morphological based identification and diversity

Members of Erysiphaceae cause powdery mildew disease on about 10,000 angiosperm species (Takamatsu et al. 2010). The Erysiphaceae are divided into five tribes and two basal genera (Cook et al. 1997). Both tree-parasitic and herb-parasitic species are included in three of the five tribes: Cystotheceae, Erysipheae and Phyllactinieae. Tree-parasitic species usually take basal positions in these tribes and herb-parasitic species have derived positions. The tribe, Golovinomycetea is a group derived from a single ancestor (Mori et al. 2000). The monophyly of the tribe is also supported by the common characteristics, i.e., ectophytic parasitism, polyascal ascomata, and Euoidium asexual morphs, with the latter producing conidia in chains without distinct fibrosin bodies. Of these five lineages, four consists of taxa infectious to dicotyledons. Blumeria graminis, which is infectious to monocotyledon plants, formed an independent lineage. Therefore, Blumeria graminis was accommodated in a monotypic tribe Blumerieae in the new system (Inuma et al. 2007).

The powdery mildew belonging to the tribe Cystotheceae have both herbaceous and woody plants as hosts and consist of three genera, Cystotheca, Podosphaera and Sawadaea, of which Cystotheca and Sawadaea are restricted to a narrow range of host families (Meeboon et al. 2013). Podosphaera consists of two sections, Podosphaera and Sphaerotheca. Section Podosphaera parasitizes woody plants (Takamatsu et al. 2000). The tribe Golovinomyceteae consists of three genera, Golovinomyces, Neoerysiphe, and Arthrocladiella. Arthrocladiella is a monotypic genus consisting of a single species A. mougeottii and has only the host genus Lycium. Neoerysiphe is also a small genus composed of four species and has about 300 herbaceous host species ranging across five plant families including Lamiaceae. Golovinomyces is a large genus comprising 27 species (Braun 1987), and it is widely distributed in the world. The tribe Phyllactinieae comprises the genera Phyllactinia, Leveillula, Pleochaeta and Queirozia which typically have hemi-endophytic (partly external and partly internal mycelia in common (Braun 1987; Liberato 2007; Liberato et al. 2006; Khodaparast et al. 2001; Ramos et al. 2013).

The tribe Erysipheae forms a separate, monophyletic clade, which is characterized by asexual morphs belonging to Oidium subgen. Pseudoidium Jacz (Takamatsu et al. 1999; Mori et al. 2000). This clade comprises Erysiphe and its sections Erysiphe, Microsphaera and Uncinula. Uncinula forestalis differs from the species of Erysiphe sect. Uncinula in having terminal, fasciculate, septate, ascoma appendages and Euoidium-like asexual morph (conidia catenate) and therefore it was placed in Caespitotheca (Takamatsu et al. 2005b). Because of the lack of asexual morphs in Uncinula septata and U. curvispora and multiseptate chasmothecial appendages arising from the upper half the fruiting body, the two species were assigned to Parauncinula (Braun and Takamatsu 2000; Takamatsu et al. 2005a). A unique taxon, Oidium phyllanthi, on Phyllanthus acidus, P. amarus and P. reticulatus produces a germination type designated as Microidium-type and was placed in a new genus Microidium (To-anun et al. 2005). With these new classifications, Erysiphales contains 17 accepted genera, 16 based on the holomorph and one on the asexual morph (Braun and Cook 2012). With the descriptions of several new species, the number of recognized powdery mildew species has increased from 515 (including 435 sexual morphs/holomorphs) in Braun (1987), to about 820 species (including about 685 sexual morphs/holomorphs) (Braun and Takamatsu 2000; Braun et al. 2002; Takamatsu et al. 2005a, b; Liberato et al. 2006; Braun and Cook 2012).

Molecular based identification and diversity

Molecular data have proven useful in reassessing species and clarifying the taxonomic significance of morphology and host data. Only a few of the described species have been reassessed using molecular data (Braun and Cook 2012). Reports began appearing in the 1990s, that used ITS and 18S rDNA sequences to infer phylogenetic relationships of Erysiphales and other major ascomycete groups (Saenz and Taylor 1999; Saenz et al. 1994). Analyses of 18S rDNA, ITS1–5.8S-ITS2, and 28S rDNA sequences led to the opinion that Erysiphales can be placed in Leotiomycetes along with Cyttariales, Helotiales, and Rhytismatales (Wang et al. 2006). Phylogenetic analyses demonstrated that Erysiphaceae formed a distinct monophyletic group (Hirata et al. 2000). Thus, Erysiphaceae is derived from a single ancestral taxon that may have acquired parasitism just once (Mori et al. 2000a; Takamatsu 2004; Wang et al. 2006). Shirouzu et al. (2020) using nrDNA and mcm7 sequence data showed that Phyllactinieae is not monophyletic. However, there is a need to re-assess the tribes in this family to establish them as subfamilies or genera. In this paper, we present a phylogenetic tree with combined ITS and LSU sequences obtained from available type material and voucher specimens (Table 8, Fig. 12). This can be used as a backbone in the identification of powdery mildew species.

Table 8 Genera in Erysiphaceae

Fig. 12

Phylogram generated from parsimony analysis based on combined ITS and LSU sequenced data Erysiphaceae. Maximum parsimony bootstrap support values greater than 60% and BYPP greater than 0.90 are indicated above the nodes. The type specimens (ex-epitypes) are in bold. The tree is rooted with Parauncinula septata

Recommended genetic markers (genus level)—ITS, LSU and SSU

Recommended genetic markers (species level)—tub2, chs, tef1

The ITS region of the precursor molecules of rRNA was revealed to form a secondary structure including several stem-loop structures, and some conserved sequences are found in the stem regions (Takamatsu et al. 1998). This makes it possible to design PCR primers that work for a wide range of the powdery mildews. Takamatsu and Kano (2001) designed four new PCR primers that are useful to determine the nucleotide sequences of the rDNA of the powdery mildews. These primers provide stability to work on a wide range of powdery mildews and specificity to eliminate contaminating DNA by PCR. Primer sets PM3/P3, ITS1/PM4, PM5/P3, and ITS1/PM6 were tested with universal primer set ITS1/ITS4 (White et al. 1990) covering all major clades of Erysiphales. Meeboon and Takamatsu (2013a) used LSU, ITS and IGS (Inter generic spacer) sequences to identify two different genetic groups of Erysiphe japonica (= Typhulochaeta japonica), powdery mildew on Quercus species based on the differences in host range. Cho et al. (2014) used ITS and 28S rDNA for the introduction of the powdery mildew species Erysiphe magnoliicola in Erysiphe sect. Microsphaera. Wang et al. (2014) also used ITS differences for phylogenetic analysis of powdery mildew disease on mulberry in Yunnan Province. Meeboon and Takamatsu (2013b) also used the 28S rDNA sequences and a combined alignment of the 28S, ITS, and IGS (Intergeneric spacer) rDNA sequences to construct a phylogeny of Erysiphe sect. Uncinula on Carpinus species and showed the cryptic species Erysiphe paracarpinicola. de Oliveira et al. (2015) used ITS sequences of Erysiphe platani on Platanus × acerifolia in Brazil as new records of taxa. Liyanage et al. (2017) used ITS, SSU and LSU sequences to identify E. quercicola infected rubber trees. Phylogenetic analyses of B. graminis based on the DNA sequences of four DNA regions, i.e. ITS, 28S rDNA, chitin synthase 1, and ß-tubulin were conducted by Inuma et al. (2007) to revealed distinct groups in the B. graminis isolates from a single host genus belonged to a single group.

83. Fomitopsis P. Karst., Meddn Soc. Fauna Flora fenn. 6: 9 (1881)

Background

Fomitopsis was established by Karsten (1881) based on four species, with F. pinicola as the generic type (Murrill 1903; Donk 1960). The genus has a cosmopolitan distribution and comprises species causing brown rot on both living and dead trees (Han et al. 2016). Fomitopsis species also contribute to the decomposition of coarse woody debris in forest communities (Gilbertson 1980; Haight et al. 2019). There are certain instances of their pathogenic role in orchards of cultivated species where they cause heart rot on Citrus (Roccotelli et al. 2014) and Prunus species (Adaskaveg 1993). A Fomitopsis sp. was also recorded in oil palm (Elaeis guineensis) as an endophyte (Rungjindamai et al. 2008; Pinruan et al. 2010).

Classification—Basidiomycota, Agaricomycetes, Incertae sedis, Polyporales, Fomitopsidaceae

Type species—Fomitopsis pinicola (Sw.) P. Karst.

Distribution—Worldwide

Disease symptoms—Fomitopsis causes brown cubical rot on both living and dead trees (Mounce 1929). The basidiospores can be dispersed by wind, or by vectors such as bark beetles (Castello et al. 1976; Pettey and Shaw 1986; Lim et al. 2005; Persson et al. 2011; Jacobsen et al. 2017; Vogel et al. 2017). Upon infecting standing trees, stumps, or logs through wounds, or through the tunnels of penetrating vectors, the fungus establishes itself in the xylem (Mounce 1929). The growth rate of Fomitopsis species in the substrata can differ depending on their ecological requirements (Markovic et al. 2011; Haight et al. 2019). When the decay starts, the wood turns yellowish-brown, which later splits into cubical fragments. The colour is generally lighter in case of F. pinicola than other agents of brown rot decay (Markovic et al. 2011). White mycelial felts can also develop in shrinkage cracks of the decayed wood (Ryvarden and Gilbertson 1993). After establishment, the perennial basidiome appears relatively rapidly (Mounce 1929, Fig. 13). The infection results in the breakage of treetops, or further infection of the base of the trees and weakening of larger roots, which may lead to eventual windthrow of standing trees.

Fig. 13

Fomitopsis pinicola a basidiomes on living European spruce, b causing brown-rot decay on narrow-leafed ash, c, d basidiomes on dead standing conifer tree, e young basidiome on hardwood log, f hyphal structure in the trama, g, h basidiospores. Scale bars: f = 20 µm, g, h = 5 µm

Hosts—The type species, F. pinicola mostly appears on gymnosperms, such as Abies, Larix, Picea and Pinus, but can also be found on angiosperms such as Acer, Alnus, Betula, Carpinus, Corylus, Elaeagnus, Fagus, Fraxinus, Malus, Populus, Prunus, Pyrus, Quercus, Salix, Sorbus, Tilia, Ulmus (Ryvarden and Gilbertson 1993; Dai 2012). The North American species in the Fomitopsis pinicola species complex have also been reported from Pseudotsuga, Sequioa and Tsuga (Haight et al. 2019). Other Fomitopsis species can be found on Ginkgo, Pinus and various angiosperm genera, such as Betula, Castanopsis, Cinamomum, Citrus, Delonix, Fagus, Eucalyptus Ligustrum, Prunus, Quercus and Tilia (Ryvarden and Gilbertson 1993; Dai 2012; Li et al. 2013; Han et al. 2016; Liu et al. 2019).

Morphological based identification and diversity

Based on morphological evidence, over 40 species were accepted in Fomitopsis (e.g. Ryvarden and Johansen 1980; Gilbertson and Ryvarden 1986; Ryvarden and Gilbertson 1993; Núñez and Ryvarden 2001; Hattori 2001). However, phylogenetic studies showed that the morphologically defined Fomitopsis was polyphyletic and taxa clustered with other brown-rot genera in the antrodia clade (Ortiz-Santana et al. 2013; Han et al. 2016). Han et al. (2016) showed that Pilatoporus and Piptoporus are synonyms of Fomitopsis sensu stricto, while the segregation of Rhodofomes was confirmed and five new genera were proposed. Fomitopsis sensu stricto is characterized by annual to perennial, mostly sessile, occasionally effused-reflexed or substipitate, soft, corky, tough to woody basidiocarps, a dimitic hyphal system with clamped generative hyphae and cylindrical to ellipsoid, hyaline, thin-walled, smooth basidiospores which are negative in Melzer’s reagent, and cause brown rot (Fig. 13).

Molecular based identification and diversity

Comprehensive multigene analyses by Han et al. (2016) accepted ten species in Fomitopsis sensu stricto. Two new Fomitopsis species were described from Brazil, F. flabellata and F. roseoalba (Tibpromma et al. 2017). Fomitopsis flabellata was transferred to Rhodofomitopsis and the new combination Fomitopsis bondartsevae was proposed (Soares et al. 2017). Mating studies and molecular phylogenetic analyses resolved four cryptic lineages in the F. pinicola species complex (Haight et al. 2016), that represents three North American species (F. mounceae, F. ochracea and F. schrenkii), and F. pinicola sensu stricto, which is restricted to Eurasia (Ryvarden and Stokland 2008; Haight et al. 2019). Three new species were proposed by Liu et al. (2019) from Australia (F. eucalypticola), Puerto Rico (F. caribensis), and China (F. ginkgonis).

The phylogenetic tree of Fomitopsis presented here is based on analyses of a combined ITS, LSU, tef1 and rpb2 sequence data (Fig. 14). In our analyses, it appears that the type of F. bondartsevae is identical to F. iberica and F. hemitephra sensu stricto (Han et al. 2016), which are grouped close to F. palustris and other species formerly discussed in Pilatoporus. Therefore, a thorough revision of the pilatoporus clade is recommended to clarify the status of these species.

Fig. 14

Phylogram generated from RAxML analysis based on combined ITS, LSU, nSSU, tef1 and rpb2 sequence data of Fomitopsis species. Related sequences were obtained from GenBank. Thirty-one strains are included in the analyses, which comprised 4143 characters including gaps. The tree was rooted with Daedalea quercina (Dai 12152) and D. dickinsii (Yuan 1090). Tree topology of the ML analysis was similar to the Bayesian analysis. ML bootstrap values ˃ 50% and BYPP ˃ 0.80 are shown respectively near the nodes

Recommended genetic marker (genus level)—LSU

Recommended genetic markers (species level)—ITS, tef1, rpb2

Accepted number of species—There are 104 epithets listed in Index Fungorum (2020). However, only 17 species have DNA sequence data (Table 9).

Table 9 DNA barcodes for accepted species of Fomitopsis

References—Li et al. (2013) (phylogeny, new species), Han et al. (2016) (phylogeny), Haight et al. 2019 (phylogeny, new species), Floudas et al. (2012) (genome, F. pinicola), Hong et al. (2017) (genome, F. palustris), Liu et al. (2019) (phylogeny, new species).

84. Ganoderma P. Karst., Revue mycol., Toulouse 3(no. 9): 17 (1881)

Background

Ganoderma was established by Karsten (1881) based on G. lucidum and characterized by double-walled basidiospores with truncate apices and ornamented endospores, and a crusty or shiny pileus surface (Moncalvo and Ryvarden 1997). This genus was divided into two subgenera, Ganoderma and Elfvingia by Karsten (1889). Various authors used different taxonomic characters for the identification of species (e.g., Murrill 1902, 1903; Atkinson 1908; Coleman 1927; Corner 1947), which resulted in an intricate taxonomy, with 344 species names in speciesfungorum.org, but an estimated 180 species (He et al. 2019) and Steyaert (1972, 1980) worked extensively on the genus and introduced many new species, transferred many to the genus and removed several synonyms. Ryvarden (1985) and Gottlieb and Wright (1999a,b) studied the macro- and micromorphology. Ganoderma presently comprises sections Amauroderma and Ganoderma, subgenera: Ganoderma and Trachyderma (Index Fungorum 2020, Wijayawardene et al. 2020).

Relevant characteristics for Ganoderma species delimitation are based on the macro and micromorphological characteristics (see in Fig. 15). The basidiomes are annual or perennial, dimidiate, sessile or substipitate to stipitate, with distinctive non-laccate (dull) or weakly to strongly laccate, glossy, shiny, smooth, spathulate, furrows, which are sulcate on the pileus surface. Some strains have several layers of thick, dull cuticles or shiny, with thin cuticle or cuticle of clavate end cells. The context is cream to dark purplish brown, brown to dark brown, sometimes spongy to firm-fibrous. Pores are 4–7 per mm, angular, entire, subcircular to circular, regular, mostly cream or white when young, light yellow to brown when mature, which are usually white to cream when fresh, turning pale yellow on drying, with bruising brown of pore surface. The tube layer is single or stratified, with pale to purplish brown, hard, and becomes woody when dry. The stipe is central or lateral when present.

Fig. 15

Morphology of Ganoderma species. a An old basidiome of Ganoderma australe, b Mature basidiome of G. casuarinicola, c, d hyphae, e tube layer hyphae, f–h Basidiospores, i Pore characteristics. Scale bars: a, b = 2 cm; c, d = 3 μm; e = 15 μm; f, g, h. = 5 μm; i = 500 μm

The Ganoderma hyphal system is di-trimitic and generative hyphae are thin-walled or occasionally thick-walled, with clamp connections. Skeletal hyphae are hyaline to brown, thick-walled, often long, unbranched. Binding hyphae are almost colourless, thin to thick-walled, branched and with clamp connections. Basidiospores are 7–30 μm long, usually broadly to narrowly ellipsoid, truncate, double-walled, and with an apical germ pore. The endosporium is brown and separated from the hyaline exosporium by inter-wall pillars, negative in Melzer’s reagent (Núñez and Ryvarden 2000; Ryvarden 2004). Basidia are broadly ellipsoid, tapering abruptly at the base, and cystidia are lacking.

Ganoderma species are widely distributed in temperate, subtropical and tropical regions, and appear to thrive in hot and humid conditions (Pilotti et al. 2004; Hapuarachchi et al. 2019a, b; Luangharn et al. 2019). Basidiomes are commonly in the form of a bracket (Pilotti et al. 2004). Ganoderma is cosmopolitan and an important wood-decaying genus. Some species of Ganoderma are pathogenic, causing root and stem rot on a variety of monocotyledons, dicotyledons and gymnosperms, including a wide range of economically important trees and perennial crops which results in the death of affected trees (Hapuarachchi et al. 2018b). Ganoderma grows as facultative parasites of trees but can also live as saprobes on rotting stumps and roots (Turner 1981; Pilotti et al. 2004). Hence, they have ecological importance in the breakdown of woody plants for nutrient mobilization. Taxa also possess effective machinery of lignocellulose-decomposing enzymes which may be useful for bioenergy production and bioremediation (Hepting 1971; Kües et al. 2015; Hyde et al. 2019).

Several Ganoderma species are prolific sources of highly active bioactive compounds such as polysaccharides, proteins, steroids and triterpenoids. These bioactive compounds show a huge structural and chemical diversity (Shim et al. 2004; Qiao et al. 2005; Wang and Liu 2008; Teng et al. 2011; De Silva et al. 2012a, b; 2013; Hapuarachchi et al. 2017; Li et al. 2018; Hyde et al. 2019). The bioactive constituents have anti-cancer, anti-inflammatory, anti-tumour, anti-oxidant, immunomodulatory, immunodeficiency, anti-diabetic, anti-viral, anti-bacterial, anti-fungal, anti-hypertensive, anti-atherosclerotic, anti-ageing, anti-androgenic, hepatoprotective and radical scavenging properties. They are also promising in neuroprotection, sleep promotion, cholesterol synthesis inhibition, preventing hypoglycemia, inhibition of lipid peroxidation/oxidative DNA damage, maintenance of gut health, prevention of obesity, and stimulation of probiotics (De Silva et al. 2012a; Hapuarachchi et al. 2016a, b, Hapuarachchi et al. 2017).

