Isolation and characterisation of novel nematophagous fungi from eggs of the cereal cyst filipjevi and identifying their secondary metabolites with nematicidal activity

Dissertation

Zur Erlangung des Akademischen Grades eines Doktors der Agrarwissenschaften (Dr. agr.)

Eingereicht am Fachbereich Ökologische Agrarwissenschaften der Universität Kassel

Von

Samad Ashrafi

February 2018

Samad Ashrafi (2018) Isolation and characterisation of novel nematophagous fungi from eggs of the cereal cyst nematode Heterodera filipjevi and identifying their secondary metabolites with nematicidal activity

A thesis in Biological control: University of Kassel

Defence date: 17.04.2018

Place: Witzenhausen, Germany

Supervisors:

Prof. Dr. Maria R. Finckh (University of Kassel)

Dr. Wolfgang Maier (Julius Kühn-Institut — Federal Research Center for Cultivated Plants)

For Saleh

Table of Contents List of Tables ...... vii List of Figures ...... viii Chapter 1. General Introduction ...... 1 1.1 Cereal cyst ...... 1 1.2 Biocontrol of nematodes ...... 3 1.3 Nematophagous fungi ...... 4 1.3.1 Nematode-trapping fungi ...... 5 1.3.2 Endoparasitic fungi ...... 6 1.3.3 Egg-parasitic fungi ...... 7 1.4 Suppression of PPNs with nematophagous fungi ...... 8 1.5 Nematophagous fungi, a source of nematicidal metabolites ...... 11 1.6 Introduction to the project...... 12 1.7 Research rationale and objectives of the project ...... 15 1.8 Refernces ...... 17 Chapter 2. Ijuhya vitellina sp. nov., a novel source for chaetoglobosin A, is a destructive parasite of the cereal cyst nematode Heterodera filipjevi ...... 22 2.1 Introduction ...... 23 2.2 Material and methods ...... 24 2.2.1 Nematode collection and materials examined ...... 24 2.2.2 Ethics statement ...... 25 2.2.3 Fungal isolation from field collected samples ...... 25 2.2.4 Pure culture based studies ...... 26 2.2.5 Pathogenicity test and Koch’s postulate ...... 26 2.2.6 Light and scanning electron microscopy ...... 27 2.2.7 DNA extraction ...... 27 2.2.8 PCR amplification and sequencing ...... 28 2.2.9 Alignment and phylogenetic reconstruction ...... 28 2.2.10 Nomenclature ...... 37 2.2.11 Metabolite profiling...... 37 2.3 Results ...... 38 2.3.1 Fungal isolation ...... 38 2.3.2 Molecular phylogenetic studies ...... 39 2.3.3 ...... 42 2.3.4 In vitro parasitism on nematode eggs and Koch’s postulates ...... 45 iv

2.3.5 Metabolite profiling...... 48 2.4 Discussion ...... 49 2.4.1 Phylogenetic analyses and systematic implications ...... 49 2.4.2 Parasitism on nematode cysts and eggs ...... 50 2.4.3 Metabolite profiling...... 51 2.5 Appendix ...... 53 2.6 References ...... 54 2.7 Supporting information ...... 60 Chapter 3. Monocillium gamsii sp. nov. and Monocillium bulbillosum: two nematode-associated fungi parasitising the eggs of Heterodera filipjevi ...... 67 3.1 Introduction ...... 68 3.2 Material and methods ...... 69 3.2.1 Sample collection and materials examined ...... 69 3.2.2 Cultural studies ...... 70 3.2.3 Pathogenicity tests against H. filipjevi ...... 70 3.2.4 Microscopy ...... 71 3.2.5 Molecular phylogenetic studies ...... 71 3.3 Results ...... 74 3.3.1 Sample collection and fungal isolation ...... 74 3.3.2 Sequence comparison and phylogenetic inference ...... 75 3.3.3 Taxonomy ...... 76 3.3.4 Development of M. gamsii in nematode eggs in vitro ...... 79 3.3.5 Parasitism of M. bulbillosum towards H. filipjevi ...... 81 3.4 Discussion ...... 81 3.5 References ...... 84 Chapter 4. Inhabiting plant roots, nematodes and — Polyphilus nom. prov., a new helotialean with two globally distributed ...... 87 4.1 Introduction ...... 88 4.2 Material and methods ...... 90 4.2.1 Origin of isolates ...... 90 4.2.2 Fungal isolation from nematode eggs ...... 90 4.2.3 Sporulation and morphology of isolates ...... 90 4.2.4 DNA extraction and amplification ...... 92 4.2.5 Phylogenetic analyses ...... 93 4.2.6 Pathogenicity against H. filipjevi ...... 93

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4.2.7 Light microscopy ...... 94 4.3 Results ...... 94 4.3.1 Sporulation and colony morphology ...... 94 4.3.2 Molecular phylogeny ...... 94 4.3.3 Isolation of fungi from H. filipjevi ...... 97 4.3.4 Pathogenicity against H. filipjevi ...... 98 4.3.5 Taxonomy ...... 98 4.4 Discussion ...... 101 4.5 References ...... 105 4.6 Supplementary ...... 109 Chapter 5. General Discussion ...... 110 5.1 Introduction ...... 110 5.2 Phylogenetic relationships of the isolated fungi ...... 112 5.3 Biology- Parasitism ...... 113 5.4 Nematophagous fungi – a treasure trove for nematicidal compounds ...... 116 5.5 References ...... 119 Summary ...... 121 Zusammenfassung ...... 123 Acknowledgements ...... 125 Eidesstattliche Erklärung ...... 127 Affidavit ...... 127

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List of Tables

Table 1.1- A taxonomic overview of the main groups of nematophagous fungi, and their modes of action ...... 10

Table 2.1- Isolates and accession numbers used in the phylogenetic analyses ...... 30

Table 3.1- Isolates and accession numbers used in the phylogenetic analyses ...... 73

Table 4.1- Culture collection numbers, collection sites, hosts, GenBank accession numbers (ITS, LSU, SSU, RPB2) and other details of isolates included in this study ...... 91

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List of Figures

Figure 1.1- life cycle of a cyst-forming nematode. A) second stage juveniles (J2s) hatch from eggs inside the cysts, leave the cysts to invade a new host, B) a J2 moving through the host root to find and establish a feeding site where it feeds and develop, C) development of J2s into either swollen females or vermiform males, D) adult males leave the roots to mate, while the females continue feeding, rupturing the root cortex, E,F) gravid females containing eggs die later to become cysts...... 4 Figure 1.2- The research strategy followed in this project...... 13 Figure 1.3- A schematic outline displaying the process of the Focused screening...... 15

Figure 2.1- Cysts and eggs of Heterodera filipjevi naturally infected with Ijuhya vitellina, and pure cultures obtained from the infected eggs. (A) Symptomatic, reddish dotted nematode cysts. (B, C) Nematode eggs accommodating reddish microsclerotia. (D, E) Microsclerotial tissue developing inside juveniles. (F) A six-month-old culture that developed from a single infected nematode egg. (G) Surface of colony showing reddish microsclerotia arranged in concentric rings. (H, I) Two-month-old cultures on PDA and CMA...... 39 Figure 2.2- Bayesian inference of phylogenetic relationships of selected taxa of the and () based on LSU sequences. Numbers above nodes are estimates of a posteriori probabilities (≥ 0.9) / NJB and MLB values (≥70%). The topology was rooted with placenta, Balansia henningsiana, B. pilulaeformis, and Hypocrella nectrioides, (Hypocreales)...... 41 Figure 2.3- Bayesian inference of infrageneric phylogenetic relationships within Ijuhya based on act, ITS, LSU, rpb1, and ß-tub sequences. Numbers above nodes are estimates of a posteriori probabilities (≥ 0.9) / NJB and MLB (≥ 70%). The topology was rooted with three distantly related ‘Ijuhya’ species (‘Ijuhya’ antillana, I. dentifera, and ‘Ijuhya’ oenanthicola)...... 42 Figure 2.4- Light micrographs of Ijuhya vitellina, formation of microsclerotia. (A-F) Transformation of hyphae into (A-D) chlamydospore or dictyochlamydospore-like structures, and (E, F) microsclerotia. (G-I) Coiling or coalescence of dictyochlamydospore-like structures. (J) Microsclerotia densely arranged in a chain. (K-N) Pigmentation first observed (K) in cell walls, and later (L-M) intensifying throughout microsclerotia. (O) A single microsclerotium inoculated on agar surface developing hyphae...... 44 Figure 2.5- SEM micrographs of microsclerotia formed by Ijuhya vitellina. (A) Filamentous hyphae developing into multicellular structures. (B) Intercalary formed dictyochlamydospores connected by hyphae (arrowed). (C) Detail of intercalary multicellular structures of microsclerotia. (D, E) Hyphae transformed into chlamydospore-like structures and microsclerotia. (F) Terminally formed microsclerotium. (G) Moniliform arrangement of microsclerotia. (H) Detail of microsclerotia illustrating a multicellular surface that forms a textura angularis...... 45 Figure 2.6- Light micrographs of the infection and colonisation process of Ijuhya vitellina in cysts and eggs of Heterodera filipjevi. (A) Symptomatic cyst, reddish-dotted due to eggs containing reddish, globose microsclerotia. (B-E) Early colonisation of nematode eggs by hyphae becoming chlamydospore- and dictychlamydospore-like to develop microsclerotia inside eggs. (F, G) Dictyochlamydospore-like structures and small microsclerotia. (H) Hyphae penetrating through the eggshell by forming appressorium-like structure (arrows). (I-K) Development of the inside nematodes eggs: (I) Formation of thick-walled hyphal cells, later (J-K) transforming into microsclerotia. The arrow in (J) points at the nematode stylet; in (K) at immature microsclerotium. (L) Egg with mature

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microsclerotium. (M-N) Near-identical cells of microsclerotium formed in (M) egg and (N) pure culture, forming a textura angulari in optical sections...... 47 Figure 2.7- Structures of chaetoglobosin A (1) and 19-O-acetylchaetoglobosin A (2)...... 48

Figure 3.1- Naturally infested cysts and eggs of Heterodera filipjevi with Monocillium gamsii, and pure cultures obtained from infected eggs. A) Field collected symptomatic cysts bearing parasitised eggs. B–E) Nematode eggs infected by M. gamsii: B, D) Nematode eggs containing microsclerotia of M. gamsii; E) An embryonated egg containing a second stage juvenile (J2) parasitised by M. gamsii (arrow points at nematode’s stylet). F) A nematode egg containing microsclerotia, and hyphae growing out of it. G–H) colony of M. gamsii grown on PDA: G) colonies developing from three individually plated infected eggs; H) A 25-d-old culture grown at 25 °C in the dark. I) The surface of a five-month-old culture detailing the sclerotioid masses covering the colony surface. Single microsclerotia can be seen as little black dots at the margin of the culture...... 74 Figure 3.2- Bayesian inference of phylogenetic relationships using four strains of the here described nematode parasite and all present in GenBank based on an alignment of ITS, LSU, rpb1, and tef sequences using GTR+I+G as nucleotide substitution model. Depicted is a 50% majority rule consensus tree derived from 3500 trees from the stationary phase of a Monte Carlo Markov Chain. A posteriori probability (BIpp) values greater than 0.95, and bootstrap values of neighbor-joining (NJBT) and maximum likelihood (MLBT) analyses greater than 0.7 are given above branches (BIpp/NJBT/MLBT). Two representatives of the Bionectriaceae, byssicola and Ijuhya vitellina, were used to root the tree...... 76 Figure 3.3- Micrographs of Monocillium gamsii. A–C) Hyphal growth by intercalary development of chlamydospore and dityochlamydospore-like structures filled with guttules. D–H) initiation of microsclerotia by interweaving or coiling of dictyochlamydospores, and growth to full size. I) Highly pigmented sclerotium at maturity displaying a textura angularis on surface view. J–N) Setae, phialides and conidia. M–O) Formation of phialides on coiling hyphal. P) Conidial heads, conidia cohering in wet heads. Q, R) SEM: Q) Phialides from with conidial heads, arising from hyphae, R) coiling hyphae (arrows), and detail of phialides bearing conidia...... 78 Figure 3.4- Cysts and eggs of Heterodera filipjevi infected by Monocillium gamsii exhibiting colonisation in vitro. A, B) infected cysts rendered black-dotted due to fungal-colonised eggs containing microsclerotia. C, D) Eggs with mature sclerotia, extracted from symptomatic cysts. E, F) Individual hyphae penetrating eggshell (arrows indicate the individual hyphae; V indicates vacuole-like structures). G–M) Fungal development inside the eggs: G, H) Earlier stages of infection in unembryonated eggs; I, J) Fungal development in the body cavity of developing juveniles where enlarged, thick-walled cells are formed and coalesce to initiate microsclerotia formation; K–M) Microsclerotia developing to full size and pigmentation. N–P) Pigmentation in microsclerotia from pale-olivaceous to darkly brown...... 80 Figure 3.5- Cysts and eggs of Heterodera filipjevi infected by Monocillium bulbillosum in vitro. A) Symptomatic cysts infected by M. bulbillosum. B) Infected egg showing early stage of fungal colonisation. C–F) Formation of microsclerotia in nematode eggs ...... 81

Figure 4.1- Colonies of Polyphilus nom. prov. species on modified Melin-Norkrans agar (MMN) A. Polyphilus sieberi nom. prov. (REF052 = DSM 106515). B. Polyphilus frankenii nom. prov. (REF052 = DSM 106520)...... 95

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Figure 4.2- Maximum Likelihood (RAXML) tree of concatenated ITS, LSU, SSU and RPB2 sequences of representative isolates of Polyphilus nom. prov. species and related fungi (best blast matches, and representative taxa of related lineages) from s. l. (). Bayesian posterior probabilities (≥ 0.90) are shown above branches and before slashes; RAXML bootstrap support values (≥ 70) are shown below branches and after slashes. Ex-type strains are shown in bold. Phialocephala fortinii (CBS 443.86) served as an outgroup. The scale bar indicates expected changes per site per branch...... 96 Figure 4.3- Maximum likelihood (RAXML) tree based on ITS sequences of representatives of Polyphilus nom. prov. species and similar sequences from GenBank. Sequences obtained in this study are shown in bold. After the accession number and sequence name, the host, isolation source and geographic origin of each sequence is shown in brackets. Bayesian posterior probabilities (≥ 0.90) are shown above branches and RAXML bootstrap support values (≥ 70) are shown below branches. Roseodiscus subcarneus (voucher D. Haelew 314a, KT92714) was used as outgroup. Abbreviations: uncultured (u.), clone (c.), isolate (is.), strain (st.), voucher (v.), surface sterilized (s. s.) ectomycorrhizal (EcM.). Sequences of uncultured fungi are marked by diamonds ()...... 97 Figure 4.4- Pathogenicity of Polyphilus sieberi nom. prov. towards Heterodera filipjevi. A–C. Fungus- infected eggs originally isolated from field-collected symptomatic cysts, detailing hyphal cells. D. Symptomatic cyst obtained from in vitro tests, containing blackish eggs infected with the fungus. E. Runner hyphae of P. sieberi nom. prov. growing in cyst mucilage. F–H. In vitro infection process of P. sieberi nom. prov. in nematode eggs: hyphae forming enlarged, thick-walled and lobate cells containing oil-like bodies. Note the pigmentation process during the early (F, G) and late (H) stages of colonization. I, J. Fungal colonization observed in slide cultures: (I) runner hyphae that penetrated the nematode egg enlarged and became filled with distinct oil-like bodies; (J) penetration of hyphae through the body cuticle and the mouth opening of a nematode juvenile, detailing the formation of enlarged hyphal cells inside the body cavity (J)...... 100

Figure 5.1- A schematice outline of results obtained during the focused screening phase of the project...... 111 Figure 5.2- Secondary metabolites from an undescribed pleosporalean fungus...... 118

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General Introduction

Chapter 1.

General Introduction

1.1 Cereal cyst nematodes

Nematodes are one of the largest groups of Metazoa occurring in freshwater, marine and terrestrial habitats. They display a great diversity in lifestyle and are either free-living or parasites. Parasitic nematodes have a wide range of hosts including plants, invertebrates, and higher including humans (Decraemer and Hunt 2013). They interact with their hosts directly as a parasite or indirectly as a vector (Thorne 1961). About 15% of the already described nematode species are plant parasitic (Decraemer and Hunt 2013; Holterman et al. 2017). Plant parasitic nematodes (PPNs) are terrestrial, widely distributed, obligate parasites attacking below and above-ground plant tissues. They represent diverse phylogenetic lineages (Holterman et al. 2017), and generally “are associated with poor growth and yield decline” (Stirling 2014). While some species such as the foliar nematode besseyi, the stem nematodes Ditylenchus spp., and the wheat seed gall nematode tritici feed on the aerial parts of plants, species such as the root-knot nematodes (Meloidogyne spp.), and the cyst-forming nematodes (e.g. Globodera and Heterodera spp.) feed exclusively in plant roots. The latter species are known to be among the most damaging and economically important PPNs (Jones et al. 2013). On the basis of their feeding mechanisms, PPNs can be categorised in three groups: ectoparasites, semi-endoparasites, and migratory endoparasites (Decraemer and Hunt 2013). The ectoparasitic nematodes are generally migratory, remain in the soil outside of the root tissues, and feed externally. Species of the genera Xiphinema and Trichodorus are some examples of this group. In semi- endoparasitic nematodes, only the anterior part of the nematode remains inside the root tissue, while the posterior part remains in soil. This life style is realised for example by Rotylenchulus species. Endoparasitic nematodes are either migratory or sedentary. Migratory endoparasites penetrate the plant roots, move and feed along the root tissue. This group includes several genera such as Anguina, Ditylenchus, and Pratylenchus. In sedentary endoparasitic nematodes, females become sedentary establishing specialised feeding sites. Cyst-forming and root-knot nematodes are examples of this group (Decraemer and Hunt 2013; Moens and Perry 2009; Perry and Moens 2011). The life cycle of PPNs typically consists of an egg stage and four juvenile stages developing into adult female and male via a series of moulting (Decraemer and Hunt 2013). Cyst nematodes are highly specialised biotrophs inducing a long-term interaction with their host plants (Turner and Subbotin 2013). They incite a specific and complex feeding structure so-called syncytium (a large and multinucleate feeding site formed by cell wall dissolution and protoplast fusion of 1

General Introduction neighbouring plant cells) in the root tissues of their host plants (Sobczak and Golinowski 2011). Their endoparasitic lifestyle is illustrated in Fig 1.1: The infective second-stage juveniles (J2s) hatch from the eggs inside a cyst and leave the cyst via natural openings (the mouth and vulval slit of the now encysted female nematode). They penetrate roots of the host plant at the elongation zone behind the growing tip using their stylet, migrate intercellularly towards the vascular cylinder, and begin feeding by establishing the syncytial cells. The juveniles undergo three moults to develop into mature males or females; the adult males leave the host root, while the females continue feeding and swell up, eventually causing the root cortex to rupture. After mating, eggs inside the female nematode develop into J2s; the non-motile and swollen female dies and its cuticle thickens and tans to form a cyst (Subbotin et al. 2010a). According to the definition given by Luc et al. (1986) a ‘heteroderid-cyst’ is a protective structure containing the nematode eggs, and ‘is derived from some or all components of the mature female body wall’, and is formed upon the death of the female. The genus Heterodera Schmidt is the type genus of the cyst-forming nematodes. It was proposed by Schmidt (1871) to accommodate Heterodera schachtii, which had been reported by Schacht (1859) from sugar beets in Germany and was the first observation of a cyst nematode parasitising its host plant. Heterodera consists of seven taxonomic groups, i.e. Afenestrata, Avenae, Cyperi, Goettingiana, Humuli, Sacchari and Schachtii, among which the members of Avenae group are known as cereal cyst nematodes (CCNs) (Subbotin et al. 2010b; Turner and Subbotin 2013). The Avenae group currently contains 10 species including the most commonly recorded and widely distributed species H. avenae, H. filipjevi and H. latipons found in most cereal-growing regions (Nicol and Rivoal 2008; Subbotin et al. 2010b; Subbotin et al. 2013). CCN infestations are a major factor responsible for yield-reduction with high economic impact in cereal producing regions (Jones et al. 2013; Nicol et al. 2011). They have especially damaging effects when combined with abiotic stresses. Therefore, the impact of CCNs can be especially severe in semi-arid regions where drought stress often occurs and can lead to significant yield reductions (Jones et al. 2013; Nicol et al. 2011). The first CCN parasitising cereals was reported in Germany by Kühn (1874), and was later described as H. avenae (Wollenweber 1924). In 1969, another CCN, H. latipons, with a Mediterranean distribution was observed on the roots of wheat and barley plants and described by Franklin (1969). During the decades of 1940s-70s, a cyst nematode species primarily identified as H. avenae was frequently reported to infect cereals in Eastern and (Subbotin et al. 2010b). It was similar to H. avenae but had some morphological differences, based on which it was described as H. filipjevi (Madzhidov 1981; Subbotin et al. 2010b). Heterodera filipjevi is prevalent in Europe, Asia and the USA (Smiley et al. 2017; Subbotin et al. 2010b; Toumi et al. 2017), and has a wide distribution in continental climates and semiarid regions (Nicol et al. 2011; Toumi et al. 2017). Despite the fact that mixed populations of CCNs are often reported in intensive cereal growing systems, certain species seem to be dominant

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General Introduction regionally leading to distinct bio-geographical patterns. In Turkey for example, H. filipjevi is the dominant CCN species in wheat fields in the Central Anatolian Plateau region (Abidou et al. 2005; Toktay et al. 2015).

1.2 Biocontrol of nematodes

The strategies to control PPNs have been reviewed comprehensively and discussed by several authors (Dababat et al. 2015; Lilley et al. 2011; Nicol and Rivoal 2008; Sikora et al. 2005; Smiley et al. 2017; Starr et al. 2002; Toumi et al. 2017). A wide variety of different approaches are applied to keep nematode populations under certain threshold levels and thus minimize their impact on cereals production. Environmentally friendly approaches include good agricultural practices, breeding and genetic resistance, and biological control methods principally focussing on regulation of pest populations by aid of ‘natural enemies’ and ‘living organisms’ (Kwenti 2017; Viaene et al. 2013). Biological control of a microorganism can be defined as: “The use of living organisms to suppress the population density or impact of a specific pest organism, making it less abundant or less damaging than it would otherwise be” (Eilenberg 2006), or as “any activity of one species that reduces the adverse effect of another” (Alston 2011). In this context, nematophagous fungi- a phylogenetically diverse group of fungi that are capable of killing nematodes- were among the first microorganisms recognised as parasites of PPNs. Julius Kühn (1877) was the first to discover a fungal species (Tarichium auxiliare Kühn) parasitising females of H. schachtii in Germany, also recognising the important role that such fungi might play in keeping the nematode numbers in check and thus helping the farmers. Today, this species is known as Catenaria auxiliaris (Kühn) Tribe, and it belongs to the fungal phylum of (Porter et al. 2011). Since then a taxonomically diverse group of fungi has been reported in association with cyst nematodes (Barron 1977; Bursnall and Tribe 1974; Kerry and Crump 1980; Szabó et al. 2012; Zhang et al. 2014). The earlier studies of nematode-associated fungi were comprehensively reviewed by Tribe (1977b) who also cited “the first parasite of a nematode ( anguillulae)” discovered by Lohde (1874). Goffart (1932) reported the first observation on the infection of the ‘oat cyst nematode’ H. avenae by an fungus currently named as Ilyonectria destructans. In the 1970-80s the parasitism of several fungal species such as C. auxiliaris, Nematophthora gynophila, Pochonia chlamydosporia, and P. rubescens towards the cereal cyst nematode H. avenae received much attention and was studied in greater detail (Kerry 1975; Kerry and Crump 1980; Kerry et al. 1982b; Willcox and Tribe 1974).

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General Introduction

Figure 1.1- life cycle of a cyst-forming nematode. A) second stage juveniles (J2s) hatch from eggs inside the cysts, leave the cysts to invade a new host, B) a J2 moving through the host root to find and establish a feeding site where it feeds and develop, C) development of J2s into either swollen females or vermiform males, D) adult males leave the roots to mate, while the females continue feeding, rupturing the root cortex, E,F) gravid females containing eggs die later to become cysts. Photos: Samad Ashrafi

1.3 Nematophagous fungi

By definition, nematophagous fungi “are microfungi that can capture, kill and digest nematodes. They use special mycelial structures, the so-called traps, or to trap vermiform nematodes or hyphal tips to attack nematode eggs and cysts before penetration of the nematode cuticle, invasion and digestion” (Nordbring-Hertz et al. 2011). They are phylogenetically diverse (Lopez-Llorca et al. 2008) and are found throughout the of fungi including , , Blastocladiomycota, and zygomycete fungi (Lopez-Llorca et al. 2008; Stirling 2014). In addition, there are also -like organisms belonging to the Oomycota (Stramenopiles) (Barron 1977; Lopez-Llorca et al. 2008; Stirling 2014) (Table 1). Nematophagous fungi attack different developmental stages of nematodes. Based on their infection mechanisms, they can be divided into

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General Introduction three groups: nematode-trapping fungi, endoparasitic fungi, and egg- and female parasitic fungi (Li et al. 2015; Nordbring-Hertz et al. 2011). Species belonging to the first two groups produce specialised morphological structures i.e. trapping organs or spores to capture and parasitise nematodes, while fungi belonging to the latter group attack and parasitise nematodes using hyphal tips or specialised infection structures known as appressoria (Jansson and Lopez-Llorca 2001; Nordbring-Hertz et al. 2011). All these morphological adaptations demonstrate a wide range of infection structures that, along with other biochemical and enzymatic mechanisms, enable nematophagous fungi to infect their hosts. The infection process mainly includes recognition, adhesion, penetration and colonisation eventually resulting in the digestion of nematodes. Recognition is a process of cell-cell communication, in which a series of physical, and enzymatic actions are involved (Li et al. 2015). Adhesive proteins such as the carbohydrate-binding proteins i.e. lectins, accumulated on fungal structures have also been reported to mediate recognition of and adhesion to the nematode surface or nematode eggshells (Li et al. 2015; Tunlid and Ahrén 2011). , lipids, and proteins e.g. collagens, are the main structural components of nematode cuticles and eggshells (Bird and McClure 1976; Perry and Trett 1986). It is assumed, that exoenzymes such as chitinases, lipases, protease, and collagenases can be secreted by nematophagous fungi to break down these distinct physical barriers in nematodes, and degrade the constituents of the cell wall resulting in nematode colonisation and digestion (Jansson and Friman 1999; Morton et al. 2004; Tunlid and Ahrén 2011). Additionally, ultrastructural studies in several nematophagous species e.g. the egg-parasites P. chlamydosporia and oviparasitica, or the endoparasite Drechmeria coniospora showed that formation of infection structures prior to penetration could facilitate fungal penetration and colonisation of nematode hosts (Dijksterhuis et al. 1990; Lopez-Llorca et al. 2002). While most of the nematophagous fungi can also survive saprotrophically, some species such as the egg parasitic fungi N. gynophila and C. auxiliaries, or the endoparasitic fungus Drechmeria coniospora are obligate parasites (Lopez-Llorca et al. 2006).

1.3.1 Nematode-trapping fungi

The nematode-trapping fungi occur in all major taxonomic groups of fungi i.e. ascomycetes, basidiomycetes and zygomycetes (Table 1), and are commonly found in soil (Nordbring-Hertz et al. 2011). According to a recent generic concept proposed by Baral et al. (2017) the majority of ascomycetous nematode-trapping fungi are placed in sensu lato, a monophyletic group within the including the genera Arthrobotrys, , Drechslerella, and Gamssylella. Nematode-trapping fungi share the ability of forming different specialised trapping organs with either adhesive or non-adhesive mycelial structures. These structures are diverse in Arthrobotrys s.l. species and include adhesive branches or networks, adhesive knobs, constricting or non-constricting rings, and 5

General Introduction three-dimensional networks. Adhesive structures are covered by a layer of adhesive polymers, which mediate attachment to the nematode cuticle (Dijksterhuis et al. 1994; Tunlid and Ahrén 2011). The constricting rings capture the nematodes mechanically by swelling the rings. Trapping structures have been used as the main criteria for genus delimitation of the asexual morph of the orbiliaceous nematode-trapping fungi. On the basis of differences in morphology of these structures Scholler et al. (1999) proposed a classification scheme and divided these Orbiliaceous fungi in four genera: Arthrobotrys (characterised by forming adhesive network), Drechslerella (by constricting rings), Dactylellina (by stalked adhesive knobs) and (by adhesive columns and unstalked knobs). A phylogenetic analysis based on genomic DNA sequences suggested that the trapping organs have evolved along two lineages, one with constricting rings and the other with adhesive structures, representing two different modes of trapping (Yang et al. 2007). Baral et al. (2017) however, reported that the capability of the formation of trapping structures in Arthrobotrys s.l. was not reproducible in pure cultures in their experiments suggesting a classification scheme based on ‘characters of sexual morphs’, molecular data, and ‘to a certain extent, on conidia morphology’. Morphogenesis in nematode-trapping fungi has been comprehensively studied demonstrating that initiation of traps in these fungi requires nematodes as external stimuli (Li et al. 2015; Nordbring-Hertz 2004; Nordbring-Hertz et al. 2011; Tunlid and Ahrén 2011). In addition to physical contact between nematode and the trapping-fungus, specific peptides e.g. nemin or some other substances such as ascaroides secreted by nematodes have also been reported to trigger trap formation (Dijksterhuis et al. 1994; Tunlid and Ahrén 2011). It can be concluded that trapping structures in Arthrobotrys s.l. species are morphological adaptations in response to the presence of nematodes, which can induce the predacious life style of these fungi (Nordbring-Hertz et al. 2011). In the absence of nematodes, this group of fungi grows saprophytically and does not develop traps spontaneously (Li et al. 2015; Nordbring-Hertz et al. 2011). In addition to Arthrobotrys s.l., few nematode-trapping fungi occur also in other fungal genera such as Nematoctonus (, Basidiomycota), and and Cystopage (, Zoopagomycota) whose trapping organs seem to be formed spontaneously (Nordbring-Hertz et al. 2011; Stirling 2014).

1.3.2 Endoparasitic fungi

Endoparasitic fungi parasitise the vermiform nematodes using their spores or conidia, which can either adhere to the host cuticle, are swallowed by the nematode, or injected into the nematode body cavity (Lopez-Llorca et al. 2006). Some examples of this group are given in Table 1. 1. Among the ascomycetous endoparasitic fungi, Drechmeria coniospora (, Hypocreales) is one of the most studied species. It produces adhesive knobs on the distal end of its conidia, by which it can attach to the nematode body cuticle and initiate nematode infection (Dijksterhuis et al. 1990). Similar to D. 6

General Introduction coniospora, Hirsutella rhossiliensis and H. minnesotensis (, Hypocreales) also produce adhesive conidia that adhere to and penetrate the nematode cuticle. In all the abovementioned cases, fungal hyphae first colonise the entire nematode body cavity, then grow out of the body cavity and sporulate (Chen et al. 2000; Shu et al. 2015; Stirling 2014). Other examples of endoparasitic fungi are species of the ascomycetous fungus Harposporium (Clavicipitaceae). These fungi produce narrowly arcuate or helicoid conidia, which can be ingested by nematodes, lodge in the digestive system of nematode, germinate, and colonise the host (Stirling 2014). Because of their narrow lumen stylet, PPNs cannot ingest the conidia of Harposporium spp. and thus cannot be infected by this group of fungi (Chen et al. 2000; Stirling 2014).

1.3.3 Egg-parasitic fungi

Egg-parasitic fungi attack the eggs and saccate females of sedentary nematodes and infect their host generally through either the development of appressoria or via simple penetration by hyphal tips (Lopez-Llorca et al. 2002; Morgan-Jones et al. 1983; 1984). Depending on the species, egg-parasitic fungi apply different modes of action to infect their nematode hosts. In general, infection initiates by hyphae coiling around or tightly adhering to the eggshells of nematode eggs. Following attachment, the hyphae penetrate into the nematode egg and the body cavity of developing juveniles (J2s), and colonise the host. Adhesion, penetration and degradation of host content may involve some mechanical, biochemical and enzymatic process (Li et al. 2015; Lopez-Llorca et al. 2008). The most studied species among others are P. chlamydosporia, Metapochonia rubescens, Metarhizium spp., (Clavicipitaceae), and Purpureocillium lilacinum (Ophiocordycipitaceae). Most representatives of egg-parasitic fungi are placed in the family Clavicipitaceae. Members of the Clavicipitaceae are well-known as plant and invertebrate pathogens (Hywel-Jones 2002). Within the Clavicipitaceae several verticillium-like anamorphs of fungal parasites of plants, fungi, insect and nematodes were formerly placed in the genus Verticillium Nees. Molecular and morphological studies by Zare et al. (2001) demonstrated that this heterogeneous group can be subdivide in four clades. Clade A comprised plant-associated species, Clade B contained entomogenous and fungicolous species, Clade C encompassed endoparasitic fungi, and clade D contained the egg-parasitic species mainly placed in the newly erected genus Pochonia (Zare et al. 2001). There are also several instances of nematode-egg parasitic species in other ascomycetous fungal families. The genus Trichoderma (, Hypocreales) comprises several species attacking eggs and females of cyst and root- knot nematodes (Szabó et al. 2012; Zhang et al. 2014). In the Bionectriaceae, Clonostachys rosea and murorum were also reported to be able to parasitise cyst nematodes (Chen and Chen 2002; Crump and Flynn 1995; Kerry et al. 1982a).

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General Introduction

The zoosporic fungus Catenaria auxiliaris (Catenariaceae, Blastocladiales) was the first parasite of cyst- nematodes to be reported (Kühn 1877; Tribe 1977a). A second Catenaria species, C. anguillulae was also reported to attack sedentary endoparasitic nematodes (Singh et al. 2013). Catenaria anguillulae is a saprotrophic species and weak parasite of nematodes while, C. auxiliaris is an obligate nematode parasite, which mainly associates with cyst-forming nematodes and destroys the young females (Kerry 1980; Stirling 2014; Tribe 1977a). It was also reported to parasitise other sedentary endoparasitic nematodes such as the reniform nematode Rotylenchulus reniformis (Castillo and Lawrence 2011). As reported by Tribe (1977a) and Wyss et al. (1992), the rhizomycelia (vegetative thallus) of C. auxiliaris and C. anguillulae containing swollen cells develop inside the nematode body. At maturity the swollen cells form “precursor sporangia”, which develop into a zoosporium or a resting sporangium. At a later developmental stage, uniflagellate zoospores form, and are released inside the female body. Under favourable conditions i.e. presence of moisture, these zoospores are able to swim and potentially infect other host substrates. Upon attachment to the nematode cuticle or eggshell, the fungal zoospores withdraw their flagellum, germinate inside the nematode, and grow to develop new sporangia(Wyss et al. 1992). Other zoosporic fungus parasitising eggs of cyst nematodes especially those of CCNs is the oomycete species Nematophthora gynophila (Leptolegniellaceae, Leptomitales) (Dackman and Nordbring-Hertz 1985; Kerry and Crump 1980). Movement of these zoosporic species i.e. C. anguillulae, C. auxiliaris, and N. gynophila highly depends on moisture, which can be a limiting factor especially in temperate or arid regions where crop damage by cyst nematodes is often severe.

