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Research Article
Monocillium gamsii sp. nov. and Monocillium bulbillosum: two nematode-associated fungi parasitising the eggs of Heterodera filipjevi
expand article infoSamad Ashrafi§, Marc Stadler|, Abdelfattah A. Dababat, Katja R. Richert-Pöggeler, Maria R. Finckh§, Wolfgang Maier
‡ Julius Kühn-Institut, Braunschweig, Germany
§ University of Kassel, Witzenhausen, Germany
| Helmholtz Centre for Infection Research GmbH (HZI), Braunschweig, Germany
¶ International Maize and Wheat Improvement Centre, Ankara, Turkey
Open Access

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

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.

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 Niesslia 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.

Materials and methods

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°10'E; altitude 985 m.a.s.l) and Haymana (39°26'N, 39°29'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 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).

Cultural studies

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).

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.

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)).

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.

Molecular phylogenetic studies

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 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: MF681481MF681514. 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).

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 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 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).

Isolates and accession numbers used in the phylogenetic analyses.

Species Isolate number Host / substrate Locality GenBank accession numbers Reference
ITS LSU rpb1 tef
Bionectria byssicola CBS 914.97–GML2665 Alchornea branches- leaf litter Uganda, Brazil AF358252 GQ506011 GQ506040 KX184977 (Hirooka et al. 2010; Moreira et al. 2016; Schroers 2001)
Hyaloseta nolinae CBS109837 Nolina micrantha, leaf litter USA, New Mexico KM231846 KM231726 KM232279 (Lombard et al. 2015)
Ijuhya vitellina DSM104494 Heterodera filipjevi, egg Turkey KY607535 KY607549 KY607576 (Ashrafi et al. 2017)
Monocillium bulbillosum CBS344.70 mouldy wallpaper Germany MF681488 MF681501 MF681513 MF681507 This study
Monocillium gamsii DSM105458 Heterodera filipjevi, egg Turkey MF681485 MF681496 MF681512 MF681506 This study
Monocillium gamsii DSM105459 Heterodera filipjevi, egg Turkey MF681483 MF681493 MF681511 MF681505 This study
Monocillium gamsii DSM105460 Heterodera filipjevi, egg Turkey MF681482 MF681492 MF681510 MF681504 This study
Monocillium gamsii DSM105461 Heterodera filipjevi, egg Turkey MF681481 MF681490 MF681509 MF681503 This study
Monocillium ligusticum CBS684.95 ectomycorrhizae of Pinus halapensis Italy MF681489 MF681502 MF681514 MF681508 This study
Nisslia exilis CBS357.70 Picea abies, bark Germany AY489718 AY489645 AY489613 (Castlebury et al. 2004)
Nisslia exilis CBS560.74 Pinus sylvestris, decayed needle England AY489720 AY489647 AY489614 (Castlebury et al. 2004)

Results

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. 1A). By dissecting the infected cysts, microsclerotia-like black bodies were found to be colonising the individual nematode eggs (Fig. 1B, C). In some infected eggs the developing juveniles were found to be entirely destroyed exhibiting an olivaceous brownish appearance (Fig. 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. 1F), and formed colonies at first white creamy, later becoming blackish dotted centrally with a general dark appearance due to the dense pigmentation (Fig. 1G).

Figure 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: 800 µm (A); 30 µm (B–E); 50 µm (F); 2 cm (H); 5 mm (I).

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. 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. 2).

Figure 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.

Taxonomy

Monocillium gamsii Ashrafi & W. Maier, sp. nov.

MycoBank No: 823248
Figs 1H, I, 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.

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.

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. 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. 4C). Similar to some naturally infected eggs, sclerotioid masses were also found in some artificially infected samples colonising almost the entire egg (Fig. 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. 4E, F). Following penetration, filamentous hyphae entirely colonised the unembryonated eggs (Fig. 4G, H) or the body cavity of the developing juveniles (Fig. 4I, J), enlarged (Fig. 4H–J), occasionally inflated, forming thick-walled, finely pigmented, and guttules-filled cells (Fig. 3J), which eventually coalesced to form discrete microsclerotia with a textura angularis appearance (Fig. 4K–M). Infection studies revealed that such microsclerotia could be formed 7–10 d after the incubation of nematode eggs with the fungus.

Figure 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 hyphae 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: 30 µm (A, H, I, K, M, O, R); 20 µm (B–G, J, L, N); 200 µm (P), 50 µm (Q).

Figure 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: 600 µm (A); 300 µm (B); 30 µm (C–O); 50 µm (P).

Pigmentation of microsclerotia occurred during fungal development from hyaline to olivaceous brown and later strongly brownish melanised cells (Fig. 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. 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.

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 (Fig. 5A–F).

Figure 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: 300 µm (A); 30 µm (B–F).

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. 2). According to phylogenetic inference, M. bulbillosum is very closely related to but separable from M. gamsii. Both 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. 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 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 profilic 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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