Current studies are identifying secondary metabolites, developing models for prediction or early detection of diseases, finding biological control methods as well as understanding genomes. Using artificial neural network spectral analyses and foliage of four disease levels, Ahmadi et al. (2017) provided an early detection method for Ganoderma basal stem rot of oil palm. Sitompul and Nasution (2020) suggested that to control Ganoderma diseases non or weakly pathogenic fungi can be considered as biological control agents. These agents could break down woody debris faster than the pathogen and occupy the same resource as the pathogen (compete for nutrients) as well as producing inhibitory secondary metabolites (Paterson 2007; Sitompul and Nasution 2020). Utomo et al. (2018) sequenced the nuclear genome of G. boninense, the main pathogen of basal stem rot, and the draft genome comprised of 79.24 megabases and 26,226 predicted coding sequences. Ramzi et al. (2019) conducted a study to understand the plant cell wall degradation and pathogenesis of G. boninense via comparative genome analysis. In their study, they found that similarly to G. lucidium, G. boninense was enriched with carbohydrate-active and cell wall degrading enzymes. Following plant-host interaction analysis, several candidate genes including polygalacturonase, endo β-1, 3-xylanase, β-glucanase and laccase were identified as potential cell wall degrading enzymes that contribute to the plant host interaction and pathogenesis. The study provided fundamental knowledge on the fungal genetic ability and capacity to secrete carbohydrate-active and cell wall degrading enzymes. Agudelo-Valencia et al. (2020) pointed out that information regarding the biotechnological importance of Ganoderma species (other than G. lucidium) is quite limited. Therefore, in their study they obtained and studied the genome of G. australe, resulting in gene prediction for the 84-megabase genome, prediction of 22,756 protein-coding genes, prediction of five putative genes and two enzyme complexes from a ganoderic acid pathway.

Most Ganoderma species are pathogenic or facultatively pathogenic, causing root and stem rot on a variety of monocotyledons, dicotyledons, and gymnosperms, including a wide range of economically important trees and perennial crops, which may result in death (Hapuarachchi et al. 2018a). Some species are saprobic and cause white-rot decay of wood (Muthelo 2009). Hence, they have ecological importance in the breakdown of woody plants for nutrient mobilization. They possess effective machinery of lignocellulose-decomposing enzymes useful for bioenergy production and bioremediation (Hepting 1971; Adaskaveg et al. 1991; Kües et al. 2015).

Classification—Basidiomycota, Agaricomycotina, Agaricomycetes, Incertae sedis, Polyporales, Ganodermataceae

Type species—Ganoderma lucidum (Curtis) P. Karst. 1881

Distribution—worldwide

Disease symptoms—basal stem, butt and root rot in economically important trees and perennial crops, especially in tropical regions. Ganoderma disease development is affected by environmental factors and tree death could be either slow or rapid depending on water availability and temperature (Coetzee et al. 2015).

Basal stem rot: Symptoms of basal stem rot disease can take several years to develop, and the presence of the pathogen is often only visible when the fungus is well-established and more than half of the tissue has been decayed. Soils with poor drainage and water stagnation during rainy seasons favour the disease (Kandan et al. 2010).

Butt rot and root rot: The primary symptoms include wilting, mild to severe, of either all leaves or just the lowest leaves in the canopy, premature death of the oldest leaves or a general decline of the tree. The advanced decay of the larger roots is evident after leaves are blown down. Decay may extend from several cms to over a metre into the lower (butt) portion of the tree, depending on the species of Ganoderma. It is quite common for basidiomes not to appear before the severe decline and death of a tree (Glen et al. 2009). Therefore, the only way to determine if Ganoderma butt rot is the cause is to cut cross-sections through the lower meter or so of the trunk after the tree is felled and examine the cross-sections for the typical pattern of rot: greatest near the soil line, decreasing in sections further from the soil line.

Ganoderma root rot may cause yellowing, wilting, or undersized leaves and dead branches. Tree vigour may decline as the decay of the sapwood advances. The first visible sign of infection is often the formation of basidiomes (solitary or in clusters) on the lower trunk and exposed root areas. There are two types: varnished and unvarnished. The upper surface of varnished fungus rot is typically red-brown with a white edge, shiny, and lacquered. Conks of the unvarnished fungus rot are brown with a white edge weathering to grey (Pilotti et al. 2004). When fresh, both have a white, porous surface on the underside. The rate of decay can lead to death in as little as 3 to 5 years from the time of infection and appears to be determined by tree vigour, which is often influenced by environmental stresses (Nirwan et al. 2016).

Hosts—Ganoderma has a wide host range, with more than 44 species from 34 potential host genera identified (Venkatarayan 1936). The root and stem rots caused by Ganoderma species, result in decreased forestry yields of e.g. Areca catechu (Palanna et al. 2020), Camellia sinensis, Cocos nucifera, (Kinge and Mih 2014), Elaeis guineensis, (Glen et al. 2009) and Hevea brasiliensis (Monkai et al. 2017) worldwide.

Pathogen biology, disease cycle and epidemiology

The fungus is spread by spores produced in the prominent basidiomes that form on the outside of the tree (conks). New spores released from the conks are dispersed throughout the summer during humid periods and infect open wounds on root flares and lower trunk areas of susceptible trees. The spores germinate, and the infection progresses to attack the sapwood of major roots and the lower tree trunk. Over the years, the number of decayed wood increases leading to dangerously soft, spongy wood in the part of the tree that functions as its anchor (Paterson 2007).

Morphology-based identification and diversity

Ganoderma species identification, limitations and their taxonomic segregation have been unclear and recently being debated (Moncalvo et al. 1995; Wang et al. 2009; Cao et al. 2012; Yao et al. 2013; Richter et al. 2015; Zhou et al. 2015a, b). Many Ganoderma collections and species have been misnamed because of the presence of heterogenic forms, taxonomic obstacles and inconsistencies in the way the genus has been subdivided (Mueller et al. 2007). Ganoderma species are genetically heterogeneous, hence a wide range of genetic variation has been reported and caused by outcrossing over generations and different geographical origins (Pilotti et al. 2004). This has led to variation in their listed morphological characteristics, even within the same species (Hong et al. 2001). Environmental factors, variability, inter hybridization and individual morphological bias, mean identification of Ganoderma species is difficult (Zheng et al. 2007a). Hence, naming a species is confused and traditional taxonomic methods based on morphology are inconclusive for establishing a stable classification system for Ganoderma species (Hseu et al. 1996; Hong et al. 2001) which in turn result in an uncertain nomenclature. This confusing situation is mainly the result of various criteria used in identification by different authors. Some authors strictly only focus on host-specificity, geographical distribution and macro morphology of basidiomes, while other authors only focus on spore characteristics as the primarily taxonomic characteristics (Sun et al. 2006; Ekandjo and Chimwamurombe 2012). Richter et al. (2015) suggested using a combination of morphological, chemotaxonomic and molecular methods to develop a more stable taxonomy for this genus.

Molecular identification and diversity

Isoenzyme analysis was the first molecular technique used to separate Ganoderma species (Park et al. 1994; Gottlieb et al. 1995, 1998; Gottlieb and Wright 1999a, b; Smith and Sivasithamparam 2000). Then, DNA sequences of the internal transcribed spacer (ITS), partial large subunit rDNA (Moncalvo et al. 1995, 2000; Cao et al. 2012; Yao et al. 2013; Richter et al. 2015) and nearly complete small subunit rDNA sequences (Hong and Jung 2004; Douanla-Meli and Langer 2009) were used. Later, multigene phylogenetic analyses with protein-coding genes such as β-tubulin (tub2), the largest subunit of RNA polymerase II gene (rpb1), the second-largest subunit of RNA polymerase II (rpb2), and translation elongation factor 1-α (tef1) were performed to resolve the taxonomic confusions within Ganoderma (Park et al. 2012; Zhou et al. 2015a, b; Hennicke et al. 2016; Jargalmaa et al. 2017). However, many problems remain in the resolution of phylogenetic relationships within the genus. As a result of the intricate taxonomy of Ganoderma, 65% of the Ganoderma sequences available in GenBank were reported to be wrongly identified or ambiguously labelled, (Jargalmaa et al. 2017). In this study, we reconstruct the phylogenetic tree based on ITS, tef1 and rpb2 sequence data (Table 10, Fig. 16).

Table 10 DNA barcodes available for Ganoderma

Fig. 16

Phylogram of 64 recognized Ganoderma species, obtained from ML of combined ITS, rpb2, and tef1 datasets. Bootstrap values from ML (left) and MP (middle) greater than 70% and BYPP, greater than 0.95, are indicated above the nodes. The tree is rooted with Coriolopsis trogii. Type specimens are indicated in bold

Recommended genetic marker (genus level)—ITS

Recommended genetic markers (species level)—rpb2, tef1

Accepted number of species—There are 456 species and infra-species epithets in Index Fungorum (2020), for 224 accepted species. However, only 64 species have DNA sequence data.

References—Coetzee et al. (2015); Xing et al. (2016, 2018); Tchoumi et al. (2019), Luangharn et al. (2019), Ye et al. (2019) (phylogeny, new species), Cabarroi-Hernández et al. (2019) (phylogeny).

85. Golovinomyces (U. Braun) V.P. Heluta, Biol. Zh. Armenii 41: 357 (1988)

Background

Braun (1978) introduced Golovinomyces as a section of Erysiphe sensu lato and Heluta (1988a) raised it to genus rank. Braun (1999) and Braun and Takamatsu (2000) accepted Golovinomyces as a distinct genus and established a new tribe, Golovinomyceteae. This is a strictly herb-parasitic genus in the Erysiphaceae. Host-parasite co-speciation was reported between Golovinomyces and Asteraceae hosts using molecular phylogenetic analyses (Matsuda and Takamatsu 2003). It was suggested that Golovinomyces first acquired parasitism on Asteraceae and then diverged to the host tribes Astereae, Cardueae, Heliantheae and Lactuceae. Bremer (1994) pointed out that Golovinomyces may have originated in South America and the geographic distribution expanded into the Northern Hemisphere. However, Takamatsu et al. (2006) suggest that Golovinomyces originated in the Northern Hemisphere, and not in South America. Fabro et al. (2008) profiled genome-wide expression on haustorium formation of G. cichoracearum in Arabidopsis. Research to understand pathogenesis towards plants has been undertaken. A draft whole genome of G. magnicellulatus, the causal agent of phlox powdery mildew was provided by Farinas et al. (2019). McKernan et al. (2020) identified 82 genes associated with resistance to G. chicoracearum, the causal agent of powdery mildew in cannabis.

Classification—Ascomycota, Pezizomycotina, Leotiomycetes, Leotiomycetidae, Erysiphales, Erysiphaceae

Type species—Golovinomyces cichoracearum (DC.) V.P. Heluta

Distribution—Worldwide (Mainly in northern hemisphere)

Disease symptoms—powdery mildew

Hosts—Has a wide range of hosts including Asteraceae, Boraginaceae, Cucurbitaceae, Malvaceae, Fabaceae, Lamiaceae, Polygonaceae, Scrophulariaceae, Solanaceae and Verbenaceae.

Pathogen biology, disease cycle and epidemiology

Discussed under Erysiphaceae.

Morphological based identification and diversity

Golovinomyces is characterized by chasmothecia with mycelioid appendages, several, mostly 2-spored asci, an asexual morph with catenescent conidia that lack fibrosin bodies, and mostly nipple-shaped appressoria (Braun 1978; Qiu et al. 2020a). Heluta (1988a) reallocated Erysiphae cichoracearum to Golovinomyces and now nearly all species of E. cichoracearum are assigned to Golovinomyces. Braun (1987) confined E. cichoracearum to powdery mildews on hosts of Asteraceae and assigned specimens on hosts belonging to other plant families to Erysiphe orontii. Braun and Cook (2012) split G. cichoracearum into several species based on molecular analyses of this complex which suggested a co-evolutionary relationship between Golovinomyces species and tribes of Asteraceae (Matsuda and Takamatsu 2003). Golovinomyces cynoglossi sensu lato, a complex of morphologically similar powdery mildews on the plant family Boraginaceae, was reassessed by Braun et al. (2018) and split into G. asperifoliorum, G. asperifolii and G. cynoglossi based on sequence analyses, biological aspects and morphological differences. Braun et al. (2019) revisited G. orontii and Qiu et al. (2020b) epitypfied and confirmed Erysiphe cucurbitacearum was a synonym of G. tabaci.

Molecular based identification and diversity

A comprehensive phylogenetic analysis by Takamatsu et al. (2013) resulted in a polyphyletic complex that split into three genetically distinct clades. Golovinomyces ambrosiae and G. spadiceus were considered as separate species by Braun and Cook (2012). However, phylogenetic analyses of ITS and 28S rDNA sequences by Takamatsu et al. (2013), including Golovinomyces species on Asteraceae, found that these two species that occur on Asian species of Eupatorium and a multitude of other hosts, including those on other plant families, formed a single large, unresolved clade (lineage III in Takamatsu et al. (2013)). The taxonomic interpretation posed a serious problem as G. ambrosiae and G. spadiceus were treated as two morphologically differentiated species. Hence, the resolution based only on ITS sequence data was considered insufficient to distinguish closely allied species. Most subsequent authors followed the taxonomic treatments in Braun and Cook (2012) and recognized G. ambrosiae and G. spadiceus as separate species, within lineage III, based on morphological differences (Qiu et al. 2020a). However, there is minimal multi loci data for the powdery mildews currently available. Most of the research involves the intra-specific genetic diversity in species such as Blumeria graminis (Walker et al. 2011), Erysiphe necator (de Oliveira et al. 2015), Golovinomyces orontii (Pirondi et al. 2015a) and Podosphaera xanthii (Pirondi et al. 2015b). Based on ITS and D1/D2 domain of 28S sequence data, Braun et al. (2019) introduced G. bolayi and G. vincae. Nayak and Bandamaravuri (2019) developed species-specific PCR primers CgF2 and CgR2 for G. orontii (the causal agent of powdery mildew in cucurbits), based on partial ITS and 5.8S rDNA, which resulted in a 233bp fragment of G. orontii.

Recommended genetic markers (genus level)—ITS, LSU

Recommended genetic markers (species level)—Comprehensive applications of multi loci approaches to solve complex taxonomic-phylogenetic problems connected with the species level classification of the powdery mildews are lacking. The phylogenetic analyses of multi loci sequence data, including ITS and LSU, IGS, tub2, chs, and consideration of morphological characters resolve species delimitation in a heterogeneous complex within Golovinomyces.

Accepted number of species—There are 81 epithets listed in Index Fungorum (2020), however, only 41 have molecular data (Table 11, Fig. 17).

Table 11 DNA barcodes for accepted species of Golovinomyces

Fig. 17

Phylogram generated from MP analysis based on combined sequences of ITS and LSU sequences of all species of Golovinomyces with molecular data. Related sequences were obtained from GenBank. Fourty-two taxa are included in the analyses, which comprise 1401 characters including gaps, of which 848 characters are constant, 392 characters are parsimony-uninformative and 161 characters parsimony-informative. The parsimony analysis of the data matrix resulted in the maximum of ten equally most parsimonious trees with a length of 927 steps (CI = 0.740, RI=0.699, RC = 0.517, HI = 0.260) in the second tree. The tree was rooted with Neoerysiphe galeopsidis (MUMH 4680). MP bootstrap support value ≥ 50% and BYPP ≥ 0.9 are shown respectively near the nodes. Ex-type strains are in bold

References—Braun (1978, 1987), Heluta (1988a, b) (morphology); Braun and Cook (2012), Takamatsu et al. (2013), Braun et al. (2019), Qiu et al. (2020a, b) (morphology and phylogeny).

86. Heterobasidion Bref., Unters. Gesammtgeb.Mykol. (Liepzig) 8: 154 (1888)

Background

Heterobasidion was introduced by Brefeld (1888) and is typified by H. annosum (≡ Polyporus annosus). Certain Heterobasidion species are important forest pathogens of the Northern Hemisphere, causing root and butt rot, mainly in conifers (Woodward et al. 1998). In coniferous plantations, Heterobasidion is one of the most widespread of wood decay agents, especially when the host is under intensive management. Heterobasidion greatly reduces site productivity and the amount of harvestable timber; estimated financial losses caused by Heterobasidion species in Europe were around 800 million euro per year (Korhonen et al. 1998; Garbelotto 2004; Asiegbu et al. 2005). On the other hand, these taxa have a relatively moderate pathogenic role in natural forest ecosystems. They affect stand species composition, density and structure, and they contribute to forest succession, nutrient recycling and even regeneration (Goheen and Otrosina 1998; Garbelotto 2004; Dai et al. 2006).

Classification—Basidiomycota, Agaricomycotina, Agaricomycetes, Incertaesedis, Russulales, Bondarzewiaceae

Type species—Heterobasidion annosum (Fr.) Bref., Unters. Gesammtgeb. Mykol. (Liepzig) 8: 154 (1888)

Distribution—North America, Europe, Asia, Australia and Oceania

Disease symptoms—There are two Heterobasidion species complexes –H. insulare sensu lato and H. annosum sensu lato—they cause the same symptoms. The H. annosum species complex is one of the major root-rot pathogenic genera of the northern temperate hemisphere (Garbelotto and Gonthier 2013; Kärhä et al. 2018). After the primary infection through stump tops, or stem and root wounds, the taxa can vegetatively infect uninjured trees (secondary infection) by the growth of the mycelium through root contacts (Rishbeth 1950, 1951a, b; Asiegbu et al. 2005; Garbelotto and Gonthier 2013). Heterobasidion could be considered both necrotrophs and saprotrophs; though some species in the H. insulare species complex (e.g. H. austral, H. araucariae) are mainly saprotrophs (Niemelä and Korhonen 1998; Dai and Korhonen 2009; Chen et al. 2014). In contrast to Europe, the pathogenicity of H. annosum sensu lato in China and Japan is uncertain; the complex seems to occur mostly on dead trees, and no symptoms of tree decline are usually visible near infected trees. These observations could be due to different, less intensive forest management strategies in the East-Asian regions, or lack of data on the butt rot symptoms (Dai et al. 2006; Tokuda et al. 2007).

The infection causes white pocket rot and heart rot in the roots and the butt of living trees (Korhonen and Stenlid 1998; Asiegbu et al. 2005). Resin, containing mycelium, may also exude from the infected roots, or the bark-scales (Rishbeth 1950). In invaded roots and the basal portions of the trunk, H. annosum sensu lato taxa colonize different plant tissues depending on the host. Heart rot is mainly caused in trees more susceptible to the colonization of the heartwood, e.g. Picea abies. In the case of Pinus, cambium and sapwood are the most severely colonized, while the sapwood of Calocedrus or Sequoiadendron trees is the most colonized (Garbelotto 2004; Asiegbu et al. 2005; Garbelotto and Gonthier 2013).

After establishment, the basidiomata of H. annosum sensu lato appear. The localization of the sporocarps is governed by the species, environmental conditions and infection strategy. Some species prefer the root collar for fruiting (H. annosum, H. irregular). Some also produce sporocarps in decay pockets in stumps and fallen trees (H. parviporum, H. abietinum and H. occidentale), or under the intact surface of stumps (H. irregulare, H. occidentale). The sporocarps are sometimes located on the higher parts of the trunk. When moisture is limited, the fungi fruit inside stumps; if the climate is moist and humid, the basidiomata can be found near the ground in the duff at the base of diseased trees. If during primary infection the stump surface is infected, the basidiomata form under an intact top layer. During active pathogenesis, if the standing trees are infected the sporocarps could be found within decay columns in the sapwood (Rishbeth 1950; Otrosina and Garbelotto 2010).

The infection kills the functioning sapwood, cambium and heart wood in the roots and at the basal portions of the trunk, resulting in white rot, reduced growth rate, crown dieback (Omdal et al. 2004), and eventually mortality and windthrow of infected trees (Rishbeth 1950; Oliva et al. 2008; Garbelotto and Gonthier 2013).