1.4 Suppression of PPNs with nematophagous fungi

Nematophagous fungi were reported to reduce nematode populations in intensive cropping systems, and are considered as key players inducing or improving soil suppressiveness to PPNs (Stirling 2014). The nematode suppression caused specifically by nematode antagonists is referred to as “specific suppressiveness” (Stirling 2014; Westphal 2005). Nematode-suppressive soils were shown to harbour various nematode-antagonistic organisms resulting in reduction of population densities of PPNs (Kerry 1988; Stirling 2014; Westphal and Becker 2001). Long-term surveys conducted in past decades demonstrated that a high abundance of egg-parasitic fungi was often found in association with low nematode population density. These soils with ‘suppressive properties’ were shown to keep the equilibrium nematode populations below an economically damaging threshold (Chen and Chen 2002; Kerry 1980; Kerry 1975; Kerry and Crump 1977; Kerry et al. 1982b). Kerry and Crump (1998) showed that the natural antagonistic potential against H. avenae correlated with the egg-parasitic fungi P. chlamydosporia and N. gynophila in “cereal growing soils” was suppressed when soils were treated with formalin. The application of formalin reduced the densities of these fungi and subsequently

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General Introduction resulted in significant increase of cyst nematode populations. If soils were not longer treated with formalin, the densities of fungal populations increased to similar levels as before the treatment, and the density of H. avenae declined to similar level as in untreated soils, accordingly (Kerry and Crump 1998). In conclusion, several decades of work strongly suggest that the persistence of natural enemies of nematodes in soil can induce natural nematode-suppression resulting in the failure of CCNs to increase their populations. The microbial communities of these types of soils can therefore be explored for microorganisms with especially high biocontrol potential towards PPNs.

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General Introduction

Table 1.1- A taxonomic overview of the main groups of nematophagous fungi, and their modes of action

Fungal class Nematoode-tapping fungi Endoparasitic fungi Egg- and female parasitic fungi

Arthrobotrys Harposporium Pochonia (adhesive networks) (ingested conidia) (appressorium)

A. oligospora H. anguillulae P. chlamydosporia

A. conoides H. cerberi P. rubescens A. musiformis A. superba Drechmeria coniospora Purpureocillium (adhesive conidia) P. lilacinum Dactylellina (appressorium) Ascomycota (adhesive knobs / non Haptocillium constricting rings) (adhesive conidia) Lecanicillium D. haptotyla H. balanoides (appressorium) L. psalliotae Drechslerella Hirsutella L. lecanii (constricting rings) (adhesive spores) D. stenobrocha H. rhossiliensis H. minnesotensis Gamsylella (adhesive branches or unstalked knobs)

Nematoctonus Nematoctonus Basidiomycota (adhesive hourglass-shaped (adhesive spores) -

knobs) N. concurrens N. concurrens N. haptocladus N. haptocladus

Stylopage hadra Zoopagomycota (adhesive hyphae) - - Cystopage cladospora (adhesive hyphae)

Catenaria Catenaria Blastocladiomycota - (zoospores) (zoospores) C. anguillulae C. anguillulae C. auxiliaris C. auxiliaris

Myzocytiopsis Nematophthora gynophila (zoospores) (zoospores) - M. glutinospora Oomycota M. vermicola M. lenticularis M. humicola Haptoglossa heterospora (gun cells, injection)

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General Introduction

1.5 Nematophagous fungi, a source of nematicidal metabolites

In the past, control of PPNs had largely relied on the application of synthetic chemicals as a common strategy for plant protection. The first attempts to use chemicals for nematode control were made by Kühn who applied a soil fumigation technique using disulfide to control H. schachtii (Thorne 1961). Since then, the use of chemical pesticides was found to be an effective method that could quickly reduce and economically control any plant-parasitic nematode. However, during the entire period when this strategy was employed, the hazards of synthetic chemicals to nature and human health were widely neglected. The lack of specificity, the high toxicity of nematicides to human health and soil biota, the long persistence in soil resulted in residues of these chemicals in crops and the food- chain, groundwater pollution, and the emergence of chemical-resistance are some of the problems caused by the use of pesticides in general (Degenkolb and Vilcinskas 2015; Ghisalberti 2002; Tanaka and Omura 1993). These adverse effects led to a raised public awareness of the problems and eventually resulted in banning the use of these chemicals by legislation in many countries. Therefore, the search for ecologically safe pest management alternatives that do not cause adverse effects on non-target organisms, brought the use of natural enemies of nematodes in focus. In this context, nematophagous fungi and their natural products are one of the major targets to be screened for biocontrol purposes. Besides their direct parasitism on nematodes, these fungi produce a wide range of secondary metabolites with nematicidal activities that can be used as an alternative for biocontrol of parasitic nematodes (Anke 2011; Anke and Sterner 1997; Degenkolb and Vilcinskas 2015; Degenkolb and Vilcinskas 2016; Li et al. 2007). Pioneering studies aiming at profiling metabolites with nematicidal activities in nematophagous or other fungi were mainly conducted in 1990s (Anke et al. 1995; Mayer et al. 1997; Ondeyka et al. 1990; Stadler et al. 1993a; Stadler et al. 1994a; Stadler et al. 1994b), and stimulated continuous search for further natural nematicidal compounds. These studies resulted in the elucidation of several natural compounds, which can be grouped on the basis of their biosynthetic origin (e.g. terrestrial or marine fungi and plants, or bacteria), biological activities, and chemical structures (Anke 2011; Anke and Sterner 2002; Degenkolb and Vilcinskas 2015; Degenkolb and Vilcinskas 2016; Ghisalberti 2002; Li et al. 2007). The chemical groups representing these natural compounds are mainly: Fatty acids, polyketides, furans, terpenoids, alkaloids, and peptide compounds including linear, cyclic and cyclodepsipeptides. A number of fatty acids with nematicidal activities were isolated from several ascomycetous and basidiomycetous species such as Aspergillus niger, pulmonarius and Hericium spp. (Stadler et al. 1994c). Among other examples of fatty acids [(Ghisalberti 2002; Li et al. 2007) and references therein], linoleic acid is an active principle of several nematode-trapping fungi (Arthrobotrys spp.),

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General Introduction which was originally isolated from A. conoides and showed significant nematicidal effects against the free living nematode (Stadler et al. 1993b). Several polyketides such as pochonins, monorden, and monocillins were isolated form P. chlamydosporia and from the hypocrealean fungus Monocillium nordinii with inhibitory activities against nematodes (Degenkolb and Vilcinskas 2015). More than 30 fungal terpenoids were isolated from ascomycetes and basidiomycetes exhibiting nematicidal activities (Ghisalberti 2002; Li et al. 2007), among which for example the oligosporon-type metabolites isolated from A. oligospora, or a macrocycleic lacton derivative isolated from a Paecilomyces sp. were found to have anthelmintic and nematicidal activities (Anke et al. 1995; Degenkolb and Vilcinskas 2015; Li et al. 2007). Li et al. (2007) reviewed 33 alkaloid compounds, which were found to have nematicidal activities against and plant parasitic nematodes. For example the pyrrolizidine alkaloid pochonicine isolated from Pochonia suchlasporia, paraherquamide A isolated from Penicillium paraherquei, and paeciloxazine isolated from a Paecilomyces sp. are known for their nematicidal and anthelmintic effects (Degenkolb and Vilcinskas 2015; Ghisalberti 2002; Kanai et al. 2004). Recently, Anke (2011) reviewed the cyclic depsipeptides, a class of metabolites known for their nematicidal activities (Ghisalberti 2002; Harder et al. 2003), isolated from several fungal sources. Examples include the lipophilic cyclodepsipeptides omphalotins including omphalotin A, a metabolite from the basidiomycete Omphalotus olearius, which was found to be very toxic towards Meloidogyne incognita (Mayer et al. 1997) and the Leucinostatins isolated from Purpureocillium lilacinum with biocidal activities against C. elegans (Park et al. 2004).

1.6 Introduction to the project

The project was initially conducted based on the results (not shown) obtained from two consecutive nematode population samplings that were conducted in wheat fields of the International Maize and Wheat Improvement Centre (CIMMYT) in Turkey. The results showed a sharp decline in the final population size of H. filipjevi suggesting the presence of soil suppressiveness against nematodes hypothetically caused by natural enemies of nematodes such as nematophagous fungi. Accordingly, the project followed a research strategy (summarised in the flowchart below, Fig 1.2), based on which cyst and egg samples of H. filipjevi were intensively screened especially for fungal species with potential nematode-parasitic interactions. At a glance it included: (1) nematode cyst sampling, and isolating nematode-associated fungi, (2) identification and molecular phylogenetic analyses of fungal isolates, (3) secondary metabolite profiling of fungal isolates of interest, and (4) evaluation of nematode antagonism of selected isolates in vitro and in the greenhouse. In general, to obtain the maximum number of fungal isolates associated with H. filipjevi, two screening approaches were applied: In the first phase of the project a ‘general screening’ was applied meaning the adult females

12

General Introduction or cysts and their contents were randomly cultured to isolate nematophagous fungi. In the second phase of the project a ‘focused screening’ was applied. This means only cysts displaying symptoms of fungal infections and unusual discolourations were selected and their fungal infected eggs were cultured individually to isolate fungi colonising the nematode eggs.

Collection of cyst samples

General screening Focused screening (field collected cysts) (only infected cysts & eggs)

Isolation of fungal species associated with wheat and cereal cyst nematodes

Metabolite profiling and In vitro and in vivo evaluation of Species identification and bioactivity assays biocontrol capacity phylogenetic studies

Characterisation of fungal Reporting the most promising Description of nematode- compounds with potential microorganisms as potential associated fungal species new nematicidal activity nematode biocontrol agents to science from cereal fields in Turkey

Figure 1.2- The research strategy followed in this project.

All cyst samples examined were collected from the experimental fields of CIMMYT-Turkey. These fields were naturally infested with H. filipjevi, and were located in two different regions (Yozgat and Haymana) in the Central Anatolian Plateau (CAP) in Turkey. During the general screening phase, various fungal isolates with more or less tight nematode- association, such as P. chlamydosporia, Paecilomyces, , and Fusarium species were identified (data not shown). In greenhouse experiments assessing the biocontrol potential of these isolates against H. filipjevi, reductions in nematode populations of up to 50% were observed (data not shown). Overall, the results obtained from the general screening phase demonstrated that nematophagous fungi were frequently present in the investigated soils. This was in accordance with our primary hypothesis on the presence of nematophagous fungi as a key component responsible for nematode suppression in the investigated fields.

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General Introduction

Close observations of the field collected cysts showed different symptoms of fungal infection. These symptoms (for example unusual discolouration or discernible fungal colonisation) were not reported in the literature from the fungal isolates we had obtained before. It suggested that these symptoms may be caused by fungal species that had not been isolated with the general screening approach. It was hypothesised that the symptom-causing fungi could be overgrown by fast growing fungi if many eggs or an entire cyst were plated on a cultural media for fungal isolation. This led us to the initiation of the focused screening phase: To obtain the maximum number of cysts, the fields were re-sampled. These cysts along with those remaining from the previous phase were individually examined under a stereomicroscope to collect only the symptomatic ones. The symptoms observed were diverse, and cysts bearing these symptoms could reproducibly be found allowing us to group them accordingly. By microscopically scrutinising the eggs of some examples of the symptomatic cysts, it was noticed that they were indeed infected by fungi. These observations were the starting point for the mentioned ‘focused screening’, in which only nematode eggs exhibiting fungal infection were scrutinised and processed for isolation of specially nematode egg-parasitic fungi, many of which proved later to be new to science. A schematic outline displaying the process of this phase of the project is shown below (Fig 1.3). Detailed data of some outcomes of this approach are presented in the further chapters.

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General Introduction

Microscopic selection of infested cysts

Fungal infested eggs

Direct DNA extraction culturing of fungi on Direct short-term from eggs, sequencing different media storage

DNA extraction and Use for further Permanent storage sequencing experiments (-140 °C)

the fungus) Comparison of results (identity of the fungus) In vitro studies and metabolite profiling Morphological and molecular identification – describing new species

Figure 1.3- A schematic outline displaying the process of the Focused screening.

1.7 Research rationale and objectives of the project

Microscopic investigations of individual cysts of H. filipjevi collected from the Anatolian research fields revealed the presence of fungal structures colonising the nematode eggs. Fungal infections in the diseased cysts caused unique symptoms (see chapters 2 – 4). It was hypothesised that there might be specific fungal species responsible for these specific symptoms. No research articles could be found reporting nematode egg-parasitic fungi causing the same infection symptoms as observed in the examined nematode eggs. Fungal infected eggs were thus the cornerstone of the project. Accordingly, a research plan was made for identifying the fungi causing the unique discolourations interpreted as disease symptoms of eggs of the cereal cyst nematode H. filipjevi. Special isolation procedures were needed to obtain the fungi in pure cultures and allow morphological and molecular identifications. In case new species would emerge from these studies, screening for secondary metabolites potentially involved in nematode killing was planned. Thus, the aims of the project were as follows:

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General Introduction

 To establish an improved screening system for nematode-parasitic fungi in nematode- suppressive soils with an emphasis on scrutinising individual infected nematode eggs  Multiphasic analyses of fungal isolates causally involved in parasitising nematode cysts and eggs: . Fungal isolation from the symptomatic nematode eggs, purification, and long-term preservation of cultures . Characterisation and taxonomic description of potentially new fungal species using morphological studies and DNA-based phylogenetic analyses  In-vitro evaluation of the biocontrol potential of identified fungal species towards eggs of H. filipjevi to assess their potential as biocontrol agents  Microscopic documentation of fungal antagonism towards eggs of H. filipjevi, and illustration of mode of actions of identified species as potential egg-parasitic fungi  Screening the identified fungal species for biologically active secondary metabolites: . Extraction and purification of secondary metabolites of fungal strains of interest by applying chromatographic methods such as preparative HPLC (High Performance Liquid Chromatography) . Selection of the optimum liquid for scale-up fermentation of cultures using liquid chromatography-mass spectrometry (LC-MS) analysis and minimum inhibitory concentration (MIC) tests . Assessing the biological activities of identified compounds towards different microorganism especially against nematodes to evaluate their potential as nematicidal substances

Structure of the presented thesis

In the chapters 2 – 4 the novel fungal species isolated from the eggs of H. filipjevi are described, their parasitism toward nematode eggs is documented, and their phylogenetic relationships are discussed. In addition, the secondary metabolites obtained from the species of interest, and their bioactivity against Heterodera filipjevi are discussed.

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1.8 References

Abidou H, El-Ahmed A, Nicol JM, Bolat N, Rivoal R, Yahyaoui A (2005) occurrence and distribution of Heterodera avenae group in Syria and Turkey. Nematol medit 33: 195-201. Alston DG (2011) General concepts of biological control. Utah State University Extension and Utah Plant Pest Diagnostic Laboratory, Logan, US, IPM-015-011. Anke H (2011) Insecticidal and nematicidal metabolites from fungi. In: Hofrichter M (Ed) Industrial Applications. Springer Berlin Heidelberg, Berlin, Heidelberg, 151-163. Anke H, Stadler M, Mayer A, Sterner O (1995) Secondary metabolites with nematocidal and antimicrobial activity from nematophagous fungi and Ascomycetes. Canadian Journal of Botany-Revue Canadienne De Botanique 73: S932-S939. Anke H, Sterner O (1997) Nematicidal metabolites from higher fungi. Current Organic Chemistry 1: 361-374. Anke H, Sterner O (2002) Insecticidal and nematicidal metabolites from fungi. In: Osiewacz HD (Ed) Industrial Applications. Springer Berlin Heidelberg, Berlin, Heidelberg, 109-127. Baral H-O, Weber E, Gams W, Hagedorn G, Liu B, Liu X, Marson G, Marvanová L, Stadler M, Weiß M (2017) Recommendations about generic names to be protected or suppressed in the Orbiliaceae (Orbiliomycetes). Mycological Progress, in press. Barron GL (1977) The nematode-destroying fungi. Canadian biological publications, Guelph, Ontario, Canada, Bird AF, McClure MA (1976) The tylenchid (Nematoda) egg shell: structure, composition and permeability. Parasitology 72: 19-28. Bursnall LA, Tribe HT (1974) Fungal parasitism in cysts of Heterodera. Transactions of the British Mycological Society 62: 595-IN596. Castillo JD, Lawrence KS (2011) First report of Catenaria auxiliaris parasitizing the reniform nematode Rotylenchulus reniformis on cotton in alabama. Plant Disease 95: 490. Chen F, Chen S (2002) Mycofloras in cysts, females, and eggs of the soybean cyst nematode in Minnesota. Applied Soil Ecology 19: 35-50. Chen S, Liu XZ, Chen FJ (2000) Hirsutella minnesotensis sp.nov., a new pathogen of the soybean cyst nematode. Mycologia 92: 819-824. Crump DH, Flynn CA (1995) Isolation and screening of fungi for the biological control of potato cyst nematodes. Nematologica 41: 628-638. Dababat AA, Imren M, Erginbas-Orakci G, Ashrafi S, Yavuzaslanoglu E, Toktay H, Pariyar SR, Elekcioglu HI, Morgounov A, Mekete T (2015) The importance and management strategies of cereal cyst nematodes, Heterodera spp., in Turkey. Euphytica 202: 173-188. Dackman C, Nordbring-Hertz B (1985) Fungal parasites of the cereal cyst nematode Heterodera avenae in southern Sweden. Journal of Nematology 17: 50-55. Decraemer W, Hunt DJ (2013) Structure and classification. In: Perry RN, Moens M (Eds) Plant nematology. CAB International, Wallingford, 3-32. Degenkolb T, Vilcinskas A (2015) Metabolites from nematophagous fungi and nematicidal natural products from fungi as an alternative for biological control. Part I: metabolites from nematophagous ascomycetes. Applied Microbiology and Biotechnology 100: 3799-3812. Degenkolb T, Vilcinskas A (2016) Metabolites from nematophagous fungi and nematicidal natural products from fungi as alternatives for biological control. Part II: metabolites from nematophagous basidiomycetes and non-nematophagous fungi. Appl Microbiol Biotechnol 100: 3813-3824. Dijksterhuis J, Veenhuis M, Harder W (1990) Ultrastructural study of adhesion and initial stages of infection of nematodes by conidia of Drechmeria coniospora. Mycological Research 94: 1-8. Dijksterhuis J, Veenhuis M, Harder W, Nordbring-Hertz B (1994) Nematophagous fungi: physiological aspects and structure-function relationships. Adv Microb Physiol 36: 111-143. Eilenberg J (2006) Concepts and visions of biological control. In: Eilenberg J, Hokkanen HMT (Eds) An Ecological and Societal Approach to Biological Control. Springer , Dordrecht, 1-11. 17

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Franklin MT (1969) Heterodera latipons n. sp., a cereal cyst nematode from the Mediterranean region. Nematologica 15: 535-542. Ghisalberti EL (2002) Secondary metabolites with antinematodal activity. In: Atta ur R (Ed) Studies in Natural Products Chemistry. Elsevier, 425-506. Goffart H (1932) Untersuchungen am Hafernematoden Heterodera schachtii Schm. unter besonderer Brücksichtigung der schleswig-holsteinischen Verhältnisse I. Arbeiten aus der Biologischen Reichsanstalt für Land- und forstwirtschaft Berlin-Dahlem 20: 1-28. Harder A, Schmitt-Wrede HP, Krucken J, Marinovski P, Wunderlich F, Willson J, Amliwala K, Holden- Dye L, Walker R (2003) Cyclooctadepsipeptides - an anthelmintically active class of compounds exhibiting a novel mode of action. International Journal of Antimicrobial Agents 22: 318-331. Holterman M, Karegar A, Mooijman P, van Megen H, van den Elsen S, Vervoort MTW, Quist CW, Karssen G, Decraemer W, Opperman CH, Bird DM, Kammenga J, Goverse A, Smant G, Helder J (2017) Disparate gain and loss of parasitic abilities among nematode lineages. PLOS ONE 12: e0185445. Hywel-Jones NL (2002) The importance of invertebrate-pathogenic fungi from the tropics. In: Watling R, Frankland JC, Ainsworth AM, Isaac S, Robinson CH (Eds) Tropical : Volume 2, micromycetes. CAB International. Jansson H-b, Friman E (1999) Infection-related surface proteins on conidia of the nematophagous fungus Drechmeria coniospora. Mycological Research 103: 249-256. Jansson H-B, Lopez-Llorca L (2001) Biology of nematophagous fungi. In: Misra JK, Bruce WH (Eds) Trichomycetes and other fungal groups. Science Publishers, Inc., Enfield (NH), USA, 145-173. Jones JT, Haegeman A, Danchin EG, Gaur HS, Helder J, Jones MG, Kikuchi T, Manzanilla-Lopez R, Palomares-Rius JE, Wesemael WM, Perry RN (2013) Top 10 plant-parasitic nematodes in molecular plant pathology. Mol Plant Pathol 14: 946-961. Kanai Y, Fujimaki T, Kochi S, Konno H, Kanazawa S, Tokumasu S (2004) Paeciloxazine, a novel nematicidal antibiotic from Paecilomyces sp. J Antibiot (Tokyo) 57: 24-28. Kerry B (1980) Biocontrol: Fungal Parasites of Female Cyst Nematodes. Journal of Nematology 12: 253- 259. Kerry B (1988) Fungal parasites of cyst nematodes. Agriculture Ecosystems & Environment 24: 293- 305. Kerry BR (1975) Fungi and the decrease of cereal cyst-nematode populations in cereal monoculture. EPPO Bulletin 5: 353-361. Kerry BR, Crump DH (1977) Observations on fungal parasites of females and eggs of the cereal cyst- nematode, Heterodera avenae, and other cyst nematodes. Nematologica 23: 193-201. Kerry BR, Crump DH (1980) Two fungi parasitic on females of cystnematodes (Heterodera spp.). Transactions of the British Mycological Society 74: 119-125. Kerry BR, Crump DH (1998) The dynamics of the decline of the cereal cyst nematode, Heterodera avenae, in four soils under intensive cereal production. Fundamental and Applied Nematology 21: 617-625. Kerry BR, Crump DH, Mullen LA (1982a) Natural control of the cereal cyst nematode, Heterodera avenae Woll., by soil fungi at three sites. Crop Protection 1: 99-109. Kerry BR, Crump DH, Mullen LA (1982b) Studies of the cereal cyst-nematode, Heterodera avenae under continuous cereals, 1975–1978. II. Fungal parasitism of nematode females and eggs. Annals of Applied Biology 100: 489-499. Khan A, Williams KL, Nevalainen HKM (2006) Infection of plant-parasitic nematodes by Paecilomyces lilacinus and lysipagum. BioControl 51: 659-678. Kühn J (1874) Über das Vorkommen von Rübennematoden an den Wurzeln der Halmfrüchte. Landw Jahrbücher 3: 47-50. Kühn J (1877) Vorläufiger Bericht über die bisherigen Ergebnisse der seit dem Jahre 1875 im Auftrage des Vereins für Rübenzucker-Industrie ausgeführten Versuche zur Ermittlung der Ursache der Rübenmüdigkeit des Bodens und zur Erforschung der Natur der Nematoden. Zeitschrift des Vereins für die Rübenzucker-Industrie des Duetschen Reich (ohne Band) 27: 452-457.

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Kwenti TE (2017) Biological Control of Parasites. In: Khater H, Govindarajan M, Benelli G (Eds) Natural Remedies in the Fight Against Parasites. InTech, Rijeka, Ch. 02. Li G, Zhang K, Xu J, Dong J, Liu Y (2007) Nematicidal substances from fungi. Recent Patents on Biotechnology 1: 212-233. Li J, Zou C, Xu J, Ji X, Niu X, Yang J, Huang X, Zhang KQ (2015) Molecular mechanisms of nematode- nematophagous microbe interactions: basis for biological control of plant-parasitic nematodes. Annu Rev Phytopathol 53: 67-95. Lilley CJ, Kyndt T, Gheysen G (2011) Nematode resistant GM crops in industrialised and developing countries. In: Jones J, Gheysen G, Fenoll C (Eds) Genomics and Molecular Genetics of Plant- Nematode Interactions. Springer Netherlands, Dordrecht, 517-541. Lohde G (1874) Über einige neue parasitische Pilze. Tageblatt Versammlung deutscher Naturforscher und Ärzte in Breslau 47: 203-206. Lopez-Llorca LV, Jansson H-B, Vicente JGM, Salinas J (2006) Nematophagous Fungi as Root Endophytes. In: Schulz BJE, Boyle CJC, Sieber TN (Eds) Microbial Root Endophytes. Springer Berlin Heidelberg, Berlin, Heidelberg, 191-206. Lopez-Llorca LV, Maciá-Vicente JG, Jansson H-B (2008) Mode of action and interactions of nematophagous fungi. In: Ciancio A, Mukerji KG (Eds) Integrated Management and Biocontrol of Vegetable and Grain Crops Nematodes. Springer Netherlands, Dordrecht, 51-76. Lopez-Llorca LV, Olivares-Bernabeu C, Salinas J, Jansson H-B, Kolattukudy PE (2002) Pre-penetration events in fungal parasitism of nematode eggs. Mycological Research 106: 499-506. Luc M, Weischer B, Stone AR, Baldwin JG (1986) On the definition of heteroderid cysts. Revue de Nématologie 9: 418-421. Madzhidov AR (1981) [Bidera filipjevi n. sp. (Heteroderina: ) in Tadzhikistan.] Izvestiya Akademii Nauk Tadzhiksköi SSR. Izvestiya Akademii Nauk Tadzhiksköi SSR, Otdelenie Biologicheskie Nauki 2: 40-44. Mayer A, Anke H, Sterner O (1997) Omphalotin, a new cyclic peptide with potent nematicidal activity from Omphalotus olearius I. Fermentation and biological activity. Natural Product Letters 10: 25-32. Moens M, Perry RN (2009) Migratory Plant Endoparasitic Nematodes: A Group Rich in Contrasts and Divergence. Annual Review of Phytopathology 47: 313-332. Morgan-Jones G, White JF, Rodriguez-Kabana R (1983) Phytonematode Pathology: Ultrastructural Studies. I. Parasitism of Meloidogyne arenaria Eggs by Verticillium chlamydosporium. Nematropica 13. Morgan-Jones G, White JF, Rodriguez-Kabana R (1984) Phytonematode pathology: Ultrastructural studies. II. Parasitism of Meloidogyne arenaria eggs and larvae by Paecilomyces lilacinus. Nematropica 14: 57-71. Morton O, Hirsch P, Kerry B (2004) Infection of plant-parasitic nematodes by nematophagous fungi – a review of the application of molecular biology to understand infection processes and to improve biological control. Nematology 6: 161-170. Nicol JM, Rivoal R (2008) Global knowledge and its application for the integrated control and management of nematodes on wheat. In: Ciancio A, Mukerji KG (Eds) Integrated management and biocontrol of vegetable and grain crops nematodes. Springer, Dordrecht, 243-287. Nicol JM, Turner SJ, Coyne DL, Nijs Ld, Hockland S, Maafi ZT (2011) Current nematode threats to world agriculture. In: Jones J, Gheysen G, Fenoll C (Eds) Genomics and Molecular Genetics of Plant- Nematode Interactions. Springer Netherlands, Dordrecht, 21-43. Nordbring-Hertz B (2004) Morphogenesis in the nematode-trapping fungus — an extensive plasticity of infection structures. Mycologist 18: 125-133. Nordbring-Hertz B, Jansson H-B, Tunlid A (2011) Nematophagous fungi. Encyclopedia of life sciences. John Wiley & Sons, Ltd. Ondeyka JG, Goegelman RT, Schaeffer JM, Kelemen L, Zitano L (1990) Novel antinematodal and antiparasitic agents from Penicillium charlesii. I. Fermentation, isolation and biological activity. The Journal of Antibiotics (Tokyo) 43: 1375-1379.

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Park JO, Hargreaves JR, McConville EJ, Stirling GR, Ghisalberti EL, Sivasithamparam K (2004) Production of leucinostatins and nematicidal activity of Australian isolates of Paecilomyces lilacinus (Thom) Samson. Letters in Applied Microbiology 38: 271-276. Perry RN, Moens M (2011) Introduction to plant-parasitic nematodes; Modes of parasitism. In: Jones J, Gheysen G, Fenoll C (Eds) Genomics and Molecular Genetics of Plant-Nematode Interactions. Springer Netherlands, Dordrecht, 3-20. Perry RN, Trett MW (1986) Ultrastructure of the eggshell of Heterodera schachtii and H. glycines (Nematoda:Tylenchida). Revue de Nematologie 9: 399-403. Porter TM, Martin W, James TY, Longcore JE, Gleason FH, Adler PH, Letcher PM, Vilgalys R (2011) Molecular phylogeny of the Blastocladiomycota (Fungi) based on nuclear ribosomal DNA. Fungal Biology 115: 381-392. Schacht H (1859) Über einige Feinde der Rübenfelder. Zeitschrift des Vereines für die Rübenzucker_Industrie im Zollverein 9: 175-179. Schmidt A (1871) Über den Rüben-Nematoden (Heterodera schachtii A.S.). Zeitschrift des Vereines für die Rübenzucker-Industrie im Zollverein 21: 1-19. Scholler M, Hagedorn G, Rubner A (1999) A reevaluation of predatory orbiliaceous fungi. II. A new generic concept. Sydowia 51: 89–113. Shu C, Lai Y, Yang E, Chen S, Xiang M, Liu X (2015) Functional response of the fungus Hirsutella rhossiliensis to the nematode, Heterodera glycines. Science -Life Sciences 58: 704-712. Sikora RA, Bridge J, Starr JL (2005) Management practices: an overview of integrated nematode management technologies. In: Luc M, Sikora RA, Bridge J (Eds) Plant parasitic nematodes in subtropical and tropical agriculture. CAB International, 793-825. Singh UB, Sahu A, Sahu N, Singh RK, Renu, Singh DK, Singh BP, Jaiswal RK, Singh DP, Rai JP, Manna MC, Singh KP, Srivastava JS, Rao AS, Prasad SR (2013) Nematophagous fungi: Catenaria anguillulae and Dactylaria brochopaga from seed galls as potential biocontrol agents of Anguina tritici and Meloidogyne graminicola in wheat (Triticum aestivum L.). Biological Control 67: 475-482. Smiley RW, Dababat AA, Iqbal S, Jones MGK, Maafi ZT, Peng D, Subbotin SA, Waeyenberge L (2017) Cereal Cyst Nematodes: A Complex and Destructive Group of Heterodera Species. Plant Disease 101: 1692-1720. Sobczak M, Golinowski W (2011) Cyst nematodes and Syncytia. In: Jones J, Gheysen G, Fenoll C (Eds) Genomics and Molecular Genetics of Plant-Nematode Interactions. Springer Netherlands, Dordrecht, 61-82. Stadler M, Anke H, Arendholz W-R, Hansske F, Anders UWE, Sterner O, Bergquist K-E (1993a) Lachnumon and lachnumol A, new metabolites with nematicidal and antimicrobial activities from the ascomycete papyraceum (Karst.) Karst. I. Producing organism, fermentation, isolation and biological activities. The Journal of Antibiotics 46: 961-967. Stadler M, Anke H, Sterner O (1993b) Linoleic acid - The nematicidal principle of several nematophagous fungi and its production in trap-forming submerged cultures. Archives of Microbiology 160: 401-405. Stadler M, Anke H, Sterner O (1994a) New nematicidal and antimicrobial compounds from the basidiomycete Cheimonophyllum candidissimum (Berk & Curt.) sing. I. Producing organism, fermentation, isolation, and biological activities. The Journal of Antibiotics 47: 1284-1289. Stadler M, Dagne E, Anke H (1994b) Nematicidal activities of two phytoalexins from Taverniera abyssinica. Planta Med 60: 550-552. Stadler M, Mayer A, Anke H, Sterner O (1994c) Fatty acids and other compounds with nematicidal activity from cultures of Basidiomycetes. Planta Med 60: 128-132. Starr JL, Bridge J, Cook R (2002) Resistance to plant-parasitic nematodes: History, current use and future potential. In: Starr JL, Cook R, Bridge J (Eds) Plant resistance to parasitic nematodes. CAB International. Stirling GR (2014) Biological control of plant-parasitic nematodes: soil ecosystem management in sustainable agriculture. CABI, Wallingford, UK, xxiv + 510 pp.

