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.
Citation: Ashrafi S, Helaly S, Schroers H-J, Stadler M, Richert-Poeggeler KR, Dababat AA, et al. (2017) Ijuhya vitellina sp. nov., a novel source for chaetoglobosin A, is a destructive parasite of the cereal cyst nematode Heterodera filipjevi. PLoS ONE 12(7): e0180032. https://doi.org/10.1371/journal.pone.0180032
Editor: Sabrina Sarrocco, Universita degli Studi di Pisa, ITALY
Received: April 14, 2017; Accepted: June 7, 2017; Published: July 12, 2017
Copyright: © 2017 Ashrafi et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: Data are available on Genbank with the accession numbers KY607531–KY607585 and KY684180–KY684193.
Funding: Funded by Deutsche Gesellschaft für Internationale Zusammenarbeit (GIZ) (https://www.giz.de/en/html/about_giz.html), Grant ID: W0267 GIZ/BMZ-Endophyte as Biocontrol International Maize and Wheat Improvement Centre (CIMMYT) (http://www.cimmyt.org) EU FP7 Project CropSustaIn, grant agreement FP7-REGPOT-CT2012-316205 Gemeinschaft der Förderer und Freunde des Julius Kühn-Instituts (GFF) for publication fee Fiat Panis Foundation, Hohenheim (http://www.stiftung-fiat-panis.de/de/) Alexander von Humboldt Foundation (https://www.humboldt-foundation.de/web/home.html) ARRS Slovenian science foundation, project J4-5527. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
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 . A diverse group of fungi has since then been described as associates of cyst nematodes . Examples include Ilyonectria destructans (Zinssm.) Rossman, L. Lombard & Crous , Pochonia chlamydosporia (Goddard) Zare & W. Gams (both Ascomycota, Hypocreales) and Nematophthora gynophila Kerry & D.H. Crump (Stramenopiles) [4, 5] that were described as parasites of Heterodera avenae Wollenweber. Kerry  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 [7–10]. 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 [11–13] including nematicidal compounds that display various anthelmintic effects . Chaetoglobosins of the cytochalasan family can have cytotoxic and inhibitory activities [15, 16] and affect insects and nematodes [17–21]. 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 [12, 13, 22–26].
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.
Material and methods
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  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 required for taxonomic and phylogenetic inferences were obtained from the Westerdijk Fungal Biodiversity Institute (Utrecht, The Netherlands).
Both the sampling sites and the nematode species were not protected and thus no permits were required.
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 method. By doing so, additional evidence was gathered to prove that the fungi isolated by culturing techniques indeed colonised the nematode eggs.
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; ), malt extract agar (MEA; Carl Roth), yeast malt agar (YMA; ), 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 .
Pathogenicity test and Koch’s postulates
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 . 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 incubated in moist glass chambers at room temperature. Developing structures were monitored and microscopically photographed regularly.
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.
Fungal mycelium was obtained from PDA and transferred to 1.5 mL microtubes. Genomic DNA was extracted with a CTAB-based method . 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 nitrogen 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.
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  and LR5  and sequenced with primers ITS1F , ITS4 , LR0R , 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  were used for amplification and sequencing of rpb1; Tact1f and Tact2r  were used for act, and T1 and T22 for ß-tub. For sequencing ß-tub, primer T222 and T224  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), 5 μl MgCl2 (2.5 mM), 5 μl dNTPs (0.2 mM of each), 2 μl of each primer (0.4 pM μl-1), 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.
Alignment and phylogenetic reconstruction
Newly generated and already published sequences were used in phylogenetic analyses (Table 1). The latter were selected according to BLASTn searches (http://blast.ncbi.nlm.nih.gov/Blast.cgi)  that used the former as queries. Representatives of Bionectriaceae and Nectriaceae were selected mainly following Hirooka et al.  and Jaklitsch and Voglmayr . 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 or their combination with rpb1 (S1 Fig) or rpb1 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  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 . 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 [46, 47]. 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 . 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 [49, 50] 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  was done in PAUP 4.0b10 in the batch file mode  applying the Kimura two-parameter model of DNA substitution  with a transition/transversion ratio of 2.0 to compute genetic distances. Support for internal nodes was estimated by 1000 bootstrap replicates  (reported as NJB values). The phylogenetic trees were visualised using FigTree v. 1.4.2 (http://tree.bio.ed.ac.uk/software/figtree).