Hosts—The host range of Heterobasidion is extremely wide. The genus has been reported from approximately 200 host species (Korhonen and Stenlid 1998). Taxa mostly occur on gymnosperms, such as Abies, Agathis, Araucaria, Calocedrus, Juniperus, Keteleeria, Larix, Picea, Pinus, Podocarpus, Pseudolarix, Pseudotsuga, Sequoia, Sequoiadendron, Thuja and Tsuga (Buchanan 1988; Corner 1989; Dai and Korhonen 2009; Otrosina and Garbelotto 2010; Garbelotto and Gonthier 2013; Garbelotto et al. 2017). Occasionally, certain Heterobasidion species grow on broad-leaved trees of various angiosperm genera (Garbelotto and Gonthier 2013; Ryvarden and Melo 2014).

Morphological based identification and diversity

There are 33 Heterobasidion epithets listed in Index Fungorum (2020). Of these, eight are related to other polypore genera, based on type studies and morphological observations (Ryvarden 1972, 1985; Buchanan and Ryvarden 1988; Dai and Niemelä1995; Hattori 2003). Besides, the taxonomic status of three further species described from Asia is unclear: viz. H. arbitrarium, H. perplexum and H. insulare (Corner 1989; Ryvarden 1989; Stalpers 1996; Hattori 2001; Dai et al. 2002; Tokuda et al. 2009). Given that no sequence data (H. arbitrarium, H. perplexum) or authentic sequences (H. insulare sensu stricto) are available for the molecular resolution, further studies are needed to clarify their status.

Formerly, Heterobasidion was considered as a group consisting of only the generic type, H. annosum and H. araucariae and H. insulare (Buchanan 1988; Chase 1989). However, mating studies on Eurasian and North American Heterobasidion collections revealed several intersterile groups, which later became the basis for designating separate taxonomic species within the H. annosum and H. insulare species complexes. Mating experiments revealed three intersterile groups of H. annosum sensu lato in Europe (Korhonen 1978b, Capretti et al. 1990) and two in North America (Otrosina et al. 1993). All intersterile groups have been recognised in the H. annosum species complex are now formally described as separate taxonomic species. European groups were described as H. abietinum, H. parviporum and H. annosum sensu stricto (Niemelä and Korhonen 1998), whereas North American groups were named H. irregulare and H. occidentale (Otrosina and Garbelotto 2010).

The mating study by Dai et al. (2002) on Asian “H. insulare” collections revealed three intersterile groups in China, which were subsequently described as Heterobasidion linzhiense (Dai et al. 2007), H. orientale and H. ecrustosum (Tokuda et al. 2009). H. australe related to the H. insulare species complex was also described from China by Dai and Korhonen (2009). Chen et al. (2014) described two further Heterobasidion species (H. amyloideum and H. tibeticum) from the eastern Himalayas based on phylogenetic evidence. These species are morphologically closely related to the members of the H. insulare species complex, but differ in presence of cystidia and amyloid skeletal hyphae in the context. The recently described H. amyloideopsis was collected in the western Himalayas (Pakistan) and formed a monophyletic group with the H. insulare species complex, sister to H. amyloideum (Zhao et al. 2017).

The main morphological characters which are used for the identification are the resupinate to pileate basidiocarps, the dimitic hyphal system with mostly simple septate generative hyphae, and the asperulate basidiospores showing no reaction in Melzer’s reagent. Besides morphology, host preference, geographical distribution, and DNA sequence data have also been used for species identification (Otrosina and Garbelotto 2010; Chen et al. 2015a).

Molecular based identification and diversity

Heterobasidion has been intensely studied by molecular methods. Sequence data are available for the majority of taxa, and molecular studies were conducted to understand the evolution (Dalman et al. 2010), mating behaviour (Gonthier and Garbelotto 2011), and pathogenicity (Liu et al. 2018a) of Heterobasidion species.

Various marker types were used to resolve the phylogeny of the H. annosum species complex, such as isoenzyme (Karlsson and Stenlid 1991a, b), AFLP (Gonthier and Garbelotto 2011) and SSR (Garbelotto et al. 2013) markers. Sequence analyses were carried out initially on nrITS and intergenic spacer regions (Kasuga and Mitchelson 1993a, b; DeScenzo and Harrington 1994), housekeeping genes (Johanesson and Stenlid 2003), peroxidase (Maijala et al. 2003) and laccase genes (Asiegbu et al. 2004), with which it was possible to distinguish four lineages (three European and one North American) within the complex (Asiegbu et al. 2005). Later, allowing the differentiation of a larger number of taxa, further nuclear genes were applied, such as the calmodulin (cam), translation elongation factor 1-α (tef1), glyceraldehydes3-phosphate dehydrogenase (gapdh), heat shock protein (hsp), glutathione-S-transferase (gst1) and transcription factor (tf) genes (Johanesson and Stenlid 2003; Ota et al. 2006; Dalman et al. 2010), as well as two mitochondrial genes, the mitochondrial ATP synthase subunit 6 (ATP6) and mitochondrial rDNA region (Linzer et al. 2008). Dalman et al. (2010) came to the conclusion, that there are two monophyletic sister clades within the H. annosum species complex, representing the Eurasian and North American species.

The protein coding largest subunit of RNA polymerase II (rpb1) and the second subunit of RNA polymerase II (rpb2) genes were used by Chen et al. (2014) and were suitable to differentiate Heterobasidion species in the H. insulare species complex. The variability of these markers was confirmed by Chen et al. (2015a) and Zhao et al. (2017) who, among other previously mentioned markers, both used the nuclear large ribosomal subunit (nrLSU) and the mitochondrial small subunit (mtSSU) sequences to their studies (Fig. 18).

Fig. 18

Members of Heterobasidion annosum species complex. a basidiome on Scots pine, b basidiome on European silver fir, c–e basidiomes on European spruce, f hyphal structure in the trama, g hyphal structure in the context, h–j basidiospores. Scale bars: f, g = 10 µm, h, j = 5 µm

In this study, we provide a phylogenetic tree (Fig. 19) based on multi-locus phylogenetic analysis of ITS–gapdh–rpb1–rpb2–tef1 sequence data. Sequences of H. arbitrarium and H. perplexum could not be analysed as they are unavailable in GenBank. Furthermore, no sequences are available for the type of H. insulare hence this species was not included in the analysis. The results provide a similar topology to those obtained by Chen et al. (2015a, b) and Zhao et al. (2017).

Fig. 19

Phylogram generated from ML analysis based on combined ITS, rpb1, rpb2, gapdh and tef1 sequence data of Heterobasidion species. Related sequences were obtained from GenBank. Fourty-four strains are included in the analyses, which comprised 4314 characters including gaps. The tree was rooted with Bondarzewia occidentalis (HHB 14803) and B. tibetica (Cui 12078). Tree topology of the ML analysis was similar to the Bayesian analysis. ML bootstrap values ˃ 50% and BYPP ˃ 0.80 are shown respectively near the nodes

Recommended genetic marker (genus level)—nLSU

Recommended genetic markers (species level) —rpb1, rpb2

Accepted number of species –There are 33 epithets in Index Fungorum (2020), however only 15 species are accepted (Table 12). Amongst these, no sequences are available for H. arbitrarum and H. insulare. Heterobasidion perplexum is not accepted in the genus, pending further studies.

Table 12 DNA barcodes for accepted species of Heterobasidion

References—Dai and Korhonen (2009) (new sp., China, morphology); Tokuda et al. (2009) (new species, East Asia); Dalman et al. (2010) (Evolution, H. annosum species complex, haplotype network); Otrosina and Garbelotto (2010) (new species, North America, biology); Garbelotto and Gonthier (2013) (biology, epidemiology, control); Chen et al. (2014) (new species, China, phylogeny); Chen et al. (2015a) (biogeography, divergence time estimation, phylogeny); Zhao et al. (2017) (new sp., Pakistan, phylogeny).

87. Meliola Fr., Syst. orb. veg. (Lundae) 1: 111 (1825)

Background

Meliola commonly known as “black mildews” or “dark mildews” is the largest genus of Meliolaceae (Hongsanan et al. 2015; Zeng et al. 2017). Fries (1825) established this genus, with the type species M. nidulans. Species in Meliola are mostly biotrophs or pathogens of living leaves and occasionally petioles, twigs, and branches (Hansford 1961; Hosagoudar 1994, 1996, 2008; Mibey and Hawksworth 1997; Old et al. 2003; Hosagoudar and Riju 2013). The phylogenetic placement of Meliola was established by using sequence data from fruiting bodies and placed in Sordariomycetes (Gregory and John 1999; Pinho et al. 2012, 2014, Hongsanan et al. 2015; Justavino et al. 2015). Meliola has been shown to be polyphyletic (Hyde et al. 2020b; Marasinghe et al. 2020; Zeng et al. 2020). There is little sequence data available in GenBank for clarifying relationships between species and establishiing host-specificity (Hongsanan et al. 2015; Zeng et al. 2017).

Classification—Ascomycota, Pezizomycotina, Sordariomycetes, Sordariomycetidae, Meliolales, Meliolaceae

Type species—Meliola nidulans (Schwein.) Cooke

Distribution—commonly found in tropical and subtropical regions (see Zeng et al. 2017)

Disease symptoms—Black mildews, forming black, radiate velvety colonies on the surface of plants.

Hosts—has a wide range of hosts (see Zeng et al. 2017)

Pathogen biology, disease cycle and epidemiology

For pathogen biology, disease cycle and epidemiology see Hongsanan et al. (2015).

Morphological based identification and diversity

Species in Meliola are characterized by forming web-like colonies on the host surface, hyphal setae developed from superficial hyphae, with hyphopodia, 2–4-spored, unitunicate asci, and 3–4-septate pigmented ascospores (Pinho et al. 2012, 2014; Hongsanan et al. 2015, 2020; Justavino et al. 2015; Hyde et al. 2020a, b; Fig. 20). Cannon and Kirk (2007) reported that the asexual morph of the genus develops from the hypha, form ampuliform hyphopodia or flask-shaped which are called “phialides” (Hongsanan et al. 2015). Conidiogenous cells formed from vegetative hyphae and small, hyaline, unicellular conidia (Cannon and Kirk 2007; Hongsanan et al. 2015). Currently, Meliola comprises over 1700 species (Zeng et al. 2017), which have mostly been introduced by host association, followed by morphology, and disease distribution (Mibey and Hawksworth 1997). Thus, species identification is almost impossible without a host name. However, the same species can be found in different hosts, but it is not clear if this is widespread (Hongsanan et al. 2015). Therefore, testing of host-specificity in Meliola is needed to establish accurate species determination.

Fig. 20

Morphology of Meliola species a Meliola thailandicum on Dimocarpus longan. b Meliola sp. on Citrus reticulata. c Meliola sp. on Citrus maxima. d Colony on the host surface. e Hyphopodia on mycelium. f Section through ascoma. g Peridium. h Setae. i Young ascus. j Mature ascus. k, l Ascospores. Scale bars: f=50 μm, g, i–l=30 μm, h=10 μm and e=5 μm

Molecular based identification and diversity

Sequence data of species in Meliola are from direct sequencing of fruiting bodies and fresh mycelium (Pinho et al. 2012, 2014; Hongsanan et al. 2015; Justavino et al. 2015; Hyde et al. 2016, 2020b). LSU and ITS sequence data placed Meliola in Sordariomycetes (Hongsanan et al. 2015, 2020; Maharachchikumbura et al. 2015, 2016; Hyde et al. 2016, 2020a, b). By adding more sequence data, Meliola was shown to be polyphyletic (Marasinghe et al. 2020; Zeng et al. 2020). A phylogenetic tree for Meliola species is presented in Fig. 21.

Fig. 21

Phylogram generated from RAxML analysis based on combined ITS and LSU sequence data of Meliola species. Related sequences were obtained from GenBank. Thirty-five strains are included in the analyses, which comprised 1655 characters including gaps. The tree was rooted with Chaetosphaeria innumera (SMH 2748). Tree topology of the ML analysis was similar to the Bayesian analysis. ML bootstrap values ≥ 50% and BYPP ≥ 0.90 are shown respectively near the nodes

Recommended genetic markers (genus level)—LSU, SSU of nrDNA

Recommended genetic marker (species level)—ITS

Accepted number of species—There are 3064 epithets listed in Index Fungorum (2020), however only 25 species have DNA molecular data (Zeng et al. 2017, Table 13).

Table 13 DNA barcodes available for Meliola

References—Cannon and Kirk (2007) (morphology); Pinho et al. (2012, 2014), Hongsanan et al. (2015, 2020), Justavino et al. (2015), Zeng et al. (2020) (morphology and phylogeny)

88. Neoerysiphe U. Braun, Schlechtendalia 3: 50 (1999)

Background

Neoerysiphe was classified in section Galeopsidis within Erysiphe. Phylogenetic analysis, however, showed Erysiphe to be polyphyletic, and Galeopsidis was raised to generic rank (Takamatsu et al. 1998; Braun 1999; Saenz and Taylor 1999). Therefore, in the current classification Neoerysiphe belongs to the tribe Golovinomyceteae.

Classification—Erysiphaceae, Erysiphales, Leotiomycetidae, Leotiomycetes, Pezizomycotina

Type species—Neoerysiphe galeopsidis (DC.) U. Braun

Distribution—Argentina, Australia, Belarus, Brazil, Bulgaria, Canada, China, Denmark, Finland, France, Germany, Hungary, India, Israel, Italy, Japan, Korea, Netherlands, Norway, Poland, Romania, Russia, Slovakia, Sweden, Switzerland, Turkey, UK, Ukraine and USA (Farr and Rossman 2020).

Disease symptoms-powdery mildew

Hosts—Neoerysiphe species have a wide host distribution infecting more than 300 species from families including Asteraceae, Acanthaceae, Bignoniaceae, Elaeocarpaceae, Lamiaceae, Rubiaceae and Verbenaceae (Amano 1986; Braun 1999; Bahcecioglu et al. 2006). In general, all species have a specific host range confined to one plant family, except N. galeopsidis which affects several species in four families (Takamatsu et al. 2008).

Pathogen biology, disease cycle and epidemiology

Discussed under Erysiphaceae.

Morphological based identification and diversity

Neoerysiphe is in the tribe Golovinomyceteae with Arthrocladiella and Golovinomyces. These genera share a common asexual morph characterized by catenate conidia without distinct fibrosin bodies (Braun 1999). Neoerysiphe is characterized by lobed appressoria and the striate surface of the conidia (Braun 1981; Cook et al. 1997; Braun and Cook 2012). Braun and Cook (2012) mentioned that 15 species of Neoerysiphe are described on different hosts belonging to 11 plant families. Of these 15 species, 11 sexual morphs and 14 asexual morphs have been identified (except N. joerstadii) (Heluta et al. 2010; Braun and Cook 2012). Striatodium is now considered as a synonym of Neoerysiphe and three species viz. N. aloysiae, N. baccharidis and N. maquii were transferred to Neoerysiphe, while Striatodium jaborosae was not transferred as its phylogenetic position are unclear (Wijayawardene et al. 2017a).

Molecular based identification and diversity

The phylogenetic placement of Neoerysiphe within Erysiphaceae has been reported in a few papers (Saenz and Taylor 1999; Mori et al. 2000; Cook et al. 2006). However, these treatments used only limited sequence data for the genus. Takamatsu et al. (2008) conducted the first comprehensive study on this genus using ITS sequence data and the divergent domains D1 and D2 of the 28S rDNA for 30 strains. In their study, the 30 taxa, clustered into three monophyletic groups that were represented by N. galeopsidis on Lamiaceae, N. galii on Rubiaceae and N. cumminsiana from Asteraceae. Takamatsu et al. (2008) used an LSU dataset to estimate the timing of divergence of Neoerysiphe. Neoerysiphe split from other genera ca 35–45 Mya and the three groups of Neoerysiphe diverged between 10 and 15 Mya in the Miocene. Heluta et al. (2010) used 65 ITS sequences in their analyses for identifying Neoerysiphe species infecting Asteraceae and Geranium in Eurasia and introduced three new species, viz. N. hiratae, N. joerstadii and N. nevoi. Gregorio-Cipriano et al. (2020) introduced a new species N. sechii causing powdery mildew on Sechium edule and S. mexicanum in Mexico. The authors mentioned that they were unable to recover DNA in pure form from some samples, as fragments of infected leaves were used during the extraction. Therefore, a specific oligonucleotide for Erysiphales at the 5= region of ITS was designed: ErysiF (5=-AGGATCATTACWGAGYGYGAG-3=) was used along with NLP1 (Limkaisang et al. 2006) to amplify a fragment of approximately 1200 bp (that included the ITS1-5.8S-ITS2 region and a section of approximately 680 nucleotides from 28S). Species used in the phylogenetic analyses done in this study are listed in Table 14 and given in Fig. 22.

Table 14 DNA barcodes available for Neoerysiphe

Fig. 22

Phylogram generated from MP analysis based on combined sequences of ITS and LSU sequences of all species of Neoerysiphe with molecular data. Related sequences were obtained from GenBank. 12 taxa are included in the analyses, which comprise 2023 characters including gaps, of which 1790 characters are constant, 167 characters are parsimony-uninformative and 66 characters parsimony-informative. The parsimony analysis of the data matrix resulted in the maximum of four equally most parsimonious trees with a length of 316 steps (CI = 0.848, RI=0.678, RC = 0.575, HI = 0.152) in the first tree. Single gene analyses were carried out and compared with each species, to compare the topology of the tree and clade stability. The tree was rooted with Golovinomyces adenophorae (MUMH144). MP bootstrap support value ≥ 50% and BYPP ≥ 0.9 are shown respectively near the nodes. Ex-type strains are in bold

Recommended genetic marker (genus level)—ITS and LSU

Recommended genetic markers (species level)—ITS

Accepted number of species—There are 16 species epithets in Index Fungorum (2020), for 15 accepted species. However, only 12 species have DNA sequence data (N. chelones, N. gnaphalii and N. rubiae do not have molecular data) (Table 14).

References—Takamatsu et al. (1998), Braun (1999), Saenz and Taylor (1999) (morphology); Heluta et al. (2010), Braun and Cook (2012), Gregorio-Cipriano et al. (2020) (morphology and phylogeny).

89. Nothophoma Qian Chen & L. Cai, Stud. Mycol. 82: 212 (2015)

Background

Nothophoma was introduced by Chen et al. (2015b) by transferring five Phoma species. Species are saprobes and pathogens. In addition, to the phytopathogens, N. gossypiicola has been isolated from clinical samples of humans in the respiratory secretion of a patient with pneumonia and a human bronchial wash sample (Valenzuela-Lopez et al. 2018). Chethana et al. (2019) showed that the comparative pathogenicity of Nothophoma species is low when compared to other opportunistic pathogens. Some species grow on other fungi or occur in soil (Boerema et al. 2004; Aveskamp et al. 2009; 2010; Chen et al. 2015b). Some Nothophoma species might be host-specific to a single plant genus or family (Aveskamp et al. 2010; Chen et al. 2015b). However, there is no study of host-specificity in Didymellaceae. Abdel-Wahab et al. (2017) identified 55 bioactive compounds from an endophyte, N. multilocularis. Of these, ten compounds showed strong antimicrobial activity in combination.