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Subbotin SA, Mundo-Ocampo M, Baldwin J (2010a) Systematics of cyst nematodes (Nematoda: Heteroderinae), Part A. Brill, Subbotin SA, Mundo-Ocampo M, Baldwin J (2010b) Systematics of cyst nematodes (Nematoda: Heteroderinae), Part B. Brill, Subbotin SA, Waeyenberge L, Moens M (2013) Molecular systematics. In: Perry RJ, Moens M (Eds) Plant nematology. CAB International, Wallingford, 40-64. Szabó M, Csepregi K, Gálber M, Virányi F, Fekete C (2012) Control plant-parasitic nematodes with Trichoderma species and nematode-trapping fungi: The role of chi18-5 and chi18-12 genes in nematode egg-parasitism. Biological Control 63: 121-128. Tanaka Y, Omura S (1993) Agroactive compounds of microbial origin. Annu Rev Microbiol 47: 57-87. Thorne G (1961) Principles of Nematology. McGraw Hill New York, Toktay H, Imren M, Ocal A, Waeyenberge L, Viaene N, Dababats A (2015) Incidence of cereal cyst nematodes in the East Anatolia Region in Turkey. Russian Journal of Nematology 23: 29-40. Toumi F, Waeyenberge L, Viaene N, Dababat AA, Nicol JM, Ogbonnaya F, Moens M (2017) Cereal cyst nematodes: importance, distribution, identification, quantification, and control. European Journal of Plant Pathology. Tribe HT (1977a) A parasite of white cysts of Heterodera: Catenaria auxiliaris. Transactions of the British Mycological Society 69: 367-376. Tribe HT (1977b) Pathology of cyst-nematodes. Biological Reviews of the Cambridge Philosophical Society 52: 477-&. Tunlid A, Ahrén D (2011) Molecular Mechanisms of the Interaction Between Nematode-Trapping Fungi and Nematodes: Lessons From Genomics. In: Davies K, Spiegel Y (Eds) Biological Control of Plant-Parasitic Nematodes:: Building Coherence between Microbial Ecology and Molecular Mechanisms. Springer Netherlands, Dordrecht, 145-169. Turner SJ, Subbotin SA (2013) Cyst nematodes. In: Perry RJ, Moens M (Eds) Plant nematology. CAB International, Wallingford, 109-141. Viaene N, coyne DL, Davies KG (2013) Biological and cultural management. In: Perry RJ, Moens M (Eds) Plant nematology. CAB International, Wallingford, 383-410. Westphal A (2005) Detection and description of soils with specific nematode suppressiveness. Journal of Nematology 37: 121-132. Westphal A, Becker JO (2001) Components of soil suppessiveness against Heterodera schachtii. Soil Biology and Biochemistry 33: 9-16. Willcox J, Tribe HT (1974) Fungal parasitism in cysts of Heterodera. I. Preliminary investigations. Transactions of the British Mycological Society 62: 585-&. Wollenweber HW (1924) Zur Kenntnis der Kartoffel-Heteroderen. Illustrierte Landwirtschaftliche Zeitung 44: 100-101. Wyss U, Voss B, Jansson H-B (1992) In vitro observations on the infection of Meloidogyne incognita eggs by the zoosporic fungus Catenaria anguillulae Sorokin. Yang Y, Yang E, An Z, Liu X (2007) Evolution of nematode-trapping cells of predatory fungi of the Orbiliaceae based on evidence from rRNA-encoding DNA and multiprotein sequences. Proceedings of the National Academy of Sciences of the United States of America 104: 8379- 8384. Zare R, Gams W, Culham A (2001) A revision of Verticillium sect. Prostrata. I. Phylogenetic studies using ITS sequences. Nova Hedwigia 71: 465-480. Zhang S, Gan Y, Xu B, Xue Y (2014) The parasitic and lethal effects of Trichoderma longibrachiatum against Heterodera avenae. Biological Control 72: 1-8.

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Chapter 2.

Ijuhya vitellina sp. nov., a novel source for chaetoglobosin A, is a destructive parasite of the cereal cyst nematode Heterodera filipjevi

Published in PLoS ONE 12(7): e0180032. https://doi.org/10.1371/journal.pone.0180032

Samad Ashrafi1,2*, Soleiman Helaly3,4†, Hans-Josef Schroers5†, Marc Stadler3, Katja R. Richert- Poeggeler1, Abdelfattah A. Dababat6, Wolfgang Maier1

1 Institute for Epidemiology and Pathogen Diagnostics, Julius Kühn-Institut (JKI) — Federal Research Centre for Cultivated Plants, Braunschweig, Germany

2 Department of Ecological Plant Protection, Faculty of Organic Agricultural Sciences, University of Kassel, Witzenhausen, Germany

3 Department Microbial Drugs, Helmholtz Centre for Infection Research GmbH (HZI), Braunschweig, Germany

4 Department of Chemistry, Faculty of Science, Aswan University, Aswan 81528, Egypt

5 Agricultural Institute of Slovenia, Ljubljana, Slovenia

6 CIMMYT (International Maize and Wheat Improvement Centre), P.K.39 06511 Emek, Ankara, Turkey

* Corresponding author

E-mail: [email protected]

† These authors contributed equally to this work

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Abstract

Cyst nematodes are globally important pathogens in agriculture. Their sedentary lifestyle and long- term association with the roots of host plants render cyst nematodes especially good targets for attack by parasitic fungi. In this context fungi were specifically isolated from nematode eggs of the cereal cyst nematode Heterodera filipjevi. Here, Ijuhya vitellina (Ascomycota, Hypocreales, Bionectriaceae), encountered in wheat fields in Turkey, is newly described on the basis of phylogenetic analyses, morphological characters and life-style related inferences. The species destructively parasitises eggs inside cysts of H. filipjevi. The parasitism was reproduced in in vitro studies. Infected eggs were found to harbour microsclerotia produced by I. vitellina that resemble long-term survival structures also known from other ascomycetes. Microsclerotia were also formed by this species in pure cultures obtained from both, solitarily isolated infected eggs obtained from fields and artificially infected eggs. Hyphae penetrating the eggshell colonised the interior of eggs and became transformed into multicellular, chlamydospore-like structures that developed into microsclerotia. When isolated on artificial media, microsclerotia germinated to produce multiple emerging hyphae. The specific nature of morphological structures produced by I. vitellina inside nematode eggs is interpreted as a unique mode of interaction allowing long-term survival of the fungus inside nematode cysts that are known to survive periods of drought or other harsh environmental conditions. Generic classification of the new species is based on molecular phylogenetic inferences using five different gene regions. I. vitellina is the only species of the genus known to parasitise nematodes and produce microsclerotia. Metabolomic analyses revealed that within the Ijuhya species studied here, only I. vitellina produces chaetoglobosin A and its derivate 19-O-acetylchaetoglobosin A. Nematicidal and nematode-inhibiting activities of these compounds have been demonstrated suggesting that the production of these compounds may represent an adaptation to nematode parasitism.

Keywords

Chaetoglobosins, cytotoxicity, new species, plant parasitic nematodes, act, ITS, LSU, rpb1, ß-tub

2.1 Introduction

Cyst nematodes are attacked by several fungal species. The first report on cyst-parasitic fungi dates back to 1877, when Julius Kühn described a Tarichium species, today known as Catenaria auxiliaris (Kuehn) Tribe, as a parasite of the sugar beet nematode Heterodera schachtii Schmidt (Kühn 1877). A diverse group of fungi has since then been described as associates of cyst nematodes (Stirling 2014).

Examples include Ilyonectria destructans (Zinssm.) Rossman, L. Lombard & Crous (Goffart 1932), Pochonia chlamydosporia (Goddard) Zare & W. Gams (both Ascomycota, Hypocreales) and 23

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Nematophthora gynophila Kerry & D.H. Crump (Stramenopiles) (Kerry 1974; Kerry and Crump 1980) that were described as parasites of Heterodera avenae Wollenweber. Kerry (1975) also demonstrated that all these species contribute to the natural suppression of nematode populations of H. avenae. Similar nematode suppressive effects were also reported from different geographical regions (Dackman and Nordbring-Hertz 1985; Kerry et al. 1982; Khan et al. 2006; Rodríguez-Kábana and Morgan-Jones 1988). These observations have increased further attention to investigate the association of fungi with cyst nematodes and their biocontrol potential. The destructive parasitism on nematodes is in some cases linked with the production of biologically active fungal secondary metabolites (Anke et al. 1995; Li et al. 2007; Stadler et al. 1994) including nematicidal compounds that display various anthelmintic effects (Anke 2011). Chaetoglobosins of the cytochalasan family can have cytotoxic and inhibitory activities (Sekita et al. 1976; Sekita et al. 1973) and affect insects and nematodes (Cutler et al. 1980; Maruyama et al. 1986; Meyer et al. 2004; Xue et al. 2012; Zheng et al. 2014). In the past decade, the use of synthetic chemical nematicides has either been reduced significantly or they were banned completely for posing health and environmental risks. Therefore, environmentally friendly and biologically effective alternatives for the control of nematode plant pests are urgently needed. They may consist in the application of whole organisms or in biologically active fungal compounds only (Anke et al. 1995; Ghisalberti 2002; Hu et al. 2013; Kundu et al. 2016; Li et al. 2007; Niu et al. 2010; Stadler et al. 1993). In this context, experimental fields of the International Maize and Wheat Improvement Centre (CIMMYT) in Turkey were screened for antagonistic fungi associated with the cereal cyst nematode Heterodera filipjevi. We report here on the discovery of a new species from the hypocrealean Bionectriaceae and describe its unique destructive parasitism on eggs of H. filipjevi. Potentially involved secondary metabolites produced by the fungus were isolated, structurally elucidated and their biological activity against nematodes was tested.

2.2 Material and methods

2.2.1 Nematode collection and materials examined

Naturally nematode-infested experimental fields of CIMMYT located in two different regions in the Central Anatolian Plateau of Turkey including Yozgat (39° 08ʼ N, 34° 10ʼ E; altitude, 985 m) and Haymana (39° 25ʼ 52ʼ N, 39° 29ʼ 44ʼ E; altitude, 1259 m) were sampled at crop maturity in 2013. Nematode cysts were extracted from rhizosphere and roots of nematode susceptible and resistant wheat varieties using the modified flotation decanting method (Coyne et al. 2007) and handpicked from the extracted suspensions under a Leitz dissecting microscope. Cysts were stored in 1.5 mL microtubes at 4˚C either in dry condition or in water for further experiments. Reference strains 24

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi required for taxonomic and phylogenetic inferences were obtained from the Westerdijk Fungal Biodiversity Institute (Utrecht, The Netherlands).

2.2.2 Ethics statement

Both the sampling sites and the nematode species were not protected and thus no permits were required.

2.2.3 Fungal isolation from field collected samples

Homogenously brown and visibly emptied cysts of H. filipjevi were separated from hereafter called symptomatic cysts showing unusual discolourations or fungal colonization. Under a laminar flow hood, symptomatic cysts were surface sterilized in 0.5% sodium hypochlorite (NaOCl) for 10 min and rinsed six times with sterile deionised water (SDW). The sterilizing effect of NaOCl was evaluated. For this, individual cysts were imprinted into potato dextrose agar medium (PDA; Merck, Germany) using a sterile forceps, and immediately transferred to new PDA plates. The control plates were incubated at room temperature and regularly monitored for contaminants for four weeks to exclude not- successfully surface-sterilised cysts from further analyses. Using a sterile forceps and an insect needle, transferred cysts were separately cut open on the fresh agar medium and eggs were dispersed on the agar plate. Cyst debris, i.e. particles of the cyst wall, was discarded and nematode eggs showing symptoms of fungal infections were rolled gently on agar surface to remove potentially occurring contaminating fungal propagules or hyphae adhering to the surface of eggs. Eggs were then transferred to a sterile watch glass containing SDW. When settled, eggs were rinsed six times by removing and replacing the majority of SDW. Eggs were then surface sterilized in 0.5% NaOCl for 2 min. The disinfection solution was removed and the eggs were washed up to six times with SDW as described above. Surface-sterilised eggs of each individual cyst were then transferred to a new agar plate using a pipette and divided in two aliquots. Eggs from one portion were individually picked up and placed on PDA amended with penicillin G (240 mg/L) and streptomycin sulphate (200 mg/L) (PDA+). Plates were incubated at room temperature and monitored regularly. Each agar plate received a maximum of 4 individually placed single eggs. Fungi emerging from these eggs were identified using morphological and molecular phylogenetic methods. In addition they were used for studying fungal-nematode interactions in vitro. For long-term maintenance, representative fungal isolates were stored cryogenically at -140°C. Individual eggs from the second portion were directly transferred into 1.5 mL microtubes for culture-independent identification. The results of species identifications obtained from the culture-independent method were then compared to the results from the culture-dependent

25

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi method. By doing so, additional evidence was gathered to prove that the fungi isolated by culturing techniques indeed colonised the nematode eggs.

2.2.4 Pure culture based studies

Sporulating structures were assessed after the isolates of the here studied fungal species were inoculated on corn meal agar (CMA; Fluka), oatmeal agar (OA; 30 g oatmeal, 18 g agar-agar, 1L deionised water), synthetic nutrient-poor agar (SNA; (Nirenberg 1976)), malt extract agar (MEA; Carl Roth), yeast malt agar (YMA; (Stadler et al. 2001)), PDA and one third strength PDA (PDA 1/3). Strains were also inoculated on PDA, CMA and SNA supplemented with sterile pieces of filter paper, carnation leaf pieces, or wheat straw. Cultures were incubated for up to 12 months at room temperature, as well as at 10, 15, 20, 25, and 30° C in dark or under different light regimes including ambient lighting, or 12h /12 h cycles of light/darkness or black light/darkness. Growth rates at various temperatures were determined by inoculating cylindrical agar plugs, 4–5 mm diam, excised from the margin of PDA cultures onto PDA (Difco). Colour changes of fungal structures formed in culture were checked in 3% watery solution of potassium hydroxide (KOH). Colour codes used in the description were determined according to Kornerup and Wanscher (1967).

2.2.5 Pathogenicity test and Koch’s postulate

The pathogenicity of the here studied fungal species was tested in vitro against cysts and eggs of H. filipjevi multiplied on wheat plants grown in steamed substrates in the greenhouse. Cysts were extracted and surface-sterilised for the experiments as described above. Three independently isolated strains of the fungus were sub-cultured on PDA+ and incubated for 3 months in 10 replicates. Ten surface-sterilised healthy cysts were then placed on top of each of the colonies. Plates were incubated at room temperature and cysts were monitored at regular intervals for fungal infection. Similar experiments were also done with surface sterilized eggs obtained from healthy cysts. To ensure that there is no contamination, eggs were individually placed on PDA and incubated at ambient temperature. Under a laminar flow hood, eggs not showing any contamination after 2 d of incubation, were individually placed on top or at the edge of 2-month-old PDA+, PDA1/3+ and SNA+ cultures. Plates were incubated at room temperature and eggs were monitored daily. The process of fungal colonisation of eggs of H. filipjevi was also studied in modified slide culture experiments (Gams et al. 1998). Single microsclerotia formed by the studied fungus (described below) were placed as inoculum in the centre of agar blocks (15×15×2 mm), and up to 20 nematode eggs were placed in their vicinity. Inoculated agar blocks were covered with sterile cover slips and slides were

26

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi incubated in moist glass chambers at room temperature. Developing structures were monitored and microscopically photographed regularly.

2.2.6 Light and scanning electron microscopy

Nematode and fungal structures were examined and photographed with a Zeiss Axioskop 2 plus compound microscope and an Olympus SZX 12 stereo microscope equipped with a Jenoptik ProgRes® digital camera supported with CapturePro 2.8 software (Jenoptik, Jena, Germany). Eggs and fungal structures were mounted in water. Cysts were photographed in a square cavity dish in water. All microscopic specimens were studied using Differential Interference Contrast (DIC) optics. Measurements are given as minimum – maximum × minimum – maximum with arithmetical means placed in brackets, followed by the number of measurements (n). For SEM, fungal structures of interest were either picked directly from the surface of colonies or collected after dissolving a small piece of agar in an agarose dissolving buffer (Zymo Research Corp., Irvine, California, USA). Structures were washed with SDW and placed on non-conductive double-sided adhesive tape on aluminium stubs. Samples were photographed using a FEI Quanta 250 scanning electron microscope (Hillsboro, Oregon, USA) at 12.5 kV in low vacuum. Images were adjusted in brightness and contrast using Adobe Photoshop software CS 5.1.

2.2.7 DNA extraction

Fungal mycelium was obtained from PDA and transferred to 1.5 mL microtubes. Genomic DNA was extracted with a CTAB-based method (Saghai-Maroof et al. 1984). Cells were disrupted by grinding using sterile micro-pestles and then lysed in 800 µl CTAB buffer at 65°C for 1 h and 300 rpm. Removal of proteins and precipitation was achieved in two steps by adding 600 µl chloroform and 350 µl isopropanol. Polar fractions were retrieved through centrifugation. DNA pellets were washed twice with 70% ethanol, re-suspended in molecular grade water or elution buffer, and stored at -20°C. Infected single eggs were transferred to 1.5 mL Eppendorf microtubes containing 5 µl SDW. Approximately 40 mg sterile silica sand and four 1 mm sterile steal beads were added. The samples were incubated in a laminar flow to evaporate the remaining water. Each sample was then frozen in liquid and then disrupted in a tissue lyser (Qiagen TissueLyser LT, Hilden, Germany) at 50 Hz for 2 min. Freezing and disruption steps were repeated three times. DNA was then extracted and purified with the Qiagen DNeasy Plant Mini kit following the manufacturer’s instructions.

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

2.2.8 PCR amplification and sequencing

Two domains of the nuclear rDNA gene cluster including the internal transcribed spacers with the 5.8S rDNA gene (ITS) and the 5’ end of the nuclear large subunit ribosomal RNA gene (LSU) were amplified with primers V9G (de Hoog and Gerrits van den Ende 1998) and LR5 (Vilgalys and Hester 1990) and sequenced with primers ITS1F (Gardes and Bruns 1993), ITS4 (White et al. 1990), LR0R (Rehner and Samuels 1994), and LR5. Three partial protein-encoding genes were amplified and sequenced including the RNA polymerase II largest subunit 1 (rpb 1), actin (act), and β-tubulin (ß-tub) genes. The primers cRPB1af and RPB1cr (Castlebury et al. 2004) were used for amplification and sequencing of rpb1; Tact1f and Tact2r (Samuels et al. 2006) were used for act, and T1 and T22 for ß-tub. For sequencing ß-tub, primer T222 and T224 (O’Donnell and Cigelnik 1997) were used as additional internal sequence primers. All PCR reactions were performed in a final volume of 50 µl containing 1 µl of template DNA and 49 µl of PCR master mix including 5 µl of 10× TrueStart, (NH4)2SO4 amended Taq Buffer (Thermo Scientific),

-1 5 µl MgCl2 (2.5 mM), 5 µl dNTPs (0.2 mM of each), 2 µl of each primer (0.4 pM µl ), and 1 Unit Taq DNA polymerase (TrueStart Hot Start, Thermo Scientific). The amplifications were carried out on a T- GRADIENT thermocycler (Biometra, Göttingen, Germany) with the following thermal programmes: 95°C (2 min) for initial denaturation followed by 40 cycles of denaturation at 95°C (30 s), annealing at 52.5°C (ITS, LSU), 54°C (rpb1), 55.5°C (β-tub), 58.5°C (act) (40 s), extension at 72°C for 100 s (ITS and LSU), 80 s (ß-tub), and 60 s (rpb1 and act), and a final extension at 72°C (10 min). PCR products were purified using the DNA Clean & ConcentratorTM-5 kit (Zymo Research Corp., Irvine, California, USA) according to the manufacturer’s instructions. The cycle sequencing products were run on an ABI 3730XL sequencing machine (Eurofins Genomics GmbH, Germany). Obtained sequences were assembled, edited and trimmed with Sequencher 5.4.1 (Gene Codes Corporation, Ann Arbor, Michigan, USA) and deposited in GenBank under accession numbers KY607531–KY607585 and KY684180–KY684193.

2.2.9 Alignment and phylogenetic reconstruction

Newly generated and already published sequences were used in phylogenetic analyses (Table2. 1). The latter were selected according to BLASTn searches (http://blast.ncbi.nlm.nih.gov/Blast.cgi) (Altschul et al. 1990) that used the former as queries. Representatives of Bionectriaceae and Nectriaceae were selected mainly following Hirooka et al. (2010) and Jaklitsch and Voglmayr (2011). Several data sets based on various combinations of ITS, LSU, rpb1, act, ß-tub sequences were compiled, aligned and analysed separately. Sequences of the LSU (Fig 2.2) or their combination with rpb1 (S1 Fig) or rpb1

28

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi and act (S2 Fig) were used for above genus level phylogenetic inferences. Species level phylogenetic inferences were based on all five generated loci. DNA sequences were aligned using the online version of Mafft v.7 (Katoh and Standley 2013) adopting the iterative refinement algorithms L-INS-I for rpb1-, act-, and ß-tub gene regions and Q-INS-i for LSU and ITS. The start and end of the alignments were trimmed manually in Se-Al v2.0 (Rambaut 1996). The alignments were deposited in TreeBASE, and are available at (http://purl.org/phylo/treebase/phylows/study/TB2:S20879). Based on these alignments, Bayesian Metropolis coupled Markov chain Monte Carlo analyses were done with MrBayes v3.2 (Huelsenbeck and Ronquist 2001; Ronquist and Huelsenbeck 2003). The general time-reversible model with the addition of invariant sites and a gamma distribution of rates across sites (GTR+I+G) was selected as the best fitting substitution model according to both the hierarchical likelihood ratio test (hLRT) and the Akaike Information Criterion (AIC) implemented in MrModeltest v2.2 (Nylander 2004). Starting with a randomly selected tree, 1.000.000 (for the two- and five-gene-data set) 2.000.000 (for the three-gene data set) and 5.000.000 generations (for the LSU data set) were run, using flat prior distributions. Trees were sampled every 500 generations and 50% majority rule consensus trees were computed and a postieriori probabilities (pp) were estimated only from trees of the plateau, and after the split frequencies had fallen below 0.01. All other trees were discarded as “burnin”. The estimations were thus based on 1600 (two- and five-gene data set), 2300 (three-gene data set) and 7000 (LSU) trees sampled. Maximum likelihood (ML) analyses were performed using RAxML 7.2.8 (Silvestro and Michalak 2012; Stamatakis 2014) implemented in Geneious 8.1.2 applying the general time-reversible (GTR) substitution model with gamma model of rate heterogeneity and 100 replicates of rapid bootstrapping (reported as MLB values). Neighbor joining analysis (Saitou and Nei 1987) was done in PAUP 4.0b10 in the batch file mode (Swofford 2002) applying the Kimura two-parameter model of DNA substitution (Kimura 1980) with a transition/transversion ratio of 2.0 to compute genetic distances. Support for internal nodes was estimated by 1000 bootstrap replicates (Felsenstein 1985) (reported as NJB values). The phylogenetic trees were visualised using FigTree v. 1.4.2

(http://tree.bio.ed.ac.uk/software/figtree).

29

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Table 2.1- Isolates and accession numbers used in the phylogenetic analyses

GenBank accession numbers Species Isolate number Host / substrate Locality Reference act ITS LSU rpb1 tub Aschersonia placenta BCC 7957 scale insect - - - DQ518753 DQ522364 - (Spatafora et al. 2007) Balansia henningsiana GAM 16112 Panicum sp. Georgia - - AY489715 AY489643 - (Castlebury et al. 2004) Balansia pilulaeformis AEG 94-2 Poaceae - - AF543788 DQ522365 - (Spatafora et al. 2007) Bionectria byssicola CBS 914.97 = G.J.S. 95-131 Branches of Alchornea Uganda GQ505962 - GQ506011 GQ506040 - (Hirooka et al. 2010) Bionectria compactiuscula CBS 592.93 = G.J.S. 93-27 - France GQ505963 - GQ506007 GQ506036 - (Hirooka et al. 2010) Bionectria epichloe CBS 118752 - - - DQ363259 - - (Jaklitsch and Voglmayr 2011) Bionectria grammicospora G.J.S. 85-218 - Indonesia - - AF193238 - - (Rossman et al. 2001) CBS 125111 Palm branch Costa Rica GQ505964 - GQ506009 GQ506038 - (Hirooka et al. 2010) Bionectria pityrodes ATCC 208842 On bark Mauritius - - AY489728 AY489658 - (Castlebury et al. 2004) Bionectria setosa CBS 834.91 Throphis racemosa Cuba - - AF210670 - (Schroers 2001) Bionectria vesiculosa HMAS 183151 Decaying leaves of a China - - HM050302 - - (Luo and dicotyledonous plant Zhuang 2010) Bryocentria brongniartii M190 - UK - - EU940125 - - (Stenroos et al. 2010) Bryocentria metzgeriae M140 - Germany - - EU940106 - - (Stenroos et al. 2010) morganii ATCC 11614 Crown canker on - - - U17409 - - (Jaklitsch and multiflora roses Voglmayr 2011) Chaetopsinectria Voucher 83362 - - - - DQ119554 - - (Jaklitsch and chaetopsinae Voglmayr 2011) Clonostachys pityrodes ATCC 208842 = G.J.S. 95-26 On bark Mauritius - - AY489728 AY489658 - (Castlebury et al. 2004) Cosmospora coccinea CBS 114050 = A.R. 2741 Inonotus nodulosus Germany GQ505967 - GQ505990 GQ506020 HM484589 (Hirooka et al. 2010; Hirooka et al. 2011)

30

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Cosmospora episphaeria G.J.S. 88-29 - - - - AY015625 - - (Jaklitsch and Voglmayr 2011) Cosmospora magnusiana CBS 129430 = A.R. 4453 Rhus typhina USA JF832441 - JF832680 JF832764 JF832839 (Hirooka et al. 2012) Cosmospora vilior CBS 126109 = G.J.S. 90-217 Xylaria sp. GQ505965 - GQ506010 GQ506039 JF832840 (Hirooka et al. 2010; Hirooka et al. 2012) Cosmospora vilior G.J.S. 96-186 - - - - AY015626 - - (Jaklitsch and Voglmayr 2011) Cosmospora viliuscula CBS 455.96 = G.J.S. 96-6 Xylaria sp. Puerto Rico GQ505966 - GQ506003 GQ506032 HM484876 (Chaverri et al. 2011; Hirooka et al. 2010) Cosmospora wegeliniana CBS 128986 = G.J.S. 93-15 France GQ505968 - GQ506006 GQ506035 HM484878 (Chaverri et al. 2011; Hirooka et al. 2010) Dialonectria episphaeria G.J.S. 10-193 stigma USA - - KC291771 KC291892 KC291932 (Herrera et al. 2013) Emericellopsis glabra CBS 125295 = A.R. 3614 Soil Mexico GQ505969 - GQ505993 GQ506023 HM484879 (Chaverri et al. 2011; Hirooka et al. 2010) Emericellopsis maritima CBS 491.71 = AFTOL-ID 999 See water Ukraine - - FJ176861 - - (Jaklitsch and Voglmayr 2011) Gliocephalotrichum ATCC 22228 Soil Louisiana, USA - - AY489732 AY489664 - (Castlebury bulbilium et al. 2004) Gliomastix masseei CBS 794.69 Dung of rabbit - - HQ232060 - - (Summerbell et al. 2011) CBS 119.67 Camarophyllus niveus Netherlands - - HQ232066 - - (Summerbell et al. 2011) Gliomastix polychroma CBS 181.27 Hevea brasiliensis, bark Sumatra - - HQ232091 - (Summerbell et al. 2011) Gliomastix roseogrisea CBS 279.79 Unknown Switzerland - - HQ232122 - - (Summerbell et al. 2011) Gliomastix tumulicola K5916-10-3 Viscous substances on Japan - - AB540476 - - (Kiyuna et al. stone wall 2011) Heleococcum aurantiacum CBS 201.35 Mushroom compost Unknown - - JX158441 - - (Grum- Grzhimaylo et al. 2013)

31

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Heleococcum japonicum CBS 397.67 = ATCC 18157 Wood panel of Abies firma Japan - - U17429 - - (Jaklitsch and Voglmayr 2011) Hydropisphaera CBS 340.77 Dead leaf of Astelia - - EU289204 - - (Platas et al. multiloculata 2008) Hydropisphaera CBS 336.77 Phormium tenax, Leaf New Zealand - EU289205 - - (Platas et al. multiseptata 2008) Hydropisphaera peziza CBS 102038 On bark Alabama - - AY489730 AY489661 - (Castlebury et al. 2004) Hydropisphaera sp. CBS 122304 = A.R. 4170 Decaying leaves of Idaho GQ505970 GQ505995 GQ506025 HM484877 (Chaverri et (fungicola) Populus trichocarpa al. 2011; Hirooka et al. 2010) Hydropisphaera suffulta CBS 136679 = Pyrenomycetes on Piper Martinique - - KU237206 - - Direct CLLMAR13069 dilatatum submission Hypocrella nectrioides G.J.S 89-104 scale insect - - - DQ518772 - - (Spatafora et al. 2007) Ijuhya antillana CBS 122797 = CLL 7321 Dead inflorescence of Martinique KY607565 KY607537 KY607552 KY607578 KY684186 This study Heliconia caribaea Ijuhya chilensis CBS 102803 Dead leaf Texas; USA KY607566 KY607538 KY607553 KY607579 KY684187 This study Ijuhya corynospora CBS 342.77 = G.J.S. 74-135 Phormium tenax, dead New Zealand KY607567 KY607539 KY607554 KY607580 KY684188 This study leaf Ijuhya dentifera CBS 574.76 = G.J.S. 74-43 Dacrydium cupressinum, New Zealand KY607568 KY607540 KY607555 KY607581 KY684189 This study bark Ijuhya faveliana CBS 133850= Palm French Guiana KY607569 KY607541 KY607556 KY607582 KY684190 This study CLLGUY12049 Ijuhya faveliana CLLG10007 Astrocarium sp., dead leaf French Guiana - - KX950705 - - (Lechat and Fournier 2017) Ijuhya fournieri CLLG12002 Astrocarium sp., dead leaf French Guiana - - KR105614 - - (Lechat et al. 2015) Ijuhya fournieri CBS 128283 = CLLG10113 Astrocarium sp., dead leaf French Guiana - - KP899118 - - (Lechat et al. 2015) Ijuhya lilliputiana CLLG12015B(LIP) Palm, dead leaves French Guiana - - KX950703 - - (Lechat and Fournier 2017) Ijuhya oenanthicola CBS 129747 = CLL10046 Oenanthe crocata France KY607570 KY607542 KY607557 KY607583 KY684191 This study Ijuhya pachydisca CLLG12001B Palm, dead leaves French Guiana - - KX950701 - - (Lechat and Fournier 2017) Ijuhya pachydisca CLLG12001B Palm, dead leaves French Guiana - - KX950704 - - (Lechat and Fournier 2017)

32

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Ijuhya paraparilis MAFF241404/TUAh52 - Japan GQ505971 - GQ506012 GQ506041 - (Hirooka et al. 2010) Ijuhya paraparilis W8063/HMAS 183506 - China - FJ969801 HM050303 - FJ969803 (Luo and Zhuang 2009) Ijuhya parilis CBS 136677 = CLL13022 Genista sp. Spain KY607571 KY607543 KY607558 KY607584 KY684192 This study Ijuhya peristomialis CBS 569.76 = G.J.S. 73-314 rachis of Cyathea New Zealand KY607572 KY607544 KY607559 KY607585 KY684193 This study dealbata Ijuhya tetraspora CBS 140721 Humulus lupulus - - KX950706 - - (Lechat and Fournier 2017) Ijuhya vitellina 72723 (36_1G) Heterodera filipjevi, egg Turkey KY607561 KY607532 KY607546 KY607574 KY684181 This study Ijuhya vitellina 72825 (37AD) Heterodera filipjevi, egg Turkey KY607562 KY607533 KY607547 KY607575 KY684182 This study Ijuhya vitellina 72934 (37T) Heterodera filipjevi, egg Turkey - KY607534 KY607548 - KY684183 This study Ijuhya vitellina DSM 104494 (41E) Heterodera filipjevi, egg Turkey KY607563 KY607535 KY607549 KY607576 KY684184 This study Ijuhya vitellina DSM 104495 (42DD) Heterodera filipjevi, egg Turkey KY607564 KY607536 KY607550 KY607577 KY684185 This study Ijuhya vitellina 12-42-1e H. filipjevi egg, uncultured Turkey - - KY607551 - - This study Ijuhya vitellina 72918 (YE3T) Heterodera filipjevi, egg Turkey KY607560 KY607531 KY607545 KY607573 KY684180 This study Kallichroma glabrum JK5123 - - - AF193233 - - (Rossman et al. 2001) Lanatonectria flocculenta CBS 126441 = G.J.S. 01-66 Bark Ecuador JF832481 - JF832713 - JF832913 (Hirooka et al. 2012) Lasionectria mantuana CBS 114291 = A.R. 4029 Decortivated wood Finland - HM484858 GQ505994 GQ506024 - (Chaverri et al. 2011; Hirooka et al. 2010) Lasionectria marigotensis CBS 131606 = Cocos nucifera, decayding French West - KR105612 KR105613 - - (Lechat and CLLGUAD11002 leaf Indies Fournier 2013) Lasionectria sylvana CBS 566.76 Cyathea smithii New Zealand - - EU289206 - - (Platas et al. 2008) Moelleriella libera P.C.672 = CUP 067773 - Honduras - - EU392594 EU392717 - (Chaverri et al. 2008) Mycoarachis inversa ATCC 22107 = A.R. 2745 Elephant dung Uganda GQ505972 - GQ505991 GQ506021 HM484882 (Chaverri et al. 2011; Hirooka et al. 2010) aurantiaca CBS 308.34 Ulmus sp. UK JF832482 - JF832682 JF832766 JF832886 (Hirooka et al. 2012) Nectria austroamericana CBS 126114 = A.R. 2808 Gleditsia triacanthos USA GQ505960 - GQ505988 GQ506016 HM484597 (Chaverri et al. 2011; Hirooka et al. 2010)

33

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Nectria balansae CBS 123351 = A.R. 4446 Coronilla sp. France GQ505977 - GQ505996 GQ506026 HM484607 (Hirooka et al. 2010; Hirooka et al. 2011) Nectria berberidicola CBS 128669 = A.R. 4662 Berberis vulgaris France JF832487 - JF832712 JF832767 JF832887 (Hirooka et al. 2012) Nectria berolinensis CBS 127382 Ribes sanguinea Austria - - HM534893 - - (Jaklitsch and Voglmayr 2014) Nectria cinnabarina CBS 255.47 = ATCC 11432 Stem of Ulmus sp. Netherlands GQ505975 - GQ505997 GQ506027 HM484832 (Hirooka et al. 2010; Hirooka et al. 2011) Nectria cucurbitula CBS 259.58 Pinus sylvestris Netherlands GQ505974 - GQ505998 GQ506028 HM484592 (Chaverri et al. 2011; Hirooka et al. 2010) Nectria cyanostoma CBS 101734 = G.J.S. 98-127 Buxus sempervirens France GQ505961 - FJ474081 GQ506017 HM484611 (Chaverri et al. 2011; Hirooka et al. 2010) Nectria haematococca CBS 114067 On bark Guyana - - AY489729 AY489660 - (Castlebury et al. 2004) Nectria lamyi CBS 115034 = A.R. 2779 Berberis vulgaris Austria HM484507 - HM484569 HM484582 HM484593 (Hirooka et al. 2011) Nectria pseudotrichia CBS 551.84 Bark Japan GQ505976 - GQ506000 GQ506030 HM484602 (Hirooka et al. 2011) Nectria pseudotrichia CBS 641.83 Wood Venezuela - - HM534899 - - (Jaklitsch and Voglmayr 2011) Nectria sinopica CBS 462.83 Hedera helix Netherlands GQ505973 - GQ506001 GQ506031 HM484595 (Hirooka et al. 2011) Nectricladiella camelliae CBS 111794 = ATCC 38571 Fruit of tree Australia - - AY793432 - - (Jaklitsch and Voglmayr 2011) Nectriopsis epimycota CBS 127459 = G.J.S. 95-94 Pyrenomycete Puerto Rico GQ505978 - GQ506008 GQ506037 - (Hirooka et al. 2010) Nectriopsis exigua CBS 126110 = G.J.S. 98-32 Myxomycete Puerto Rico GQ505979 - GQ505986 GQ506014 HM484883 (Chaverri et al. 2011; Hirooka et al. 2010) Nectriopsis violacea CBS 424.64 Fuligo septica Germany - - AY489719 AY489646 - (Castlebury et al. 2004) vasinfecta JP963 - - - - U17406 - - (Jaklitsch and Voglmayr 2011)

34

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Neonectria coccinea CBS 119158 = G.J.S. 98-114 Fagus sp. Germany KC660422 - KC660620 KC660672 KC660727 (Hirooka et al. 2013) Neonectria ditissima CBS 100316 Malus domestica Ireland HM352880 - HM364311 HM364330 HM352864 (Chaverri et al. 2011) Neonectria punicea MAFF241548 =TPP-h328 Twigs Japan KC660372 - KC660569 KC660637 KC660713 (Hirooka et al. 2013) Neonectria veuillotiana CBS 125114 = G.J.S.92-24 Fagus sylvatica France GQ505980 - GQ506005 GQ506034 JQ394725 (Hirooka et al. 2010; Salgado- Salazar et al. 2012) Neonectria westlandica CBS 112464 = G.J.S. 83-156 Dacrydium cupressinum New Zealand GQ505959 - GQ505987 GQ506015 HM484610 (Hirooka et al. 2011) Nigrosabulum globosum ATCC 22102 Cow dung USA: - - AF096195 - - (Jaklitsch and Wyoming Voglmayr 2011) Ochronectria calami CBS 125.87 On palm Indonesia - - AY489717 AY489644 - (Castlebury et al. 2004) Ochronectria thailandica MFLUCC 15-0140 - - - KU564069 - - Direct submission Ophionectria trichospora CBS 109876 = G.J.S. 01-206 Bark Cameroon - - AF543790 AY489669 HM484886 (Chaverri et al. 2011) Paracylindrocarpon aloicola CBS 141300 = CPC 27362 Leaves and twigs of Aloe South - - KX228328 - - (Crous et al. sp. 2016) Persiciospora africana ATCC64691 Forest soil Botswana - - AY015631 - - (Jaklitsch and Voglmayr 2011) Pleonectria aquifolii CBS 307.34 Ilex aquifolium UK JF832444 - JF832718 JF832792 JF832842 (Hirooka et al. 2012) Pleonectria coryli CBS 129358 = A.R. 4583 Corylus avellana France JF832476 - JF832740 JF832797 JF832872 (Hirooka et al. 2012) Protocreopsis korfii CBS138733 = CLLM14077 - - KT852955 Direct submission Protocreopsis pertusa C.T.R. 72-184 Decaying palm Venezuela GQ505981 - GQ506002 - - (Hirooka et al. 2010) Pseudonectria CBS 128674 = A.R. 4592 Pachysandra sp. USA JF832512 - JF832715 JF832791 JF832909 (Hirooka et pachysandricola al. 2012) Pseudonectria rousseliana CBS 114049 Buxus sempervirens, leaf Spain - - U17416 AY489670 - (Castlebury et al. 2004) Roumegueriella rufula G.J.S.91-164 Globodera rostochiensis - - - EF469082 EF469099 - (Sung et al. 2007) Roumegueriella rufula CBS 346.85 Globodera rostochiensis Switzerland - - GQ505999 GQ506029 - (Hirooka et al. 2010) Selinia pulchra A.R. 2812 Cow dung Argentina GQ505982 - GQ505992 GQ506022 HM484884 (Chaverri et al. 2011;

35

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Hirooka et al. 2010) Stephanonectria keithii CBS 114057 Eleagnus sp., bark France - - AY489727 AY489657 - (Castlebury et al. 2004) Stilbocrea macrostoma CBS 114375 = G.J.S. 73-26 Geniostoma ligustifolia New Zealand - - AY489725 AY489655 - (Castlebury et al. 2004) Stilbocrea macrostoma G.J.S. 02-125 - Sri Lanka GQ505983 - GQ506004 GQ506033 - (Hirooka et al. 2010) Stromatonectria caraganae CBS 125579 Branches of Caragana Austria - HQ112288 HQ112288 - HQ112289 (Jaklitsch and spp. Voglmayr 2011) Stromatonectria caraganae CBS 127387 Branches of Caragana Austria - HQ112287 HQ112287 - - (Jaklitsch and spp. Voglmayr 2011) Verrucostoma freycinetiae MAFF240100/h523 Freycinetia boninensis Japan GQ505984 - GQ506013 GQ506018 HM484885 (Chaverri et al. 2011; Hirooka et al. 2010) Viridispora alata CBS 125123 = A.R. 1770 Bark Madeira GQ505985 - GQ505989 GQ506019 JF832912 (Hirooka et al. 2012) Viridispora diparietispora CBS 102797=ATCCMYA627 Crataegus crus-galli USA - - AY489735 AY489668 - (Castlebury et al. 2004) AEG: A. E. Glenn personal collection; ATCC: American Type Culture Collection, Manassas, VA, USA; A.R.: Amy Y. Rossman, USDA-ARS MD USA; BCC: BIOTEC Culture Collection, Pathum Thani, Thailand; CBS: Westerdijk Fungal Biodiversity Institute (Utrecht, The Netherlands); C.L.L.:Christian Lechat, Ascofrance, Villiers en Bois, France.; C.T.R.: Clark T. Rogerson, The New York Botanical Garden, NY, USA; DSM: The open collection of the Leibniz-Institut DSMZ- Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH; GAM: Julian H. Miller Mycological Herbarium Athens, GA, USA; G.J.S.: Gary J. Samuels, USDA-ARS MD USA; MAFF: MAFF Genebank, National Institute of Agrobiological Sciences, Ibaraki, Japan; HMAS: The mycological Herbarium, Institute of Microbiology, Chinese Academy of Sciences, China; PC: Herbier Cryptogamique, Départment de Systématique et Évolution, Muséum National d’Histoire Naturelle, Paris, France.

36

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

2.2.10 Nomenclature

The electronic version of this article in Portable Document Format (PDF) in a work with an ISSN or ISBN will represent a published work according to the International Code of Nomenclature for algae, fungi, and plants, and hence the new names contained in the electronic publication of a PLOS article are effectively published under that Code from the electronic edition alone, so there is no longer any need to provide printed copies. In addition, new names contained in this work have been submitted to MycoBank from where they will be made available to the Global Names Index. The unique MycoBank number can be resolved and the associated information viewed through any standard web browser by appending the MycoBank number contained in this publication to the prefix http://www.mycobank.org/MB/. The online version of this work is archived and available from the following digital repositories: PubMed Central, LOCKSS.

2.2.11 Metabolite profiling

2.2.11.1 Fermentation and extraction of cultures Three different liquid media (Q6/2, YM and ZM; (Kuhnert et al. 2015)) were used for the initial screening of fermentation in 500 mL Erlenmeyer flasks each containing 200 mL medium. Inoculum consisted of few 5-mm-diam. culture discs of strain DSM 104495 excised from PDA. The submerged cultures were incubated in the dark at 23°C and 140 rpm and harvested two days after sugars were depleted. Secondary metabolites were extracted from both mycelium and culture filtrate following methods described by Kuhnert et al. (2015). For large-scale fermentation, the above mentioned strain was inoculated into 3 L of the selected medium (see below) and processed for extraction similarly.

2.2.11.2 Selection of the liquid culture for scale-up culturing Minimum inhibitory concentration (MIC) tests were performed to determine the optimum medium with the highest antimicrobial activity in crude extracts. The cultural medium showing highest MIC activity was chosen for scale-up fermentation, accordingly. The crude extracts obtained from the examined cultural media i.e. Q6/2, YM and ZM were tested following Chepkirui et al. (2016).

2.2.11.3 Isolation of secondary metabolites The EtOAc organic extract (110 mg) was dissolved in MeOH and purified using preparative RP-HPLC

[column 250 × 20 mm, Kromasil C18, 7 µm; equipped with a Kromasil C18 pre-column 50 x 20 mm, 7

µm]. Solvent A: H2O; solvent B: acetonitrile; gradient: 50% B increasing to 80% B in 40 min, increasing to 100% B in 5 min, holding at 100% B for 10 min; flow rate 20 mL/min, UV detection at 230, 254, and 325 nm], yielded 2.3 mg of compound 1 and 2 mg of compound 2. The compounds were eluted at 32 min and 43 min, respectively. 37

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

2.2.11.4 Structure elucidation 1D and 2D NMR spectra were recorded on a Bruker Avance III 700 spectrometer with a 5 mm TXI cryoprobe (1H 700 MHz, 13C 175 MHz) spectrometer; optical rotations were measured on a Perkin- Elmer 241 polarimeter. All HPLC-MS analyses were performed on Agilent 1260 Infinity Systems with diode array detector and C18 Acquity UPLC BEH column (2.1 x 50 mm, 1.7 μm) from Waters with the gradient described by Helaly et al. (2016), combined with ion trap MS (amazon speed, Bruker); and HR- ESIMS spectra on a time-of-flight (TOF) MS (Maxis, Bruker). Chemicals and solvents were obtained from AppliChem GmbH, Avantor Performance Materials, Carl Roth GmbH & Co. KG and Merck KGaA in analytical and HPLC grade.

2.2.11.5 Nematode bioassays and cytotoxicity A biological assay was conducted to evaluate nematicidal activity of pure compounds against Caenorhabditis elegans and H. filipjevi according to Helaly et al. (2016). Surface-sterilised cysts of H. filipjevi propagated in the greenhouse were incubated in sterile tap water under aseptic conditions for nematode hatching. The freshly hatched second stage juveniles (J2) were collected and used for the experiment. Caenorhabditis elegans was cultivated as described by Helaly et al. (2016). The number of nematodes was adjusted to 600/mL of J2 of H. filipjevi and 600/mL of adults and juveniles of C. elegans in sterile tap water. The assays were carried out in 24-well microtiter plates. Each well received 1 mL of nematode suspension. The compounds were tested against nematodes at the concentrations of 100, 50, 20 and 10 µg/mL in MeOH. Each treatment included three replications. Ivermectin (Sigma- Aldrich) was used as positive and MeOH in DMSO (v/v) as negative control. Nematodes were monitored for 30 min after inoculation and afterwards plates were incubated at 24 °C for 18 h.

Cytotoxicity (IC50) of compounds was tested against different cell lines as described by Richter et al. (2016).

2.3 Results

2.3.1 Fungal isolation

Among other nematode egg colonizing fungi to be described elsewhere, Ijuhya vitellina, newly described below, was encountered. It rendered nematode cysts collected from fields in Turkey reddish dotted upon microscopic examination (Fig 2.1A). Reddish dots inside cysts consisted of nematode eggs each containing one or few microsclerotia (Figs 2.1B and 1C). In some infected eggs, microsclerotial tissues were found developing inside juveniles (Figs 2.1D and 1E). When inoculated on agar medium (PDA), mycelium emerged from symptomatic nematode eggs and developed reddish orange, brick red or reddish brown cultures (Fig 2.1F). Predominantly globose or ellipsoidal, reddish microsclerotia were

38

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi formed on the surface of (Fig 2.1G) or submerged in the medium. Microsclerotia formed in vitro resembled the structures encountered in nematode eggs. Cultures growing from individually inoculated nematode eggs or in vitro-produced microsclerotia developed slowly and reached a diameter of 2.8 cm within 3 months (Figs 2.1F and 1H and 1I). The sterility check revealed no fungal growth from the examined cysts.

Figure 2.1- Cysts and eggs of Heterodera filipjevi naturally infected with Ijuhya vitellina, and pure cultures obtained from the infected eggs. (A) Symptomatic, reddish dotted nematode cysts. (B, C) Nematode eggs accommodating reddish microsclerotia. (D, E) Microsclerotial tissue developing inside juveniles. (F) A six-month- old culture that developed from a single infected nematode egg. (G) Surface of colony showing reddish microsclerotia arranged in concentric rings. (H, I) Two-month-old cultures on PDA and CMA. Scale bars: A = 0.5 mm, B = 30 µm, C-E = 50 µm, F = 1 cm (also applying for H, I), G = 400 µm.

2.3.2 Molecular phylogenetic studies

2.3.2.1 DNA sequence comparisons and culture-independent identification LSU sequences were obtained for all 14 studied strains of the fungus. Four of these LSU sequences were obtained from environmental specimens (individual nematode eggs showing reddish 39

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi microsclerotia) were identical to those retrieved from pure cultures. One (2, 2, 3) nucleotide substitutions were observed among retrieved act (ITS, ß-tub, rbp1) sequences. Most substitutions were observed in strain 37AD. BLASTn searches indicated relatedness of the encountered fungus to the Bionectriaceae. More specifically, the LSU sequence of I. vitellina was most similar to that of Ijuhya paraparilis and its ITS sequence to that of Stromatonectria caraganae according to initial searches in GenBank.

2.3.2.2 Phylogenetic reconstructions Alignment of 112 LSU sequences representing 100 taxa of Nectriaceae and Bionectriaceae was 860 sites long. That of 66 combined LSU (844 sites) and rpb1 (682 sites) sequences comprised 60 taxa, and that of 70 combined LSU (848), act (607) and rpb1 (684) sequences represented 64 taxa. One alignment of 15 strains representing 10 species of the genus Ijuhya only was based on all five gene regions sampled: act (624), ITS (936), LSU (819), rpb1 (712) ß-tub (875). Strains of the nematode parasite were highly supported as a monophyletic species group in all analyses, and clustered within a highly supported clade together with other species of the bionectriaceous genus Ijuhya (Fig 2.2 and S1 and S2 Figs) including I. peristomialis, a later synonym of I. vitrea, which is the type species of Ijuhya (Rossman et al. 1999). Relatedness of Ijuhya with selected taxa of the Bionectriaceae, including Bionectria, the type genus of this family and various others was also highly supported. Phylogenetic analyses based on LSU only, LSU/rpb1 and LSU/rpb1/act suggested that Ijuhya oenanthicola, I. dentifera and I. antillana are distantly related to Ijuhya sensu stricto. Instead, they display closer phylogenetic affinities to Lasionectra and Ochronectria (Fig 2.2 and S1 and S2 Figs). All five loci were sampled for the Ijuhya species available as cultures to us. Hypotheses on the intra- generic phylogenetic relationships of representatives of Ijuhya were derived using these sequences. In addition two specimens of I. paraparilis from GenBank, for which three of these five gene regions were available, were added to this data set. Based on the results of the larger phylogenies, the ‘Ijuhya’ species distantly related to the Ijuhya sensu stricto were used as outgroup here. Two highly supported subclades are suggested for the in-group of genus Ijuhya, of which one includes the type species I. peristomialis, and in addition I. chilensis, I. faveliana, I. paraparilis, and I. parilis (Fig 2.3). The other subclade includes I. vitellina and its closest sister species, I. corynospora (Fig 2.3). This is in concordance with the phylogenetic hypotheses obtained from the larger two- and three-gene data sets (S1 and S2 Figs). The two isolates of I. paraparilis originating from Japan and China, did not form a species clade and are thus unlikely conspecific.

40

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Figure 2.2- Bayesian inference of phylogenetic relationships of selected taxa of the Bionectriaceae and Nectriaceae (Hypocreales) based on LSU sequences. Numbers above nodes are estimates of a posteriori probabilities (≥ 0.9) / NJB and MLB values (≥70%). The topology was rooted with Aschersonia placenta, Balansia henningsiana, B. pilulaeformis, and Hypocrella nectrioides, (Hypocreales).

41

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Figure 2.3- Bayesian inference of infrageneric phylogenetic relationships within Ijuhya based on act, ITS, LSU, rpb1, and ß-tub sequences. Numbers above nodes are estimates of a posteriori probabilities (≥ 0.9) / NJB and MLB (≥ 70%). The topology was rooted with three distantly related ‘Ijuhya’ species (‘Ijuhya’ antillana, I. dentifera, and ‘Ijuhya’ oenanthicola).

2.3.3 Taxonomy

Ijuhya vitellina Ashrafi, W. Maier & Schroers, sp. nov., Mycobank MB 821493 Holotype for Ijuhya vitellina (here designated) (MB 821493): Turkey, Yozgat, experimental wheat field: dried culture on SNA with carnation leaf pieces, originating from an individual egg from a cyst of Heterodera filipjevi, isolated by Samad Ashrafi, August 2013 (B 70 0016479, deposited at the herbarium of the Botanic Garden and Botanical Museum Berlin-Dahlem). Ex-holotype strain: DSM 104494, deposited in the open collection of the Leibniz-Institut DSMZ- Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH). Ex-holotype sequences: act: KY607563; ITS: KY607535; LSU: KY607549; rpb1: KY607576; tub2: KY684184. Additional material examined, from the same location: DSM 104495 (dried culture, B 70 0016480), GenBank accession number: act: KY607564; ITS: KY607536; LSU: KY607550; rpb1: KY607577; tub2: KY684185. For other material studied, see Table 2. 1. Etymology: From Latin vitellus meaning egg yolk, referring to the colour and shape of microsclerotia formed by the species in nematode eggs. 42

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Naturally infected eggs typically accommodating one, occasionally two globose to subglobose, fulvous (brownish-yellow, reddish yellow, yolk-coloured) multicellular microsclerotia with a textura angularis appearance, similar to microsclerotia formed in culture. Colonies slow in growth; at 20°C on PDA, 6.5–9 mm diam. after 7 d, 13–16 mm after 14 d; optimum temperature for growth around 25°C, 9–12 mm (7 d), 16–19 mm (14 d); at 30°C 7–8 mm (7 d), 10–13 mm (14 d); no growth observed at 35°C. Colony reverse on OA after 21 d reddish orange (7A7), brick red to burnt Sienna (7B–D7–8) to dark brown (7F7–8) in central parts of colonies; on CMA and SNA covered with carnation leaf pieces Sahara (6C5), brick red or burnt Sienna (7B–D7–8); on up to 12 months old PDA caramel brown to brownish orange (6C6–8), cognac brown (6E7), brownish orange, brick red, copper red (7C–E7–8), or dark brown (7F6). Colony surface of similar pigmentation as reverse, granular because of solitary, gregarious or clustered microsclerotia formed on the surface of or submerged in the agar media, often arranged in concentric rings. Aerial mycelium on PDA within 3 wk not observed or sparsely to abundantly produced in sectors, white, felty to wet-cottony, on SNA and SNA with carnation leaf pieces absent or present as occasionally formed solitary, erect, typically apically coiling hyphae. Conidiophores and conidia not observed. Microsclerotia typically ellipsoidal to cylindrically oblong, sometimes globose, orange to brownish orange or brick red, not changing colour in KOH, on 6 wk-old OA 23–51 × 26–66 (36 × 44) µm (n= 52), on 6 wk-old PDA 25–46 × 32–58 (35 × 43) µm (n= 32), on ca. 12 months-old PDA cultures 26–58 × 34–77 (42 × 52) µm (n= 63). Cells of microsclerotia angular, forming a textura angularis in surface and subsurface view, variable in size, on 21 d old OA colonies 3.5–7.0 × 5.0–9.0 (5.0 × 7.0) (n= 41), on 21 d old PDA colonies 3.1–6.0 × 4.0–8.0 (5.0 × 6.0) µm (n= 50), on 12 month-old PDA colonies 3.0–7.0 × 4.5–11.0 (5.0 × 7.0) µm (n= 74) µm; walls first hyaline, later orange to brownish orange, 1.0–2.0 µm (n= 70). Microsclerotia developing from intercalary cells of hyphae or terminally at side branches, solitary or moniliform, first chlamydospore or dictyochlamydospore-like (Figs 2.4A-4D and 5A-5C), cells angular, pigmented, first pale luteous, then brick reddish (Figs 2.4K-4N), filled with small guttules. Dictyochlamydospore-like structures may enlarge through coiling or expand to form globose to ellipsoidal microsclerotia (Figs 2.4E and 4F and 5D-5F) formed solitary, in chains (Figs 2.4J and 5G), or clusters. Globose and ellipsoidal microsclerotia developing in culture reminiscing microsclerotial structures of the fungus encountered in field-collected nematode eggs. Culturing of single microsclerotia directly extracted from nematode eggs, or retrieved from one-year old wheat straw cultures resulted in fungal growth (Fig 2.4O). Teleomorph not observed.

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Figure 2.4- Light micrographs of Ijuhya vitellina, formation of microsclerotia. (A-F) Transformation of hyphae into (A-D) chlamydospore or dictyochlamydospore-like structures, and (E, F) microsclerotia. (G-I) Coiling or coalescence of dictyochlamydospore-like structures. (J) Microsclerotia densely arranged in a chain. (K-N) Pigmentation first observed (K) in cell walls, and later (L-M) intensifying throughout microsclerotia. (O) A single microsclerotium inoculated on agar surface developing hyphae. A-I, K-N: from PDA, J: from CMA, O: from PDA 1/3. Scale bars: (A, C, E-I, K, L, O) = 30 µm; B, J = 50 µm; D = 200 µm; (M, N) = 10 µm.

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Figure 2.5- SEM micrographs of microsclerotia formed by Ijuhya vitellina. (A) Filamentous hyphae developing into multicellular structures. (B) Intercalary formed dictyochlamydospores connected by hyphae (arrowed). (C) Detail of intercalary multicellular structures of microsclerotia. (D, E) Hyphae transformed into chlamydospore- like structures and microsclerotia. (F) Terminally formed microsclerotium. (G) Moniliform arrangement of microsclerotia. (H) Detail of microsclerotia illustrating a multicellular surface that forms a textura angularis. Scale bars: A = 100 µm; (B, E-H) = 50 µm; (C, D) =30 µm.

2.3.4 In vitro parasitism on nematode eggs and Koch’s postulates

Ijuhya vitellina infected eggs of H. filipjevi in vitro. Eggs in healthy cysts placed on colonies of I. vitellina became parasitized by hyphae within four weeks (Fig 2.6A). Hyphal cells inside eggs enlarged and 45

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi destroyed unembryonated eggs or developing juveniles (Fig 2.6B). Eggs were entirely occupied by compartmentalised thick-walled hyphae (Fig 2.6C), and guttules-filled moniliform chlamydospores (Fig 2.6D) that eventually developed into microsclerotia (Figs 2.6E-6G). Such microsclerotia were observed within 4–5 weeks after cysts or single eggs were placed on fungal colonies; they measured 26–66 x 31– 75 (41 x 49) µm (n=97) in eggs of 3-month-old infected cysts. Slide culture based observations revealed that individual eggs were infected within two weeks by hyphae that emerged from microsclerotia used as inoculum. Infection started with individual hyphae or appressorium-like structures that penetrated the eggshell and cuticle of developing juveniles (Fig 2.6H). Following penetration, similar infection processes and structures as described above, including swollen hyphal cells, thick-walled and multicellular structures filled with guttules, and subglobose or ellipsoidal microsclerotia were observed (Figs 2.6I-6L). Microsclerotia developing inside artificially infected eggs (Figs 2.6M and 6N) appearing textura angularis (cf. Fig 2.5H for details) were indistinguishable from those encountered in field-collected cysts (cf. Fig 2.1B and 1C).

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Figure 2.6- Light micrographs of the infection and colonisation process of Ijuhya vitellina in cysts and eggs of Heterodera filipjevi. (A) Symptomatic cyst, reddish-dotted due to eggs containing reddish, globose microsclerotia. (B-E) Early colonisation of nematode eggs by hyphae becoming chlamydospore- and dictychlamydospore-like to develop microsclerotia inside eggs. (F, G) Dictyochlamydospore-like structures and small microsclerotia. (H) Hyphae penetrating through the eggshell by forming appressorium-like structure (arrows). (I-K) Development of the fungus inside nematodes eggs: (I) Formation of thick-walled hyphal cells, later (J-K) transforming into microsclerotia. The arrow in (J) points at the nematode stylet; in (K) at immature microsclerotium. (L) Egg with mature microsclerotium. (M-N) Near-identical cells of microsclerotium formed in (M) egg and (N) pure culture, forming a textura angulari in optical sections. Material obtained from (B-G, M) infected cysts directly placed and incubated on fungal colony, (H-L) slide cultures, (N) OA. Scale bars: A = 300 µm; (B-N) = 30 µm.

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

2.3.5 Metabolite profiling

2.3.5.1 LC-MS analysis of the crude extracts Crude extracts obtained from Q6/2 medium showed highest antimicrobial activity. Therefore, this medium was chosen for up-scaling purposes. HPLC-UV chromatogram of the crude extract of I. vitellina revealed two major peaks at retention times 10.8 and 12.1 min. The peak at 10.8 min showed

+ - + molecular ion peaks at m/z 529.2 [M+H] , 527.2 [M-H] and 511.2 [M+H-H2O] . Accordingly, the molecular mass of compound 1 was determined as 528.2 g/mol. Similarly, the molecular mass of compound 2 was determined as m/z 570.2 g/mol (S3-S5 Figs). After scale-up fermentation, the crude extract was purified as described in the experimental section and obtained pure compounds were submitted to HRESIMS and NMR analysis for structure elucidation.

2.3.5.2 Structure determination of chaetoglobosins

HRMS analysis of compounds 1 and 2 revealed the molecular formulae C32H36N2O5 and C34H38N2O6, respectively. Comprehensive analysis of the 1D and 2D NMR data of 1 and 2 indicated that compound 1 is chaetoglobosin A (Sekita et al. 1973; Silverton et al. 1976) while compound 2 is its 19-O- acetylchaetoglobosin A (Probst and Tamm 1981) (Fig 2.7). The detailed description of the structure elucidation is included in the supporting information (S1 Text and S2 Text and S6 Fig).

Figure 2.7- Structures of chaetoglobosin A (1) and 19-O-acetylchaetoglobosin A (2).

2.3.5.3 Screening of Ijuhya spp. for chaetoglobosins and other secondary metabolites Chaetoglobosin A and its derivatives could not be found in eight other Ijuhya species. In the range of the retention times of the isolated chaetoglobosins (ca. 10-12 min) no related chaetoglobosins with similar masses and UV/Vis spectra (Sekita et al. 1982; Spöndlin and Tamm 1988) were detected.

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

2.3.5.4 Microtiter plate assay for nematicidal activities Chaetoglobosin A and 19-O-acetylchaetoglobosin A caused a temporary immobilisation of C. elegans and the second stage juveniles of H. filipjevi at 50 and 100 µg/mL. The immobilisation rate was higher at 100 µg/mL and for chaetoglobosin A. Both nematode species were immobilised shortly after having been exposed to the solutions. No nematicidal activities of the tested compounds were observed.

2.3.5.5 Cytotoxicity bioassay

Chaetoglobosin A (1) inhibited cell lines L929, KB3.1, PC-3 and HUVEC with IC50 values in µg/mL of 1.6, 0.15, 0.42 and 0.78, respectively. The acetyl derivative (2) showed inhibition against the same cell lines with IC50 values of 0.7, 0.19, 0.7 and 0.25 µg/mL, respectively.

2.4 Discussion

2.4.1 Phylogenetic analyses and systematic implications

Ijuhya vitellina is inferred as a new species on the basis of comparative morphological and molecular phylogenetic evidences. Phylogenetically, and supported by DNA sequences of five gene regions, the fungus occupies a distinct and highly supported monophyletic species clade nesting in the Ijuhya core group of the Bionectriaceae. Classification of the new species in genus Ijuhya is, however, purely based on phylogenetic evidence. While all other Ijuhya species are teleomorphically typified and largely characterized by morphological characters of ascomata and (Rossman et al. 1999; Samuels 1976; Samuels 1988), the teleomorph of I. vitellina is unknown. Nematode associated life-style has never been described for Ijuhya species before. Other members of the genus have so far been found on plant substrata. Ijuhya vitellina differs most clearly from other Ijuhya species by the formation of brightly coloured, orange to reddish microsclerotia that have not been described for any of the other Ijuhya species. Further phylogenetic analyses on the basis of alternative taxon selections and additional data are thus required to confirm monophyly of I. vitellina with Ijuhya or resolve I. vitellina outside Ijuhya sensu stricto rendering the description of an additional genus necessary. Ijuhya vitellina, based on our taxon sampling, is most closely related to I. corynospora described from dead leaves of Phormium tenax in New Zealand (Fig 2.2). However, also in this case no correlating characters supporting this sister group relationship can be found. Neither were chlamydospores or microsclerotia described for I. corynospora sensu stricto (Samuels 1978) nor did it parasitise nematode eggs in our in vitro experiments. Also I. vitellina does form chaetoglobosins, while no such metabolites were encountered in I. corynospora or in any other of the here studied Ijuhya species. Phylogenetic analyses suggest that I. antillana, I. dentifera, and I. oenanthicola are only distantly related to Ijuhya sensu stricto and not part of the genus. Morphological characters described for I. antillana (Lechat and

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

Courtecuisse 2010) and I. oenanthicola (Lechat and Hairaud 2012) conform well to the concept of the Nectria sylvana group (Samuels 1976), for which Lasionectria became the generic depository (Rossman et al. 1999). Specifically, fasciculate hyphal clusters formed by perithecia of I. antillana (Fig. 1, a, b, d in Lechat and Courtecuisse (2010)) and I. oenanthicola (Plate 1, Fig. A–C in Lechat and Hairaud (2012)) and occurrence of 1-septate striate ascospores are similar to those seen in Lasionectria sensu stricto. Accordingly, I. antillana and I. oenanthicola are combined into Lasionectria (Appendix).

2.4.2 Parasitism on nematode cysts and eggs

Our observations from field collected nematode cysts and various in vitro infection studies showed that I. vitellina parasitises cereal cyst nematodes. Several ascomyceteous fungal species have been reported to parasitise plant parasitic nematodes including cyst nematodes (Chen and Dickson 2004; Siddiqui and Mahmood 1996; Stirling 2014; Yang et al. 2012). Within the Bionectriaceae, to our knowledge only two species, Clonostachys rosea and Gliomastix murorum, were described as antagonists of animal- and plant parasitic nematodes (Ahmed et al. 2014; Kerry et al. 1982; Zhang et al. 2008). Hyphae of I. vitellina penetrate nematode eggs either directly or by developing an appressorium-like structure. We purport that hyphae of I. vitellina may similarly enter nematode cysts or juveniles. Upon penetration the fungus forms hyphae inside eggs as is also reported for other cyst nematode parasitic fungi (Stirling 2014). Penetration may apply through mechanic or chemical mechanisms (Dijksterhuis et al. 1991; Jansson and Lopez-Llorca 2001; Leger et al. 1991; Lopez-Llorca et al. 2002). Interactions involving chemical mechanisms are based on enzymes whose activities allow the penetration of the multilayered eggshell that in cyst nematodes mainly consists of chitin and lipids (Bird and McClure 1976; Curtis et al. 2011; Morton et al. 2004; Perry and Trett 1986; Stirling 2014). Hyphal penetrations by I. vitellina may thus involve similar strategies to invade eggs, juveniles and cysts of H. filipjevi. Hyphae coiling around and penetrating nematode eggs and filling the content of eggs have been described previously in various cases (Nordbring-Hertz et al. 2011). It is possible that hyphae directly use nutritional resources provided by nematode eggs for hyphal growth and mycelium development. The behaviour of I. vitellina inside nematode eggs differs drastically. Hyphae immediately swell and become transformed into globose or ellipsoidal dictyochlamydospores that develop into microsclerotia. Cells in these structures are angular and filled with guttules. Microsclerotia are rigid and readily resist mechanical manipulations. Additionally, guttules forming inside microsclerotia could contain lipid-like compounds for the storage of energy or may provide protection against desiccation (Jansson and Friman 1999). Thus, these structures might play an important role in the survival of I. vitellina, for example during drought stress or other harsh environmental conditions. In addition, I. vitellina may not only recruit nutrients from nematode eggs for hyphal development, but inhabit these 50

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi eggs inside cysts for protection and long-time survival. This is a plausible explanation as also healthy eggs of H. filipjevi can survive several years inside cysts. Empirical support for this hypothesis comes from the observation that we were (still) able to isolate I. vitellina from infected eggs after field- collected cysts were kept for several months at 4 °C. The formation of microsclerotia in I. vitellina could be considered as the start and the end of the fungus’ development, at least with respect to those parts of its life cycle that were studied here. Once formed within nematode eggs, microsclerotia may remain inactive but produce newly emerging hyphae when the life-cycle of this species is newly initiated under favourable environmental conditions. Guttules similar to those formed by I. vitellina have been suggested to serve as energy reservoirs in some other nematode parasitic fungi, e.g., the trap-forming Arthrobotrys species (Baral et al. 2017) and could also be involved in the parasitism of nematodes (Nordbring-Hertz et al. 2011). It was reported that guttules may contain linoleic acid as a compound responsible for nematode killing (Anke et al. 1995). Dijksterhuis et al. (1994) also suggested that microbodies, e.g., lipid organelles, present infection-related features in nematophagous fungi. However, their exact functions have not been fully elucidated. This is the first time that fungal survival structures were encountered inside nematode eggs and, accordingly, a new mode of fungus nematode interaction is described herewith. Cysts provide a protected environment for nematode eggs where biotic and abiotic stresses are significantly reduced and eggs survive several years. Such niche might thus be a suitable environment for a nematophagous fungus where it may produce equally long-living survival structures. This situation, along with the presence of mucilaginous content of cysts, might even accelerate the fungal growth and provide optimum conditions for the fungus to colonise the entire cyst cavity and parasitise the eggs. In all cysts collected in fields, however, only a fraction of cysts carried eggs infected with microsclerotia. It is possible that I. vitellina survives in a dormant stage as microsclerotia inside cysts and that it emerges from individually parasitised eggs at favorable conditions, e.g., at times juveniles hatch from non-infected eggs within the same cyst. Microsclerotia encountered in culture have similar shapes and sizes as those I. vitellina forms in nematode eggs, either in vitro or in field collected cysts. The same applies for the cells of these microsclerotia. Accordingly this suggests that in field and in vitro encountered microsclerotia are homologous. If formed in culture they may therefore mimic the egg- parasitising habit of I. vitellina in nature. Absence of such microsclerotia in other closely related species of Ijuhya could therefore suggest that I. vitellina is the only nematode parasitising species of this genus.