Classification—Ascomycota, Pezizomycotina, Dothideomycetes, Pleosporomycetidae, Pleosporales, Didymellaceae

Type species—Nothophoma infossa (Ellis & Everh.) Qian Chen & L. Cai

Distribution—Argentina, China, Italy, India, Korea, Netherlands, Spain, Tunisia, Ukraine, United States

Disease symptoms—brown spot of fruits, leaf spots, shoot canker, stem cankers

Leaf spot produced by Nothophoma anigozanthi is elliptical to circular and black. Nothophoma pruni and N. quercina develop small, dark red or purple pinpoint lesions (Chethana et al. 2019). Liu et al. (2018b) identified N. quercina infection on ornamental crab-apple. Symptoms on the trunk appear as warts, the periderm around warts can become cracked, and the bark is roughened with a scaly periderm. During dry weather, these cankers expand and coalesce (Liu et al. 2018b; Fig. 23). Nothophoma quercina develops shoot necrosis, stem browning, and wilted leaves on Chaenomeles sinensis (Yun and Oh 2016). In Tunisia, shoot blights caused by N. quercina were observed with diffuse cankers with gummosis on buds (Triki et al. 2019).

Fig. 23

Nothophoma quercina on Malus micromalus a Malus micromalus (Crab-Apple tree). b Canker on the trunk. c, d appearance of conidiomata on trunk. e longitudinal section through conidiomata. f cross-section of conidiomata g, h conidiogenous cells. i, j conidia. k upper view on PDA. l reverse view on PDA. Scale bars: d=1000 μm e, f = 50μm g–j 10 μm

Hosts— Has a wide range of hosts including Anigozanthos flavidus, Anigozanthus maugleisii, Arachis hypogaea, Chaenomeles sinensis, Gossypium sp., Fraxinu spennsylvanica, Malus micromalus, Microsphaera alphitoides, Olea europaea, Phellodendrona murense, Pistacia vera, Prunu savium, Prunus dulcis, Spiraea salicifolia, Quercus sp. and Ziziphus jujube (Babaahmadi et al. 2018; Chen et al 2015b, 2017; Chethana et al. 2019; Jianyu et al. 2016; Liu et al. 2018b; Moral et al. 2017, 2018; Soleimani et al. 2018; Triki et al. 2019; Valenzuela-Lopez et al. 2018; Yun and Oh 2016; Zhang et al. 2020).

Morphological based identification and diversity

This genus was introduced by Chen et al. (2015b) based on molecular data to delineate a more natural classification for the Ascochyta-Didymella-Phoma species complex (Chen et al. 2015b; Fig. 23). Species produce elongate, barrel-shaped, olivaceous brown chlamydospores in chains (Chen et al. 2015b). However, there is little morphological variation among species (Valenzuela-Lopez et al. 2018).

Molecular based identification and diversity

Species identification is based on multi-locus sequence phylogeny. Phylogenetic analyses of combined LSU, ITS, tub2 and rpb2 sequence data resulted in several new species being added to this genus by Chen et al. (2015b), Abdel-Wahab et al. (2017), Valenzuela-Lopez et al. (2018), Chethana et al. (2019), Marin-Felix et al. (2019) and Zhang et al. (2020). Here we provide an updated phylogenetic tree for this genus (Fig. 24).

Fig. 24

Phylogram generated from maximum likelihood analysis based on combined LSU, ITS, tub2 and rpb2 sequence data of Nothophoma species. Related sequences were obtained from GenBank. Seventeen strains are included in the combined sequence analyses. Phoma herbarum (CBS 615.75) and Vacuiphoma bulgarica (CBS 357.84) was used as the outgroup taxa. The best scoring RAxML tree with a final likelihood value of − 5537.646741 is presented. The matrix had 284 distinct alignment patterns, with 12.23% of undetermined characters or gaps. Estimated base frequencies were as follows: A = 0.238395, C = 0.241637, G = 0.276596, T = 0.243371; substitution rates AC = 0.975188, AG = 4.004775, AT = 1.500008, CG = 0.519461, CT = 10.843965, GT = 1.000000; gamma distribution shape parameter a = 1.764918. ML bootstrap support value ≥ 50% and BYPP ≥ 0.95 are shown respectively near the nodes. Ex-type strains are in bold

Recommended genetic markers (genus level)—LSU, ITS

Recommended genetic markers (species level)—tub2, rpb2

Since the colony morphology and other morphological features in Didymellaceae often overlap, initial species identification is recommended with LSU and ITS sequence data using all type species in Didymellaceae. Once the genus is identified as Nothophoma, the phylogenetic analysis could be done with LSU, ITS, tub2, and rpb2 sequence data.

Accepted number of species—There are 12 species in Index Fungorum (2020) with DNA sequence data (Table 15).

Table 15 DNA barcodes available for Nothophoma

References—Chen et al. (2015b), Abdel-Wahab et al. (2017), Valenzuela-Lopez et al. (2018), Chethana et al. (2019), Marin-Felix et al. (2019), Zhang et al. (2020) (morphology and phylogeny)

90. Phellinus Quél., Enchir. fung. (Paris): 172 (1886)

Background

Phellinus was introduced by Quélet (1886) with P. igniarius (≡ Boletus igniarius) as its type species (Murrill 1903) and is placed in Hymenochaetaceae (He et al. 2019). Traditionally, most poroid Hymenochaetaceae were placed in Phellinus, which has been characterized by a dimitic hyphal system and perennial habit of the basidiomata (Gilbertson 1979; Larsen and Cobb-Poulle 1990; Ryvarden and Gilbertson 1994; Núñez and Ryvarden 2000). However, phylogenetic studies revealed that the morphologically defined Phellinus sensu lato had polyphyletic origins within the Hymenochaetoid clade, and most species previously classified as Phellinus are now members of various segregate genera (e.g. Wagner and Fischer 2001, 2002; Jeong et al. 2005; Dai 2010; Rajchenberg et al. 2015; Drechsler-Santos et al. 2016). According to the most narrowly defined generic concept, Phellinus sensu stricto is limited to the P. igniarius species complex (Fischer and Binder 2004), which includes species causing a delignifying trunk rot mostly on various deciduous trees in temperate areas (Brazee 2015; Zhou et al. 2016). Based on a wider generic concept, several morphologically similar species described from East Asia, Africa or America are considered as part of Phellinus sensu stricto (Decock et al. 2006; Yombiyeni et al. 2011; Cui and Decock 2013; Bian et al. 2016; Campos-Santana et al. 2016; Vlasák and Vlasák 2017; Salvador-Montoya et al. 2018; Zhu et al. 2018). In this study, we follow the broader concept of classification of Phellinus sensu stricto, pending further studies.

Classification—Agaricomycotina, Basidiomycota, Agaricomycetes, Incertae sedis, Hymenochaetales, Hymenochaetaceae

Type species—Phellinus igniarius (L.) Quél., Enchir. fung. (Paris): 177 (1886)

Distribution—If the wider generic concept of Phellinus were accepted it would be a globally distributed genus, with certain species found in East Asia, Europe, North America (Dai 2010; Cui and Decock 2013; Brazee 2015; Zhou et al. 2016; Vlasák and Vlasák 2017; Zhu et al. 2018), Central- and South America (Decock et al. 2006; Campos-Santana et al. 2016; Salvador-Montoya et al. 2018) and Africa (Yombiyeni et al. 2011; Cloete et al. 2016). However, the members of the P. igniarius species complex are known only from the Northern Hemisphere (Brazee 2015; Zhou et al. 2016).

Disease symptoms—Members of Phellinus produce white rot, decaying polysaccharides and delignifying the substrata (Niemelä 1974, 1977; Ryvarden and Gilbertson 1994; Wagner and Fischer 2002; Decock et al. 2006; Cui and Decock 2013; Brazee 2015; Cloete et al. 2016, de Campos-Santana et al. 2016). The rot could be localized in the trunk as a column of decay (Brazee 2015), in both fallen and in standing dead trunks (Niemelä 1977; Campos-Santana et al. 2016). Branches of living trees (Niemelä 1974; Decock et al. 2006), dead, fallen, corticated branches and logs (Niemelä 1972) and dead stumps (Niemelä 1972; Decock et al. 2006; Campos-Santana et al. 2016) are colonized and decayed. The fungus penetrates the heartwood, causing heartrot (Niemelä 1974; Larsson et al. 2006), sometimes extending into the sapwood (Niemelä 1977; Larsson et al. 2006). Decay characteristics (i.e. colour, fragility and fragmentation) vary between species (Niemelä 1972, 1974, 1977; Yombiyeni et al. 2011; Luna et al. 2012; Campos-Santana et al. 2016). Pathogenic species, such as P. tremulae or P. resupinatus are usually associated with other basidiomycete species, pathogenic bacteria and basal fungi (Kallio 1972; Cloete et al. 2016). Phellinus tremulae is a common and harmful pathogen of aspen (Populus species), penetrating the heartwood along dead branches (Niemelä 1974), but is also capable of spreading through the sapwood (Larsson et al. 2006), forming conks around the decayed tissues (Jones 1998; Fig. 25). Phellinus resupinatus is also a factor of Esca disease, causing white rot and decline of the cordons in vineyards (Cloete et al. 2016), besides other symptoms caused by this disease (Jayawardena et al. 2019a).

Fig. 25

Phellinus igniarius. a causing white-rot decay on willow b–d basidiomes on living willow e, f hymenial setae g tramal skeletal hyphae h basidiospores, Scale bars: e–h = 10 µm

Hosts—Most species in the P. igniarius species complex are specialized to a single or few angiosperm genera (Fischer and Binder 1995; Zhou et al. 2016), and only P. piceicola has been reported from gymnosperms (Cui and Dai 2012). Species of the P. igniarius species complex have been recorded from various host genera, such as Acer, Alnus, Arctostaphylos, Betula, Carpinus, Fagus, Fraxinus, Laburnum, Picea, Populus, Prunus, Salix, Sorbus and Tilia (Tomšovský et al. 2010; Brazee 2015; Zhou et al. 2016). The members of other Phellinus sensu stricto lineages are known from several additional angiosperm genera, such as Artemisia, Astronium, Caesalpinia, Carya, Castanopsis, Dimorphandra, Minquartia, Morus, Sacaglottis, Schinopsis, Quercus and Vitis (Lombard and Larsen 1985; Decock et al. 2006; Yombiyeni et al. 2011; Cui and Decock 2013; de Campos-Santana et al. 2016; Vlasák and Vlasák 2017; Salvador-Montoya et al. 2018).

Morphological based identification and diversity

Phellinus in a wider sense is morphologically heterogenous. The main features of the P. igniarius species complex are the crusted pileal surface (except resupinate species), the hymenial setae arising from the subhymenium (except specimens of “P. pseudoigniarius”, see Dai and Yang 2008; Zhou et al. 2016), and the colourless, inamyloid, indextrinoid and weakly cyanophilous basidiospores (Wagner and Fischer 2001; Dai 2010; Zhou et al. 2016). In many cases, the species separation in the complex is difficult when solely based on morphological characters (Sell 2008). Host preference is also widely used for delimiting species (Tomšovský et al. 2010).

Similar to members of the P. igniarius species complex, other Phellinus species also have perennial basidiomata, but differ in having distinctive macroscopical features (e.g. size and shape of pores, rimose surface, cracked basidiocarps, absence of pileus crust, see Dai et al. 2008; Bian et al. 2016; Cloete et al. 2016; Vlasák and Vlasák 2017) or microscopic characteristics (e.g. hyphal structure, the shape of setae, basidiospore reaction in chemical solutions). For example, P. bicuspidatus is unique in having a monomitic hyphal system with short bicuspid setae (Lombard and Larsen 1985; Cloete et al. 2016). Members of the P. ellipsoideus group are well-characterised by their weakly dextrinoid basidiospores and hooked hymenial setae (Zhu et al. 2018).

There are several “Phellinus” species which have been described solely on morphological features. The status of these species should be critically re-evaluated based on molecular evidence. Amongst these, certain species (e.g. P. deuteroprunicola, P. eugeniae, P. formosanus, P. livescens, P. prunicola, P. setulosus, P. tenuiculus, P. wahlbergii) may belong to Phellinus sensu stricto (Gilbertson 1979; Chang 1995; Chang and Chou 1999, 2000; Robledo et al. 2003; Wang et al. 2011; Rajchenberg et al. 2015; Campos-Santana et al. 2016), but further studies are required to confirm their placements.

Molecular based identification and diversity

In early molecular studies, the restriction fragment length polymorphism (RFLP) data of enzymatically amplified rDNA was used by Fischer (1995) to study the taxonomy of P. igniarius and its closest relatives in Europe. Later, single nuclear genes (ITS, Fischer and Binder 2004), or combined datasets (ITS-tef1, Tomšovský et al. 2010; Zhou et al. 2016) were used to investigate the species boundaries and phylogenetic relationships within the P. igniarius species complex. Phylogenetic analyses by Brazee (2015) used ITS, LSU, tef1 and rpb2, with isolates representing 13 species-level lineages in the complex. Zhou et al. (2016) distinguished 15 species, five of which are described as new from China and the USA. Based on our multigene analysis (Fig. 26), 16 species can be found in the P. igniarius species complex, distributed throughout the Northern Hemisphere. Amongst these, ten species are known from eastern Asia, eight from Europe and seven from North America.

Fig. 26

Phylogram generatedfrom RAxML analysis based on combined ITS, LSU, Tef1-α and rpb2 sequence data of Phellinus species. Related sequences were obtained from GenBank. Fifty-five strains are included in the analyses, which comprised 3170 characters including gaps. The tree was rooted with Phellinopsis conchata (DLL2009-149 and L-7601). Tree topology of the ML analysis was similar to the Bayesian analysis.ML bootstrap values >50% and BYPP >0.80% are shown respectively near the nodes

Phellinus caribaeo-quercicola was the first species described from the “P. ellipsoideus group” based on molecular evidence (Decock et al. 2006). The nLSU-based phylogenetic analysis of Decock et al. (2006), have shown that P. caribaeo-quercicola grouped close to the P. igniarius species complex and some other Phellinus species (viz. P. bicuspidatus, P. chaquensis and P. spiculosus). Later molecular taxonomic studies used combined datasets of various nuclear markers. The combined analyses of nITS, nLSU, tef1 and rpb2 have confirmed the phylogenetic position of P. caribaeo-quercicola and five morphologically similar species have been accepted in Phellinus sensu stricto (Yombiyeni et al. 2011; Cui and Decock 2013; Campos-Santana et al. 2016; Zhu et al. 2018). Currently, this later group consists of six species and mostly has tropical or subtropical distributions (Zhu et al. 2018).

Recommended genetic marker (genus level)—LSU

Recommended genetic markers (species level)—ITS, tef1, rpb2

Accepted number of species—There are 479 epithets listed in Index Fungorum (2020. However, most of the species belong to other poroid Hymenochaetaceae genera, such as Fomitiporia, Fomitiporella, Fulvifomes, Fuscoporia, Nothophellinus, Phellinidium, Phellinopsis, Phellinotus, Phellopilus, Phylloporia, Porodaedalea, Sanghuangporus and Tropicoporus (Wagner and Fischer 2001, 2002; Niemalä et al. 2001; Dai 2010; Drechsler-Santos et al. 2016; Rajchenberg et al. 2015; Zhou et al. 2016). Based on molecular data, 30 species are accepted in Phellinus sensu stricto, from among 16 species in the P. igniarius species complex (Table 16; Fig. 26).

Table 16 DNA barcodes available for Phellinus

References—Tomšovský et al. (2010) (phylogeny, P. igniarius species complex, Europe), Brazee (2015) (phylogeny, P. igniarius species complex, North America), Zhou et al. (2016) (phylogeny, P. igniarius species complex), Zhu et al. (2018) (phylogeny, P. ellipsoideus group)

91. Pseudoseptoria Speg., Anal. Mus. nac. B. Aires, Ser. 3 13: 388 (1910) [1911]

Background

Spegazzini (1910) introduced Pseudoseptoria as an asexual genus typified with Pseudoseptoria donacicola. Wijayawardene et al. (2012) placed the genus under Ascomycota, genera incertae sedis. Quaedvlieg et al. (2013) placed the genus in Dothioraceae and this was accepted by Thambugala et al. (2014). With LSU sequence data, Crous et al. (2017) placed Pseudoseptoria to Saccotheciaceae. Wijayawardene et al. (2017a, b, 2018, 2020) accepted this placement. Species of Pseudoseptoria are recorded as pathogens on Poaceae (Quaedvlieg et al. 2013), impairing the photosynthetic process resulting in yield loss.

Classification—Ascomycota, Pezizomycotina, Dothideomycetes, Dothideomycetidae, Dothideales, Saccotheciaceae

Type species—Pseudoseptoria donacicola Speg.

Distribution—Australia, Canada, India, Italy, New Zealand, Poland, Russia, UK and USA (Dennis1986; Ginns 1986; French 1989; Pennycook 1989; Merezhko 1991; Cunnington 2003; Mulenko et al. 2008; Kamal 2010; Farr and Rossman 2020).

Disease symptoms—halo spot, leaf blotch and stem speckle

Halo spot: Elliptical, tan to brownish-grey spots (<10mm long) with a dark border surrounded by a prominent yellow halo that can be observed on the leaf blade, sometimes covering the entire leaf blade. In older lesions, small pycnidia may be visible (Slopek and Labun 1992; Carmona et al. 1996; Murray et al. 2013).

Leaf blotch: Brown flecks and frog-eye spots on leaf blades can be observed in early spring, which enlarges to straw-coloured blotches scattered with minute pycnidia. These spots may drop out, leaving holes (Horst 2013).

Stem speckle: The disease occurs in the leaves, sheaths, culms, and head spikes. The lesions are rectangular, ash white, (1-2 mm long) with a brown, thin border. The lesion is delimited by leaf veins and becomes distinct with a clear boundary. The conidia formed on the lesions disperse by wind and rain.

Pathogen biology, disease cycle and epidemiology

The pathogen is dispersed through spores in rain splash. Infection requires an extended period of wetness. Spore germination and infection occur optimally at temperatures between 15 and 25 °C. Spores produced in overwintering crop debris serve as sources of primary inocula (Sinclair and Dhingra1995). Further studies are needed regarding the disease mechanisms and disease cycle.

Hosts—members of Poaceae are susceptible: Alopecurus pratensis, Arrhenatheru melatius, Arundo donax, Bromus species, Dactylis glomerata, Danthonia spicata, Elymus alaskanus, Festu carubra, Hordeum vulgare, Panicum virgatum, Phleum species, Phragmites australis and Poa species (Ginns 1986; Pennycook 1989; Shivas1989; Greuter et al. 1991; Merezhko 1991; Roane and Roane 1996; Gravert and Munkvold 2002; Mulenko et al. 2008; Farr and Rossman 2020)

Morphological based identification and diversity

The genus is characterized by immersed, branched, septate, pale brown mycelium, pycnidial, solitary or linearly aggregated, immersed, brown, globose, unilocular, thin-walled conidiomata of walls of pale brown cells of textura angularis with distinct, central, circular ostioles. Conidiogenous cells are discrete, determinate or indeterminate, hyaline, smooth, ampulliform with a prominent cylindrical papilla and falcate. Conidia are fusoid, hyaline, aseptate, guttulate, smooth and thin-walled, and acutely rounded at each end (Sutton 1980; Quaedvlieg et al. 2013).

Molecular based identification and diversity

Quaedvlieg et al. (2013) revised the Septoria and septoria-like genera based on morphology and multi loci analyses and introduced two new species. Phylogenetic analysis conducted by Crous et al. (2017) was based only on LSU sequence data. In our analysis, we used LSU, ITS and rpb2 and obtained the same topology (Fig. 27).

Fig. 27

Phylogenetic tree generated by maximum likelihood analysis of combined LSU, ITS and rpb2 sequence data. Fourteen strains are included in the analyses, which comprised 2207 characters including gaps. The tree was rooted with Elsinoe veneta (CBS 164.29) and Elsinoe phaseoli (CBS165.31). Tree topology of the ML analysis was similar to the BYPP analysis. The best scoring RAxML tree with a final likelihood value of − 7330.368152 is presented. The matrix had 500 distinct alignment patterns, with 29.98% of undetermined characters or gaps. Estimated base frequencies were as follows: A = 0.250948, C = 0.235140, G = 0.278289, T = 0.235623; substitution rates AC = 2.242594, AG = 3.171649, AT = 1.699638, CG = 1.459509, CT = 8.754890, GT = 1.000000; gamma distribution shape parameter α = 0.601361 ML bootstrap values ≥ 65% and BYPP ≥ 0.90 are shown respectively near the nodes

Recommended genetic marker (genus level)—LSU

Recommended genetic markers (species level)—LSU, ITS and rpb2

Accepted number of species—There are eight epithets listed in Index Fungorum (2020). However, only three species have DNA sequence data (P. collariana, P. donacis and P. obscura) (Table 17).

Table 17 DNA barcodes available for Pseudoseptoria

References—Sutton (1980) (morphology); Quaedvlieg et al. (2013), Crous et al. (2017) (morphology and phylogeny)

92. Stemphylium Wallr., Flora Cryptogamica Germaniae 2: 300 (1833)

Background

Stemphylium mainly comprises saprobes or weak plant pathogens (Woudenberg et al. 2017). However, some species are primary pathogens causing leaf blight on various crops, resulting in yield and economic losses (Hanse et al. 2015; Brahmanage et al. 2018). The asexual morph is a dematiaceous hyphomycete while the sexual morph was previously defined as Pleospora sensu stricto (Inderbitzin et al. 2009; Woudenberg et al. 2017). Rossman et al. (2015) recommended the use of Stemphylium over Pleospora which has been followed by various authors (Hongsanan et al. 2017, 2020; Wijayawardene et al. 2018, 2020). Stemphylium is one of the most important moulds human allergens in the USA (Gutiérrez-Rodríguez et al. 2011). Brahmanage et al. (2018) discussed the pathogenicity, disease severity, distribution and molecular phylogenetic affinities of pathogenic isolates of Stemphylium.

Stemphylium leaf blight caused by S. versicarum was identified as an emerging disease in New York, USA. Sharma et al. (2020b) provided two genome resources for two S. versicarum isolates from leaf blight of onion. Genomic data allows for an understanding of the population biology, fungicide resistance, as well as development of control strategies against the disease. Pathogenesis related 511 secreted proteins were predicted from S. lycopersici by Zeng et al. (2018) which helps in understanding the roles of proteins in host penetration and tissue necrosis. Stemphylium loti secretes Tenuazonic acid, inhibiting the plant plasma membrane H+-ATPase, which results in membrane potential depolarization and eventually necrosis (Bjørk et al. 2019). Su et al. (2019) fine-mapped the tomato grey spot resistance gene Sm, in a 185kb region through a map-based cloning strategy. Leach et al. (2020) identified a relationship between thrips (Thrips tabaci) and S. vesicarium in the development of Stemphylium leaf blight in onion.

Classification—Ascomycota, Pezizomycotina, Dothideomycetes, Pleosporomycetidae, Pleosporales, Pleosporaceae

Type species—Stemphylium botryosum Wallr.

Distribution—worldwide

Disease symptoms—Gray spot, Stemphylium leaf blight (Leaf spot, defoliation, curling and bending of the leaf margins and stems)

Initial symptoms of the leaves are small, irregular, brown spots. Generally, the spots gradually lighten and eventually become greyish as they become necrotic and dry. When severe, yellow spots can be seen throughout all leaves of the plant and the heavily infected leaves die (Basallote-Ureba et al. 1999, Crous et al. 2016; Brahmanage et al. 2018).

Hosts—Species are pathogenic on a wide range of hosts including Amaryllidaceae, Asparagaceae, Fabaceae, Malvaceae, Poaceae, Rosaceae and Solanaceae

Pathogen biology, disease cycle and epidemiology

Species can survive as saprobes on crop residues, soil, plant debris and on many alternative hosts and ascospores become the primary inocula in the following season. Once the disease is established during favourable conditions, conidial production in primary lesions may occur, dispersing spores to healthy plants by wind and rain splashing. Environmental factors such as temperature and moisture are key factors in disease development. Seedlings of plants can transmit the diseases if they become infected in the nursery (Basallote-Ureba et al. 1998, 1999; Boshuizen et al. 2004; Zheng et al. 2010; Blancard 2012). However, to date, diseases and epidemiology such as factors affecting the disease development, interactions with different hosts and genetics of host resistance are poorly studied (Das et al. 2019).

Morphological based identification and diversity

Species can be distinguished from other hyphomycetes in Pleosporaceae forming phaeodictyospores, based on percurrent proliferation of its conidiophores and apically swollen conidiogenous cells (Köhl et al. 2009). Simmons (1967) established criteria for morphological identification of various Stemphylium species and introduced Pleospora herbarum as the sexual morph of the type species Stemphylium botryosum. However, Simmons (1985) subsequently reclassified and reported Pleospora tarda as the sexual morph of Stemphylium botryosum and Pleospora herbarum as the sexual morph of Stemphylium herbarum (Moslemi et al. 2017). Morphological features, such as size and time of pseudothecial maturation, conidiophores and conidia and ascospore shape and size can be considered as important characteristics in species identification (Câmara et al. 2002; Fig. 28).

Fig. 28

Stemphylium sp. a Ascomata on host b Vertical section through an ascoma c immature and mature asci d Pseudoparaphyses e Ascospores f Ascospores in Indian ink. Scale bars: b = 50 μm, c–f = 10 μm

Köhl et al. (2009) and Woudenberg et al. (2017) pointed out that the lack of (ex-) type material of species and morphology-based species identifications without molecular evidence make it difficult in determining correct species nomenclature. Therefore, relying on morphological characters alone in identifying species is not recommended.

Molecular based identification and diversity

ITS (rDNA) and glyceraldehyde-3-phosphate dehydrogenase (gapdh) sequences were used by Câmara et al. (2002) to confirm the monophyly of Stemphylium. In the extensive study of 110 Stemphylium strains from various hosts and DNA sequence data of ITS, gapdh and tef1 loci and the intergenic spacer between vmaA and vpsA, Inderbitzin et al. (2009) identified 23 representatives derived from type strains, while 40 strains remained unnamed. Woudenberg et al. (2017) revised the genus and accepted 28 species, synonymizing 22 names and proposing two new combinations based on combined analyses of the ITS, gapdh and cmdA gene regions. Marin-Felix et al. (2019) introduced three new species (S. rombundicum, S. truncatulae and S. waikerieanum), while Brahmanage et al. (2018) introduced S. dianthi based on multi loci phylogeny. In this study, we reconstruct the phylogeny based on combined ITS, gapdh and cmdA sequence data (Fig. 29).

Fig. 29

Phylogram generated from MP analysis based on combined sequences of ITS, gapdh and cmdA sequences of all species of Stemphylium. Related sequences were obtained from GenBank. Thirty three taxa are included in the analyses, which comprise 1936 characters including gaps, of which 1355 characters are constant, 271 characters are parsimony-uninformative and 310 characters parsimony-informative. The parsimony analysis of the data matrix resulted in the maximum of four equally most parsimonious trees with a length of 1112 steps (CI = 0.660, RI=0.721, RC = 0.476, HI = 0.340) in the first tree. Single gene analyses were carried out and compared with each species, to compare the topology of the tree and clade stability. The tree was rooted with Alternaria abundance (CBS 534.83). MP bootstrap support value ≥ 50% and BYPP ≥ 0.9 are shown respectively near the nodes. Ex-type strains are in bold

Recommended genetic marker (genus level)—ITS

Recommended genetic markers (species level ) —cmdA, gapdh

Accepted number of species—There are 207 epithets listed in Index Fungorum, however only 32 species have DNA sequence data (Table 18).

Table 18 DNA barcodes available for Stemphylium

References—Simmons (1967), Köhl et al. (2009) (morphology); Câmara et al. (2002), Inderbitzin et al. (2009), Moslemi et al. (2017), Woudenberg et al. (2017), Brahmanage et al. (2019), Marin-Felix et al. (2019) (morphology and phylogeny)

93. Thyrostroma Höhn., Sitzungsberichte der Kaiserlichen Akademie der Wissenschaften Math.-naturw. Klasse Abt. I 120: 472 (1911)

Background

Thyrostroma belongs to Dothidotthiaceae of Pleosporales in Dothideomycetes, Ascomycota (Hongsanan et al. 2020). Thyrostroma was established by Höhnel (Höhnel 1911) and is typified by T. compactum. Thyrostroma had been treated as a synonym of Coryneum, Stegonsporium, Stigmina, and Thyrococcum, Thyrostromella and Wilsonomyces (Höhnel Höhnel 1911; Morgan-Jones 1971; Sutton and Pascoe 1989; Sutton 1997; Index Fungorum 2020). Thyrostroma has been reported as the asexual morph of Dothidotthia based on the production of a hyphomycete state in culture (Ramaley 2005), however, there is no phylogenetic evidence to support this link. With new morphological information and phylogenetic analyses, Thyrostroma and Dothidotthia species were retained in separate genera (Crous et al. 2016; Marin-Felix et al. 2017; Senwanna et al. 2019). Thyrostroma species are pathogens, saprobes or endophytes associated with canker, dieback and leaf spots in terrestrial habitats (Yuan and Old 1990; Marin-Felix et al. 2017; Senwanna et al. 2019). Species of Thyrostroma have been recorded from various plants, however, host-specificity and pathogenic capacity of Thyrostroma has not yet been clarified.

Classification—Ascomycota, Pezizomycota, Dothideomycetes, Pleosporomycetidae, Pleosporales, Dothidotthiaceae

Type species—Thyrostroma compactum (Sacc.) Höhn

Distribution—Australia, Iran, Korea, Russia, USA, Uzbekistan

Disease Symptoms—Thyrostroma canker, dieback and leaf spots (Fig. 30a, b)

Fig. 30

a, b Symptoms on Ulmus pumila caused by Thyrostroma ulmicola (MFLU 16-1622); c, d Sporodochia on the host surface. e Section of sporodochium. f Conidiogenesis and conidiogenous cells. g–l Conidia. m Germinated conidium. Scale bars: d = 1000 µm, e = 200 µm, f–m = 30 µm

Hosts—Pathogens of Acanthophyllum sp., Astragalus sp., Capparis parvifiora, Celtis occidentalis, Cornus officinalis, Echinops sp., Elaeagnus angustifolia, Ephedra equisetina, Eucalyptus mannifera subsp. maculosa, Franseria sp., Halimodendron halodendron, Lycium barbarum, Morus alba, Robinia pseudoacacia, Sambucus caerulea, Styphnolobium japonicum, Tilia cordata, Ulmus pumila (Farr and Rossman 2020).

Morphological based identification and diversity

Thyrostroma species can be differentiated using conidial dimensions and septation in aged conidia and molecular phylogeny (Crous et al. 2016; Marin-Felix et al. 2017; Senwanna et al. 2019; Fig. 30). Senwanna et al. (2019) reported the sexual morph of Thyrostroma in T. ulmicola for the first time. The sexual morph is characterized by pseudothecial, immersed, erumpent or superficial, uniloculate or multiloculate ascostromata, globose to subglobose ascomata, a two-layered peridium, bitunicate, clavate asci, fusiform to ellipsoidal, 1-septate, ascospores.

Molecular based identification and diversity

In the past, there has been no comprehensive phylogenetic study in Thyrostroma and consequently, its taxonomy was and still is mostly based on morphological characters. Based on LSU sequence data, Thyrostroma clustered in a well-supported clade within the Dothidotthiaceae (Marin-Felix et al. 2017; Crous et al. 2019). The asexual morph and sexual morph relationship were resolved by Senwanna et al. (2019) by molecular evidence. To achieve correct generic and species identification and taxonomic placement, phylogenetic studies using LSU, SSU, ITS, and tef1 were performed (Senwanna et al. 2019). This study reconstructs the phylogeny using a combined LSU, SSU, ITS, and tef1 sequence dataset (Fig. 31). The topology is in accordance with Marin-Felix et al. (2017), Senwanna et al. (2019) and Hyde et al. (2020b).

Fig. 31

Phylogenetic tree generated by maximum likelihood analysis of LSU, SSU, ITS and tef1 sequence data of Thyrostroma species. Related sequences were obtained from GenBank. The tree was rooted with Dothidotthia robiniae (MFLUCC 16-1175), D. symphoricarpi (CPC 12929) and Wilsonomyces carpophilus CBS 147.36). Tree topology of the ML analysis was similar to the Bayesian analysis. The best scoring RAxML tree with a final likelihood value of − 5556.187049 is presented. The matrix had 170 distinct alignment patterns, with 18.30% of undetermined characters or gaps. Estimated base frequencies were as follows: A = 0.243922, C = 0.240705, G = 0.270231, T = 0.245142; substitution rates AC = 3.570314, AG = 6.771467, AT 4.177691, CG = 1.603201, CT = 31.935571, GT = 1.000000; gamma distribution shape parameter α = 0.602378. Maximum likelihood bootstrap support values greater than 60% and Bayesian posterior probabilities ≥ 0.95 (BYPP) are indicated above the nodes. Ex-type (ex-epitype) and voucher strains are in bold

Recommended genetic marker (genus level)—LSU

Recommended genetic markers (species level)—ITS, tef1, rpb2 and tub2

LSU, ITS and tef1 are the common genetic markers used in the identification of Thyrostroma species. Combined LSU, SSU, ITS and tef1 genes provide a satisfactory resolution for resolving species. Based on the comparison of ITS and tef1gene regions, most species in Thyrostroma are not significantly different from one another, therefore, Senwanna et al. (2019) suggested that rpb2, tub2 are reliable genes for distinguishing species within Thyrostroma.

Accepted number of species—There are 27 epithets in Index Fungorum (2020), however only 13 species have DNA sequence data (Table 19).

Table 19 DNA barcodes available for Thyrostroma

References—Höhnel (1911), Yuan and Old (1990), Ramaley (2005) (morphology); Marin-Felix et al. (2017), Crous et al. (2016, 2019), Senwanna et al. (2019) (morphology and phylogeny).

94. Wojnowiciella Crous, Hern.-Restr.& M.J. Wingf., Persoonia 34, 201 (2015)

Background

Wojnowiciella was introduced by Crous et al. (2015) to include Wojnowiciella eucalypti which exhibited somewhat similar morphological characteristics to Wojnowicia, such as setose pycnidia, with ampulliform, enteroblastic, phialidic conidiogenous cells, but differed with apapillate conidiomata lacking setae and having dark brown conidia.

Classification—Ascomycota, Pezizomycota, Dothideomycetes, Pleosporales, Phaeosphaeriaceae

Type species—Wojnowiciella eucalypti Crous, Hern.-Restr. & M.J. Wingf

Distribution—Australia (Hernandez-Restrepo et al. 2016), China (Crous et al. 2015, Giraldo et al. 2017), Colombia (Crous et al. 2015, Giraldo et al. 2017), New Zealand (Crous et al. 2019), South Africa and Western Cape (Crous et al. 2016)

Disease symptoms—Leaf spots

Most species are reported as saprobes with the exception of Wojnowiciella cissampeli, W. eucalypti and W. vibruni which were isolated from leaves and twigs of Cissampelos capensis, Eucalyptus and Viburnum utile respectively (Hernandez-Restrepo et al. 2016). Their pathogenicity or disease symptoms are not indicated clearly and there is a need to establish pathogenicity of these species.

Hosts—Cissampelos capensis, Dactylis sp., Eucalyptus grandis, Rosa sp., Leptocarpus sp., Lonicera sp., Spartium sp. and Viburnum utile (Farr and Rossman 2020).

Morphological based identification and diversity

Wojnowiciella was introduced to include species that were phylogenetically distinct but morphologically similar to Wojnowicia (Crous et al. 2015). Wojnowiciella is characterized by apapillate conidiomata without setae and dark brown conidia. Some species of Wojnowiciella also produce hyaline microconidia. Karunarathna et al. (2017) first reported the sexual morph of W. dactylidis. Phookamsak et al. (2019) transferred Wojnowicia rosicola to Wojnowiciella rosicola based on morphology and phylogenetic analyses.

Molecular based identification and diversity

Wojnowiciella is a well-supported genus in the family Phaeosphaeriaceae (Phookamsak et al. 2019). A combined multiloci phylogeny of LSU, SSU, tef1 and ITS is used in placing species of Wojnowiciella within Phaeosphaeriaceae. To identify species within the genus ITS, LSU, rpb2 and tef1 are used (Marin-Felix et al. 2019; Phookamsak et al. 2019). Here we provide an updated phylogenetic tree for this genus (Fig. 32).

Fig. 32

Phylogram generated from MP analysis based on combined sequences of ITS, LSU and tef1 sequences of all the accepted species of Wojnowiciella. Related sequences were obtained from GenBank. Ten taxa are included in the analyses, which comprise 2460 characters including gaps. Single gene analyses were carried out and compared with each species, to compare the topology of the tree and clade stability. The tree was rooted with Galiicola baoshanensis (HKAS102234). The best scoring RAxML tree with a final likelihood value of − 6772.195394 is presented. The matrix had 261 distinct alignment patterns, with 0.96% of undetermined characters or gaps. Estimated base frequencies were as follows: A = 0.230657, C = 0.279364, G = 0.252128, T = 0.237852; substitution rates AC = 1.388608, AG = 2.845402, AT = 2.389715, CG = 0.838197, CT = 7.220493, GT = 1.000000; gamma distribution shape parameter a = 0.650385. ML and MP bootstrap support value ≥ 50% are shown respectively near the nodes. Ex-type strains are in bold

Recommended genetic markers (genus level)—LSU

Recommended genetic markers (species level)—ITS, rpb2, tef1

Accepted number of species—Nine species are accepted with molecular data (Table 20).

Table 20 Details of Wojnowiciella, isolates used in the phylogenetic analyses

References—Crous et al. (2015), Karunarathna et al. (2017), Marin-Felix et al. (2019), Phookamsak et al. (2019) (morphology and phylogeny)

Updated genera

The following genera are updated due to the addition of many new species during recent years.

95. Cladosporium Link, Mag. Gesell. naturf. Freunde, Berlin 7: 37 (1816) [1815]

Cladosporium Link, Mag. Gesell. naturf. Freunde, Berlin 7: 37 (1816) [1815]

Background

Cladosporium belongs to Cladosporiaceae in the order Capnodiales (Hyde et al. 2013). Established in 1816 with C. herbarum as type species, Cladosporium is one of the largest genera of dematiaceous hyphomycetes. Davidiella was erected by Braun et al. (2003) to accommodate the sexual morph of Cladosporium sensu stricto. Davidiella was therefore recognized as a synonym of Cladosporium as Cladosporium has priority over Davidiella at generic rank, and is also the more commonly used name in literature (Bensch et al. 2012). Therefore, Cladosporiaceae took preference over Davidiellaceae (Bensch et al. 2012). Cladosporium species have a worldwide distribution and can be easily spread in the environment, because of their small conidia. Cladosporium includes many important pathogens causing leaf spots and stem rots of many plant hosts. For example, Cladosporium fulvum is the causal agent of tomato leaf mold (van Kan et al. 1991). Cladosporium species have been recorded as endophytes and may have a positive effect, for example, C. sphaerospermum was isolated from the roots of Glycine max which can promote its growth (Hamayun et al. 2009). Some species, such as C. herbarum, are also known as common contaminants in clinical laboratories and cause allergic lung disease (de Hoog et al. 2000). Several species were also isolated from human respiratory samples (Sandoval-Denis et al. 2016). Thirteen species are fungicolous (Heuchert et al. 2005; Sun et al. 2019) and have the potential for biological control in agriculture and forestry (Torres et al. 2017).