2.4.3 Metabolite profiling

Ijuhya vitellina is reported here as a novel source of chaetoglobosin A. The vast majority of chaetoglobosins (A, B, C, D, E, F, G, and J) and their respective derivatives have mostly been isolated from the fungus Chaetomium globosum (Kawahara et al. 2013; Sekita et al. 1976; 1977; Sekita et al. 51

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

1973). Chaetoglobosin C is also produced by Penicillium aurantiovirens (Springer et al. 1976), and chaetoglobosin K was first extracted from Diplodia macrospora (Cutler et al. 1980). Interestingly, no such chaetoglobosins were encountered in the other, closely related Ijuhya species including I. chilensis, I. corynospora, I. faveliana, I. parilis, and I. peristomialis. Whether chaetoglobosin A and its acetyl derivative play a role in nematode egg parasitism can be inferred only with uncertainty. A temporary inhibition of mobility was observed when the two chaetoglobosins were tested in vitro against C. elegans and H. filipjevi. Chaetoglobosins affect and inhibit polymerization of actin and can degrade microfilaments (Löw et al. 1979; Maruyama et al. 1986). This might explain our observation of the effect of chaetoglobosin A and its derivative 19-O-acetylchaetoglobosin A on paralyzing the tested nematodes at 50 and 100 µg/mL. However, at higher concentrations (300 µg/mL) chaetoglobosin A was reported to have toxic effects and caused nematode mortality (Hu et al. 2013). Thus, chaetoglobosins produced by I. vitellina may have a function in the described parasitism of nematode eggs.

Acknowledgements

This work was partially funded by the Deutsche Gesellschaft für Internationale Zusammenarbeit (GIZ). Additional support of CIMMYT, the Julius Kühn-Institut (JKI), and the Helmholtz Centre for Infection Research (HZI) to SA is gratefully acknowledged. Special thanks go to the Fiat Panis Foundation, Hohenheim, for a “PhD completion stipend” to SA, and to the Gemeinschaft der Förderer und Freunde des Julius Kühn-Instituts (GFF) for additional support including publication fees. We would like to thank Anke Brisske-Rode for her skilful technical support of the molecular studies and Katrin Balke for her help in culture maintaining. We also thank Christian Richter, Felix Kaspar and Simone Heitkämper for their contribution to the chemical analyses. SH is grateful for the financial support from the Alexander von Humboldt Foundation. Collaboration between the Institute for Epidemiology and Pathogen Diagnostics of JKI and Agricultural Institute of Slovenia was initiated within FP7 Project CropSustaIn, grant agreement FP7-REGPOT-CT2012-316205. HJS acknowledges support from the ARRS Slovenian science foundation through project J4-5527.

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

2.5 Appendix

Lasionectria (Sacc.) Cooke, Grevillea 12: 111. 1884. Holomorphs of species described in Lasionectria are characterized by perithecia often showing triangular fascicles of densely packed hyphae that emerge from outer perithecial wall regions and 1- septate ascospores (Rossman et al. 1999). Structures illustrated for Ijuhya antillana (Lechat and Courtecuisse 2010) and Ijuhya oenanthicola (Lechat and Hairaud 2012) are conform with the generic concept of Lasionectria. Both species are phylogenetically closely related with Lasionectria mantuana (S1 and S2 Figs), which is the type species of genus Lasioinectria. Accordingly the following combinations are suggested: Lasionectria antillana (Lechat & Courtec.) Schroers, Ashrafi, W. Maier comb. nov., Mycobank MB 821498. Basionym, Ijuhya antillana Lechat & Courtec., Mycotaxon 113: 444. 2010. Mycobank MB516744. Lasionectria oenanthicola (Lechat & Hairaud) Schroers, Ashrafi, W. Maier comb. nov., Mycobank MB 821499. Basionym, Ijuhya oenanthicola Lechat & Hairaud, Mycotaxon 119: 249. 2012. Mycobank MB561714.

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

2.6 References

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2.7 Supporting information

S1 Figure- Bayesian inference of phylogenetic relationships of selected taxa of the Bionectriaceae and Nectriaceae (Hypocreales) based on LSU and rpb1 sequences. Numbers above nodes are estimates of a posteriori probabilities greater than 0.94 / NJB and MLB values greater than 70%. The topology was rooted with Aschersonia placenta, Balansia henningsiana, B. pilulaeformis, and Moelleriella libera (Hypocreales). Two highly supported subclades are suggested for the in-group of genus Ijuhya, of which one includes I. peristomialis, I. chilensis, I. faveliana, I. paraparilis, and I. parilis. The other subclade includes I. vitellina and its closest sister species, I. corynospora. The distantly related I. antillana and I. oenanthicola are inferred as phylogenetic relatives of Lasionectria mantuana. 60

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

S2 Figure- Bayesian inference of phylogenetic relationships of selected taxa of the Bionectriaceae and Nectriaceae (Hypocreales) based on act, LSU, and rpb1 sequences. Numbers above nodes are estimates of a posteriori probabilities greater than 0.94 / NJB and MLB values greater than 70%. The topology was rooted with Aschersonia placenta, Balansia henningsiana, B. pilulaeformis, and Moelleriella libera (Hypocreales). Two highly supported subclades are suggested for the in-group of genus Ijuhya, of which one includes I. peristomialis, I. chilensis, I. faveliana, I. paraparilis, and I. parilis. The other subclade includes I. vitellina and its closest sister species, I. corynospora. The distantly related I. antillana and I. oenanthicola are inferred as phylogenetic relatives of Lasionectria mantuana.

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

S3 Figure- LCMS Chromatogram for the crude extract of Ijuhya vitellina. Peaks represent chaetoglobosin A (1) and 19-O-acetylchaetoglobosin A (2); Insertion is the UV-VIS spectrum of chaetoglobosin A (1).

S4 Figure- Mass spectrum of chaetoglobosin A (1).

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

S5 Figure- Mass spectrum of 19-O-acetylchaetoglobosin A (2).

S1 Text. Structure determination of chaetoglobosins

Compound 1 was obtained as yellow powder. Its molecular formula was determined as C32H36N2O5 by the HRESIMS which showed ion peak at m/z 529.2699 [M+H]+ (calcd 529.2697). The 1H NMR spectrum of 1 exhibited signals for 10 aromatic/olefinic protons, seven methines, two methylens and four methyls. Two additional signals were assigned to hetero protons suggested the presence of two NH and/or OH groups in 1. The 13C NMR showed 32 carbon resonances, of which 23 were observed in

DEPT spectrum indicating 9 non-protonated among them 3 carbonyls (δC 201.7, 196.7 and 172.9) ( S1 Table). The planar structure of compound 1 was deduced from the 2D NMR data including COSY and HMBC spectra. COSY correlation between H-4´/H-5´/H-6´/H-7´ together with HMBC correlations from H-4´to C-6´/C-7a´/C-3´; H-7´ to C-5´/C-3a´; H-2´ to C-3a´/C-7a´ and from NH-1´ to C- 2´/C-3´/C-3a´/C-7a´ revealed the presence of an indole moiety in compound 1 (S6 Fig). In addition, an isoindole derivative was assigned by HMBC correlations network from NH-2 to C-1/C-3/C-4/C-9; a group of HMBC correlations from H-4 to C-1/C-3/C-5/C-6/C-8/C-9; H3-10 to C-4/C-5/C-6; H3-12 to C- 5/C-6/C-7; COSY correlation between H-7 and H-8 together with HMBC correlations from H-7 to C-6/C- 9 and from H-8 to C-7/C-9/C-1 allowed the construction of 6,6a-dimethylhexahydro-1aH-oxireno[2,3- f]isoindol-3(4H)-one. The above mentioned structure units of compound 1 was connected by methylene group assigned by means of COSY correlation from H-3 to H-10 and a series of HMBC correlations from H-10 to C-3/C-4/C-2´/C-3´/C-4´ as well as correlations from H-2´ to C-10. Finally, a chain of 11-carbon atoms was determined in the same manner by COSY and HMBC correlations, whereas COSY correlations were determined for a series of protons from H-13 to H-17 and HMBC correlations from H3-16´ to C-15/C-16/C-17; from H3-18´ to C-17/C-18/C-19. Furthermore, COSY correlation between the two olefinic protons H-21 and H-22 with HMBC from H-21 to the carbonyl

63

Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi carbon C-23 and to C-9 and from H-4 to C-23; H-7 to C-14, H-13 to C-7/C-8 allowed the construction of 13-membered ring at C-8/C-9 of the isoindole moiety.

Compound 2 was isolated as a yellow powder. It has the molecular formula of C34H38N2O6 deduced from the HRESIMS data that showed a peak at m/z 571. 2803 [M+H]+ (calcd 571.2803). Its UV spectrum was similar to that of chaetoglobosin A and the molecular mass difference of 42 Da suggested that 2 was likely an acetyl derivative of 1. The 1H and 13C NMR spectra of 2 were very similar to those of 1,

13 but the C NMR spectrum showed two additional carbon resonances at δC 170.2 and δC 20.8, further supporting the presence of an additional acetyl group. The presence of the acetyl group was determined by HMBC correlation from H3-25 (δH 2.18) to the carbonyl carbon C-24 and its position was determined by the HMBC correlation from H-19 (δH 5.91, δC 83.3) to C-24.

S6 Figure- 2D NMR assignment of chaetoglobosin A (1) and 19-O-acetylchaetoglobosin A (2). HMBC (arrows) and COSY (bold bonds) correlations.

S2 Text. Spectroscopic data for chaetoglobosin A (1) and 19-O-acetylchaetoglobosin A (2) Chaetoglobosin A (1):

20 + Yellow powder; [] D - 165° (c 0.1, MeOH); NMR data see Table S1; MS data m/z 529.21 [M+H] , 511.22

+ - - + [M+H-H2O] , 527.20 [M-H] , 573.15 [M-H+HCOOH] ; HRESIMS m/z 529.2699 [M+H] (calcd 529.2697 for C32H37N2O5).

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

19-O-acetylchaetoglobosin A (2):

25 1 Yellow powder; [] D - 147° (c 0.1, MeOH); H NMR (700 MHz, CDCl3) δH 1.02 (d, J=6.88 Hz, H-16´), 1.25 (d, J=7.31 Hz, H-11), 1.30 (s, H-12), 1.47 (d, J=1.29 Hz, H-18´), 1.86 (m, H-5), 2.05 (dt, J=13.77, 10.97 Hz, H-15a), 2.15 (dd, J=9.68, 4.95 Hz, H-8), 2.20 (s, H-25), 2.27 (m, H-15b), 2.46 (m, H-16), 2.67 (dd, J=14.63, 7.74 Hz, H-10a), 2.80 (d, J=5.16 Hz, H-7), 2.91 (m, H-10b), 2.93 (m, H-4), 3.80 (dt, J=7.21, 3.50 Hz, H-3), 5.20 (ddd, J=15.06, 10.76, 3.87 Hz, H-14), 5.71 (dd, J=9.03, 1.29 Hz, H-17), 5.74 (s, NH-2), 5.93 (s, H-19), 6.09 (ddd, J=15.17, 9.79, 1.29 Hz, H-21), 6.39 (br d, J=16.78 Hz, H-22), 7.01 (d, J=2.15 Hz, H-2´), 7.15 (dd, J=6.88, 7.31 Hz, H-5´), 7.21 (dd, J=6.88, 8.17 Hz, H-6´) 7.37 (d, J=8.17 Hz, H-7´), 7.50 (d, J=7.74 Hz, H-4´), 7.58 (d, J=16.78 Hz, H-22), 8.20 (br s, NH-1´); 13C NMR (176MHz, CHLOROFORM-d) δ = 196.8 (C, C-23), 195.0 (C, C-20), 173.3 (C, C-1), 170.1 (C, C-24), 142.6 (CH, C-17), 136.3 (C, C-7a´), 134.8 (CH, C-22), 133.4 (CH, C-14), 133.2 (CH, C-21), 128.3 (CH, C-13), 127.9 (C, C-18), 127.0 (C, C-3a´), 123.5 (CH, C-2´), 122.6 (CH, C-5´), 120.1 (CH, C-6´), 118.3 (CH, C-4´), 111.7 (CH, C-7´), 110.2 (C, C-3´), 83.3 (CH,

C-19), 63.2 (C, C-9), 62.3 (C, C-7), 57.9 (C, C-6), 52.4 (C, C-3), 48.6 (CH, C-8), 47.4 (CH, C-4), 41.5 (CH2, C-

14), 36.3 (CH, C-5), 34.1 (CH2, C-10), 32.2 (CH, C-16), 20.84 (CH3, C-25), 20.81 (CH3, C-16´), 19.9 (CH3, C-

+ + 12), 13.5 (CH3, C-11), 11.7 (CH3, C-18´); MS data m/z 571.22 [M+H] , 511.22 [M+H-AcOH] , 569.15 [M-

- - + H] , 615.12 [M-H+HCOOH] ; HRESIMS m/z 571.2803 [M+H] (calcd 571.2803 for C34H39N2O5).

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Ijuhya vitellina sp. nov. a parasite of Heterodera filipjevi

S1 Table. NMR spectroscopic data for chaetoglobosin A (1) (700 MHz, CDCl3) 1 Pos. δC, type δH, mult. ( J in Hz) HMBC 1 172.9, C - - 2 - 5.73, s 1, 9, 3, 4 3 52.6, CH 3.79, m 1, 4, 5 4 47.2, CH 3.05, dd (4.73, 3.44) 1, 3, 5, 6, 8, 9, 23 5 36.2, CH 1.86, m 3, 4, 6, 9, 11, 12 6 58.0, C - - 7 62.4, CH 2.81, d (4.73) 6, 8, 9, 12, 13 8 48.9, CH 2.15, dd (9.90, 4.73) 7, 9, 1, 13, 14 9 63.2, C - - 2.63, dd (14.63, 8.17) 10 34.4, CH2 2.97, dd (14.63, 3.87) 2´, 3´, 3a´, 3, 4

11 13.6, CH3 1.27, d (7.31) 4, 5, 6

12 19.8, CH3 1.31, s 5, 6, 7 13 128.2, CH 6.07, dd (15.27, 9.90) 7, 8, 15 14 133.8, CH 5.24, ddd (15.17, 10.86, 4.09) 8, 15, 16 2.05, dd (11.35, 13.44) 15 41.8, CH2 2.28, m 13, 14, 16, 16´, 17

16 32.1, CH2 2.47, m 15, 16´, 17, 18

16´ 21.0, CH3 1.02, d (6.9) 15, 16, 17 17 140.4, CH 5.63, dd (9.03, 1.29) 15, 16, 18, 19 18 132.3, C - -

18´ 10.6, CH3 1.34, d (1.29) 17, 18, 19 19 81.8, CH 5.06, s 17, 18´, 20, 21 20 201.7, C - - 21 131.6, CH 6.51, br d (16.78) 19, 21, 22 22 136.4, CH 7.77, br d (16.53) 20, 21, 22, 9 23 196.7, C - - 1´ - 8.14, br s 2´, 3´, 3a´, 7a´ 2´ 123.2, CH 6.98, d (2.15) 3´, 3a´, 7a´, 10 3´ 110.4, C - - 3a´ 126.8, C - - 4´ 118.3, CH 7.50, d (7.74) 6´, 7a´, 3´ 5´ 120.0, CH 7.16, dd (7.42, 6.88) 3a´, 7´ 6´ 122.5, CH 7.22, dd (7.64, 6.88) 4´, 7a´ 7´ 111.6, CH 7.38, d (8.17) 5´, 3a´ 7a´ 136.3, C - -

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Monocillium gamsii sp. nov. and Monocillium bulbillosum parasites of Heterodera filipjevi

Chapter 3.

Monocillium gamsii sp. nov. and Monocillium bulbillosum: two nematode-associated fungi parasitising the eggs of Heterodera filipjevi

Published in MycoKeys 27: 21-38 (2017) https://doi.org/10.3897/mycokeys.27.21254

Samad Ashrafi ‡§, Marc Stadler |, Abdelfattah A. Dababat ¶, Katja R. Richert-Pöggeler ‡, Maria R. Finckh §, Wolfgang Maier ‡

‡ Institute for Epidemiology and Pathogen Diagnostics, Julius Kühn-Institut (JKI)–Federal Research Centre for Cultivated Plants, Braunschweig, Germany

§ Department of Ecological Plant Protection, Faculty of Organic Agricultural Sciences, University of Kassel, Witzenhausen, Germany

| Department Microbial Drugs, Helmholtz Centre for Infection Research GmbH (HZI), Braunschweig, Germany

¶ CIMMYT (International Maize and Wheat Improvement Centre), P.K.39 06511 Emek, Ankara, Turkey

Corresponding author

Samad Ashrafi ([email protected] – [email protected])

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Monocillium gamsii sp. nov. and Monocillium bulbillosum parasites of Heterodera filipjevi

Abstract

Monocillium gamsii sp. nov. (Ascomycota, Hypocreales, Niessliaceae) isolated from eggs of the cereal cyst nematode Heterodera filipjevi is described and illustrated based on morphological and molecular phylogenetic evidence. The new taxon discovered in wheat fields in Turkey destructively parasitises nematode eggs. The infected eggs were readily colonised by the fungus, which produced microsclerotia. The fungus could be grown on artificial media and the parasitism of M. gamsii towards H. filipjevi was reproducible in vitro. Hyphae penetrating the nematode eggs entirely colonised the embryo, developed into multicellular chlamydospore and dictyochlamydospore-like structures eventually forming microsclerotia. Molecular and morphological differences and similarities between M. gamsii and its phylogenetically related species are discussed. Monocillium bulbillosum was found to be closely related to the new species. The pathogenicity of M. bulbillosum against H. filipjevi was also assayed in vitro because of its sister group relationship to M. gamsii revealing that this species was also capable of colonising eggs of H. filipjevi.

Keywords

Egg-parasitic fungi, Niessliaceae, new species, plant parasitic nematodes, taxonomy, molecular phylogeny, ITS, LSU, rpb1, tef

3.1 Introduction

Various fungi have been reported as natural enemies of plant parasitic nematodes (PPN) (Nordbring- Hertz et al. 2011; Siddiqui and Mahmood 1996; Stirling 2014). A group of these fungi infect females and egg contents of endoparasitic nematodes such as cyst nematodes (Kerry 1988; Rodríguez-Kábana and Morgan-Jones 1988), which are biotrophic plant pathogens establishing a long-term parasitic interaction with their host plants. The unique sedentary life style of this group of PPN render them especially vulnerable of being colonised by their natural enemies (Lopez-Llorca et al. 2008). Cyst nematodes are globally distributed and were the first group of PPN reported to be parasitised by fungi (Kühn 1877), which spurred investigations to find additional nematode-antagonistic fungi ever since [(Tribe 1977) and references therein]. Most egg-parasitic fungi belong to the ascomyceteous Hypocreales, e.g. Pochonia chlamydosporia (Goddard) Zare & W. Gams, Metapochonia rubescens (Zare, W. Gams & López-Llorca) Kepler, S.A. Rehner & Humber, Lecanicillium lecanii (Zimm.) Zare & W. Gams, Metarhizium spp., Purpureocillium lilacinum (Thom) Luangsa-ard, Houbraken, Hywel-Jones & Samson, and Trichoderma spp. (Kerry and Hirsch 2011; Khan et al. 2006; Szabó et al. 2012; Zhang et al. 2014). In contrast, none of the second important group of nematode-antagonistic Ascomycota, the Orbiliomycetes (Baral et al. 2017) has been reported to parasitise nematode cysts and eggs. 68

Monocillium gamsii sp. nov. and Monocillium bulbillosum parasites of Heterodera filipjevi

Grant and Elliott (1984) reported Monocillium sp. parasitising the cysts of the soybean cyst nematode Heterodera glycines. This is so far the only report on Monocillium antagonising a plant parasitic nematode. The genus Monocillium Saksena 1955 was emended and placed in the Niessliaceae by Gams (1971), and Monocillium spp. were regarded as the asexual morphs of the hypocrealean genus Auersw. 1869. However, the types of both genera have not yet been connected conclusively by elucidation of the life cycle or by molecular data, hence we hesitate to regard these genera as synonymous and treat them as separate taxonomic entities for the time being. The genus Monocillium currently comprises eighteen species (http://www.mycobank.org/quicksearch.aspx) and is defined by showing acremonium-like morphology, but is characterised by unbranched conidiophores with phialides having thickened walls in the lower part. The known species were isolated from soil, plant materials such as dead leaves and wood, but also from other fungi, and building material (such as wall paper). Among all Monocillium species described so far (Barron 1961; Gams 1971; Gams and Turhan 1996; Girlanda and Luppi-Mosca 1997; Ramaley 2001) M. curvisetosum W. Gams & Turhan is the only species which was originally isolated from aphids as an unusual host for this genus. However its potential parasitic association with its host has not yet been reported. Egg-parasitic fungi attacking cyst nematodes have repeatedly been isolated from all agricultural soils in various geographic regions (Chen and Chen 2002; Dababat et al. 2015). Experimental wheat fields of the International Maize and Wheat Improvement Centre (CIMMYT) in Turkey, where a significant reduction in population size of the cereal cyst nematode Heterodera filipjevi had been observed between two consecutive years (unpublished data), were sampled to isolate and study fungal candidates that could be causally involved in this drop of the nematode population size. Here we report a so-far undescribed hypocrealean species which destructively parasitised the eggs of H. filipjevi. The antagonistic interaction of this fungus with the nematode eggs was studied based on in vitro tests. We also report the antagonistic potential of M. bulbillosum as the most closely related species to the herein described fungus, towards the eggs of H. filipjevi.

3.2 Material and methods

3.2.1 Sample collection and materials examined

Cysts of H. filipjevi were collected from experimental wheat fields of CIMMYT in the Central Anatolian Plateau of Turkey in 2013. The fields located in Yozgat (39° 08’ N, 34° 10ʼ E; altitude 985 m.a.s.l) and Haymana (39° 26’ N, 39° 29ʼ E, altitude 1260 m.a.s.l) were naturally nematode infested. The samples including soil and roots were collected at random from the rhizosphere of wheat plants at the end of the growing season. Cysts were extracted from the collected samples using the modified flotation decanting method (Coyne et al. 2007). From the extracted suspensions, cysts were manually collected 69

Monocillium gamsii sp. nov. and Monocillium bulbillosum parasites of Heterodera filipjevi under a dissecting microscope and stored in 1.5 ml microtubes at 4 ˚C either in dry condition or in sterile tap water until further use. For taxonomic and phylogenetic inferences, additional fungal strains were obtained from the Westerdijk Fungal Biodiversity Institute (formerly CBS-KNAW, Utrecht, Netherlands).

3.2.2 Cultural studies

3.2.2.1 Fungal isolation from eggs of Heterodera filipjevi The field-collected cysts of H. filipjevi were scrutinised by using a dissecting microscope to separate symptomatic cysts showing defined discolourations or bearing discernible hyphae, from healthy- looking (i.e. homogeneously brown) or empty cysts. Symptomatic cysts were selected, surface- sterilised in 5% sodium hypochlorite (NaOCl), and dissected to collect their egg contents. Only the nematode eggs showing symptoms of fungal infection were processed for fungal isolation and culture- dependent species identification. A portion of the fungal infected eggs were additionally used for culture-independent identification. The methods applied here, have been described in greater detail in Ashrafi et al. (2017).

3.2.2.2 Growth rate studies Growth rates were determined at various temperatures from 15 to 35 °C at 5 °C intervals in the dark or in ambient conditions by placing agar disks (5 mm diam.), excised from the margin of a young potato dextrose agar (PDA) culture onto five replicate plates of PDA, cornmeal agar (CMA), oatmeal agar (OA; 30 g oatmeal, 18 g agar-agar, 1L deionised water), synthetic nutrient-poor agar (SNA; Nirenberg (1976)), and malt extract agar (MEA). The colony diameter was measured weekly for a 3 week period. Colour changes of fungal structures formed in culture were checked using 3% potassium hydroxide (KOH) watery solution.

3.2.3 Pathogenicity tests against H. filipjevi

The antagonistic potential of the below described species and M. bulbillosum, respectively, was assessed towards H. filipjevi in vitro as previously described (Ashrafi et al. 2017). Briefly, healthy cysts and eggs were surface-sterilised and placed either on or at the margin of the growing mycelium of one- month-old PDA or 2% water agar (WA) cultures of the two fungal species. To document the process of colonisation of eggs of H. filipjevi by the new fungal species, a slide culture technique was also performed using PDA 1/3 strength (compare Ashrafi et al. (2017)).

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Monocillium gamsii sp. nov. and Monocillium bulbillosum parasites of Heterodera filipjevi

3.2.4 Microscopy

Nematode eggs and fungal structures were examined and photographed by a Zeiss Axioskop 2 plus compound microscope and an Olympus SZX 12 stereo microscope equipped with a Jenoptik ProgRes® digital camera. Images were recorded using CapturePro 2.8 software (Jenoptic, Jena, Germany). Nematode eggs colonised by fungi, and fungal structures were mounted in water or lactic acid and photographed. Cysts were photographed in water in a square cavity dish (40×40×16 mm). To illustrate different stages of fungal development and fungal colonisation of nematode eggs, slide cultures were prepared (Gams et al. 1998) and then photographed. Nomarski Differential Interference Contrast (DIC) optic was used for observation and measurements. All measurements were taken in water, and are given as x1–x2 (x3 ± SD), with x1 = minimum value observed, x2 = maximum value observed, x3 = average, and standard deviation (SD), followed by the number of measurements (n). Scanning electron microscopy was performed on a Quanta 250 scanning electron microscope (FEI Deutschland GmbH, Frankfurt, Germany). Fungal structures of interest were obtained from a one- month-old OA culture grown at 23 °C in the dark and directly analysed using environmental scanning electron microscopy (ESEM). For the experiment, pressures between 410 and 490 Pa at 4 °C were employed. For cooling the sample chamber was equipped with a Peltier stage. Fungal mycelia with abundant conidia were placed on non-conductive double-sided adhesive discs on a flat specimen stub and positioned on the Peltier stage for cooling. Images were taken at acceleration voltage of 12.5 kV. Scanning speed was 60 µsec. For imaging of beam sensitive fungal structures, the scanning modus was changed to 3 µsec with 20-fold line integration. Images were adjusted in brightness and contrast using Adobe Photoshop software CS 5.1.

3.2.5 Molecular phylogenetic studies

3.2.5.1 DNA extraction, PCR amplification and DNA sequencing Fungal genomic DNA was isolated from mycelia grown on PDA using a modified CTAB method, and from individual nematode eggs infected by fungi using the Qiagen DNeasy Plant Mini Kit (Qiagen, Hilden, Germany) as reported in Ashrafi et al. (2017). For each specimen, four nuclear loci were amplified: The internal transcribed spacers including the 5.8S rDNA gene (ITS) using the primers ITS1F (Gardes and Bruns 1993) and ITS4 (White et al. 1990); the 5’ end of the ribosomal large subunit (LSU) DNA with the primers LROR (Rehner and Samuels 1994) and LR5 (Vilgalys and Hester 1990); partial RNA polymerase II largest subunit 1 (rpb1) using the primers cRPB1af and RPB1cr (Castlebury et al. 2004); and partial translation-elongation factor 1-α (TEF) using the primers EF1-983f and EF1-2218r (Castlebury et al. 2004). All PCR reactions were performed as described previously (Ashrafi et al. 2017) with the following thermal programmes: 95 °C (2 min) for

71

Monocillium gamsii sp. nov. and Monocillium bulbillosum parasites of Heterodera filipjevi initial denaturation followed by 40 cycles of denaturation at 95 °C (30 s), annealing at 52 °C (ITS), 51 °C (LSU), 54 °C (rpb1), and 60 °C (TEF) (40 s), extension at 72 °C (1 min for ITS, LSU and rpb1, and 1 min and 20 sec for TEF), and a final extension at 72 °C (10 min). Amplicons were purified using the DNA Clean & ConcentratorTM-5 kit (Zymo Research Corp., Irvine, California, USA) and sequenced by Eurofins Genomics GmbH, (Ebersberg, Germany) with the same primers as used for PCR amplification. Obtained sequences were assembled, edited and trimmed with Sequencher 5.4.1 (Gene Codes Corporation, Ann Arbor, Michigan, USA) and deposited in GenBank under the following accession numbers: MF681481– MF681514. The sequences generated were compared to sequences available in GenBank using a BLASTn search (http://blast.ncbi.nlm.nih.gov/Blast.cgi) (Altschul et al. 1990).

3.2.5.2 DNA sequence alignment and phylogenetic inference The newly generated sequences together with closely related sequences selected as revealed by BLASTn searches were used for phylogenetic analyses (Table 3. 1). The sequences were aligned using the online version of Mafft v.7 (Katoh and Standley 2013). All sequences were aligned using the iterative refinement methods: Sequences of the rpb1 and TEF gene regions were aligned using the algorithms implemented in L–INS–i, while LSU and ITS were aligned applying the Q–INS–i algorithm. Only the start and end of the alignments were trimmed manually in Se-Al v2.0 (Rambaut 1996). The following phylogenetic analyses were applied: a Bayesian method of phylogenetic inference using Metropolis Coupled Monte Carlo Markov chains (MC3) as implemented in the computer program MrBayes v3.2 (Huelsenbeck and Ronquist 2001; Ronquist and Huelsenbeck 2003). We used MrModeltest v2.2 (Nylander 2004) to determine the best fitting DNA substitution model for the Bayesian approach. Both the hierarchical likelihood ratio test (hLRT) and the Akaike Information Criterion (AIC) selected the general time reversible model of DNA substitution with gamma distributed substitution rates and invariate sites (GTR+I+G) as the best fitting model for all individual data sets and was implemented for the analyses accordingly. For the Bayesian analyses four incrementally heated simultaneous Monte Carlo Markov chains were run with 2.000.000 generations using random starting trees and flat prior distributions. Trees were sampled every 500 generations resulting in a total of 4001 sampled trees. A 50% majority rule consensus tree was computed only from trees of the plateau, and if, additionally, the split frequencies were below 0.01. Thus, 501 trees were discarded as “burnin”. Maximum likelihood (ML) analyses were performed using RAxML 7.2.8 (Silvestro and Michalak 2012; Stamatakis 2014) implemented in Geneious 8.1.2 applying the general time-reversible (GTR) substitution model with gamma model of rate heterogeneity and 1000 replicates of rapid bootstrapping. Neighbor-joining (NJ) analyses (Saitou and Nei 1987) was done in PAUP 4.0b10 in the batch file mode (Swofford 2002) applying the Kimura two-parameter model of DNA substitution (Kimura 1980) with a transition/transversion ratio of 2.0 to compute genetic distances. Support for

72

Monocillium gamsii sp. nov. and Monocillium bulbillosum parasites of Heterodera filipjevi internal nodes was estimated by 1000 bootstrap replicates (Felsenstein 1985). Two members of Bionectriaceae, Bionectria byssicola (Berk. & Broome) Schroers & Samuels and Ijuhya vitellina Ashrafi, W. Maier & Schroers, were selected as outgroup to root the trees. The phylogenetic trees were visualised using FigTree v. 1.4.2 (http://tree.bio.ed.ac.uk/software/figtree).

Table 3.1- Isolates and accession numbers used in the phylogenetic analyses

Isolate Host / GenBank accession numbers Species Locality Reference number substrate ITS LSU rpb1 tef Bionectria CBS 914.97– Alchornea Uganda- AF358252 GQ506011 GQ506040 KX184977 (Hirooka et byssicola GML2665 branches- leaf al. 2010; litter Moreira et al. 2016; Schroers 2001)

Hyaloseta CBS109837 Nolina USA, KM231846 KM231726 KM232279 - (Lombard nolinae micrantha, leaf New et al. 2015) litter Mexico Ijuhya DSM104494 Heterodera Turkey KY607535 KY607549 KY607576 - (Ashrafi et vitellina filipjevi, egg al. 2017) Monocillium CBS344.70 mouldy Germany MF681488 MF681501 MF681513 MF681507 This study bulbillosum wallpaper Monocillium DSM105458 Heterodera Turkey MF681485 MF681496 MF681512 MF681506 This study gamsii filipjevi, egg Monocillium DSM105459 Heterodera Turkey MF681483 MF681493 MF681511 MF681505 This study gamsii filipjevi, egg Monocillium DSM105460 Heterodera Turkey MF681482 MF681492 MF681510 MF681504 This study gamsii filipjevi, egg Monocillium DSM105461 Heterodera Turkey MF681481 MF681490 MF681509 MF681503 This study gamsii filipjevi, egg Monocillium CBS684.95 ectomycorrhizae Italy MF681489 MF681502 MF681514 MF681508 This study ligusticum of Pinus halapensis Nisslia exilis CBS357.70 Picea abies, bark Germany - AY489718 AY489645 AY489613 (Castlebury et al. 2004) Nisslia exilis CBS560.74 Pinus sylvestris, England - AY489720 AY489647 AY489614 (Castlebury decayed needle et al. 2004)

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Monocillium gamsii sp. nov. and Monocillium bulbillosum parasites of Heterodera filipjevi

3.3 Results

3.3.1 Sample collection and fungal isolation

Among the field-collected samples, a high proportion of cysts was found containing blackish bodies resembling microsclerotia-like structures upon microscopy (Fig 3.1A). By dissecting the infected cysts, microsclerotia-like black bodies were found to be colonising the individual nematode eggs (Fig 3.1B, C). In some infected eggs the developing juveniles were found to be entirely destroyed exhibiting an olivaceous brownish appearance (Fig 3.1D, E). Eggs were colonised by one or occasionally two microsclerotia. When cultured on PDA, hyphae grew out of the microsclerotia of the infected eggs (Fig 3.1F), and formed colonies at first white creamy, later becoming blackish dotted centrally with a general dark appearance due to the dense pigmentation (Fig 3.1G).

Figure 3.1- Naturally infested cysts and eggs of Heterodera filipjevi with Monocillium gamsii, and pure cultures obtained from infected eggs. A) Field collected symptomatic cysts bearing parasitised eggs. B–E) Nematode eggs infected by M. gamsii: B, D) Nematode eggs containing microsclerotia of M. gamsii; E) An embryonated egg containing a second stage juvenile (J2) parasitised by M. gamsii (arrow points at nematode’s stylet). F) A nematode egg containing microsclerotia, and hyphae growing out of it. G–H) colony of M. gamsii grown on PDA: G) colonies developing from three individually plated infected eggs; H) A 25-d-old culture grown at 25 °C in the dark. I) The surface of a five-month-old culture detailing the sclerotioid masses covering the colony surface. Single microsclerotia can be seen as little black dots at the margin of the culture. Scale bars: A = 800 µm; B–E = 30 µm; F = 50 µm; H = 2 cm; I = 5 mm. 74

Monocillium gamsii sp. nov. and Monocillium bulbillosum parasites of Heterodera filipjevi

3.3.2 Sequence comparison and phylogenetic inference

The DNA sequences of four different gene regions obtained from the examined specimens of the here described nematode-parasitic fungus were either identical (in TEF and RPB1), or nearly identical (1 base pair (bp) substitution in LSU, and up to 2 bp substitutions in ITS). The most similar DNA sequences found in GenBank using BLASTn searches belonged to Hyaloseta nolinae, the sexual morph of Monocillium nolinae, and shared similarities of 96% in the ITS region, 99% in the LSU, and 89% in rpb1. A similar BLASTn search in MycoBank showed identities of the ITS sequence of 96.6% with M. bulbillosum, 93.9% with H. nolinae and 92.7% with Niesslia exosporioides, and of the LSU sequence of 99.5% with H. nolinae, and 99% with both M. bulbillosum and Niesslia exosporioides suggesting a close relationship with the representatives of the Niessliaceae. Fungal DNA could also be directly isolated and sequenced from individual eggs displaying the typical symptoms of fungal infection. These DNA sequences were identical to the sequences retrieved from pure cultures supporting the conspecificity of the symptom-causing structures within the egg with the isolated pure cultures derived from the eggs. The final combined ITS, LSU, rpb1 and tef dataset comprised 11 strains representing 7 species with a total alignment length of 2949 bp (603 ITS, 797 LSU, 649 rpb1, 900 tef). The topologies of the phylogenetic trees were identical using Bayesian inference (Fig 3.2), neighbor-joining or maximum likelihood (not shown). The four sequenced strains of the here described nematode egg-colonising fungus were recovered as a highly supported monophyletic group with a close sister group relationship to M. bulbillosum and with H. nolinae as the next-closest relative. In the second monophyletic clade of Niessliaceae, two strains of the type species of Niesslia, N. exilis, proved to be paraphyletic with respect to M. ligusticum (Fig 3.2).