There have been studies towards understanding the genetic components of Cladosporium. Cladosporium fulvum is an important model species in the plant pathology study. Iakovidis et al. (2020) reported classical mapping strategies for loci of tomato that response to sequence-monomorphic effector Ecp5. Convergent evolution could be used for choosing different functional genes according to individual plant breeding needs. Ge et al. (2019) showed that Cladosporium species have the potential to be used in industrial processes. They identified a new glucose oxidase gene CtgoxB from C. tianshanense and suggested this could be a candidate for the aquatic feed and detergent industries. Transcriptome and proteome analyses of C. fulvuim showed that 14 out of 59 predicted proteases are expressed during in vitro and in planta, of which nine belong to serine proteases and the rest belong to metallo and aspartic proteases (Jashni et al. 2019). This study also confirmed the presence of six proteases at proteome level during the infection.

Grinn-Gofroń et al. (2019) developed and evaluated the models of forecasting possibilities of airborne spore concentrations in 18 sites in six countries across Europe. The study revealed the possibility of reliable prediction of fungal spore levels using gridded meteorological data. They concluded that these forecasting models can be used in the more timely and efficient management of phytopathogenic and of human allergic diseases. An environmentally isolated strain of C. sphaerospoermum substantially enhanced plant growth, early flowering and increase in crop yield after exposure in vitro (Li et al. 2019). Pan et al. (2020) identified four new hybrid polyketides (Cladosin L-O) from C. shaerospermum which showed strong cytotoxicity, antifungal activity and moderate antibacterial activity.

Classification: Ascomycota, Pezizomycotina, Dothideomycetes, Pleosporomycetidae, Capnodiales, Cladosporiaceae

Type species–Cladosporium herbarum (Pers.) Link

Distribution– Worldwide

Disease symptoms–Leaf spots, leaf blight, discolourations, necrosis, or shot-hole symptoms, on stems and fruits, rots

Hosts– Cladosporium species occur on a wide range of host plants including Asparagaceae, Asteraceae, Fabaceae, Myrtaceae, Orchidaceae, Poaceae, Solanaceae and Vitaceae (Farr and Rossman 2020). Some species can be hyperparasites of insects and fungi (Heuchert et al. 2005; Islam et al. 2019; Sun et al. 2019; Abdel-Baky 2000). These species can cause allergies in humans such as sneezing, hives and also can cause eye, ear and sinus infections (de Hoog et al. 2000).

Pathogen biology, disease cycle and epidemiology

Cladosporium survives in the soil or on plant debris and produce spores during humid weather. Fungal spores germinate under high humidity and cool to warm temperatures. Wind, rain and irrigation splash, workers, tools, and insects readily disseminate spores (Jordan et al. 1990; Lan and Scherm 2003; Liu et al. 2019).

Morphological based identification and diversity

The asexual morph of Cladosporium species is characterized by a unique coronate structure of the conidiogenous loci and conidia, consisting of the central convex dome surrounded by a raised periclinal rim (Bensch et al. 2012; Fig. 33), while ascomata of sexual morphs are identical to those of Mycosphaerella (sect. Tassiana) (Braun et al. 2003). Historically, all types of dematiaceous hyphomycetes with amero- to phragmosporous conidia formed in acropetal chains had been assigned to Cladosporium sensu lato, resulting in the complication to resolve a natural classification of Cladosporium. Various mycologists proposed natural genetic circumscriptions of Cladosporium (David 1997; Braun et al. 2003; Aptroot 2006). David (1997) found the unique structure of conidiogenous loci and conidial hila using scanning electron microscopy. Based on the genetic circumscriptions, some cladosporioid groups, such as Fusicladium being non-coronate (Schubert et al. 2003), have been excluded from Cladosporium s. str. Various Cladosporium species have been re-examined based on the new generic concepts (Schubert and Braun 2004, 2005a, b, 2007; Schubert 2005; Schubert et al. 2006; Braun and Schubert 2007; Braun et al. 2008). A polyphasic approach revealed three major species complexes within Cladosporium, viz. C. cladosporioides, C. herbarum and C. sphaerospermum (Schubert et al. 2007; Dugan et al. 2008; Bensch et al. 2010; Bensch et al. 2015). A modern monograph of the genus treated 993 names of Cladosporium sensu lato, of which 169 were recognized in Cladosporium sensu stricto and others remain doubtful (Bensch et al. (2012).

Fig. 33

Cladosporium cladosporioides. a Conidiomata. b, c, e Macro- and micronematous conidiophores and conidia chains. d. Secondary ramoconidia. f. Conidia. Scale bars: b, c, e, f = 50 µm, d–g=10 µm

Molecular based identification and diversity

The first molecular examination of Cladosporium-like hyphomycetes based on ITS and SSU was carried out by Braun et al. (2003), who confirmed the strong heterogeneity. A new genus Davidiella was established to accommodate the sexual morphs of Cladosporium sensu stricto species which were previously assigned in Mycosphaerella. Aptroot (2006) made a better circumscription of Davidiella after he found species of Davidiella have ascospores with irregular cellular inclusions, which are absent in Mycosphaerella. Schoch et al. (2006) studied the phylogenetic relationships of 96 taxa of the Dothideomycetes using LSU, SSU, tef1 and rpb2 gene data. Davidiella and its Cladosporium asexual morphs were assigned to the family Cladosporiaceae in the order Capnodiales, together with Mycosphaerellaceae. Crous et al. (2007) delimited Cladosporium from morphologically similar genera using their morphology and DNA phylogeny based on LSU. Several species were transferred to new genera such as Hyalodendriella, Ochrocladosporium, Rachicladosporium, Rhizocladosporium, Toxicocladosporium and Verrucocladosporium. Furthermore, C. castellanii was confirmed as a synonym of Stenella araguata, while the type species of Stenella resided in Teratosphaeriaceae instead of Mycosphaerellaceae. Schubert et al. (2007) performed a comprehensive study of the C. herbarum species complex based on both morphology and phylogenetic analysis with five combined genes. Bensch et al. (2010) carried out species and ecological diversity within the C. cladosporioides species complex. More than 200 isolates belonging to the C. cladosporioides species complex were examined and analyzed on the basis of ITS, actand tef1 gene regions. A comprehensive monograph of Cladosporium sensu lato was provided by Bensch et al. (2012) based on morphology and combined ITS, act and tef1 sequence data. In their study, 993 names assigned to Cladosporium sensu lato are treated and 169 names were recognized in Cladosporium sensu stricto. Bensch et al. (2015) introduced the three major species complexes in Cladosporium, i.e. C. cladosporioides, C. herbarum and C. sphaerospermum, and 19 new species were described. Razafinarivo et al. (2016) introduced a new species C. lebrasiae from milk bread rolls in France, Ma et al. (2017) introduced six new soil-inhabiting Cladosporium species from plateaus in China. Bensch et al. (2018) studied Cladosporium species from indoor environments and introduced 16 new species. Several new Cladosporium species including Cladosporium omanense (Halo et al. 2019), C. passiflorae and C. passifloricola (Rosado et al. 2019) have been introduced more recently. In this study, we reconstruct the phylogeny of Cladosporium based on ITS, tef1 and act sequenced data (Table 21; Fig. 34).

Table 21 DNA barcodes available for Cladosporium

Fig. 34

Phylogram generated from Maximum Likelihood analysis based on ITS, tef1 and act sequenced data. Bootstrap support values ≥ 75% and Bayesian posterior probabilities ≥ 0.95 are given near the nodes. The ex-type (ex-epitype) and voucher strains are in bold. The tree is rooted with Toxicocladosporium banksiae CBS 128215

Recommended genetic marker (genus level)—ITS and LSU

Recommended genetic markers (species level)—act and tef1 (in a few cases tub2)

Accepted number of species–There are 844 epithets listed in Index Fungorum (2020), however, 138 species have DNA sequence data.

References–David (1997), Aptroot (2006), Schubert and Braun (2004, 2005a, b, 2007), Schubert (2005), Schubert et al. (2006), Braun and Schubert (2007), Braun et al. (2008) (morphology), Braun et al. (2003), Schoch et al. (2006), Bensch et al. (2010, 2012, 2015), Ma et al. (2017) (morphology and phylogeny)

96. Colletotrichum Corda, in Sturm, Deutschl. Fl., 3 Abt. (Pilze Deutschl.) 3(12): 41 (1831)

Background

Colletotrichum was introduced by Corda (1831), belonging to Glomerellaceae (Glomerellales, Sordariomycetes), and is the sole member of this family (Maharachchikumbura et al. 2015, 2016; Hyde et al. 2020b). Species may occupy different lifestyles, ranging from necrotrophy to hemibiotrophy as well as endophytism (Crouch et al. 2014). Colletotrichum species are important plant pathogens in both tropical and temperate regions on many economically important crops (Hyde et al. 2009a, b, 2014; Cannon et al. 2012; Jayawardena et al. 2016b, c). Based on recognized scientific and economic importance this genus was voted the eighth most important plant pathogenic group in the world (Dean et al. 2012). Colletotrichum species have been identified as endophytes (Manamgoda et al. 2013; Tao et al. 2013; Hyde et al. 2014; Jayawardena et al. 2016c) and some are saprobes on dead plant material (Photita et al. 2005; Jayawardena et al. 2016b). A few species have been identified to be pathogenic to humans (C. coccodes, C. dematium, C. gloeosporioides (Natarajan et al. 2013)) and on insects (C. fioriniae (Damm et al. 2012b)). Colletotrichum species are cosmopolitan in distribution and show a diverse hosts association (Sharma et al. 2015). A host plant genus can be infected by many Colletotrichum species (Silva et al. 2012; Jayawardena et al. 2016c), and on the contrary, a single species of Colletotrichum can infect many host plants (Damm et al. 2012a, b; Weir et al. 2012).

Correct species identification is important to understand the species diversity, plant pathology and quarantine, concerning human infections, agriculture, bio-control, plant breeding, whole-genome sequencing, developing and maintaining knowledge databases, bioprospecting and understanding the evolutionary history (Jayawardena et al. 2016a). Due to a small number of distinctive morphological characters available for identification, misidentification of these species is frequent. Misapplication and misidentification of species are also due to the misunderstanding of their host-specific nature, ambiguous species boundaries and incorrect sequences (Cannon et al. 2012; Hyde et al. 2014; Jayawardena et al. 2016a). Therefore, having a stable taxonomy for the identification of these species is a significant practical concern (Shenoy et al. 2007). To establish a natural classification system, researchers strongly recommend the use of geographical, ecological, morphological and genetic data (Cai et al. 2009; Sharma and Shenoy 2016).

Species of Colletotrichum are extensively studied as model organisms (Cannon et al. 2012; Hyde et al. 2014). This enables the researchers to understand the pathogen variation, infection mechanism, evolution and population dynamics. Pathogenicity genes of C. graminicola, C. higginsianum and C. orbiculare have been studied (Huser et al. 2009; O’Connell et al. 2012). Asakura et al. (2009) discovered the importance of the pexophagy factor ATG26 for appressorium formation. A total of 28 genome projects that include 25 different Colletotrichum species can be found; 15 of these strains are still at the annotation stage and 13 are now at the ‘Fungal Standard Draft’ stage (Carbú et al. 2019). These genomes will allow further analysis of species diversity and evolutionary mechanisms and may serve as a foundation for genetic analysis that leads to a greater understanding of interactions between plants and fungal pathogens (Meng et al. 2020). Baroncelli et al. (2016) studied four strains of C. acutatum and illustrated the plasticity of Colletotrichum genomes and showed that major changes in host range are associated with relatively recent changes in gene content. A genome of C. fructicola from apple in China was compared with its reference genome, which identified a number of strong duplication/loss events at key phylogenetic nodes (Liang et al. 2018). Gan et al. (2019) provided the updated genome for C. orbiculare and also provided three draft genomes for C. trifolli, C. sidae and C. spinosum. Colletotrichum higginsianum has a compartmentalized genome consisting of gene-sparse, transposable elements dense regions with more effector candidate genes and gene-dense, TE-sparse regions harbouring conserved genes which help the pathogen to generate genomic diversity (Tsushima et al. 2019). Comparative genome analysis indicated that there is a rapid evolution of pathogenicity genes in C. tanaceti (Lelwala et al. 2019).

Species of Colletotrichum can be used as biocontrol agents and as biocatalysts (C. dematium, C. gloeosporioides, C. graminicola, C. lindemuthianum, C. orbiculare, C.theobromicola, C. trifoli (Jayawardena et al. 2016b)). Jayawardena et al. (2016b) discussed the importance of secondary metabolites produced by species with relation to pathogenesis, medicines, disease control and toxins.

Classification—Ascomycota, Pezizomycotina Sordariomycetes, Hypocreomycetidae, Glomerellales, Glomerellaceae

Type species—Colletotrichum lineola Corda, in Sturm, Deutschl. Fl., 3 Abt. (Pilze Deutschl.) 3(12): 41 (1831)

Distribution—Worldwide

Disease symptoms—Anthracnose disease, red rot, crown and stem rots, ripe rot, seedling blights and brown blotch.

Anthracnose disease symptoms include defined, often sunken necrotic spots on leaves, stems, flowers or fruits and may show a lot of variation depending on the host (35a−e).

Hosts—Pathogens on many host families including, Amaryllidaceae, Amaranthaceae, Anacardiaceae, Annonaceae, Apiaceae, Apocynaceae, Araceae, Araliaceae, Arecaceae, Asparagaceae, Asteraceae, Bignoniaceae, Campanulaceae, Caricaceae, Crassulaceae, Cucurbitaceae, Euphorbiaceae, Fabaceae, Iridaceae, Lamiaceae, Lauraceae, Malvaceae, Melastomataceae, Menispermaceae, Moraceae, Myrtaceae, Oleaceae, Olivaceae, Orchidaceae, Passifloraceae, Pinaceae, Piperaceae, Plumbaginaceae, Poaceae, Podocarpaceae, Polemoniaceae, Proteaceae, Ranunculaceae, Rosaceae, Rubiaceae, Rutaceae, Solanaceae, Theaceae and Vitaceae.

Pathogen biology, disease cycle and epidemiology

For Colletotrichum biology, disease cycle and epidemiology see Cannon et al. (2012) and De Silva et al. (2017).

Morphological based identification and diversity

Due to the overlapping morphological characters, species delimitation based on morphology alone is hardly possible (Jayawardena et al. 2016b; Marin-Felix et al. 2017; Fig. 35f–l).

Fig. 35

Colletotrichum sp. a Chilli anthracnose symptoms b Crown rot of strawberry c Leaf blight of Cannas sp. d, e strawberry anthracnose symptoms f, g Conidiomata and spore mass h Conidiophore i Setae j Conidiogenous cells and curved conidia k Conidia of C. gloeosporioides l Appressoria. Phylogram generated from RAxML analysis based on combined sequences of ITS and LSU of Mucor and Backusella species. Eighty-one taxa were used for the analysis, which consisted 1256 characters including gaps. The tree is rooted using Backusella lamprospora (CBS 195.28), and B. grandis (CBS 186.87). Likelihood of the best scoring ML tree was -16174.718247. The concatenated matrix contained 657 distinct alignment patterns with 22.27% of undetermined characters or gaps. Estimated base frequencies were as follows; A = 0.302146, C = 0.168433, G = 0.219872, T = 0.309549; substitution rates AC = 0.776817, AG = 3.154911, AT = 1.674079, CG = 0.632500, CT = 4.808262, GT = 1.000000; gamma distribution shape parameter α = 0.291909. Type species in the dataset are indicated using T. Maximum likelihood RAxML bootstrap support value ≥ 70% and Bayesian posterior probabilities ≥ 0.70 are shown near the nodes (ML/BYPP). – indicates bootstrap support values lower than 70% and * shows unrecovered branching

Molecular based identification and diversity

Cai et al. (2009) proposed the use of a polyphasic approach with multi-loci sequence analyses combined with geographical, ecological and morphological data for reliable species delimitation. Application of this polyphasic approach resulted in the delimitation of almost 200 species, most of them belonging to species complexes such as acutatum, boninense and gloeosporioides. There is no universal set of loci to use when identifying Colletotrichum species. Cannon et al. (2012), Damm et al. (2012a, b, 2013, 2014, 2019, Liu et al. 2016) used ITS, gapdh, chs, act, his and tub2 (with some also gs or cal) for studying species within the acutatum, boninense, dematium, destructivum, gigasporum, orbiculare, spaethianum and truncatum species complexes, while Weir et al. (2012) additionally applied gs, cal and sod2 within the gloeosporioides species complex. Hyde et al. (2014), Jayawardena et al. (2016b), Marin-Felix et al. (2017) used ITS, gapdh, chs, act and tub2 to differentiate the species. Using five loci for the whole genus gave similar results to 6-7 loci used for the whole genus. In contrast, Crouch et al. (2009b) applied ITS, sod2, apn2 and mat1/apn2, to study the graminicola and caudatum species complexes. Use of ApMat locus to delimit the species within gloeosporioides species complex was emphasized by Silva et al. (2012) and Sharma et al. (2015) as it provides a higher resolution when compared to previously used loci. However, studies by Liu et al. (2015, 2016) revealed that using this locus with other loci would provide a satisfactory species delimitation within the gloeosporioides species complex. For species delimitation, application of the Genealogical Concordance Phylogenetic Species Recognition (GCPSR) has proven to be a powerful tool (Cai et al. 2009). Coalescent-based species delimitation methods can also be used to infer the dynamic of divergence, evolutionary process and the relationships among species (McCormack et al. 2009, Liu et al. 2016).

Jayawardena et al. (2016b) provided the accepted species for the genus with backbone trees for each species complex and with notes for each accepted species. De Silva et al. (2017) reviewed the lifestyles and how this can be applied to plant biosecurity. Ariyawansa et al. (2015), Yan et al. (2015), Li et al. (2016), De Silva et al. (2017), Hyde et al. (2020a, b), Marin-Felix et al. (2017), Tibpromma et al. (2017, 2018), Samarakoon et al. (2018), Bhunjun et al. (2019) have introduced new species based on morphology, phylogeny and GCPSR evidence. Damm et al. (2019) introduced three new species complexes namely, dracaenophilum, magnum and orchidearum based on morphology and phylogeny. Cabral et al. (2020) based on pathological, morphological, cytogenomic, biochemical and molecular data, assigned the previously known C. kahawae subsp. ciggario as a new species, C. ciggario. At present, based on multi-loci phylogeny there are 14 species complexes. With 59 new species having been added since the last treatment (Marin-Felix et al. 2017), here we present an analysis using five loci (Table 21) for all Colletotrichum species (Fig. 36). From the studies conducted on this genus, it is clear that the resolution of species differs depending on both locus and species. Therefore, to select a better genetic marker and the best secondary barcoding gene/genes is still an ongoing process.

Fig. 36

Phylogram generated from MP analysis based on combined sequences of ITS, gapdh, chs, act and tub2 sequences of all species of Colletotrichum with molecular data. Related sequences were obtained from GenBank. Two hundred and fourty nine taxa are included in the analyses, which comprise 2296 characters including gaps, of which 868 characters are constant, 295 characters are parsimony-uninformative and 1133 characters parsimony-informative. The parsimony analysis of the data matrix resulted in the maximum of ten equally most parsimonious trees with a length of 10088 steps (CI = 0.283, RI=0.840, RC = 0.237, HI = 0.717) in the third tree. Single gene analyses were carried out and compared with each species, to compare the topology of the tree and clade stability. The tree was rooted with Monilochaetes infuscans (CBS 869.96) and M. populi (CBS 139623). MP bootstrap support value ≥ 50% and BYPP ≥ 0.9 are shown respectively near the nodes. Ex-type strains are in bold

Recommended genetic marker (genus level)—ITS

Recommended genetic markers (species level)—act, apmat, apn2, cal, chs-1, gapdh, gs, his, mat1/apn2, sod2, and tub2.

Accepted number of species—There are 903 epithets listed in Index Fungorum (2020), however, 247 species with molecular data are treated as accepted (Table 22).

Table 22 DNA barcodes available for Colletotrichum, isolates used in the phylogenetic analyses

References—Cai et al. (2009) (polyphasic approach); Hyde et al. (2009a, b) (morphology and accepted species); Cannon et al. (2012) (A review and an updated account of the genus); Crouch et al. (2009a, b, c, 2014), Damm et al. (2009, 2012a, b, 2013, 2014, 2019), Weir et al. (2012) (morphology and phylogeny); Hyde et al. (2014), Jayawardena et al. (2016b), Marin-Felix et al. (2017) (accepted number of species).

97. Mucor Fresen., Beitr. Mykol. 1: 7 (1850)

Background

Mucor belongs to the order Mucorales, which is among one of the most studied groups of early diverging lineages of fungi. The genus has the largest number of species within the order and half of the sequences submitted to GenBank for Mucorales are of Mucor (Hoffmann et al. 2013; Spatafora et al. 2016; Hyde et al. 2014; Nguyen and Lee 2018). Mucor belongs to the phylum Mucoromycota, subphylum Mucoromycotina, class Mucoromycetes, order Mucorales and family Mucoraceae (Wijayawardene et al. 2018, 2020). It was described by Fresenius in 1850 and the type species is Mucor mucedo. Recent molecular studies of mucoralean species have indicated that Mucor is polyphyletic (Nguyen et al. 2017). However, even with definite results showing the polyphyly of Mucor, few clear lineages within Mucor are recognized. Some of these lineages share innate characteristics, such as sporangium size and branching of tall sporangiophores and the morphology is still widely used in current taxonomy (Walther et al. 2013). Analysis of internal transcribed spacer (ITS) and large subunit (LSU) rDNA sequence data of several mucoralean species, showed that some Mucor species with curved sporangiophores grouped with species of Backusella and hence was transferred to Backusella (Walther et al. 2013; Nguyen et al. 2017). Mucor species are commonly isolated from soil, dung, insect, and fruits (Benny 2008). Some species are of biotechnological importance such as biofuel, enzyme, terpernoid production and biotransformation while other species cause mucoromycosis in immunosuppressed humans (Nguyen et al. 2017; Steve et al. 2018; Morin-Sardin et al. 2017). Comparative analyses of five Mucor species based on their lifestyles (M. fuscus and M. lanceolatus (used for cheese production), M. circinelloides and M. racemosus (opportunistic pathogens) and M. endophyticus (an endophyte)) revealed the core transcriptome comprising 5566 orthogroups included genes potentially involved in secondary metabolism. Due to the wide taxonomic range investigated, the five transcriptomes also displayed specificities that can be linked to the different lifestyles, such as differences in the composition of transcripts identified as virulence factors or carbohydrate transporters. Research on this genus has changed its course to identify the link between genetic and biological data, especially in terms of lifestyle and adaptations to a given habitat (Lebreton et al. 2019) (Figs. 37, 38).

Fig. 37

Phylogram generated from RAxML analysis based on combined sequences of ITS and LSU of Mucor and Backusella species. Eighty-seven taxa were used for the analysis, which consisted of 1264 characters including gaps. The tree is rooted using Backusella lamprospora (CBS 195.28), and B. grandis (CBS 186.87). Likelihood of the best-scoring ML tree was − 17553.567209. The concatenated matrix contained 716 distinct alignment patterns with 21.98% of undetermined characters or gaps. Estimated base frequencies were as follows: A = 0.302082, C = 0.168706, G = 0.219403, T = 0.309809; substitution rates AC = 0.749467, AG = 2.977575, AT = 1.651634, CG = 0.631954, CT = 4.647089, GT = 1.000000; gamma distribution shape parameter α = 0.302610. The type species are in bold. Scale bar indicates the number of substitutions per site. ML bootstrap support values greater than 70% are shown near the nodes

Fig. 38

Disease cycle of Phytophthora infestans (redrawn from Bengtsson 2013)

Classification—Zygomycota, Mucoromycotina, Mucoromycetes, Mucorales, Mucorineae, Mucoraceae

Type species—Mucor mucedo Fresen.

Distribution—Worldwide

Disease symptoms—Mucor rot and soft rot

Mucor species especially M. fragilis, M. irregularis, M. piriformis and M. racemosus often cause postharvest diseases such as Mucor rot and soft rot. The initial symptoms of Mucor rot are similar to plant diseases caused by green mold, blue mold, and sour mold. The infected tissue becomes soft and watery. The lesions turn light to dark brown and as the infection progresses, white or shiny grey sporangiophores form at the lesions. Fungal growth spreads across the whole host and masses of sporangiophores bearing black to pale brown sporangia are observed. Decaying fruits become “juicy” within which are abundant spores of the fungus (Li et al. 2014; Saito et al. 2016). Ito et al. (1979) found that three species of fruit flies namely Certitis capitata, Dacus cucurbitae and D. dorsalis, can transmit Mucor rot in guava.

Soft rot caused by Mucor racemosus results in water-soaked appearance followed by a softening of the infected part. When the disease progresses growth of white mycelium and brownish to grey sporangia can be observed. Finally, the infected tissue is broken down and disintegrates in a watery rot (Kwon and Hong 2005; López et al. 2016).

Hosts—Wide host range including, Actinidia deliciosa, Citrus reticulata, Dioscorea species, Fragaria × ananassa, Mangifera indica, Manihot esculenta, Prunus species, Psidium guajava, Solanum melongena, Solanum lycopersicum and Vitis species (Farr and Rossman 2020).

Pathogen biology, disease cycle and epidemiology

The pathogen reproduces asexually. Mucor rot often develops by infecting punctured wounds and cracks on the surface of the fruit, stem end or calyx of the host. In the early stages of the infection, the fruit becomes soft and appears water-soaked. The lesions formed are quasicircular or irregular, light to dark brown and the sporangiophores protrude through the wounds (Kwon and Hong 2005; Saito et al. 2016; Michailides and Spotts 1990). As the infection advances, the infected part disintegrates into a watery rot and the infection spreads and extends to all extremities of the fruit or even the surface of the container. The infected part is covered with a large mass of mycelium with erect sporangiophores and sporangia (Saito et al. 2016; Michailides and Spotts 1990). When tested, rotten apple and pear by some Mucor species release an alcoholic odour while Mucor rot in peaches and nectarines caused by M. piriformis emits a pleasant aromatic odour. At an advanced stage, Mucor rot can be distinguished from other rots caused by Rhizopus or Gilbertella. Differences are observed in the mycelial character, growth, sporangiophores and sporangia. For Mucor rot, erect, white or yellowish sporangiophore with grey to black sporangia is observed which covers the decay lesion densely. However, for Rhizopus rot, the mycelia are interwoven with stolons with dark sporangiophores and black sporangia. The sporangial wall eventually dries and falls apart while in Mucor rot, the sporangia absorb water from the sporangial wall which dissolves (Michailides and Spotts 1990).

Morphology- based identification and diversity

Mucor is characterized by fast-growing colonies. The sporangiophores are simple or branched without basal rhizoids. However, under some conditions, they form rhizoids. These species normally form globose sporangia, containing the columella and spores. The sporangium is non-apophysate with pigmented and ornamented zygosporangial walls. Arthrospores, chlamydospores, and zygospores may be produced by some species. The zygospores lack appendaged suspenders and broad aseptate or sparsely septate hyphae are commonly found in Mucor species (Nguyen et al. 2016). When spores from sporangia are released, a remaining collarette is observed. The sporangiospores are round or slightly elongated (Larone 1995; Sutton et al. 1998; de Hoog et al. 2000). With 76 accepted species, the genus is the largest and most studied group in Mucorales (Walther et al. 2019).

Molecular identification and diversity

The present taxonomy of Mucor is mostly based on morphological characters and interfertility tests. The genus was previously diagnosed using biological species recognition and morphological species recognition (Schipper 1973; Hermet et al. 2012). However, identification often fails with only morphology hence phylogenetic species recognition has been used to resolve species (Taylor et al. 2000). The use of multi-gene (ITS, tef1 and act) phylogenetic analysis showed that Mucor is not monophyletic (Nguyen et al. 2017). An extensive study by Walther et al. (2013), using about 400 Mucor strains, led to a refinement in the classification of Mucor species. Phylogeny-based on 28S rDNA led to the transfer of some species to different groups and it was shown that some of these groups intermingled with other genera, such as Chaetocladium and Helicostylum, which do not belong to Mucoraceae. The use of five markers (ITS, rpb1, tsr1, mcm7 and cfs) phylogeny by Wagner et al. (2019), combined with phenotypic studies, mating tests and the determination of the maximum growth temperatures revealed 16 phylogenetic species of which 14 showed distinct phenotypical traits and were recognized as discrete species.

Recommended genetic markers (genus level)—LSU and SSU

Recommended genetic markers (species level)—ITS and rpb 1

Accepted number of species—There are 735 species epithets in Index Fungorum (2020), however only 76 species have DNA sequence data (Table 23) (Walther et al. 2019).

Table 23 DNA barcodes available for Mucor

References—Larone (1995), Sutton et al. (1998), de Hoog et al. (2000) (morphology), Nguyen et al. (2016, 2017), Walther et al. (2013, 2019), Wagner et al. (2019) (morphology and phylogeny).

98. Phytophthora de Bary, J. Roy. Agric. Soc. England, ser. 2 12: 240 (1876)

Background

Phytophthora is classified in the kingdom Straminipila within the diploid, alga-like Oomycetes in the Stramenopile clade of the Kingdom Chromista (Cavalier-Smith 1986; Dick 1995; Yoon et al. 2002; Wijayawardene et al. 2020). Phytophthora consists of about 130 described species with many important plant pathogens. The Oomycota are biologically different from main fungal groups within the Kingdom Fungi (Corliss 1994; Cavalier-Smith 1998). For example, their cell walls are made primarily of cellulose instead of chitin as in most fungi and they cannot synthesize β-hydroxysterols, which is vital for synthesizing hormones that regulate sexual reproduction (Hyde et al. 2014). Another important difference is that oomycetes are diploid throughout their life cycle. One similarity between Phytophthora species and Eumycotan fungi is that they both produce hyphae.

Classification—Oomycota, Peronosporales, Peronosporacae

Type species—Phytophthora infestans (Mont.) de Bary

Distribution—worldwide

Disease symptoms—blight, canker, dieback, root rots and wilt

Species can have a large impact on agriculture (e.g. Phytophthora infestans, potato late blight), arbiculture (e.g. Phytophthora ramorum, sudden oak death) and whole ecosystems (e.g. Phytophthora cinnamomi in Australia). Phytophthora species damage plants by killing the tissues and resulting necrosis can be seen in leaves, stems or roots. Some species can cause multiple symptoms on a single host, or cause different symptoms on different hosts (Jung and Blaschke 1996).

Blight: Initial symptom is the development of a “water-soaked” appearance, which progresses into brown or black irregular-shaped spots or wedge-shaped lesions. These lesions are usually not surrounded by a yellow halo (Babadoost 2004; Pande et al. 2011; Ali et al. 2017).

Canker: A dark discoloured necrotic lesion in the inner bark of a tree can be seen often on the stem or branches. However, generally, cankers are visible once the outer bark is removed. Cankers are often seen with a reddish-brown liquid that oozes through the bark (Davidson et al. 2002; Jung et al. 2018).

Dieback: Death of shoot tips, twigs and branch tips can be observed. The infection progresses towards the main stem accompanied by a loss of foliage (Kuske and Benson 1983; Akilli et al. 2013).

Decline and Death: This is a gradual process that will take place over several years. Plants fail to grow and the canopy becomes thin due to loss of foliage. Then the whole canopy or sections of the canopy may die (Marais 1980; Belisario et al. 2004; González et al. 2020).

Rot: Dark discoloured rotten tissues that are common on roots, but sometimes extend above the soil surface. However, collar rot occurs at the base of the trunk and extends just below the soil line (Jung and Blaschke 1996; Graham et al. 2011; Summerell and Liew 2020).

Wilting: This is the first above-ground symptom of root rot. Foliage becomes flaccid due to lack of water intake (Vettraino et al. 2009; Xiong et al. 2019).

Phytophthora causes disease in important agricultural and ecological plants. Phytophthora infestans was responsible for the Irish potato famine from 1845 to 1852, causing the death of over 1 million people. Phytophthora ramorum has resulted in the death of millions of coast live oak, tanoak and Japanese larch trees, thus altering the forest ecosystems in California and Oregon, USA (Goheen et al. 2002; Rizzo et al. 2002, 2005).

Hosts—Phytophthora agathidicida (commonly known as kauri dieback), which causes kauri death, is considered as one of the world’s most feared fungi (Hyde et al. 2018a). An extensive survey in previously unexplored ecosystems such as natural forests (Rea et al. 2010; Vettraino et al. 2011; Jung et al. 2011, 2017; Reeser et al. 2013), streams (Reeser et al. 2007; Bezuidenhout et al. 2010; Yang et al. 2016; Brazee et al. 2017), riparian ecosystems (Brasier et al. 2003, 2004; Hansen et al. 2012), and irrigation systems (Hong et al. 2010, 2012; Yang et al. 2014a, b) has led an exponential increase in the number of species.

Pathogen biology, disease cycle and epidemiology

Morphological based identification and diversity

Species-level classification is based on the morphological characterization of reproductive structures including the sporangium (asexual) and oospore (sexual) as well as the production of chlamydospores (Martin et al. 2012). Characteristics that are important for species classification include the diameter of the oogonium and oospore, thickness of the oospore wall, whether or not the oospore fills the oogonium, ornamentation on the oogonial wall, and mode of attachment of the antheridium (Hyde et al. 2014). Identification and classification of Phytophthora species into morphological groups based on several characteristics was initially based on the key provided by Waterhouse (1963), which was later updated by Stamps et al. (1990).

Molecular based identification and diversity

Phytophthora has been historically placed in the Pythiales with Pythium and related genera, however recent phylogenetic analysis with the large (LSU) or small (SSU) rDNA sequences or cox2 gene has indicated a closer relationship with downy mildew and white rusts (Albugo.) in the Peronosporales (Beakes and Sekimoto 2009; Thines et al. 2009). Additional multigene analyses are vital to clarify the relationship between the Peronosporales and Pythium. Early efforts focusing on the phylogenetic relationships in Phytophthora used nuclear-encoded rDNA, primarily the ITS region (Crawford et al. 1996; Cooke and Duncan 1997; Förster et al. 2000). The first comprehensive study was based on the phylogenetic study of the ITS region (Cooke et al. 2000). The study by Kroon et al. (2004) was based on analysis using two nuclear (tef1, tub2) and two mitochondrial (cox1 and nad1) genes. Subsequent phylogenetic analysis was based on sequences of seven nuclear genetic markers (60S ribosomal protein L10, tub2, enolase, heat shock protein90, large subunit rDNA, TigA gene fusion and tef1) which divided the species into 10 well-supported clades (Blair et al. 2008). The phylogenetic study by Martin et al. (2014) was based on seven nuclear and four mitochondrial genes (cox2, nad9, rps10 and secY). More recently, an extensive study of the genus by Yang et al. (2017) was based on sequences of seven nuclear genetic markers as in Blair et al. (2008).

The number of described species in Phytophthora was approximately 55 in 1999, but since then there has been a significant increase in the number of species nearly doubling the number of described species to 105 (Brasier 2007), and over 128 species (Hyde et al. 2014). Additional species have since been described, for example, P. cocois (Weir et al. 2015), P. crassamura (Scanu et al. 2015), P. attenuata, P. xheterohybrida, P. xincrassata (Jung et al. 2017) bringing the total to over 150 species (Jung et al. 2019). The phylogenetic tree constructed is presented in Fig. 39 and the accepted species are given in Table 24.

Fig. 39

Maximum likelihood of Phytophthora based on the concatenated seven nuclear genetic markers (60S Ribosomal protein L10 (60S), beta-tubulin (tub), elongation factor 1 alpha (tef1), enolase (Enl), heat shock protein 90 (hsp90), 28S ribosomal DNA (28S), and tigA gene fusion protein (TigA)). ML bootstrap support values over 60% are indicated and BYPP ≥ 0.90 are shown respectively near the nodes. The type species are in bold. Scale bar indicates number of substitutions per site. The tree was rooted with Phytopythiumvexansand Pythium undulatumas as the our group. Likelihood of the best scoring ML tree was − 114471.902046. Estimated base frequencies were as follows: A = 0.216570, C = 0.275568, G = 0.312230, T = 0.195632; substitution rates AC = 0.414835, AG = 1.176570, AT = 0.600142, CG = 0.970565, CT = 5.227735, GT = 1.000000

Table 24 DNA barcodes available for Phytophthora

Recommended genetic markers (genus level)—LSU, SSU and cox2

Recommended genetic markers (species level)—LSU, tub2 and cox2

Accepted number of species– There are 317 epithets listed in Index Fungorum (2020), however only 162 species have DNA sequence data (Table 24).

References—Waterhouse (1963), Stamps et al. (1990) (morphology); Crawford et al. (1996), Cooke and Duncan (1997), Cooke et al. (2000), Förster et al. (2000), Brasier (2007), Blair et al. (2008) (morphology and phylogeny); Hyde et al. (2014) (phylogeny and accepted species) (Fig. 40).

Fig. 40

Disease cycle of a Pythium species (redrawn from van West et al. 2003)

99. Pythium Pringsh., Jb. wiss. Bot. 1: 304 (1858)

Background

Pythium is the largest and most comprehensively studied genus in Pythiaceae sensu lato, order Peronosporales sensu lato, class Peronosporomycetes, phylum Oomycota, and kingdom Straminipila (Beakes et al. 2014). Pringsheim (1858) described the genus. However, the initial classification of Pythium has changed many times based on several studies using morphological characteristics (Uzuhashi et al. 2010). Pythium comprises of more than 230 extant species (Hyde et al. 2014), however, identification of species has always been problematic due to limited morphological characters, difficulty in isolating some taxa and lack of molecular data for certain species (Lévesque and de Cock 2004).

Classification—Oomycota, Pythiales, Pythiaceae

Type species—Pythium monospermum Pringsh. (Pringsheim 1858)

Distribution—worldwide

Disease symptoms—generally cause rot of fruit, roots and stem including pre- or post-emergence damping-off of seeds and seedlings.

Pythium causes crown and root rot in mature plants, where plants suddenly wilt during warm and sunny weather and when plants have their first heavy fruit load. Often, upper leaves of infected plants wilt in the day and recover overnight. However, plants eventually die (Craft and Nelson 1996; Postma et al. 2000). The first symptoms of Pythium root infections include stunting. In the root system, initial symptoms are brown to dark-brown lesions on root tips and feeder roots. As the disease progresses, symptoms are soft, brown, stubby roots and lack of feeder roots. In larger roots, the outer root tissue or cortex peels away, leaving the string-like vascular bundles underneath (Postma et al. 2000; Moorman et al. 2002; Al-Mahmooli et al. 2015). Pythium rot also occurs in the crown at the stem base. In cucumber, diseased crowns turn orange-brown, often with a soft rot at the base, while in strawberry seedling roots have dark brown, water-soaked rot and rotten crowns (Columbia and English 1988; Ishiguro et al. 2014). Several species of Pythium cause blight of turfgrass, which initially appears as “greasy” water-soaked areas, but later turn brown and grey (Vencelli and Powell 2008).