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Figure 3.2- Bayesian inference of phylogenetic relationships using four strains of the here described nematode parasite and all Niessliaceae present in GenBank based on an alignment of ITS, LSU, rpb1, and tef sequences using GTR+I+G as nucleotide substitution model. Depicted is a 50% majority rule consensus tree derived from 3500 trees from the stationary phase of a Monte Carlo Markov Chain. A posteriori probability (BIpp) values greater than 0.95, and bootstrap values of neighbor-joining (NJBT) and maximum likelihood (MLBT) analyses greater than 0.7 are given above branches (BIpp/NJBT/MLBT). Two representatives of the Bionectriaceae, Bionectria byssicola and Ijuhya vitellina, were used to root the tree.

3.3.3 Taxonomy

Monocillium gamsii Ashrafi & W. Maier, sp. nov. MycoBank No: MB 823248 (Figs 3.1H, 1I & Fig 3.3) Holotype Turkey, Yozgat, experimental wheat field: dried culture on PDA, originating from an individual egg from a cyst of Heterodera filipjevi, isolated by Samad Ashrafi, August 2013, dried culture on PDA, deposited at the herbarium of the Botanic Garden and Botanical Museum Berlin-Dahlem: B700016491. Ex- holotype strain: DSM 105458, deposited in the open collection of the Leibniz-Institut DSMZ Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH, GenBank accession numbers: ITS: MF681485; LSU: MF681496; rpb1: MF681512; tef: MF681506.

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Additional material examined From the same location: DSM 105459 (dried culture on PDA, B700016492), GenBank accession number: MF681483 (ITS), MF681493 (LSU), MF681511 (rpb1), MF681505 (tef); DSM 105460 (dried culture on PDA, B700016493), GenBank accession number: MF681482 (ITS), MF681492 (LSU), MF681510 (rpb1), MF681504 (tef); DSM 105461, GenBank accession number: MF681481 (ITS), MF681490 (LSU), MF681509 (rpb1), MF681503 (tef); and CBS 141176. Etymology In honour and memory of Prof Walter Gams for his outstanding works on the genera Monocillium and Niesslia. Diagnosis Naturally occurring infected eggs often accommodating one subglobose, strongly pigmented, dark brownish microsclerotium. Description Colonies slow-growing, at 20 °C on PDA reaching 10–12 mm diam. (7d), 19–22 mm (14 d), 25–32 (21 d); optimum temperature for growth 25 °C, 14–16 mm (7 d), 22–25 mm (14 d), 31–34 mm (21d); at 30 °C 10–11 mm (7d), 15–17 (14 d), 22–25 mm (21 d), no growth observed at 35 °C; optimum temperature for growth in other examined cultural media 25 °C, after 21 d reaching 31–32 mm diam. (CMA), 36–40 mm (MEA), 40–50 mm (OA), 32–40 mm (SNA); colonies on PDA finely wrinkled, slightly elevated centrally, first pale creamy, later centrally becoming dotted, greyish-brown to fuscous black due to formation of darkly pigmented microsclerotia, margins and reverse pale creamy. Vegetative hyphae hyaline, thin-walled, forming strands or coils, often with dictyochlamydospore-like structures, occasionally bearing setae with elongate, ellipsoid tips, variable in size. Chlamydospores or dictyochlamydospores mostly developing intercalary, filled with small guttules, gradually pigmented, turning brownish firstly at cell walls, interweaving to form microsclerotia. Cells of microsclerotia angular, pigmented, first pale olivaceous brown filled with guttules, later dark brown, forming a textura angularis in surface view. Guttules often absent in mature and strongly melanised sclerotial cells. Microsclerotia later covering the entire colony, developing sclerotioid masses, not changing colour in KOH. Phialides often separated from hyphae by a basal septum, thick-walled in the lower part, the wall thickening distinct at about 1/3 to 1/2 of the total length from the base, thin-walled from ca. midpoint extending to the tip, occasionally slightly inflated in the middle part, gradually tapering to the tip, 21– 39 µm (28.7 ± 4.4) in length, 1.0–2.1 µm (1.4 ± 0.2) wide at the base (n = 90), solitary, arising directly from hyphae or hyphal rope, occasionally arising from hyphal coils surrounding several conidia. Conidiogenesis abundant, conidia hydrophilic, adhering in watery droplets, oblong, rarely clavate or ampulliform, one-celled, smooth-walled, 4.1–7.4 × 1.4–2.9 µm (4.9 ± 0.6 × 2.1 ± 0.3) (n = 250). Sexual morph not observed.

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Figure 3.3- Micrographs of Monocillium gamsii. A–C) Hyphal growth by intercalary development of chlamydospore and dityochlamydospore-like structures filled with guttules. D–H) initiation of microsclerotia by interweaving or coiling of dictyochlamydospores, and growth to full size. I) Highly pigmented sclerotium at maturity displaying a textura angularis on surface view. J–N) Setae, phialides and conidia. M–O) Formation of phialides on coiling hyphal. P) Conidial heads, conidia cohering in wet heads. Q, R) SEM: Q) Phialides from mycelium with conidial heads, arising from hyphae, R) coiling hyphae (arrows), and detail of phialides bearing conidia. A–I from PDA 1/3 strength; J–P from PDA: Q, R from OA. Scale bars: (A, H, I, K, M, O, R) = 30 µm; (B–G, J, L, N) = 20 µm; P = 200 µm, Q = 50 µm. 78

Monocillium gamsii sp. nov. and Monocillium bulbillosum parasites of Heterodera filipjevi

3.3.4 Development of M. gamsii in nematode eggs in vitro

Monocillium gamsii infected cysts and eggs of H. filipjevi in vitro. Initial indications of infection were observed in healthy nematode cysts placed on the fungal colonies within 2–3 weeks (Fig 3.4A, B). The fungus rendered the homogenously brown healthy looking cysts black-dotted, bearing a strong resemblance to the naturally infected cysts collected from fields. By dissecting the symptomatic cysts, nematode eggs were found to be colonised with darkly pigmented spherical to subglobous microsclerotia formed inside the body cavity of the developing juveniles (Fig 3.4C). Similar to some naturally infected eggs, sclerotioid masses were also found in some artificially infected samples colonising almost the entire egg (Fig 3.4D). In the slide cultures, fungal infection of eggs was initiated by individual hyphae directly penetrating the eggshell and body cuticle of developing juveniles (Fig 3.4E, F). Following penetration, filamentous hyphae entirely colonised the unembryonated eggs (Fig 3.4 G, H) or the body cavity of the developing juveniles (Fig 3.4I, J), enlarged (Fig 3.4H–J), occasionally inflated, forming thick-walled, finely pigmented, and guttules-filled cells (Fig 3.3J), which eventually coalesced to form discrete microsclerotia with a textura angularis appearance (Fig 3. 4K–M). Infection studies revealed that such microsclerotia could be formed 7–10 d after the incubation of nematode eggs with the fungus. Pigmentation of microsclerotia occurred during fungal development from hyaline to olivaceous brown and later strongly brownish melanised cells (Fig 3.4N–P). Microsclerotia developing inside the artificially infected eggs displayed a textura angularis and were indistinguishable from those found in the naturally infected samples. At the early stages of development, microsclerotial cells were often filled with guttules (oil-like bodies), which were not observed in the mature microsclerotia. At the early stages of fungal infection (up to 10 d after inoculation), some vacuole-like structures were observed inside the eggs along the body cavity of developing juveniles (cf. Fig 3.4E, F) with a glistening reflexive appearance, which were not observed at later stages of development, or in the field collected samples containing mature sclerotia.

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Figure 3.4- Cysts and eggs of Heterodera filipjevi infected by Monocillium gamsii exhibiting colonisation in vitro. A, B) infected cysts rendered black-dotted due to fungal-colonised eggs containing microsclerotia. C, D) Eggs with mature sclerotia, extracted from symptomatic cysts. E, F) Individual hyphae penetrating eggshell (arrows indicate the individual hyphae; V indicates vacuole-like structures). G–M) Fungal development inside the eggs: G, H) Earlier stages of infection in unembryonated eggs; I, J) Fungal development in the body cavity of developing juveniles where enlarged, thick-walled cells are formed and coalesce to initiate microsclerotia formation; K–M) Microsclerotia developing to full size and pigmentation. N–P) Pigmentation in microsclerotia from pale- olivaceous to darkly brown. Scale bars: A = 600 µm; B = 300 µm; C–O = 30 µm; P = 50 µm.

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3.3.5 Parasitism of M. bulbillosum towards H. filipjevi

The antagonistic potential of M. bulbillosum was also examined against H. filipjevi in vitro. Eggs of H. filipjevi were infected by M. bulbillosum in the course of 2–4 weeks. The infection symptoms were similar to the symptoms described for M. gamsii. Monocillium bulbillosum rendered cysts black dotted, containing eggs colonised with microsclerotia. In early stages of infection, eggs were entirely colonised with filamentous hyphae which later developed into microsclerotia with a textura angularis on the surface (Figs 3.5A–F).

Figure 3.5- Cysts and eggs of Heterodera filipjevi infected by Monocillium bulbillosum in vitro. A) Symptomatic cysts infected by M. bulbillosum. B) Infected egg showing early stage of fungal colonisation. C–F) Formation of microsclerotia in nematode eggs (arrow points at nematode’s stylet). Scale Bars: A = 300 µm; B–F = 30 µm.

3.4 Discussion

The results obtained from comparative morphological characteristics and molecular phylogenetic inference using four gene regions, suggested M. gamsii as a new species. Within the genus Monocillium, only M. bulbillosum, M. curvisetosum, M. indicum and M. ligusticum have been reported to form (micro-) sclerotia in culture, as is also the case in M. gamsii. In the (micro-) sclerotia forming group, M. gamsii can be separated from M. indicum by producing conidia cohering in watery heads instead of dry conidia. Monocillium curvisetosum produces dry and globose conidia, while M. gamsii produces oblong and watery conidia. The new taxon differs from M. ligusticum by having much shorter phialides: 21–39 µm vs (35–) 40–70 (–140) µm (Girlanda and Luppi-Mosca 1997). The difference between these two species is also strongly supported by sequence comparison (Fig 3.2). According to phylogenetic inference, M. bulbillosum is very closely related to but separable from M. gamsii. Both

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M. gamsii and M. bulbillosum form microsclerotia in culture, however M. bulbillosum forms mainly bulbillose and individually distinct microsclerotia while these structures in M. gamsii are mostly confluent and non-separable. Monocillium gamsii grows slightly faster in comparative growth tests on PDA. In addition, M. gamsii forms setae, and its conidia are clearly longer than those of M. bulbillosum (4.1–7.4 × 1.4–2.9 µm vs 2.9–3.5 × 1.8–2.1 µm). They also differ clearly by the habitat they were originally isolated from. While M. gamsii was isolated from the eggs of nematodes in a semiarid region in the Central Anatolian plateau of Turkey, M. bulbillosum was isolated only once from wall paper in Kiel Germany (Gams 1971). Interestingly though, M. bulbillosum was also able to parasitise eggs of H. filipjevi in our in vitro assays and formed microsclerotia in the infected eggs in a similar manner as M. gamsii. Hyaloseta nolinae (asexual morph: Monocillium nolinae) was included in this study according to a BLASTn search in GenBank, showing a high sequence similarity with the sequences of M. gamsii as query. In the phylogenetic analyses presented here it forms a highly supported monophyletic group with M. gamsii and M. bulbillosum (Fig 3.2). In contrast to M. gamsii, M. nolinae (the asexual morph of H. nolinae) does not form microsclerotia in culture. Hyaloseta Ramaley 2001 was described as a monotypic genus from Asparagaceae (formerly Agavaceae) in New Mexico developing both conidia and ascomata on the fibrous leaves of its host (Ramaley 2001). According to the limited phylogenetic evidence presented here M. gamsii and M. bulbillosum could be transferred to the holomorph genus Hyaloseta. However, the differential characters used to define Hyaloseta in comparison to Niesslia are subtle. Therefore, as long as an extensive molecular phylogenetic analysis of all representatives of Niesslia and Monocillium is pending, it seems less disruptive to place the new species in the ‘anamorph genus’ Monocillium. It is intriguing that microsclerotia, which represent the main symptoms of fungal infection of nematode cysts and eggs in both M. bulbillosum and M. gamsii were readily reproduced in fungal pure cultures and were also formed in artificially infected nematode eggs. Apart from the essential role of conidia in fungal reproduction and dispersal, it seems that microsclerotia also play an important part in the developmental cycle of these fungi, at least with respect to those parts of their life cycle that could be assayed in vitro here and during which it interacts with nematodes. Monocillium gamsii was found in field-collected dried cysts in the semiarid Central Anatolian Plateau. In nature, fungal sclerotia are generally considered as resting structures by which the fungus may tolerate abiotic stresses like dessication, and can thus survive until favourable conditions return. Support for this hypothesis comes from the observation that we were able to isolate M. gamsii from field-collected cysts obtained by culturing the microslerotium-containing infected eggs that had been kept for more than one year in dry conditions either at 4 °C or at room temperature. Furthermore, nematode cysts are protective structures in which nematode eggs can survive for many years in soil in the absence of host plants or

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Monocillium gamsii sp. nov. and Monocillium bulbillosum parasites of Heterodera filipjevi in adverse environments. By colonising the cyst contents, i.e. the mucilaginous matrix and the eggs, the egg-colonising fungi for example M. gamsii and M. bulbillosum, may thus benefit from this “specific” niche where they may have equivalent prolonged-survival conditions. Our microscopic observations of the in vitro tests revealed that M. gamsii is capable of destructively and quickly parasiting the nematode eggs within the cysts first by penetrating the eggshell, followed by prolific formation of microsclerotia. We did not observe formation of any specific infecting structure like in the case of the recently described cyst and egg-parasitic fungus Ijuhya vitellina which developed appressoria (Ashrafi et al. 2017). Incubation of cysts on the colony of M. bulbillosum demonstrated that this species can also parasitise the nematode eggs in a similar manner, and even form microsclerotia. These observations suggest that both species may be candidates for nematode biocontrol.

Acknowledgements

This work was finacially supported by the Federal Ministery for Economic Cooporation and Development, Germany, Grant ID: W0267 GIZ/BMZ-Endophyte as Biocontrol. Additional support of the International Maize and Wheat Improvement Centre (CIMMYT) and the Julius Kühn-Institut are gratefully acknowledged. Special thanks go to the Fiat Panis Foundation, Ulm, Germany, for a “PhD completion stipend” to SA. We thank Dr Thomas Kühne for carefully reading the manuscript, and Anke Brisske-Rode and Katrin Balke for technical support in the molecular studies and culture maintaining.

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3.5 References

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Chapter 4.

Inhabiting plant roots, nematodes and truffles — Polyphilus nom. prov., a new helotialean genus with two globally distributed species

Submitted to Mycologia

Samad Ashrafia,b*, Dániel G. Knappc*, Damien Blaudezd, Michel Chalote,f, Jose G. Maciá-Vicenteg,h, Imre Zagyvai, Abdelfattah A. Dababatj, Wolfgang Maiera, Gábor M. Kovácsc aInstitute for Epidemiology and Pathogen Diagnostics, Julius Kühn-Institut (JKI) — Federal Research Centre for Cultivated Plants, Braunschweig, Germany; bDepartment of Ecological Plant Protection, Faculty of Organic Agricultural Sciences, University of Kassel, Witzenhausen, Germany; cDepartment of Plant Anatomy, Institute of Biology, Eötvös Loránd University, Pázmány Péter sétány 1/C, H-1117 Budapest, Hungary; dUniversité de Lorraine, UMR CNRS 7360 Laboratoire Interdisciplinaire des Environnements Continentaux, Faculté des Sciences et Technologies, BP 70239, F-54506 Vandoeuvre- lès-Nancy, France; eUniversité de Bourgogne Franche-Comté, UMR CNRS 6249 Laboratoire Chrono- Environnement, Pôle Universitaire du Pays de Montbéliard, 4 place Tharradin, BP 71427, F-25211, Montbéliard, France; fUniversité de Lorraine, Faculté des Sciences et Technologies, BP 70239, F-54506, Vandoeuvre-lès-Nancy, France; gInstitute of Ecology, Evolution and Diversity, Goethe Universität Frankfurt, Max-von-Laue-Str. 13, 60438 Frankfurt am Main, Germany; hIntegrative Fungal Research Cluster (IPF), Frankfurt am Main, Germany; iNefag Rt. Nagykunsági Forestry and Wood Industry Rt., Szolnok, Hungary; jCIMMYT (International Maize and Wheat Improvement Centre), P.K.39 06511 Emek, Ankara, Turkey * equally contributed

Keywords ; Hyaloscyphaceae; taxonomy; systematic; endophyte; lifestyle

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Polyphilus nom. prov., a new helotialean genus

Abstract

Fungal root endophytes, including the common form-group of dark septate endophytes (DSEs), represent different taxonomic groups and potentially diverse life strategies. In this study, we investigated two unidentified helotialean lineages found previously in a study of DSE fungi of semiarid , from several other sites, and collected recently from a pezizalean ascoma and from eggs of the cereal cyst nematode Heterodera filipjevi. The taxonomic positions and phylogenetic relationships of 21 isolates with different hosts and geographic origins were studied in detail. Four loci (nrDNA ITS1-5.8S-ITS2 (ITS), partial 28S nrDNA (LSU), partial 18S nrDNA (SSU), and partial RNA Polymerase II Second Largest Subunit (RPB2)) were amplified and sequenced for molecular phylogenetic analyses. Analyses of similar ITS sequences found in public databases revealed that the two lineages are globally distributed and have been detected in several biomes in different geographic regions. The host interaction of isolates from nematodes was examined using in vitro bioassays, which revealed that the isolates could penetrate the nematode cysts and colonize the eggs of H. filipjevi as observed in field-collected samples. This is the first report of a DSE as well as the first helotialean fungal species colonizing the eggs of a plant-parasitic nematode. Neither conidiomata and conidia nor ascomata formation could be detected in any of the isolates from various sources. Based on the results of morphological and molecular phylogenetic analyses, these isolates were found to represent a distinct lineage within the Helotiales in the Hyaloscyphaceae. For this lineage, we propose here the new genus Polyphilus nom. prov. represented by two new species, P. sieberi nom. prov. and P. frankenii nom. prov.

4.1 Introduction

Fungal endophytes asymptomatically colonize plant tissues for a period of their life cycle (Petrini 1991; Saikkonen et al. 1998). They belong to different taxonomic groups and might represent diverse trophic strategies (Porras-Alfaro and Bayman 2011; Rodriguez et al. 2009). Dark septate endophytes (DSEs) compose a form-group of root endophytic fungi with melanized, septate intraradical hyphae (Jumpponen and Trappe 1998). DSE fungi can be found in several orders of Ascomycotina such as Helotiales, and Pleosporales (See Andrade-Linares and Franken (2013); Sieber and Grünig (2013)). DSEs belonging to Helotiales (class Leotiomycetes) are among the most widespread and common root endophytes, like the members of the well-known Phialocephala fortinii s.l.–Acephala applanata species complex (PAC) and several Cadophora species (Grünig et al. 2008; Sieber and Grünig 2013). Helotiales is the largest and most diverse in the Leotiomycetes, with approximately 395 genera belonging to 13 families (Wang et al. 2006). Helotialean fungi show diverse life styles: they can be 88

Polyphilus nom. prov., a new helotialean genus saprobes, parasites of different organisms, endophytes colonizing a broad spectrum of plants, or plant mutualist symbionts forming ectomycorrhizas or ericoid mycorrhizas, as in the case of Pezoloma ericae (syn. Rhizoscyphus ericae) (Wang et al. 2006). Although most helotialean species produce small apothecia, root endophytes have no clear teleomorph connections (Wang et al. 2006). Novel helotialean genera and species are continuously being described in association with a variety of plant hosts and organs; for example, endophytes of pine foliage and shoots (Yuan and Verkley 2015) or associated with horsetail shoots (Baral and Haelewaters 2015), and DSE fungi in the roots of grasses and pines (Walsh et al. 2015; 2014). Two unidentified helotialean phylotypes were detected in a study about root-colonizing fungi from semiarid sandy grasslands of Hungary (Knapp et al. 2012). One lineage, referred to as the “DSE-2 group”, comprised 11 strains isolated from various host plants. The group was considered a DSE group based on in vitro re-synthesis tests, in which the isolates colonized roots, formed microsclerotia and caused no visible negative effect on the plant (Knapp et al. 2012). The other unidentified helotialean root-associated fungus was represented by only one isolate, REF050, classified in the DSE-5 group. This fungus was not considered a DSE based on previous criteria (Knapp et al. 2012). At the time of that study, no nrDNA ITS1-5.8S-ITS2 (ITS barcode) or nrDNA 28S (LSU) sequences were found in public databases with high BLAST similarity to those from REF050. In contrast, sequences of the DSE-2 group were highly similar or identical to sequences of fungi mainly originating from soil and roots of a variety of plants from different continents (Knapp et al. 2012). Later, Glynou et al. (2016) isolated endophytes from roots of Microthlaspi spp. from a wide range of localities in Europe and Turkey, and found some isolates with ITS sequences highly similar to those of DSE-2. These isolates were classified within the operational taxonomic unit (OTU) OTU037, which was sixth in frequency out of the 44 helotialean OTUs obtained in the study (Glynou et al. 2016). Lacercat-Didier et al. (2016) isolated endophytes from ectomycorrhizal poplar roots collected from a metal-contaminated site in France, some of which also showed high sequence similarities to both the DSE-2 clade and the DSE-5 lineage. All those isolates were able to colonize roots of hybrid Populus in re-synthesis experiments. The clade including these and other sequences was described as a “new endophytic putative species belonging to the Helotiales” (Lacercat-Didier et al. 2016). Further strains with high ITS sequence similarity to the DSE-2 clade were isolated from two additional sources: the inner tissue of a truffle ascoma collected from a Hungarian similar to the habitats of the original DSE-2 isolates, and eggs of the cereal cyst nematode (CCN) Heterodera filipjevi originating from the Central Anatolian Plateau of Turkey. We studied the isolates mentioned above to assess the taxonomic novelty of the strains and to determine their phylogenetic relationships. To accomplish this, we carried out a polyphasic taxonomic study using multilocus phylogenetic analyses and morphological comparisons. The results reinforced

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Polyphilus nom. prov., a new helotialean genus our original assumptions that these fungi belong to novel taxa; thus, we describe here a new genus with two novel species. In addition, given the natural occurrence of isolates inside nematode eggs, we examined their interaction with nematodes using in vitro bioassays.

4.2 Material and methods

4.2.1 Origin of isolates

A total of 21 isolates belonging to the DSE-2 group and two isolates belonging to the DSE-5 group sensu Knapp et al. (2012) originating from different hosts and geographic regions (Table 4. 1) were examined in the present study. The sampling sites were previously described in Kovács and Szigetvári (2002), Knapp et al. (2012), Glynou et al. (2016), Lacercat-Didier et al. (2016), and Ashrafi et al. (2017). Pezizalean truffle ascomata (Hydnobolites cf. sp.) found by a trained truffling dog were collected under Festuca, Stipa, Juniperus, Populus, and Salix plants on a semiarid sandy grassland close to Csévharaszt, Hungary during autumn 2015. In the laboratory, the ascomata were carefully washed and broken in half in a sterile box. Small (2–3 mm) tissue pieces from the freshly broken surfaces were placed on modified Melin-Norkrans medium (MMN, Marx (1969)) and kept in the dark at room temperature. After 10 d, small pieces of the mycelia growing out from the tissues were transferred onto new medium. Two isolates with similar colony morphology originating from two different tissue parts were kept for further characterization.

4.2.2 Fungal isolation from nematode eggs

Cysts of H. filipjevi were collected from the rhizosphere of wheat plants. Cysts were extracted from root and soil samples using the modified flotation method (Coyne et al. 2007), directly handpicked from the extracted suspensions under a dissecting microscope, and stored at 4 °C until use. The cysts were examined using a dissecting microscope to separate healthy (homogenously brown) or empty cysts from symptomatic cysts (i.e. cysts bearing black eggs). Fungi were isolated from symptomatic cysts as described in Ashrafi et al. (2017).

4.2.3 Sporulation and morphology of isolates

To induce sporulation, isolates were cultured on MMN, malt extract agar (MEA), potato dextrose agar (PDA), minimal media (MM) (de Vries et al. 2004), Murashige-Skoog agar [MS, (Murashige and Skoog 1962), M5524, Sigma-Aldrich Co. LLC., USA)], synthetic nutrient-poor agar (SNA), oatmeal agar (OA) and cornmeal agar (CMA) (see Crous et al. (2009)), in 9-cm Petri dishes. All cultures were incubated at room temperature in the dark. The majority of strains were kept at low temperature (5–

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Table 4.1- Culture collection numbers, collection sites, hosts, GenBank accession numbers (ITS, LSU, SSU, RPB2) and other details of isolates included in this study

Isolation GenBank accession numbers Species Isolate Strain numbers 1 Collection site Cordinates Host source IST LSU SSU RPB2 Polyphilus sieberi REF051a Fülöpháza, Hungary N 46°52' E 19°25' Fumana procumbens roots JN859271 JN859470 MG719711 MG719732 nom. prov. REF052a DSM 106515 (ex- Fülöpháza, Hungary N 46°52' E 19°25' Asclepias syriaca roots JN859272 MG719692 MG719712 MG719733 type) REF053a Tatárszentgyörgy, N 47°03.5' E Ailanthus altissima roots JN859273 MG719693 MG719713 MG719734 Hungary 19°24.3' REF054a Fülöpháza, Hungary N 46°52' E 19°25' Helianthemum ovatum roots JN859274 MG719694 MG719714 MG719735 REF056a DSM 106516 Fülöpháza, Hungary N 46°52' E 19°25' Juniperus communis roots JN859276 MG719695 MG719715 MG719736 REF057a Fülöpháza, Hungary N 46°52' E 19°25' J. communis roots JN859277 MG719696 MG719716 MG719737 REF058a Tatárszentgyörgy, N 47°03.5' E J. communis roots JN859278 MG719697 MG719717 MG719738 Hungary 19°24.3' REF059a Fülöpháza, Hungary N 46°52' E 19°25' J. communis roots JN859279 JN859472 MG719718 MG719739 M03b DSM 106519 Evin-Malmaison, N 50°43.6’ E 3°01.3’ Populus deltoides x P. roots MG719684 MG719698 MG719719 MG719740 France nigra P02b Pierrelaye, France N 49°02.8’ E 2°17.6’ P. deltoides x P. nigra roots MG719685 MG719699 MG719720 MG719741 P06b Pierrelaye, France N 49°02.8’ E 2°17.6’ P. deltoides x P. nigra roots MG719686 MG719700 MG719721 MG719742 17Ac DSM 106517 Yozgat, Turkey N 39° 08ʼ E 34°10ʼ Heterodera filipjevi cyst/egg MG719687 MG719701 MG719722 MG719743 17Cc Yozgat, Turkey N 39° 08ʼ E 34°10ʼ H. filipjevi cyst/egg MG719688 MG719702 MG719723 MG719744 P1582d DSM 106518 Diviya, Bulgaria N 42°34.2' E Microthlaspi erraticum roots KT268876 MG719703 MG719724 MG719745 22°41.4' P2020e Challement, France N 47°18.0' E 3°35.4' Microthlaspi sp. roots KT269283 MG719704 MG719725 MG719746 P2262e Vivonne, France N 46°24.6' E 0°13.2' M. perfoliatum roots KT269499 MG719705 MG719726 MG719747 P2418e Vivonne, France N 46°24.6' E 0°13.2' M. perfoliatum roots KT269651 MG719706 MG719727 MG719748 TT3Af Csévharaszt, N 47°17.2' E Hydnobolytes cf. sp. ascoma MG719689 MG719707 MG719728 MG719749 Hungary 19°23.8' TT3Bf Csévharaszt, N 47°17.2' E Hydnobolytes cf. sp. ascoma MG719690 MG719708 MG719729 MG719750 Hungary 19°23.8' P. frankenii nom. REF050a DSM 106520 (ex- Bugac, Hungary N 46°39.1' E J. communis roots JN859270 MG719709 MG719730 MG719751 prov. type) 19°36.4' V16b DSM 106521 Villerupt, France N 49°48.1’ E 5°93.5’ Pinus sylvestris roots MG719691 MG719710 MG719731 MG719752 1DSM: DSMZ-Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH, Braunschweig, Germany; a Isolates collected by Dániel G. Knapp and Gábor M. Kovács; b Isolates collected by Damien Blaudez and c d e Michel Chalot; Isolates collected by Samad Ashrafi; plants collected by Tahir Ali and Sebastian Ploch, isolates obtained by Kyriaki Glynou and Jose G. Maciá-Vicente; Isolates collected by K. Glynou and J.G. Maciá- f Vicente; Fruit-bodies collected by Imre Zagyva, isolates gained by DG. Knapp and GM. Kovács.

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6 °C) for 11 months. Isolates were also cultured on autoclaved stems of stinging nettle (Urtica dioica), as sexual morphs of another DSE could be induced on these (see Knapp et al. (2015)). Horsetail (Equisetum arvense) shoots were also used, because a phylogenetically related species was described from this substrate (see Baral and Haelewaters (2015)). Colony blocks of representative isolates were floated in sterile water for 10 wk. Colony morphology was assessed on MMN and MEA media. Agar plates were inoculated in three replicates with 4-mm diameter mycelial plugs of each isolate in 6-cm Petri dishes at room temperature and the colonies were examined after 3 wk.

4.2.4 DNA extraction and amplification

Genomic DNA was extracted from fungal mycelia using a modified CTAB method (Kovács et al. 2001; Murray and Thompson 1980) or the E.Z.N.A. ® Fungal DNA Mini Kit (OMEGA Bio-tek, Norcross, Georgia) following the manufacturer’s instructions. To extract fungal DNA from nematode eggs, single eggs were transferred to 1.5-ml Eppendorf microtubes containing 5 µl DNA-free water, 40 mg sterile silica sand and four steel beads. Each sample was first air-dried under a laminar flow, frozen by dipping the tubes in liquid nitrogen, and homogenized at 50 Hz for 2 min (Qiagen TissueLyser LT, Hilden, Germany). Freezing and homogenization were repeated three times. DNA was then extracted using the Qiagen DNeasy Plant Mini kit following the manufacturer’s instructions. The nrDNA ITS1-5.8S-ITS2 (ITS), partial 28S nrDNA (LSU), partial 18S nrDNA (SSU) and partial RNA Polymerase II Second Largest Subunit (RPB2) regions of the isolates were amplified using DreamTaq® polymerase (Thermo Fisher Scientific, Lithuania). The ITS and LSU regions of nrDNA were amplified using the primer pair ITS1F (Gardes and Bruns 1993) and ITS4 (White et al. 1990), and the primers LR0R (Rehner and Samuels 1994) and LR5 (Vilgalys and Hester 1990), respectively, as described previously (Kovács et al. 2008). The primers NS1 and NS4 (White et al. 1990) were used to amplify the SSU region, and the partial RPB2 region was amplified using the primer pair RPB2-6F and RPB2-7R (Liu et al. 1999). Sequencing of the amplicons was carried out with the primers used for amplification by LGC GmbH (Berlin, Germany), Eurofins Genomics (Ebersberg, Germany) and GATC Biotech (Konstanz, Germany).

The sequences were compiled from electropherograms using the Pregap4 and Gap4 tools of the STADEN software package (Staden et al. 2000) and SEQUENCHER 5.4 (GeneCodes Corporation, Ann Arbor, MI), respectively, and deposited in GenBank (MG719684–MG719752). The sequences obtained were compared with sequences in public databases using BLASTn searches (http://blast.ncbi.nlm.nih.gov/Blast.cgi) (Altschul et al. 1990).

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4.2.5 Phylogenetic analyses

Three datasets were prepared for phylogenetic analyses: (i) an order-level dataset was used to study the phylogenetic position of the isolates among related helotialean taxa; (ii) a second dataset was established to study the intra-group phylogenetic relationships of the isolates studied here; (iii) the third dataset included representative ITS sequences of our isolates and similar sequences obtained from public databases via a BLAST search. Alignments of our sequences and the sequences of corresponding entries from GenBank were assembled using the E-INS-i method in MAFFT version 7 (Katoh and Standley 2013). The alignments were checked and edited with MEGA6 (Tamura et al. 2013) and deposited in TREEBASE (S22024; www.treebase.org). For the three datasets, multi-locus phylogenetic Bayesian analyses were performed with MRBAYES 3.1.2 (Ronquist and Huelsenbeck 2003) using the GTR+G nucleotide substitution model. Four Markov chains were run for 10 000 000 generations and sampled every 1000 generations with a burn-in value set at 6000 sampled trees. Topological convergence was checked by

AWTY online (Nylander et al. 2008). Maximum likelihood (ML) phylogenetic analyses were carried out with the RAXMLGUI 1.3 (Silvestro and Michalak 2012) implementation of RAxML (Stamatakis 2014). The GTR+G nucleotide substitution model was used for the partitions with ML estimation of base frequencies, and a ML bootstrap analysis with 1000 replicates was used to test the support for the branches. The phylogenetic trees were visualised and edited using MEGA6 (Tamura et al. 2013). The ITS, LSU, SSU and RPB2 sequences were used for order-level phylogenetic analyses, and considered as four partitions. These four regions were used to determine the intra-group phylogenetic relationships of our isolates with the methods described above. To improve phylogenetic resolution, indels in the ITS region were coded (Nagy et al. 2012) using the simple indel coding algorithm (Simmons et al. 2001; Young and Healy 2003) with the program FASTGAP (Borchsenius 2009). In the Bayesian analysis, the two-parameter Markov (Mk2 Lewis) model was used for the indel partition of the dataset and the GTR+G model for the nucleotide partitions. In the ML analyses, in addition to the nucleotide partitions (GTR+G), the indel data were treated as binary data (BIN). The third dataset comprised only ITS sequences of representative DSE-2 isolates and similar GenBank sequences; the parameters and methods used for phylogenetic analyses were the same as mentioned above.

4.2.6 Pathogenicity against H. filipjevi

To assess the pathogenicity of the isolates (17A and 17C) originating from nematode cysts against H. filipjevi, in vitro tests were performed to fulfill Koch's postulates. For this, healthy cysts or eggs were placed either directly on or at the edge of fungal colonies, or a modified version of Riddell’s slide

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Polyphilus nom. prov., a new helotialean genus culture technique (Gams et al. 1998) was applied to test the ability of the obtained isolates to colonize H. filipjevi eggs. The details of the tests are described in Ashrafi et al. (2017).

4.2.7 Light microscopy

Symptomatic cysts and eggs of H. filipjevi were mounted in water and examined with an Olympus SZX 12 dissecting microscope and a Zeiss Axioskop 2 plus compound microscope using Nomarski Differential Interference Contrast (DIC); both were equipped with a Jenoptik ProgRes® digital camera (Jenoptik, Jena, Germany). Nematode cysts and eggs were photographed in water. Microscopic photographs were recorded using the CAPTUREPRO 2.8 software (Jenoptik), and the images were processed using the Adobe PHOTOSHOP software CS 5.1.

4.3 Results

4.3.1 Sporulation and colony morphology

All isolates showed similar colony morphology, growth characteristics and color. The colonies on MMN media were slow-growing, radially striate, and bright cream to white; no exudates were present (Fig 4.1). Numerous media and culture conditions were tested and different autoclaved plant organs were inoculated to induce sporulation. Neither conidiomata nor conidia nor ascomata formation could be detected. As the cultures aged, spherical monilioid cells formed in the colonies on MMN. No sporulation of the strains was observed in any of the media or conditions tested, or in the four different laboratories where the strains were isolated, maintained and kept e.g. at 4–7 C for 6–18 months before this study.

4.3.2 Molecular phylogeny

Based on the results of the order-level phylogenetic analyses, the clade containing the isolates studied here was unambiguously grouped in the order Helotiales, within the poorly resolved family Hyaloscyphaceae s.l. (Fig 4.2). The fungi grouped with different incertae sedis clades and several distinct genera (Fig 4.2). The clade comprising our isolates formed a monophyletic group with Belonioscyphella hypnorum, sp., C. albidolutea, Leohumicola sp., L. minima, L. verrucosa, sp., P. staphyleae, Rodwayella citrinula, Rommelaarsia flavovirens, Roseodiscus rhodoleucus, R. subcarneus, Tricladium angulatum and Urceolella carestiana. However, this clade only received strong support by Bayesian inference, and was not significantly supported by bootstrapping in the RAXML analysis.

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Figure 4.1- Colonies of Polyphilus nom. prov. species on modified Melin-Norkrans agar (MMN) A. Polyphilus sieberi nom. prov. (REF052 = DSM 106515). B. Polyphilus frankenii nom. prov. (REF052 = DSM 106520).

The intra-group phylogenetic analyses of the isolates studied here revealed two well-supported clades (Supplementary Fig 1). The first clade comprised the majority of isolates (19) with moderate intra-clade heterogeneity (Supplementary Fig 1). The second clade included two isolates, REF050 and V16. The difference in the ITS and RPB2 sequences of the two clades was 2.8–3.3% and 3.1–3.3%, respectively. The results of the molecular phylogenetic analyses reinforced our hypothesis that this clade represents a novel helotialean genus, in which we describe two species. Several identical or nearly identical ITS sequences deposited in public databases were found during similarity searches of the ITS sequences of our strains. These similar sequences represent uncultured fungi and isolates derived mainly from soil and roots of various host plants (Fig 4.3). Both clades of our isolates grouped with sequences originating from different geographic and climatic regions and from different continents (Fig 4.3). The first clade contained the majority of our isolates, including those that originated from nematode eggs and truffles. These grouped with sequences obtained mainly from plant-associated fungi (Fig 4.3). The second clade, comprising our REF050 and V16 isolates, included sequences originating mostly from plant-associated fungi, soil samples, and stromata of the entomopathogenic fungus Ophiocordyceps sinensis (Fig 4.3).

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Figure 4.2- Maximum Likelihood (RAXML) tree of concatenated ITS, LSU, SSU and RPB2 sequences of representative isolates of Polyphilus nom. prov. species and related fungi (best blast matches, and representative taxa of related lineages) from Hyaloscyphaceae s. l. (Helotiales). Bayesian posterior probabilities (≥ 0.90) are shown above branches and before slashes; RAXML bootstrap support values (≥ 70) are shown below branches and after slashes. Ex-type strains are shown in bold. Phialocephala fortinii (CBS 443.86) served as an outgroup. The scale bar indicates expected changes per site per branch.

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4.3.3 Isolation of fungi from H. filipjevi

Isolates 17A and 17C were obtained from naturally infected eggs of H. filipjevi. In infected cysts, the fungus caused black discoloration, which could clearly be seen using a stereomicroscope. Infected cysts contained mostly dark brown eggs, which were entirely colonized by distinctly septate, moniliform, thick-walled, and strongly melanized hyphae (Fig 4.4A, B). Some infected eggs displayed no discoloration and were found to be colonized by hyaline hyphae containing numerous oil-like bodies (Fig 4.4C). When cultured on PDA, mycelium emerged from infected eggs and formed green-grayish to olivaceous gray cultures.

Figure 4.3- Maximum likelihood (RAXML) tree based on ITS sequences of representatives of Polyphilus nom. prov. species and similar sequences from GenBank. Sequences obtained in this study are shown in bold. After the accession number and sequence name, the host, isolation source and geographic origin of each sequence is shown in brackets. Bayesian posterior probabilities (≥ 0.90) are shown above branches and RAXML bootstrap support values (≥ 70) are shown below branches. Roseodiscus subcarneus (voucher D. Haelew 314a, KT92714) was used as outgroup. Abbreviations: uncultured (u.), clone (c.), isolate (is.), strain (st.), voucher (v.), surface sterilized (s. s.) ectomycorrhizal (EcM.). Sequences of uncultured fungi are marked by diamonds (). The scale bar indicates expected changes per site per branch. 97

Polyphilus nom. prov., a new helotialean genus

4.3.4 Pathogenicity against H. filipjevi

In vitro tests revealed that isolates 17A and 17C could penetrate the nematode cysts and colonize the eggs of H. filipjevi (Fig 4.4D). Cysts placed directly on the fungal mycelium were entirely colonized by ‘runner hyphae’ growing in the cyst mucilage (Fig 4.4E). The colonization of eggs of the incubated cysts or eggs placed on or in the vicinity of the fungal mycelium was initiated by the formation of enlarged, irregularly lobed and hyaline or lightly pigmented hyphae containing many oil-like bodies; these hyphae eventually occupied the entire host body cavity (Fig 4.4F, G). Infected eggs became pigmented upon fungal development, from pale greenish gray to greenish brown (Fig 4.4F, H). The artificially infected eggs resembled naturally infected samples (Fig 4.4A). Slide culture-based observations revealed that eggs were infected within 7–10 d after placing them on the margin of a growing mycelium. Infection started with individual hyphae penetrating the egg and nematode body cavity. Following penetration, the hyphae developed and formed enlarged cells filled with oil-like structures and destroyed the egg content (Fig 4.4I, J). Accumulation of oil-like structures was generally more extensive in the hyaline hyphae than in the pigmented ones. The fungus could also enter the body cavity of juvenile nematodes through natural openings such as the mouth aperture (Fig 4.4J).

4.3.5 Taxonomy

Polyphilus D.G. Knapp, Ashrafi, W. Maier & Kovács, nom. prov. Figs 4.1–4 Typification: Polyphilus sieberi nom. prov. Ashrafi, D.G. Knapp, W. Maier & Kovács. Etymology: Polyphilus nom. prov., from the ancient Greek words “poly” (more than one, much and many) and “philia” (brotherly love, like, having an affinity for something), referring to the fact that the strains of the species have been isolated from many different sources and geographic regions. Notes: The genus Polyphilus nom. prov. contains species associated with plant roots as endophytes, plant parasitic nematodes and truffles. Isolates can be collected from surface-sterilized roots, surface- sterilized nematode eggs, and truffle ascomata, and can be cultured and maintained on general media. These fungal isolates do not produce fruiting bodies or spores of any kind. Colony morphology is quite similar among Polyphilus nom. prov. spp. including shape, growth characteristics, color and absence of sexual and asexual spores (Fig 4.1). Colonies are radially striate, bright grey to white and slow-growing (Fig 4.1). Isolates were collected from surface-sterilized roots of common juniper (Juniperus communis), sprawling needle sunrose (Fumana procumbens), common milkweed (Asclepias syriaca), tree of heaven (Ailanthus altissima), Helianthemum ovatum, hybrid poplar (Populus deltoides × P. nigra), scots pine (Pinus sylvestris), Microthlaspi perfoliatum, M.

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Polyphilus nom. prov., a new helotialean genus erraticum, Microthlaspi sp., from surface-sterilized eggs of the CCN Heterodera filipjevi, and from a truffle (cf. Hydnobolites sp.) ascoma. Phylogenetically, Polyphilus nom. prov., currently comprising two new species described below, forms a well-supported monophyletic group that clusters incertae sedis within Hyaloscyphaceae.

Polyphilus sieberi Ashrafi, D.G. Knapp, W. Maier & Kovács, nom. prov. Figs 4.1–4 Typification: HUNGARY. Fülöpháza: Kiskunság National Park, in roots of Asclepias syriaca, Jul 2008, D.G. Knapp & G.M. Kovács, REF052 (holotype BP109051, deposited to the herbarium of the Hungarian Natural History Museum). Ex-type culture DSM 106515. GenBank: ITS = JN859272; LSU = MG719692; SSU = MG719712; RPB2 = MG719733. Etymology: We name this species in honor of Dr. Thomas N. Sieber, who has contributed significantly to our knowledge on root endophytes and DSE fungi. Description: Colonies on MMN slow-growing, 2.4–3.4 mm diam after 21 d at room temperature in the dark; raised, surface smooth or with sparse aerial mycelium, even margins, colony radially striate and wrinkled, bright grey to white, creamy, no exudates, no medium staining; pale amber reverse; cultures sterile (Fig 4.1). Colonies on MEA slow-growing, 2–2.8 mm diam after 21 d in the dark; centrally elevated, dense aerial mycelium, colony radially striate and wrinkled with lobate margins, light grey to smoke, no exudates; iron gray reverse; cultures sterile. P. sieberi nom. prov. (DSM 106515 = REF052) differs from one of its closest phylogenetic neighbors, Urceolella carestiana (TNS-F18014), by unique fixed alleles in the ITS, LSU and RPB2 loci based on alignments of each locus deposited in TreeBASE as study S22024: ITS positions: 83 (deletion), 171 (T), 415 (C), 424 (G), 449 (T), 488 (A); LSU positions: 445 (C), 520 (C); RPB2 positions: 124 (G), 295 (C), 313 (C), 397 (C), 445 (T), 616 (T), 700 (A). Other specimens examined: HUNGARY, Fülöpháza, in roots of Fumana procumbens, Jul 2008, D.G. Knapp & G.M. Kovács REF051. Tatárszentgyörgy, in roots of Ailanthus altissima, REF053. Fülöpháza, in roots of Helianthemum ovatum, Apr 2008, REF054. in roots of Juniperus communis, REF056 (= DSM 106516). Jul 2008, REF057. Tatárszentgyörgy, REF058. Fülöpháza, REF059. in fruiting bodies of cf. Hydnobolites sp., Nov 2015, TT3A. ibid., TT3B. FRANCE, Evin-Malmaison, in roots of Populus deltoides × P. nigra hybrid, Jun 2011, D. Blaudez & M. Chalot, M3 (= DSM 106519). Pierrelaye, P02. ibid., P06. TURKEY, Yozgat, in eggs of Heterodera filipjevi, Aug 2013, S. Ashrafi, 17A (=DSM 106517). ibid., 17C. BULGARIA, Diviya, in roots of Microthlaspi erraticum, May 2013, K. Glynou & J.G. Maciá-Vicente, P1582 (= DSM 106518). FRANCE, Challement, in roots of Microthlaspi sp., P2020. Vivonne, in roots of Microthlaspi perfoliatum, P2262. ibid., P2418.

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Figure 4.4- Pathogenicity of Polyphilus sieberi nom. prov. towards Heterodera filipjevi. A–C. Fungus-infected eggs originally isolated from field-collected symptomatic cysts, detailing hyphal cells. D. Symptomatic cyst obtained from in vitro tests, containing blackish eggs infected with the fungus. E. Runner hyphae of P. sieberi nom. prov. growing in cyst mucilage. F–H. In vitro infection process of P. sieberi nom. prov. in nematode eggs: hyphae forming enlarged, thick-walled and lobate cells containing oil-like bodies. Note the pigmentation process during the early (F, G) and late (H) stages of colonization. I, J. Fungal colonization observed in slide cultures: (I) runner hyphae that penetrated the nematode egg enlarged and became filled with distinct oil-like bodies; (J) penetration of hyphae through the body cuticle and the mouth opening of a nematode juvenile, detailing the formation of enlarged hyphal cells inside the body cavity (J). (arrowed: nematode stylet). Scale bars: A, C, F–J = 30 µm; B, E = 10 µm; D = 0.6 mm.

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Polyphilus frankenii D.G. Knapp, Ashrafi, W. Maier & Kovács, nom. prov. Figs 4.1–3 Typification: HUNGARY. Fülöpháza: Kiskunság National Park, in roots of Juniperus communis, Apr 2008, D.G. Knapp & G.M. Kovács, REF050 (holotype BP109050, deposited to the herbarium of the Hungarian Natural History Museum). Ex-type culture DSM 106520. GenBank: ITS = JN859270; LSU = MG719709; SSU = MG719730; RPB2 = MG719751. Etymology: We name this species in honor of Dr. Philipp Franken, who has contributed significantly to our knowledge on root endophytes and DSE fungi and on their interaction with plants. Description: Colonies on MMN slow-growing, 2.5 mm diam after 21 d at room temperature in the dark; raised, surface smooth or with sparse aerial mycelium, even margins, colony radially striate and wrinkled, light grey to white, no exudates, no staining of the medium; pale amber reverse; cultures sterile (Fig 4.1). Colonies on MEA slow-growing, 2.2 mm diam after 21 d in the dark; centrally elevated, dense aerial mycelium, colony radially striate and wrinkled with lobate margins, light gray to smoke, no exudates; iron gray reverse; cultures sterile. P. frankenii nom. prov. (DSM 106520 = REF050) differs from P. sieberi nom. prov. (DSM 106515 = REF052) by unique fixed alleles in the ITS, LSU, SSU and RPB2 loci based on alignments of each locus deposited in TreeBASE as study S22024: ITS positions: 115 (T); 116 (T), 159 (T), 187 (C), 191 (T), 222 (A), 424 (C), 430 (G), 455 (T), 493 (A), 499 (A), 500 (C), 517 (T); LSU positions: 471 (C), 545 (T), 546 (C); RPB2 positions: 54 (T), 71 (C), 80 (A), 126 (G), 147 (G), 156 (C), 180 (T), 276 (A), 297 (C), 315 (C), 348 (C), 399 (C), 444 (G), 447 (A), 588 (G), 618 (G), 699 (C), 702 (A), 770 (C), 783 (A), 785 (A), 787 (A), 792 (G). Other specimens examined: FRANCE, Villerupt, in roots of Pinus sylvestris, Nov 2011, D. Blaudez & M. Chalot, V16 (= DSM 106521).

4.4 Discussion

The two species of the new genus described here belong to Helotiales and, phylogenetically, appear nested in a diverse group in the Hyaloscyphaceae s.l. (Fig 4.2). The new taxa are grouped within a clade that comprises most of the taxa of “Clade-9” sensu Han et al. (2014), which contained Cistella sp., C. albidolutea, Psilachnum sp., P. staphyleae, Rodwayella citrinula, Urceolella carestiana and U. crispula. However, in our analyses U. crispula did not group with members of Clade-9, but formed a clade with herbarum and populinum, which belonged to “Clade-2” in Han et al. (2014). Clade- 9 received strong support in Han et al. (2014), but in our analysis the group was only supported by Bayesian inference when supplemented with further genera (Belonioscyphella, Leohumicola, Rommelaarsia, Roseodiscus). The representatives belonging to Clade-9 (Han et al. 2014) were collected

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Polyphilus nom. prov., a new helotialean genus from various materials and do not exhibit any unifying morphological characteristics. In contrast, the concepts of the individual genera within this group are based on teleomorph morphology (e.g. Han et al. (2009)). Species within the clade with Polyphilus nom. prov. spp. were isolated from diverse substrates: the Psilachnum species were collected from tree leaves (Han et al. 2014), Urceolella carestiana was collected from the rachis of a maiden fern (Thelypteris nipponica) (Han et al. 2014) and Cistella sp. was obtained from a corncob (Zea mays). Both Cistella albidolutea and Rodwayella citrinula were isolated from the sheath of a Carex plant. Belonioscyphella hypnorum is a necrotrophic parasite of various mosses and develops apothecia mostly on stems in the upper part of shoots (Huhtinen et al. 2010). Roseodiscus species can be isolated from mosses and horsetail ferns in bogs and swamps and might be both parasitic and saprotrophic (Baral and Krieglsteiner 2006). Rommelaarsia flavovirens produces apothecia on dead stems of Equisetum (Baral and Haelewaters 2015). Leohumicola species are soil fungi isolated from heated soils in Canada (Hambleton et al. 2005). Tricladium angulatum is a freshwater fungus isolated from foam (Baschien et al. 2006) that represents a distinct clade among Tricladium species (Baschien et al. 2006; Campbell et al. 2009) and seems to have affinity to Hyaloscyphaceae s.l. (Campbell et al. 2009). Only Leohumicola and the Polyphilus nom. prov. spp. were mainly isolated from soils or underground materials in semiarid regions; all other representatives of the clade were isolated from aboveground parts of land plants and from humid habitats. Thus, the species of this clade tend to occur in humid to wet habitats in swamps, wet soils and even in freshwater, inhabiting mosses, ferns, horsetails, sedges and water plants (Baral and Haelewaters 2015; Baral and Krieglsteiner 2006; Campbell et al. 2009; Han et al. 2014). In contrast, the majority of Polyphilus nom. prov. isolates described in this study were collected in semiarid regions such as the isolates from roots and truffles in sandy grasslands, from roots of semiarid poplar stands and from CCN eggs in the semiarid Central Anatolian plateau. However, the sequences from public databases clustering with the two Polyphilus nom. prov. species described here represent mainly root-inhabiting fungi with more diverse habitat and host origins (Fig 4.3). There has been a continuous increase in taxon numbers in Helotiales, but the last comprehensive order-level analysis was published more than a decade ago (Wang et al. 2006). Although family-level analyses and studies have been carried out (e.g. Han et al. (2014)) and nomenclatural recommendations and clarifications have been published (Johnston et al. 2014), the statement in the 10th edition of the Dictionary of the Fungi (Kirk et al. 2008) that “the taxonomy of the order is unsettled, and molecular data are inadequate to elucidate relationships in many cases” seems to be still valid. Although P. sieberi nom. prov. and P. frankenii nom. prov. differed significantly in each locus examined, the morphology of the two species was highly similar. They shared growth characteristics and could not produce spores on the different media and tissues tested, like the majority of DSE fungi (Sieber

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Polyphilus nom. prov., a new helotialean genus and Grünig 2013). Spherical monilioid cells developed as the colonies of both species aged, which was also reported previously (Lacercat-Didier et al. 2016). The new species P. frankenii nom. prov. was represented by two isolates originating from plant roots. Based on the results of a previous re-synthesis experiment, REF050 isolated from surface-sterilized roots of Juniperus communis was not considered a DSE (Knapp et al. 2012). The fungi represented by ITS sequences forming a clade with P. frankenii nom. prov. originated from several continents and mainly from roots, whereas only a few were isolated from soil directly. In contrast, the isolates belonging to P. sieberi nom. prov. were considered DSEs based on in vitro colonization tests (Knapp et al. 2012), and were even more diverse in origin. Other root-associated P. sieberi nom. prov. isolates originated from poplar (Lacercat-Didier et al. 2016) and Microthlaspi spp. (Glynou et al. 2016), respectively. Isolates originating from a truffle ascoma also belonged to P. sieberi nom. prov. Truffles can be complex microhabitats hosting bacteria, yeasts and filamentous fungi that can be isolated from healthy and intact ascomata (Pacioni and Leonardi 2016). For example, (Pacioni et al. 2007) isolated filamentous fungi from fresh and healthy truffles and found several ascomycetes and basidiomycetes, including some entomopathogenic and endophytic species. Different genera and species of fungal entomopathogens and parasites of other invertebrates have been reported as naturally occurring fungal endophytes (Sikora et al. 2008; Vega 2008). Several insect- parasitic fungi such as Acremonium, Beauveria and Metarhizium species have been detected in healthy organs of different plant species (Barelli et al. 2016; Foulon et al. 2016; Vega 2008). These fungi mainly belong to the order Hypocreales, which contains endophytic and insect-parasitic representatives sharing common ancestors (Gao et al. 2011; Spatafora et al. 2007). It was also reported that the colonization of plants by particular endophytic fungi can improve the defense of the host toward nematodes (Schouten 2016), and provide nutrient benefits to the host plant (Barelli et al. 2016; Behie et al. 2017; 2012). Although the interactions of class-1 and class-2 endophytes sensu Rodriguez et al. (2009) with nematodes were discussed previously (Rodriguez et al. 2009; Schouten 2016), no nematode-parasitic fungus has yet been reported from the class-4 endophytes comprising DSE fungi. Thus, to our knowledge P. sieberi nom. prov. is the first DSE, as well as the first helotialean fungal species, reported to colonize the eggs of a plant-parasitic nematode. Nematode cysts protect the nematode eggs from adverse environmental conditions. DSEs are considered abundant in habitats with harsh environmental conditions including alpine, arctic, arid and semiarid regions (Mandyam and Jumpponen 2005; Rodriguez et al. 2009). For soil fungi, cysts or truffle ascomata may serve as microenvironments where they have better conditions for survival. The interior mucilaginous matrix of cyst nematodes has been reported to contain some glycoconjugates and lectins (Nour et al. 2003), the latter of which is assumed to play a role in nematode recognition by fungi (Li et al. 2015). The mucilage complex of plant roots was also reported to contain carbohydrates that are

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Polyphilus nom. prov., a new helotialean genus involved in water storage, and thus protecting the roots and perhaps root-colonizing fungi from desiccation (Barrow 2003). It is hypothesized that the colonization of cyst mucilage by P. sieberi nom. prov. can help the fungus to avoid potential desiccation. Observations from our in vitro experiments showed that isolates belonging to P. sieberi nom. prov. could entirely colonize the cyst cavity by growing “runner hyphae” into the cyst mucilage. After penetration of the egg, the hyphal cells differentiated into distinct moniliform, thick-walled cells filled with oil-droplet structures, which were absent in the runner hyphae. These structures might be involved in nutrient acquisition and translocation from the host (Barrow 2003; Jumpponen and Trappe 1998), as well as in nematode-killing and degradation of nematode body components (Nordbring-Hertz et al. 2011). Nematode colonization by P. sieberi nom. prov. might therefore provide advantages for nematode host plants, by directly killing eggs and thus reducing the parasite load or by transferring nutrients from the degraded eggs and juveniles to the plant as was reported for the endophytic fungus Metarhizium robertsii and insects (Behie et al. 2012). Polyphilus nom. prov., the novel genus described here, is a globally distributed Hyaloscyphaceae genus derived from soils that is primarily associated with plant roots based on the specimens isolated here and on the conspecific or congeneric fungal sequences identified from public databases. Nevertheless, two distinct isolation sources—nematode cysts and a truffle ascoma—suggest that these fungi might also colonize other organisms in soil. The ecological functions of these interactions as well as possible multitrophic interactions should be tested in experimental systems.

Acknowledgemnets

This work was supported by the Hungarian Scientific Research Fund (NKFIH/OTKA K109102), the Deutsche Gesellschaft für Internationale Zusammenarbeit (GIZ) and the International Maize and Wheat Improvement Centre (CIMMYT). The project was also supported by the French ANR Research Program [grant number ANR BIOFILTREE 2010-INTB-1703-01 to MC] and by the Agence de l’Environnement et de la Maitrise de l’Energie (ADEME) [grant number PROLIPHYT 1172C0053 to MC]. JGMV acknowledges LOEWE (Landes-Offensive zur Entwicklung Wissenschaftlich-ökonomischer Exzellenz) of the state of Hesse for financial support within the framework of the Cluster for Integrative Fungal Research (IPF). Special thanks go to the Fiat Panis Foundation, Germany (stiftung-fiat-panis.de) for a “PhD completion stipend” to SA. We thank Anke Brisske-Rode and Katrin Balke for technical assistance and Dr. Csaba Locsmándi and Dr. Andrey Yurkov, curators of the Hungarian Natural History Museum (BP) and the Leibniz Institute DSMZ-German Collection of Microorganisms and Cell Cultures (DSMZ), respectively, for their help. We appreciate Dr. Shaun R. Pennycook’s help in nomenclatural questions.

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White TJ, Bruns T, Lee S, Taylor J (1990) Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: Innis M, Gelfand D, Sninsky J, White T (Eds) PCR Protocols: a guide to methods and applications. Academic Press, San Diego, 315-322. Young ND, Healy J (2003) GapCoder automates the use of indel characters in phylogenetic analysis. BMC Bioinformatics 4: 6. Yuan Z, Verkley GJM (2015) Pezicula neosporulosa sp. nov. (Helotiales, Ascomycota), an endophytic fungus associated with Abies spp. in China and Europe. Mycoscience 56: 205-213.

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4.6 Supplementary

Supplementary Figure 1- Maximum Likelihood (RAXML) tree of Polyphilus nom. prov. species inferred from concatenated ITS, LSU, SSU and RPB2 sequences with coded indels for the ITS region. Bayesian posterior probabilities (≥ 0.90) are shown above branches and before slashes; RAXML bootstrap support values (≥ 70) are shown below branches and after slashes. Type strains are shown in bold. Roseodiscus subcarneus (voucher 314a) served as outgroup. The scale bar indicates expected changes per site per branch.

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Chapter 5.

General Discussion

5.1 Introduction

Cyst nematodes were the first plant parasitic nematodes (PPNs) that had been screened for nematophagous fungi. In 1877, Kühn found Catenaria auxiliaris parasitising the sugar beet cyst nematode Heterodera schachtii (Kühn 1877). While discussing the potential impact of Catenaria auxiliaris on the nematode, Kühn (1881) coined the nematophagous fungi “hilfreiche Hilfstruppen” (valuable auxiliary troops) against nematode infection, stating that “nematodes also have natural enemies” which could help controlling the negative effects of PPNs. Later, these pioneering observations drew extensive attention to the potential of nematophagous fungi for the biocontrol of nematodes (Tribe 1977). Although many nematophagous fungi attacking eggs of cyst-forming nematodes (Chen and Chen 2002; Jatala 1986; Siddiqui and Mahmood 1996; Stirling 2014; Tribe 1977) were found over several decades of research in this field, new discoveries can still be made. The outcome of the study presented here with the finding of several new fungi, which can destructively parasitise nematode eggs, demonstrated this. To achieve the aims of the project two screening approaches termed as ‘general’ and ‘focused’ screening were applied with a great emphasis on focused screening. The results of the first phase of the study (general screening) were not presented here since the main interest during the project was on the focused screening and the first three publications dealt with the results obtained from this approach accordingly. The fungi isolated in the general screening phase were well-known species that had previously been demonstrated to associate with nematodes as egg-parasitic fungi, e.g. Alternaria sp., Fusarium oxysporum, Paecilomyces fumosoroseus, Pochonia chlamydosporia, Purpureocillium lilacinum, and Trichoderma species. The number fungi obtained in this screening phase suggests the presence of a rich mycobiota associating with wheat roots and nematode eggs. These species can be considered as one key component of soil suppressiveness against Heterodera filipjevi in the studied fields. The results obtained from the second phase of the project showed that using a screening approach for nematophagous fungi differing radically from standard screening methods can yield a quite different microbial community. In the focused screening approach, introduced here, single nematode eggs showing symptoms of fungal infection were cultured separately to isolate the fungal species, which had already colonised each individual egg. It was assumed that culturing of a single surface-sterilised infected egg would theoretically yield only one fungal strain. In short, this technique creates a process during which the growth and purity of any fungus growing from each infected egg can easily be 110

General Discussion monitored regardless whether it is a slow or a fast growing strain. Fig 5.1 gives an overview of the main results obtained by scrutinising individual fungal infected eggs as the cornerstone for this research. In the dissertation, the three species Ijuhya vitellina, Monocillium gamsii, and Polyphilus sieberi nom. prov. were described. The two natural compounds chaetoglobosin A and its derivative 19-O- acetylchaetoglobosin A were identified, and their impact against H. filipjevi was studied. In addition to the results discussed here, three more pleosporalean fungal species were isolated, which are currently under taxonomic circumscription and metabolite profiling. The reproducibility of this method could be evaluated for other nematode species as well as in other geographical settings. Recently, by applying this method we could isolate another potentially new pleosporalean species (data in process) from the eggs of a population of the sugar beet cyst nematode Heterodera schachtii in Germany. This is an intriguing result, because H. schachtii as an economically important PPN in sugar beet growing systems has been monitored for fungi associated with its females and eggs since Kühn’s original finding.

Figure 5.1- A schematice outline of results obtained during the focused screening phase of the project.

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5.2 Phylogenetic relationships of the isolated fungi

The newly found egg-parasitising fungi discussed here broaden the spectrum of potential biocontrol agents attacking the eggs of cyst nematodes. Several examples of egg-parasitic fungi are found in major fungal taxonomic groups, but the highest diversity of these fungi is found in Ascomycota. The species obtained during this project accordingly all belong to different phylogenetic lineages within the Ascomycota: Ijuhya vitellina (Bionectriaceae) and Monocillium gamsii (Niessliaceae) are representatives of the Hypocreales, while Polyphilus sieberi nom. prov. (Hyaloscyphaceae) is a member of the Helotiales and additionally belongs to the root-inhabiting form group of the “dark septate endophytes” (DSEs). Three more hitherto undescribed pleosporalean species could be isolated including two other DSEs. These results are additional evidence suggesting that a broad phylogenetic range of fungal species can attack eggs of cyst-forming nematodes, but ascomycetes seem to be clearly dominating. Within the Bionectriaceae, there are a few examples of nematode-parasitising fungi (Ahmed et al. 2014; Chen and Chen 2002). However, within the genus of Ijuhya, I. vitellina is the only nematophagous species described so far. Given its phylogenetic position on a branch with relatively large genetic distance, its unique biology and the secondary compound production it could in future potentially be grouped in a genus of its own, because all these characters set it apart from the other representatives of the genus. Currently, we wanted to avoid the potential instabilities of a monotypic genus. In the Niessliaceae, M. gamsii and M. bulbillosum are the only species with a nematode egg-parasitic lifestyle, but otherwise are typical representatives of the family and genus. The primary studies on P. sieberi nom. prov. (Knapp et al. 2012) examined its capacity for endophytic colonization of its original host plants (Asclepias syriaca, Juniperus communis), and reported the fungus as a DSE. Endophytic fungi colonise the plant tissues and grow internally within the root, stem or leaves without causing any apparent symptoms of disease or harm to their host plants (Rodriguez et al. 2009; Schulz et al. 2015). A comprehensive review of fungal endophytes suggests four classes of endophytic fungi delineating the main criteria of each class (Rodriguez et al. 2009). This classification scheme categorises the fungal endophytes into clavicipitaceous and non-clavicipitaceous endophytes, and classifies them based on their phylogenetic relationship, lifestyle and ecological interaction. Accordingly, the class 1 merely contains clavicipitaceous endophytes representing “a small number of phylogenetically related clavicipitaceous species that are fastidious in culture and limited to some cool- and warm-season grasses” (Rodriguez et al. 2009). The main group of entomopathogenic and nematode-parasitic fungi with endophytic lifestyle in planta belong here (Rodriguez et al. 2009). The entomopathogenic fungus Metarhizium, as well as some egg-parasitic fungi such as Lecanicillium lecanii and Pochonia chlamydosporia are some examples of this group of endophytes (Behie et al.

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2012; Lopez-Llorca et al. 2006). The other three classes proposed are non-clavicipitaceous which are “highly diverse, representing a polyphyletic assemblage of primarily ascomycetous fungi with diverse and often poorly defined or unknown ecological roles” (Rodriguez et al. 2009). Arthrobotrys spp. are examples of nematophagous species with an endophytic lifestyle that are placed in the non- clavicipitaceous class 2 endophytes. So far, there is no nematode parasitic fungus reported from class 3 endophytes. In class 4 endophytes accommodating DSEs, P. sieberi nom. prov. is the first species reported to have a multifunctional lifestyle parasitising eggs of a plant parasitic nematode.