Several Pythium species are capable of causing fruit rot in numerous crops (Martin and Loper 1999). Pythium fruit rot is commonly known as a cottony leak or watery rot and occurs during wet weather or in poorly drained areas of fields (Ho and Abd-Elsalam 2020; Sharma et al. 2020a). Initial symptoms of the fruit rot are brownish, water-soaked lesions that quickly become large, watery, soft and rotten. The rot generally begins on the parts of fruit in contact with the soil. In cucumber, a brown to dark green blister can be seen on fruit before they become watery and rot. Later, white cottony mycelium can be seen on rotten tissues, especially during humid weather. Pythium fruit rot is most severe in poorly-drained fields during wet weather. The disease can render fruit unmarketable (Ho 2009; Sharma et al. 2020a).

Pre-emergence damping-off causes seeds and young seedlings to rot before they emerge from the growing medium in greenhouses, while post-emergence damping-off kills newly emerged seedlings. In the latter, the pathogen causes a water-soaked, soft brown lesion at the stem base, near the soil line, that pinches off the stem causing the seedling to topple over and die (Weiland et al. 2012).

Hosts—Pythium has a wide range of hosts including species of Cucurbitaceae and Poaceae, Ananas comosus, Arachis hypogaea, Brassica sp., Carica papaya, Beta vulgaris, Daucus carota subsp. sativus, Dendrobium sp, Solanum sp. and Zingiber officinale. Some species are pathogens of algae, fungi, other oomycetes, nematodes, insects, animals and humans (Van der Plaäts-Niterink1981; Czeczuga et al. 2005; Kawamura et al. 2005; Hwang et al. 2009; Li et al. 2010; Weiland et al. 2012; Ho 2013; Hyde et al 2014). Several species inhabit different soils in cultivated and uncultivated fields including forest (Uzuhashi et al 2010). Pythium arrhenomanes, P. dissotocum, P. elongatum, P. myriotylum, and P. spinosum are important pathogens of rice seedlings (Hendrix and Campbell 1973; Hsieh 1978; Ventura et al. 1981; Chun and Schneider 1998; Eberle et al. 2007; Kreye et al. 2009; Oliva et al. 2010; Banaay et al. 2012; Van Buyten and Höfte 2013). Pythium insidiosum causes pythiosis in mammals including humans (van der Plaäts-Niterink 1981; de Cock et al. 1987). Some species target below-ground plant parts and some species can cause fruit rot, however, some Pythium species can also benefit plants as endophytes by acting as biocontrol agents (Benhamou et al. 1997) and by stimulating plant growth (Martin and Loper 1999; Mazzola et al. 2002).

Pathogen biology, disease cycle and epidemiology

Pythium species grow and colonize a plant by producing hyphae which extract nutrients from the host. Once the hyphae from opposite mating types meet, they produce thick-walled oospores which serve as overwintering structures. Upon germination, an oospore may produce more hyphae, or develop a zoosporangium, which produces motile zoospores that swim to and infect plants. Zoosporangia can also germinate and directly infect plants (Ho 2009; van West et al. 2003).

Morphological based identification and diversity

Pythium has hyaline hyphae which are coenocytic without cross septa (van der Plaäts-Niterink 1981). Filamentous and globose sporangia are present, and zoospores develop in a vesicle, which is formed at the tip of a discharge tube from a sporangium. After fertilization with paragynous or hypogynous antheridia, oospores are formed in smooth or ornamented oogonia. The oospore can fill the whole organism or can have space between the walls of the oogonia and oospore. The process of zoospore formation within a vesicle is a characteristic feature of the genus, which distinguishes it from morphologically similar genera such as Phytophthora and Halophytophthora. However, the formation of zoospores is similar to Lagenidium, which features endobiotic and holocarpic features not observed in Pythium (Dick 2001). Species delimitation based on morphological characteristics such as shape and size of sporangia and oogonia is difficult as these characteristics are often shared among different species.

Molecular based identification and diversity

Lévesque and de Cock (2004) separated the genus into 11 clades (A-K) using phylogenies of ITS and 28S. Clade K, which includes P. vexans was transferred to a new genus Phytopythium with Phytopythium sindhum as type species (Bala et al. 2010), while the remaining clades can be divided into two groups: species with filamentous sporangia (clades A-D) and species with globose sporangia (clades E–J). Identification of Pythium isolates to species level is recommended based on cox1 and ITS gene regions. The use of ITS region alone cannot accurately identify all Pythium species. Several species are indistinguishable based on both ITS and cox1 sequences. Lévesque and de Cock (2004) provided the first extensive study of Pythium, accepting 116 species. Additional species have recently been described for example P. alternatum (Rahman et al. 2015), P. biforme, P. brachiatum, P. junctum, P. utonaiense (Uzuhashi et al. 2015), P. cedri (Chen et al. 2017), P. heteroogonium, P. longipapillum, P. oryzicollum (Salmaninezhad and Mostowfizadeh-Ghalamfarsa 2019). Currently, there are more than 130 accepted species in the genus (Arafa et al. 2020). The phylogenetic tree constructed is presented in Fig. 41 and the information of species are given in Table 25.

Fig. 41

Maximum likelihood of Pythium species based on the concatenated SSU, ITS, LSU, cox2 and tub2 regions. The maximum parsimonious dataset consisted of 528 constant, 71 parsimony-informative and 556 parsimony-uninformative characters. The parsimony analysis of the data matrix resulted in the maximum of ten equally most parsimonious trees with a length of 1637 steps (CI = 0.200, RI = 0.737, RC = 0.147, HI = 0.800). ML and MP bootstrap support values over 60% and BYPP ≥ 0.90 are indicated. Type strains are in bold and the 11 clades (A–K) are indicated. Scale bar indicates number of substitutions per site. The tree was rooted with Lagenidium giganteum (CBS 580.84) and Lagenidium sp. (DAOM 242348 and CBS 127283). Likelihood of the best scoring ML tree was − 28453.969593. Estimated base frequencies were as follows: A = 0.264872, C = 0.163069, G = 0.213432, T = 0.358627; substitution rates AC = 1.219282, AG = 3.062456, AT = 3.113530, CG = 0.855790, CT = 4.379562, GT = 1.000000

Table 25 DNA barcodes available for Pythium

Recommended genetic markers (generic level within Pythium sensu lato)—18S (small subunit, SSU) and 28S (large subunit, LSU) nuclear rRNA genes

Recommended genetic markers (sub-generic, inter- and intra-specific level)—The internal transcribed spacers (ITS including ITS1, 5.8S rRNA, and ITS2), cytochrome c oxidase subunit 2 (cox2)

Accepted number of species—There are 330 epithets listed in Index Fungorum (2020), however only 157 species have DNA sequence data (Table 25).

References—van der Plaäts-Niterink (1981), Dick (2001) (morphology), Lévesque and de Cock (2004), Hyde et al. (2014), Arafa et al. (2020) (phylogeny and accepted species numbers)

100. Rhizopus Ehrenb., Nova Acta Phys.-Med. Acad. Caes. Leop.-Carol. Nat. Cur. 10: 198 (1821)

Background

Rhizopus is classified in the subphylum Mucoromycotina, class Mucoromycetes, order Mucorales and family Rhizopodaceae (Wijayawardene et al, 2018, 2020). The genus is one of the most diverse and constitutes an important genus within the order Mucorales. Rhizopus species are common post-harvest pathogens of fruits, vegetables, crops and stored foods, while some Rhizopus species are human pathogens. Rhizopus arrhizus and Rhizopus microsporus can cause mucoromycosis in immunocompromised humans (Yildirim et al. 2010; Benedict and Brandt 2016). Morphology-based (size of sporangia and sporangiophores, and rhizoids) and physiology-based (growth temperature) identification and classification grouped the genus in three groups: R. microsporus, R. stolonifer, and R. arrhizus (syn: R. oryzae) (Schipper 1984). Schipper (1984) and Schipper and Stalpers (1984), provided the first significant monographs of Rhizopus. Fundamental morphological-based identification was provided which is still widely used in current taxonomic classification for Rhizopus (Schipper 1984; Schipper and Stalpers 1984; Hartanti et al. 2015). The inclusion of DNA-based phylogenetic tools has resulted in significant changes in the taxonomic classification (Vebliza et al. 2018). With the implementation of molecular-based identification, Abe et al. (2006, 2010), Zheng et al. (2007b), and Liu et al. (2007) provided significant contributions in the classification of Rhizopus. Briefly, in current taxonomy Rhizopus arrhizus is a synonym of R. oryzae, R. reflexus to R. lyococcus and Amylomyces rouxii is a synonym of Rhizopus arrhizus (Liu et al. 2007; Hyde et al. 2014; Vebliza et al. 2018) (Fig. 42).

Fig. 42

Phylogenetic tree generated by maximum likelihood analysis of combined ITS- LSU-SSU sequence data of Rhizopus, Backusella and Mucor species. Twenty-six taxa containing 2600 characters including gaps were used in the phylogenetic analysis. The tree was rooted using Backusella circina (CBS 128.70) and Mucor indicus (CBS 226.29). The best scoring RAxML tree with a final likelihood value of − 11138.113172 is presented. The matrix contained 761 distinct alignment patterns, with 41.20% of undetermined characters or gaps. Estimated base frequencies were as follows: A = 0.293528, C = 0.180262, G = 0.236482, T = 0.289728; substitution rates AC = 0.787730, AG = 2.127667, AT = 1.579251, CG = 0.715792, CT = 3.683423, GT = 1.000000; gamma distribution shape parameter α = 0.181944.ML bootstrap support values greater than 70% are shown near the nodes. The type species are in bold. Scale bar indicates the number of substitutions per site

Phylogenomic approaches have the potential to provide a clear understanding of the inter-relationships of species (Gryganskyi et al. 2018). In recent revisions, data from whole-genome sequencing have been used (Gryganskyi et al. 2018). Phylogenetic analysis based on a dataset of 192 orthologous protein-coding genes extracted from whole-genome sequencing of representative species provided a robust phylogeny and tree topology for Rhizopus. The phylogenetic analysis resulted in similar tree topology obtained from studies which utilize ITS and pyrG genes or 76 orthologous proteins from the genomes (Liu et al. 2007; Chibucos et al. 2016). In brief, R. microsporus is suggested to be a monophyletic sister clade to other Rhizopus clades, R. stolonifer was found to be sister to R. arrhizus and R. delemar and these four species are monophyletic (Gryganskyi et al. 2010, 2018).

A comparative analysis of the mating-type locus across Rhizopus revealed that its structure is flexible even between different species in the same genus, but shows similarities between Rhizopus and other mucoralean taxa. Variation of the genome size was also noted to be approximately three-fold within a species which are induced by changes in transposable element copy numbers and genome duplications (Gryganskyi et al. 2018). Bruni et al. (2019) successfully adapted the CRISPR-Cas 9 technique for inducing pyrF gene-specific mutations in two strains of R. delemar, the causative agent of mucoromycosis. This new tool is suggested to be useful in investigating the pathogenesis mechanisms of R. delemar and also generating specific mutants of Mucorales fungi.

Classification—Mucoromycota, Mucoromycotina, Mucoromycetes, Mucorales, Rhizopodaceae

Type species—Rhizopus stolonifera (Ehrenb.) Vuill. 1902

Distribution—worldwide

Disease symptoms—Rhizopus blight, Rhizopus head rot and Rhizopus soft rot

Rhizopus blight: Rhizopus blight can affect flowers, leaves, and stems. When infected, the plant shows symptoms such as soft and mushy brown rot. The rot produces white mycelia with black sporangia and the abundant mycelia projects a ‘bearded’ appearance. Spores of the fungus can be spread by water and air. The mode of infection is similar to bacterial soft rot in which enzymes secreted by the fungus causes cell deterioration of the host tissue. The fungi require high temperatures, high humidity and weakened host tissues or wounds (Hartley 1992).

Rhizopus head rot on sunflowers: Rhizopus head rot may be caused by several Rhizopus species such as Rhizopus arrhizus, R. microsporus and R. stolonifer (Markell et al. 2015). Historically, Rhizopus head rot was deemed as a minor disease. However, recent surveys have shown their severity. Initial signs of Rhizopus head rot are dark spots of different sizes on different types of wounds on the plant. Soft watery rot appears on the infected fruit which often turns dark brown and extends to the back of the flower head, sepals and peduncles as the disease progresses. The infected sunflower receptacle disintegrates and becomes soft and pulpy. Infection by Rhizopus causes the head to shrivel and dry. Morphological characteristics are mycelial strands bearing sporangiophore and sporangia which are seen as the disease advances (Markell et al. 2015; Zhou et al. 2018). These whiskers are tufts of hyphae containing numerous sporangia and generally appear around lenticels or breaks in the periderm. Sometimes hyphae may not be visible on the outside of the root but can be viewed by pulling apart the infected tissue, giving it a stringy appearance (Clark et al. 2013).

Rhizopus soft rot: Common causative agents of Rhizopus soft rot are Rhizopus stolonifer and Rhizopus oryzae. The disease is considered as one of the most common and destructive postharvest diseases in many plants such as sweet potato (Ipomoea batatas), potato (Solanum tuberosum) and tomato (Solanum lycopersicum). The most frequent mode of infection is wounds and injuries present on the plants. Studies have also shown that the type of wounding and storage time have a significant impact on the susceptibility of infection by Rhizopus species (Scruggs and Quesada-Ocampo 2016). Earliest symptoms of infections are soft water-soaked lesions. The disease spreads across the wounded area and progresses to the extremities of the substrate. Hyphae soon develop on the rotten tissues and produce grey sporangiophores which subsequently bear sporangia (Khokhar et al. 2019). Whiskers are characteristics features of Rhizopus soft rot and have been reported in the case of soft rot on sweet potatoes (Clark et al. 2013; Scruggs and Quesada-Ocampo 2016) (Table 26).

Table 26 DNA barcodes available for Rhizopus

Hosts—Wide range of hosts including species of Allium, Ananas, Brassica, Cucumis, Cucurbita, Fragaria, Lycopersicon, Phaseolus, Pisum and Solanum (Farr and Rossman 2020)

Pathogen biology, disease cycle and epidemiology

The pathogen reproduces asexually. Spores of Rhizopus species are commonly found in the air and can survive easily on crop debris, fruits, vegetables, and even on tools and equipment. Factors such as the Rhizopus species, type of fruit, stage of maturity of the plant and fruit or the storage will have a slight difference in the disease cycle. Rhizopus stolonifer, as well as the other species causing post-harvest diseases such as Rhizopus soft rot, require wound injuries, cracks or any mechanical damage for entry (Hartley 1992; Bautista-Baños et al. 2014; Scruggs and Quesada-Ocampo 2016). Infection and colonization are highly dependent on the enzymes produced by the fungi. To establish within the host, Rhizopus species produce numerous enzymes, including amylase, pectinase, and cellulase that can damage cell walls and permit host colonization (Ogundero 1988; Tang et al. 2012). This results in the softening of the host tissue; one of the symptoms of the disease (Nelson 2009; Kwon et al, 2012; Bautista-Baños et al. 2014; Feliziani and Romanazzi 2016). During initial stages of infection, Rhizopus rot appears as water-soaked areas and in the case of Rhizopus stolonifer, the rot also exudes clear leachate. In the case of Rhizopus soft rot caused by R. oryzae in banana, the symptoms and disease cycle are similar to R. stolonifer (Kwon et al. 2012). In Okinawan sweet potatoes, the disease causes a soft and moist appearance and a stringy flesh during the initial stages and as the disease progresses, the tissue of the sweet potato turns brownish and eventually black (Nelson 2009). In the case of R. stolonifer, the fungal mycelia quickly spread across the infection site. The sporangia formed are normally black and the whole plant is covered by fungal mycelia (Bautista-Baños et al. 2014). The enzymes exuded from the pathogen generally liquefy the internal tissues, for an example in sweet potato parenchyma of the root becomes liquefied, leaving the periderm and outer fibres of the root intact (Scruggs and Quesada-Ocampo 2016). The disease becomes more severe in warm, humid environments (Zoffoli and Latorre 2011). Avoidance of Rhizopus species is difficult due to their ubiquitous nature; therefore, sanitation and storing produce under unfavourable disease conditions is the key to control this pathogen.

Morphology- based identification and diversity

Rhizopus is normally distinguished by rhizoids, stolons and single or branched sporangiophores (Vebliza et al. 2018). Identification of species takes into account the growth temperature, size of sporangiophore and sporangium and the branching of rhizoids (Abe et al. 2007). The white mycelia consist of coenocytic hyphae which bear the sporangiophore with normally black sporangia. These taxa are fast-growing and form rhizoids at the base of sporangiophores. The sporangium contains a columella and spores (Bullerman 2003). During the sexual stage, there is the formation of zygospores and chlamydospores can also be seen during the growth of the fungi (Bullerman 2003; Abe et al. 2007).

Molecular identification and diversity

Traditionally, Rhizopus species were classified using morphological characters such as the shape and size of the structures (chlamydospores, rhizoids, sporangiophores and columellae) and physiological features such as optimal growth conditions. Current classification and taxonomic grouping follow that of Schipper (Schipper 1984). Schipper classified Rhizopus into three groups namely R. microsporus, R. stolonifer and R. arrhizus based on the physiological factors and morphology (Abe et al. 2010; Gryganskyi et al. 2018). Later, studies such as Abe et al. (2006), Liu et al. (2007), Zheng et al. (2007a, b), Abe et al. (2010) implemented molecular phylogeny using DNA sequence data in the classification of these fungi. With novel approaches used, the classification proposed by Schipper was found to agree with some recent studies while others divided the genus into ten species and seven varieties or eight species. Zheng et al. (2007b) used zygospore formation, and molecular systematic morphological characters, mating compatibility, physiology and molecular systematic to accept the division of the genus in ten species and seven varieties. Abe et al. (2010) also used the rDNA ITS gene region together with actin-1 and tef1, to reorganize the proposed taxonomy into eight species instead of ten species. One important data provided by this study was the problematic rDNA ITS region of R. americanus. It was discovered that R. sexualis var. americanus has three rDNA ITS gene regions which are distinct from each other. However, Liu et al. (2007) were not able to obtain all three rDNA ITS gene region instead they were able to amplify only one ITS region which was similar to that of Rhizopus oryzae. So, this led to the conclusion that R. americanus was phylogenetically different from R. sexualis.

Genetic markers (species and genus level)—ITS and rpb1

Genetic markers (higher-level phylogeny)—SSU, LSU and act

Accepted number of species—There are 152 species epithets in Index Fungorum (2020), however only 12 species have DNA sequence data (Table 25).

References—Bullerman (2003), Abe et al. (2007) (morphology); Abe et al. (2010), Gryganskyi et al. (2018), Vebliza et al. (2018) (morphology and phylogeny)

Discussion

This is the fourth in the One Stop Shop series focusing on providing a stable platform for the taxonomy of plant pathogenic fungi and fungus-like organisms. These series aim to provide updated backbone trees and information regarding plant pathogens in one place for ease of access. Databases play an important role in aggregating the scattered data into an easily accessible form and many of the pathogenic genera were annotated in the UNITE database (Nilsson et al. 2014). However, this database mainly focused on ITS region rather than the protein-coding gene regions. There are very few databases dedicated to identity the plant pathogens and related fungi-like organisms. We have been trying to provide a stable and updated taxonomy for 97 genera and three families since 2014, which are listed in Table 1. All this information is available in http://www.onestopshopfungi.org.