5.3 Biology- Parasitism

Cyst nematodes are sedentary parasites and establish a long-term biotrophic interaction with their host plants. During development, the nematode head region remains in the cortical plant cells to feed, while the rest of the female body ruptures the root. The gravid females later die and become cysts that can remain in soil for many years. This situation causes a long-term exposure of the cyst nematodes to the soil microbial community, which may provide opportunities for nematode-antagonistic fungi to parasitise females and nematode eggs. In general, the cyst structure protects the nematode eggs from harsh environmental conditions (Luc et al. 1986). Thus, for fungi that are capable of parasitising nematode cysts and their eggs, cysts may serve as “specific” niches where the fungi may have equivalent prolonged survival chances. In addition, inside the cysts, eggs are embedded in a mucilaginous matrix, which has been reported containing lipids, glycoconjugates and lectins (Nour et al. 2003). The latter might play a role in nematode recognition by nematophagous fungi (Li et al. 2015). As discussed in chapter (5), the mucilaginous matrix on plant roots was also reported to contain some carbohydrates that may protect them from desiccation by functioning as a water reservoir [(Barrow 2003) and references therein], especially in semiarid regions, where drought-stressed conditions are common and cyst nematodes and DSEs are abundant (Jumpponen and Trappe 1998; Mandyam and Jumpponen 2005; Rodriguez et al. 2009). Considering all these points, it may be hypothesised that colonisation of the cyst mucilage matrix by these fungi may: i) sustain the colonising fungi by providing a favourable micro-environment and nutrient source, and protecting the fungi from potential drought stress, while growing towards the eggs. Colonisation of the mucilage matrix of cysts was documented for P. sieberi nom. prov. showing the growth of ‘runner hyphae’ throughout the cyst mucilage; ii) accelerate fungal growth resulting in a faster colonisation of nematode eggs. This is also supported by our observations in the infection studies, during which eggs inside cysts were infected faster than individual eggs under the same incubation parameters. Although the cysts of nematodes are to some extent ‘easy targets’ to be colonised, the nematophagous fungi still need to overcome several physical and chemical obstacles before the can colonise nematode

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General Discussion eggs. There are three natural opening on a cyst i.e. the mouth opening, the secretory-excretory pore, and the vulval slit, which provide a natural pathway for fungi to enter the cyst cavity. Despite these natural pathways to the inside of the cyst, the nematophagous fungi may still need to penetrate the cyst wall to reach inside the cyst cavity. Inside the cyst, the fungus can presumably use the mucilage matrix as an intermediate nutrient source until encountering the eggs. The eggshell of cyst nematodes is a multilayered structure which mainly consists of chitin and lipids (Perry and Trett 1986). As discussed in chapter 2, to penetrate the eggs, the egg-parasitic fungi need to overcome this physical barrier. To do so, the nematophagous fungi may employ a combination of morphological adaptations, e.g. the formation of an appressorium and enzymatic activities (Li et al. 2015; Lopez-Llorca et al. 2008). In nematophagous and entomopathogenic fungi, physical penetration is commonly assisted with several enzymatic processes to lyse chitin, lipids and other structural components of the eggshell or host body cuticle (Dijksterhuis et al. 1991; Leger et al. 1991; Lopez-Llorca et al. 2002b) as main barriers to fend-off infection. The observations presented in this thesis showed that regardless of their different phylogenetic origin M. gamsii and P. sieberi nom. prov. penetrated the eggshells and cuticle of developing juveniles in a similar way using a ‘simple’ hyphal penetration, while in addition to direct penetration I. vitellina formed an appressorium-like structure to enter the nematode egg. Beside the difference in mechanical penetration strategies, it is presumed that these three fungi can possibly apply different enzymatic strategies, assumingly lipidases, chitinases, and proteases, which enable degradation of eggshells, and the body cuticle of developing juveniles. Further microscopic observations, especially electron microscopic studies are needed to help to document and understand the penetration process detailing the morphological adaptations to nematode infection. Detailed studies on potential enzymatic activities involved in infection may also shed some light on the mode of action of these newly found egg-parasitic fungi. Within the nematophagous fungi, host recognition and infection is a cell-cell communication process in which several genes are involved to facilitate the adherence to the eggshell or the host body cuticle (Li et al. 2015). Therefore, detailed molecular biological studies involving also genomic and transcriptomic approaches may also give insights into the infection strategies of the here- studied fungi. Inside the nematode eggs, hyphae of the hypocrealean fungi I. vitellina, M. gamsii and its sister species M. bulbillosum convert into chlamydospore-like structures and form microsclerotia, multicellular structures, which are assumed to function as survival structures. As discussed in chapter (2) microsclerotia formed inside the eggs contained oil-like droplets or guttules that became abundantly present during their formation and could potentially function as energy reservoirs. It can be hypothesized that formation of microsclerotia is a unique mode of survival in the nematophagous fungi studied here that helps the fungi survive in the dormant stage and tolerate abiotic stresses while

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General Discussion inhabiting the nematode eggs during adverse environmental conditions. On the basis of the results obtained, formation of microsclerotia was a common mode of reproduction in the naturally or artificially infected egg samples, as well as in cultural media where the pure fungi were maintained: Whenever a single microsclerotium isolated from any of the abovementioned sources was cultured on fungal growth media, it produced newly emerging hyphae later converting into new microsclerotia which were identical with the examined ones. This suggests that microsclerotia are physiologically functional as they are responsive to favourable environmental condition for generating new hyphae. Taken together the formation of these structures represents a unique stage of the life cycle and development of these newly found egg-parasitic fungi. Like the other three abovementioned fungi, P. sieberi nom. prov. also formed hyphae, which became enriched in oil-like droplets upon colonization of the eggs and developing juveniles of H. filipjevi. The function of these organelles is neither fully characterised nor understood yet. However, it was suggested that they might be involved in energy storage and nutrient acquisition (Barrow 2003), in degradation of nematode eggshells and cuticles, and in nematode killing (Anke et al. 1995; Dijksterhuis et al. 1994; Nordbring-Hertz et al. 2011) (See chapter 2). Recently, Behie and colleagues (Behie et al. 2017; 2012) reported reciprocal actions between plants and the entomopathogenic fungus Metarhizium through which the fungus translocates insect-derived nitrogen to the host plant and in exchange receives photosynthates. Some nematophagous fungi such as Arthrobotrys spp., Lecanicillium lecanii, Pochonia chlamydosporia, and Purpureocillium lilacinum were also reported to colonise the plant roots endophytically (Bordallo et al. 2002; Hirsch et al. 2000; Lopez-Llorca et al. 2002a; Schouten 2016). The results obtained in this project suggest that P. sieberi nom. prov. is a multitrophic species (endophyte of plants, parasite of nematodes and probably an accidental commensal of truffle; See chapter 5) which may be related to a multifunctional lifecycle (Barelli et al. 2016) i.e. a potential mutualistic-parasitic interaction with plants and nematodes. It is thought that by parasitising the nematode eggs inside a protected microenvironment – a cyst – these fungi could overcome the adverse conditions in the absent of a host plant. But if the intact juveniles hatch and leave the cyst to attack new plants, there would be no more nutrient sources available for the fungus and the fungus would need to use another host, e.g. a plant, to survive. If the host i.e. wheat or other cereals is available, the fungus could endophytically colonise the plant which hypothetically can result in a symbiotic interaction. This hypothesized symbiosis may have a role in plant protection and nutrition by killing the plant parasitic nematodes by either direct colonization or by producing specific secondary metabolites, which can be involved in nematode inhibition.

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5.4 Nematophagous fungi – a treasure trove for nematicidal compounds

The (elimination of) hazards caused by synthetic nematicides to the environment and to the health of humans are reasons to search for new alternatives for control of parasitic nematodes. This opened up a new research field to exploit the natural enemies of nematodes as biocontrol agents, by using either the whole organism or their natural products against pathogens. In addition, increasing resistances of various pathogens to some natural compounds with medicinal, veterinary and plant protection applications are newly emerging problems that intensify the search for new natural sources supplying ‘safe’ and new bioactive compounds (Berdy 2012). Continuous screening for new nematode-antagonistic microorganisms as a source for biologically active metabolites can potentially result in the discovery of novel natural products with nematicidal effects. This seems even more promising when it is considered that about 45% of the 60-80 thousand known microbial metabolites derived from different microorganisms are produced by fungi (Berdy 2012), and that only about 3-8% of 2.2 to 3.8 million fungal taxa estimated to exist have been identified so far (Hawksworth and Lucking 2017). The present study showed that application of improved screening methods with a specific focus on diseased nematodes is a promising approach to find new nematophagous fungi even from widely distributed and intensively studied plant parasitic nematodes, which in addition also produce interesting novel bioactive metabolites. As summarized in Fig 5.1, the ongoing metabolite profiling of the newly described, and so-far unknown fungi obtained in the project, has resulted not only in finding new resources for biologically important compounds (Ijuhya vitellina as a new source for chaetoglobosin A- see chapter 2), but also in the discovery of new natural products. Also, four new cyclodepsipeptide compounds and the known compound Arthrichitin (a cell wall active and antifungal compound) was isolated from a so-far undescribed Ophiosphaerella (Pleosporales-Ascomycota) species, among which two new compounds showed nematicidal activity against H. filipjevi (data in process). The ongoing screening of M. gamsii also provided promising crude extracts, which are currently processed to be purified, identified and bio-assayed for potential biological activities. It is noteworthy that some Monocillium species e.g. M. nordinii were extensively screened for secondary metabolites resulting in the isolation of bioactive natural products such as monorden and monocillins (Ayer et al. 1980). The possibility of discovery of new secondary metabolites from M. gamsii is encouraging to make more attempts for metabolite profiling of all other fungal species, which have been isolated during this project. Berdy (2012) suggests that microorganisms ‘living in unique surroundings or extreme environments’ are promising resources for the discovery of new secondary metabolites. As it was discussed in the previous chapters, cysts of nematodes are specific (micro-)environments where diverse microbial 116

General Discussion communities live in association with nematode eggs. The finding of six new fungal species from the cereal cyst nematode H. filipjevi in just two experimental wheat fields in Turkey adds a significant amount of diversity to the already known microbial communities of these microhabitats. These findings suggest that cyst nematodes are a promising resource for finding new bioactive natural products. There are many geographic regions that have not yet been extensively explored for nematode antagonistic microorganisms, neither have many host species been sampled accordingly. This can be because of several difficulties, such as financial limitations, lack of human resources and facilities, or difficult accessibility. The here obtained results suggest that exploring additional regions where cyst nematodes have been reported to exist, provides a unique opportunity to find cysts bearing potentially new microorganisms with nematode-antagonistic properties. Among other types of microorganisms producing bioactive compounds, endophytic microorganisms such as endophytic fungi are important sources for new natural compounds (Berdy 2012; Nicoletti and Fiorentino 2015; Schulz et al. 2002). Endophytic fungi have extensively been screened as promising sources for bioactive secondary metabolites (Nicoletti and Fiorentino 2015; Schulz et al. 2008), and a majority of them are capable of producing metabolites with biological activity (Berdy 2012). As mentioned earlier, a ‘multifunctional lifestyle’ was reported for the entomopathogenic fungi Metarhizium and Beauveria which parasitise their insect hosts and simultaneously interact endophytically with their host plants (Barelli et al. 2016). It can be speculated that like in this ‘endophytic insect-pathogenic’ lifestyle, an endophytic nematode-parasitic life style may also exist for the ‘endophytic nematophagous fungi’. And these potential endophytes may be involved in metabolic interactions with their host plants and host nematodes. Support for this hypothesis is the isolation of a profile of novel secondary metabolites (Fig 5.2) from an aforementioned dark septate endophytic pleosporalean species that was found parasitising eggs of H. filipjevi. The endophytic lifestyle and potentially mutualistic interaction of the fungi found in this project with wheat or other cereals hosts of H. filipjevi can be experimentally examined. Also, the secondary metabolites involved in this tripartite relationship can be further examined for their beneficial effects to ‘their producers’ as well as to their host plants.

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Figure 5.2- Secondary metabolites from an undescribed pleosporalean fungus.

If to quote Joan Bennett in Keller et al. (2011), “we may not understand their biological functions yet, but we can predict that such functions exist because secondary metabolites would not be biosynthesized unless they benefit their producers”. It could be hypothesized that if a multifunctional (endophytic-parasitic) life style exists for these fungi, their secondary metabolites could functionally be involved in a tripartite interaction (nematode-fungus-plant) via different ways: i) Directly, by biocidal activity against nematodes. This would result in improving plant health by reducing the biotic stress caused by nematode invasion. ii) Indirectly, to serve as “chemical interface” (Berdy 2012) between plants and nematodes. This may lead to improve the plant defence mechanisms against nematode parasitism through induced resistance, or by translocating nematode-derived nutrients to the host plant. In exchange the fungus may also acquire some plant-derived nutrients (Behie et al. 2017). Millions of species are estimated to exist (Hebert et al. 2016) while only a minority of them have yet been identified. This inspiring biodiversity in nature provides ample opportunities to discover and identify new microorganisms whose capabilities in control of diseases and pathogens can be exploited to improve the quality of life.

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5.5 References

Ahmed M, Laing MD, Nsahlai IV (2014) Use of Clonostachys rosea against sheep nematodes developing in pastures. Biocontrol Science and Technology 24: 389-398. Anke H, Stadler M, Mayer A, Sterner O (1995) Secondary Metabolites with Nematocidal and Antimicrobial Activity from Nematophagous Fungi and Ascomycetes. Canadian Journal of Botany 73: S932-S939. Ayer WA, Lee SP, Tsuneda A, Hiratsuka Y (1980) The isolation, identification, and bioassay of the antifungal metabolites produced by Monocillium nordinii. Canadian Journal of Microbiology 26: 766-773. Barelli L, Moonjely S, Behie SW, Bidochka MJ (2016) Fungi with multifunctional lifestyles: endophytic insect pathogenic fungi. Plant Molecular Biology 90: 657-664. Barrow JR (2003) Atypical morphology of dark septate fungal root endophytes of Bouteloua in arid southwestern USA rangelands. Mycorrhiza 13: 239-247. Behie SW, Moreira CC, Sementchoukova I, Barelli L, Zelisko PM, Bidochka MJ (2017) Carbon translocation from a plant to an insect-pathogenic endophytic fungus. Nature Communications 8: 14245. Behie SW, Zelisko PM, Bidochka MJ (2012) Endophytic Insect-Parasitic Fungi Translocate Nitrogen Directly from Insects to Plants. Science 336: 1576-1577. Berdy J (2012) Thoughts and facts about antibiotics: Where we are now and where we are heading. J Antibiot 65: 385-395. Bordallo JJ, Lopez-Llorca LV, Jansson HB, Salinas J, Persmark L, Asensio L (2002) Colonization of plant roots by egg-parasitic and nematode-trapping fungi. New Phytologist 154: 491-499. Chen F, Chen S (2002) Mycofloras in cysts, females, and eggs of the soybean cyst nematode in Minnesota. Applied Soil Ecology 19: 35-50. Dijksterhuis J, Harder W, Wyss U, Veenhuis M (1991) Colonization and digestion of nematodes by the endoparasitic nematophagous fungus Drechmeria coniospora. Mycological Research 95: 873- 878. Dijksterhuis J, Veenhuis M, Harder W, Nordbring-Hertz B (1994) Nematophagous fungi: physiological aspects and structure-function relationships. Adv Microb Physiol 36: 111-143. Hawksworth DL, Lucking R (2017) Fungal Diversity Revisited: 2.2 to 3.8 Million Species. Microbiol Spectr 5. Hebert PD, Hollingsworth PM, Hajibabaei M (2016) From writing to reading the encyclopedia of life. Philosophical Transactions of the Royal Society B: Biological Sciences 371. Hirsch PR, Mauchline TH, Mendum TA, Kerry BR (2000) Detection of the nematophagous fungus Verticillium chlamydosporium in nematode-infested plant roots using PCR. Mycological Research 104: 435-439. Jatala P (1986) Biological control of plant-parasitic nematodes. Annual Review of Phytopathology 24: 453-489. Jumpponen ARI, Trappe JM (1998) Dark septate endophytes: a review of facultative biotrophic root- colonizing fungi. New Phytologist 140: 295-310. Keller NP, Bennett J, Turner G (2011) Secondary metabolism: Then, now and tomorrow. Fungal Genetics and Biology 48: 1-3. Knapp DG, Pintye A, Kovacs GM (2012) The dark side is not fastidious - dark septate endophytic fungi of native and invasive plants of semiarid sandy areas. PLoS One 7: e32570. Kühn J (1877) Vorläufiger Bericht über die bisherigen Ergebnisse der seit dem Jahre 1875 im Auftrage des Vereins für Rübenzucker-Industrie ausgeführten Versuche zur Ermittlung der Ursache der Rübenmüdigkeit des Bodens und zur Erforschung der Natur der Nematoden. Zeitschrift des Vereins für die Rübenzucker-Industrie des Deutschen Reichs 27: 452-457. Kühn J (1881) Die Ergebnisse der Versuche zur Ermittelung der Ursache der Rübenmüdigkeit und zur Erforschung der Natur der Nematoden. Berichte aus dem physiologischen Laboratorium und der Versuchsanstalt des landsoirthschaftlichen Instituts der Uniuersitat Halle 3: 1-153. 119

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Leger RJS, Roberts DW, Staples RC (1991) A model to explain differentiation of appressoria by germlings of Metarhizium anisopliae. Journal of Invertebrate Pathology 57: 299-310. Li J, Zou C, Xu J, Ji X, Niu X, Yang J, Huang X, Zhang KQ (2015) Molecular mechanisms of nematode- nematophagous microbe interactions: basis for biological control of plant-parasitic nematodes. Annu Rev Phytopathol 53: 67-95. Lopez-Llorca LV, Bordallo JJ, Salinas J, Monfort E, Lopez-Serna ML (2002a) Use of light and scanning electron microscopy to examine colonisation of barley rhizosphere by the nematophagous fungus Verticillium chlamydosporium. Micron 33: 61-67. Lopez-Llorca LV, Jansson H-B, Vicente JGM, Salinas J (2006) Nematophagous Fungi as Root Endophytes. In: Schulz BJE, Boyle CJC, Sieber TN (Eds) Microbial Root Endophytes. Springer Berlin Heidelberg, Berlin, Heidelberg, 191-206. Lopez-Llorca LV, Maciá-Vicente JG, Jansson H-B (2008) Mode of action and interactions of nematophagous fungi. In: Ciancio A, Mukerji KG (Eds) Integrated Management and Biocontrol of Vegetable and Grain Crops Nematodes. Springer Netherlands, Dordrecht, 51-76. Lopez-Llorca LV, Olivares-Bernabeu C, Salinas J, Jansson H-B, Kolattukudy PE (2002b) Pre-penetration events in fungal parasitism of nematode eggs. Mycological Research 106: 499-506. Luc M, Weischer B, Stone AR, Baldwin JG (1986) On the definition of heteroderid cysts. Revue de Nématologie 9: 418-421. Mandyam K, Jumpponen A (2005) Seeking the elusive function of the root-colonising dark septate endophytic fungi. Studies in Mycology 53: 173-189. Nicoletti R, Fiorentino A (2015) Plant bioactive metabolites and drugs produced by endophytic fungi of Spermatophyta. Agriculture 5: 918. Nordbring-Hertz B, Jansson H-B, Tunlid A (2011) Nematophagous fungi. Encyclopedia of Life Sciences. John Wiley & Sons, Ltd. Nour SM, Lawrence JR, Zhu H, Swerhone GDW, Welsh M, Welacky TW, Topp E (2003) Bacteria associated with cysts of the soybean cyst nematode (Heterodera glycines). Applied and Environmental Microbiology 69: 607-615. Perry RN, Trett MW (1986) Ultrastructure of the eggshell of Heterodera schachtii and H. glycines (Nematoda:Tylenchida). Revue de Nematologie 9: 399-403. Rodriguez RJ, White JF, Arnold AE, Redman RS (2009) Fungal endophytes: diversity and functional roles. New Phytologist 182: 314-330. Schouten A (2016) Mechanisms Involved in Nematode Control by Endophytic Fungi. Annual Review of Phytopathology 54: 121-142. Schulz B, Boyle C, Draeger S, Rommert AK, Krohn K (2002) Endophytic fungi: a source of novel biologically active secondary metabolites. Mycological Research 106: 996-1004. Schulz B, Draeger S, dela Cruz Thomas E, Rheinheimer J, Siems K, Loesgen S, Bitzer J, Schloerke O, Zeeck A, Kock I, Hussain H, Dai J, Krohn K (2008) Screening strategies for obtaining novel, biologically active, fungal secondary metabolites from marine habitats. Botanica Marina, 219-234 pp. Schulz B, Haas S, Junker C, Andree N, Schobert M (2015) Fungal endophytes are involved in multiple balanced antagonisms. Current Science 109: 39-45. Siddiqui ZA, Mahmood I (1996) Biological control of plant parasitic nematodes by fungi: A review. Bioresource Technology 58: 229-239. Stirling GR (2014) Biological control of plant-parasitic nematodes: soil ecosystem management in sustainable agriculture. CABI, Wallingford, UK, xxiv + 510 pp. Tribe HT (1977) Pathology of cyst-nematodes. Biological Reviews of the Cambridge Philosophical Society 52: 477-&.

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Summary

Parasitism caused by cereal cyst nematodes (CCNs) is a major limiting biotic factor in cereal cropping systems. CCNs attack cereal crops and can lead to significant yield reductions. Field observations in experimental wheat fields of the International Maize and Wheat Improvement Centre (CIMMYT) in Turkey had revealed a sharp decline in nematode populations of the cereal cyst nematode Heterodera filipjevi. Microscopic studies of the cyst samples obtained from these fields regularly demonstrated the presence of fungi colonising nematode cysts and eggs. It was hypothesised that these or other fungi could be involved in the reduction of the population size of H. filipjevi. The present study therefore aimed at (i) isolating fungi from cysts of H. filipjevi; (ii) specifically isolating symptom-causing fungi from single eggs by establishing a specific isolation technique; (iii) classifying the fungal isolates of interest by morphological studies and multi-locus molecular phylogenetic analyses; (iv) studying the nematode- fungus interaction microscopically; (v) fulfilling Koch’s postulates for the fungi isolated from the nematodes; (vi) isolating and identifying secondary metabolites produced by the fungal isolates of interest. Two screening approaches were applied to isolate a maximum number of fungi associated with nematode eggs: In a general screening, field-collected cyst samples and their contents were cultured without prior selection. In a focused screening approach, only those nematode eggs exhibiting symptoms of fungal infection were individually cultured. This approach resulted in finding six new fungal species, three of which are described here: Ijuhya vitellina sp. nov., Monocillium gamsii sp. nov., and Polyphilus sieberi nom. prov. Ijuhya vitellina, M. gamsii belong to the families of Bionectriaceae and Niessliaceae, respectively and are the first reports of nematophagous fungi within the genera Ijuhya and Monocillium. The fungal genus Polyphilus nom. prov. was proposed to accommodate two species (P. sieberi nom. prov. and P. frankenii nom. prov.), of which only P. sieberi was isolated from nematode eggs. This is the first case where a dark septate endophyte (DSE) was shown to parasitise nematode eggs. In addition to the species described here, three hitherto undescribed pleosporalean species were isolated, two of which were also DSEs. All the new fungal species could be successfully re-isolated from the eggs of artificially infected nematode cysts in vitro, and Koch’s postulates were thus fulfilled. Additionally, their antagonistic interactions with nematode eggs were documented in detail using light microscopy: After penetrating the eggshell and the body cuticle of developing juveniles inside the eggs, hyphae of I. vitellina and M. gamsii developed and formed microsclerotia. These microsclerotia were rich in oil-like droplets and germinated on artificial media. They are interpreted as long-term survival structures. Parasitism of nematode eggs caused by P. sieberi as a DSE suggests a multifunctional lifestyle that could potentially benefit the plant host by killing nematodes as well as by transferring nutrients from colonised nematodes. 121

Summary

The isolated fungi were screened for secondary metabolites, and their chemical structures were elucidated. Chaetoglobosin A and its derivative 19-O-acetylchaetoglobosin A were isolated from I. vitellina, which is a new source for this compound with broad bioactivity that also showed inhibiting activity against the second-stage juvenile of H. filipjevi. These compounds were not present in the plant-associated species of the genus Ijuhya investigated here. From a so-far undescribed nematode- egg parasitic pleosporalean species several novel compounds were identified. Two of these are cyclodepsipeptides, a class of compounds known for their anthelmintic effects. Taken together, these results suggest that nematode-antagonistic fungi represent a promising source for finding new natural compounds with nematicidal activity. The discovery of several novel fungal species is intriguing given that cyst nematodes have been intensively screened for nematophagous fungi since the pioneering studies of Julius Kühn for over 150 years. The new findings became possible by applying a new isolation technique that can be used in future to discover even more fungi from other plant parasitic nematodes or in other geographic regions.

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Summary

Zusammenfassung

Die Getreidezystennematoden sind ein wichtiger limitierender biotischer Faktor im Getreideanbau. Sie parasitieren die Getreidepflanzen und verursachen dadurch signifikante Einbußen in der Getreideproduktion. Beobachtungen in Versuchsweizenfelder des „International Maize and Wheat Improvement Centre“ (CIMMYT) in der Türkei zeigten eine starke Reduktion in der Population der Getreidezystennematoden zwischen zwei aufeinanderfolgenden Probenahmen. Mikroskopische Untersuchungen der Nematodenzysten von diesen Versuchsfeldern ergaben, dass die Cysten und Eier häufig von Pilzen kolonisiert wurden. Diese Beobachtungen führten zu der Hypothese, dass Pilze für die beobachtete Reduktion der Population von Heterodera filipjevi verantwortlich sein könnten. Die vorliegende Studie verfolgte deshalb die folgenden Ziele: 1. Pilze sollten von den Zysten von H. filipjevi isoliert werden. 2. Mit Hilfe einer neu etablierten Isolationsmethode sollten spezifisch die symptomverursachenden Pilze von einzelnen Eiern isoliert werden. 3. Die isolierten Pilze sollten durch morphologische und molekularphylogenetische Analysen, die auf mehreren Genregionen beruhten, klassifiziert werden. (4) Die Nematoden-Pilz-Interaktion sollte mikroskopisch untersucht werden. (5) Die Kochschen Postulate sollten für die isolierten Pilze erfüllt werden. (6) Von den nachgewiesenermaßen parasitischen Pilzen produzierte Sekundärmetaboliten sollten isoliert und charakterisiert werden. In der Studie wurden zwei verschiedene Isolationsmethoden angewandt, um eine möglichst große Zahl von Pilzen, die mit Nematodeneiern assoziiert sind, zu isolieren. In der sogenannten „Allgemeinen Untersuchung“ wurden in den Versuchsfeldern gesammelte Cysten zufällig ausgewählt und kultiviert. In der sogenannten „Spezifischen Untersuchung“ wurden nur Nematodeneier, die Symptome einer pilzlichen Infektion zeigten, einzeln auf Nährmedium kultiviert. Mit dieser Methode wurden sechs neue Arten gefunden, von denen drei in der vorliegenden Arbeit beschrieben werden: Ijuhya vitellina sp. nov., Monocillium gamsii sp. nov. und Polyphilus sieberi nom. prov. Ijuhya vitellina und M. gamsii gehören zur Familie der Bionectriaceae bzw. der Niessliaceae. Beide Arten sind die ersten nematophagen Arten innerhalb ihrer Gattungen. Die Gattung Polyphilus nom. prov. wurde eingeführt mit den zwei Arten P. sieberi nom. prov. und P. frankenii nom. prov., von der allerdings nur P. sieberi von Nematodeneiern isoliert wurde. Dies ist gleichzeitig das erste Mal, dass ein Pilz der Formgruppe „Dunkle septierte Wurzelpilze (DSE)“ als Nematodeneierparasit nachgewiesen wurde. Zusäztlich zu den drei hier neu beschriebenen Arten, wurden drei weitere bislang noch unbekannte Arten isoliert, die zur Ordnung der Pleosporales gehören und von denen zwei ebenfalls zu den DSEs gehören. Die drei neuen von Nematodeneiern isolierten Pilzarten konnten in vitro wiederum gesunde Nematodeneier und –cysten infizieren und von diesen re-isoliert werden, wodurch die Kochschen Postulate erfüllt wurden. Ihre antagonistische Interaktion mit Nematodeneiern wurden mittels Lichtmikroskopie im

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Summary

Detail untersucht: Hyphen von I. vitellina und M. gamsii penetrieren zunächst die Eischale, dann die Kutikula des darin befindlichen Nematodenemybryos und bilden schließlich Mikrosklerotien. Diese Mikrosklerotien sind reich in ölreichen Vesikeln und keimen auf künstlichen Nährmedien und werden als „Langzeit-Überdauerungsstrukturen“ interpretiert. Das Parasitieren von Nematodeneiern durch den als DSE bestätigten P. sieberi zeigt, dass dieser einen komplexen Lebenszyklus aufweist: Möglicherweise nutzt er einer besiedelten Pflanze, indem Nematoden an der Wurzel parasitiert und Nährstöffe aus den Nematoden an die Pflanze transloziert werden. Die isolierten Pilze wurden auf Sekundärmetabolite untersucht und ihre chemische Struktur aufgeklärt. Chaetoglobosin A und sein Derivat 19-O-acetylchaetoglobosin A wurden aus I. vitellina isoliert, welche eine neue Quelle für diese Naturstoffe darstellt, und die in vitro hemmende Aktivität gegen Juvenile des zweiten Stadiums (J2-Juvenile) von H. filipjevi zeigten. Diese Biomoleküle kommen nur in I. vitellina vor aber nicht in den pflanzenasoziierten Arten der Gattung Ijuhya. Einige neue Naturstoffe wurden von einer bislang unbeschriebenen pleosporalen Art identifiziert, von denen zwei Cyclodepsipeptide sind, einer Gruppe von Naturstoffen die unter anderem für ihre anthelminthischen Eigenschaften bekannt sind. Die Isolierung von einigen neuen Pilzarten aus Weizenzystennematoden ist erstaunlich, insbesondere vor dem Hintergrund, dass Zystennematoden schon seit über 150 Jahren, seit den bahnbrechenden Untersuchungen von Julius Kühn, sehr intensiv auf potentiell nematodenparasitische Pilze untersucht wurden. Diese neuen Entdeckungen wurden durch die Entwicklung einer neuen Isolationsmethode ermöglicht, die in Zukunft genutzt werden kann, um weitere bislang unbekannte Pilze von anderen pflanzenparasitischen Nematoden oder anderen geographischen Regionen zu entdecken.

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Acknowledgements

Acknowledgements

Finally, it is time to acknowledge personages and institutions. Without their help this work could not have been successfully completed. The project established a strong collaborative link between the International Maize and Wheat Improvement Centre (CIMMYT-Turkey), the Julius Kühn-Institut (JKI) and the Helmholtz Centre for Infection Research (HZI). I would like to thank all these research institutions for facilitating this research. I gratefully acknowledge CIMMYT, the Deutsche Gesellschaft für Internationale Zusammenarbeit (GIZ), the Fiat Panis Foundation, Ulm, JKI, and the Gesellschaft der Freunde und Förderer des JKI (GFF) for providing me with doctoral scholarship and financial support. I would like to express my sincere respect and gratitude to my doctoral supervisor Prof. Dr. Maria Finckh for the invaluable guidance, persistent support, and for generously providing every possible help during this dissertation. I am grateful to her for accepting me as a PhD candidate, and giving me the freedom to choose and pursue my research interests. It was an honour to accomplish this project under her supervision. I am sincerely grateful and would like to express my deepest gratitude to Dr. Wolfgang Maier, the direct supervisor of my project, for the supervision of the this project, the invaluable advice, the freedom and long-time support he generously gave me to follow up my research idea. I thank him for the tremendous help and immense amount of time he spent for sustained discussions we had during this time. His daily advice is a reminder of how his great scientific knowledge could lead me to achieve the aims of the project. I would like to express my special thanks to Prof. Dr. Marc Stadler, the head of the department of Microbial drugs at Helmholtz Centre for Infection Research (HZI), whose great knowledge and long experience on fungi and their secondary metabolites inspired me to start learning about fungal secondary metabolites. Discussions with him on the project were indispensable. My special thanks go to the doctoral committee as well as to the members of the examination commission for their time to read this dissertation and evaluate this project. I am sincerely grateful to Dr. Abdelfattah A. Dababat at CIMMYT-Turkey who generously supported me during the entire project. His initial field observations were the starting point of this collaborative research project. I would like to express my sincere gratitude to Prof. Dr. Thomas Kühne the former director of the Institute for Epidemiology and Pathogen Diagnostics of JKI for the continuous support, and encouragement to accomplish this work.

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Acknowledgements

My sincere thanks also go to Dr. Björn Niere who provided an opportunity to join his working group for some time. Without his support, it would have been difficult to continue this project. I extend my gratitude to Dr. Hans-Josef Schroers for sharing his vast mycological knowledge, and his insightful comments and recommendations on this project. I am deeply grateful to Dr. Zahra Tanha Maafi, the head of the nematology department at the Iranian Research Institute for Plant Protection (IRIPP), and Prof. Dr. Gerrit Karssen, the head of the nematology department at the National Plant Protection Organisation of the Netherlands (NPPO), for all their support during this time. Their scientific life has always inspired me to learn. My gratitude goes to my friends and colleagues who directly or indirectly helped me to complete my work: Dr. Gábor M. Kovács, Dr. Dániel G. Knapp, Dr. Andrea Fadani, Julia Kreiss, Dr. Adnan Šišić, James Mania Mwangi, Dr. Christian Richter, Dr. Soleiman Helaly, Somayyeh Sedaghatjoo, Katrin Balke, Mascha Hoffmeister, Ahmed Gomaa, Olivera Topalović, and Dr. Gül Erginbas-Orakci. Special thanks to Anke Brisske Rode for her skilful technical assistance. I am deeply grateful to my parents for all their support. I would like to thank my siblings Soheila Ashrafi and Somayeh Ashrafi for supporting me throughout my work, also for supporting and taking care of my parents in my absence. I am genuinely grateful to Azam Abdollahi and Farideh Madani, and Javad Farhoomand Tehrani for their generosity, unstinting support, and kind care. Special thanks to my dear friends Nasir Sahabi and Roshanak Bagheri for their help throughout the completion of my work. I would like to acknowledge with gratitude the continuous support of Farnoosh Farhoomand Tehrani, my wife, who patiently tolerated all difficulties we faced together during this time. She is an inextricable part of the story.

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Affidativ

Eidesstattliche Erklärung

„Hiermit versichere ich, dass ich die vorliegende Dissertation selbständig, ohne unerlaubte Hilfe Dritter angefertigt und andere als die in der Dissertation angegebenen Hilfsmittel nicht benutzt habe. Alle Stellen, die wörtlich oder sinngemäß aus veröffentlichten oder unveröffentlichten Schriften entnommen sind, habe ich als solche kenntlich gemacht. Dritte waren an der inhaltlichen Erstellung der Dissertation nicht beteiligt; insbesondere habe ich nicht die Hilfe eines kommerziellen Promotionsberaters in Anspruch genommen. Kein Teil dieser Arbeit ist in einem anderen Promotions- oder Habilitationsverfahren durch mich verwendet worden.“

Affidavit

“I herewith give assurance that I completed this dissertation independently without prohibited assistance of third parties or aids other than those identified in this dissertation. All pages that are from published or unpublished writings, either words-for-words or in paraphrase have been clearly identified as such. Third parties were not involved in the drafting of the materials contents of this dissertation; most specifically I did no employ the assistance of a dissertation advisor. No part of this thesis has been used in another doctoral or tenure process.”

Braunschweig, 2 February 2018

Samad Ashrafi

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