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.
Fermentation and extraction of cultures.
Three different liquid media (Q6/2, YM and ZM; ) 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. . For large-scale fermentation, the above mentioned strain was inoculated into 3 L of the selected medium (see below) and processed for extraction similarly.
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. .
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.
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. , 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.
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. . 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. . 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. .
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 1A). Reddish dots inside cysts consisted of nematode eggs each containing one or few microsclerotia (Fig 1B and 1C). In some infected eggs, microsclerotial tissues were found developing inside juveniles (Fig 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 1F). Predominantly globose or ellipsoidal, reddish microsclerotia were formed on the surface of (Fig 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 (Fig 1F, 1H and 1I). The sterility check revealed no fungal growth from the examined cysts.
(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.
Molecular phylogenetic studies
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 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.
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 and S1 and S2 Figs) including I. peristomialis, a later synonym of I. vitrea, which is the type species of Ijuhya . 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 and S1 and S2 Figs).
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).
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 3). The other subclade includes I. vitellina and its closest sister species, I. corynospora (Fig 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.
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 1.
Etymology: From Latin vitellus meaning egg yolk, referring to the colour and shape of microsclerotia formed by the species in nematode eggs.
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 (Fig 4A–4D and Fig 5A–5C), cells angular, pigmented, first pale luteous, then brick reddish (Fig 4K–4N), filled with small guttules. Dictyochlamydospore-like structures may enlarge through coiling or expand to form globose to ellipsoidal microsclerotia (Fig 4E and 4F and Fig 5D–5F) formed solitary, in chains (Fig 4J and Fig 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 4O). Teleomorph not observed.
(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.
(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.
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 6A). Hyphal cells inside eggs enlarged and destroyed unembryonated eggs or developing juveniles (Fig 6B). Eggs were entirely occupied by compartmentalised thick-walled hyphae (Fig 6C), and guttules-filled moniliform chlamydospores (Fig 6D) that eventually developed into microsclerotia (Fig 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.
(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.
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 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 (Fig 6I–6L). Microsclerotia developing inside artificially infected eggs (Fig 6M and 6N) appearing textura angularis (cf. Fig 5H for details) were indistinguishable from those encountered in field-collected cysts (cf. Fig 1B and 1C).
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.
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 [16, 83] while compound 2 is its 19-O-acetylchaetoglobosin A  (Fig 7). The detailed description of the structure elucidation is included in the supporting information (S1 Text and S2 Text and S6 Fig).
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 [85, 86] were detected.
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.
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 ascospores [82, 87, 88], 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). 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  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  and I. oenanthicola  conform well to the concept of the Nectria sylvana group , for which Lasionectria became the generic depository . Specifically, fasciculate hyphal clusters formed by perithecia of I. antillana (Fig 1A, 1B and 1D in Lechat and Courtecuisse ) and I. oenanthicola (Plate 1, Fig. A–C in Lechat and Hairaud ) 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).
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 [2, 92–94]. Within the Bionectriaceae, to our knowledge only two species, Clonostachys rosea and Gliomastix murorum, were described as antagonists of animal- and plant parasitic nematodes [7, 95, 96].
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 . Penetration may apply through mechanic or chemical mechanisms [97–100]. 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 [2, 101–104]. 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 . 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 . 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 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  and could also be involved in the parasitism of nematodes . It was reported that guttules may contain linoleic acid as a compound responsible for nematode killing . Dijksterhuis et al.  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.
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 [15, 16, 109, 110]. Chaetoglobosin C is also produced by Penicillium aurantiovirens , and chaetoglobosin K was first extracted from Diplodia macrospora . 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 [17, 112]. 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 . Thus, chaetoglobosins produced by I. vitellina may have a function in the described parasitism of nematode eggs.
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 . Structures illustrated for Ijuhya antillana  and Ijuhya oenanthicola  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.
S1 Fig. 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.
S2 Fig. 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.
S3 Fig. 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 Fig. Mass spectrum of chaetoglobosin A (1).
S5 Fig. Mass spectrum of 19-O-acetylchaetoglobosin A (2).
S6 Fig. 2D NMR assignment of chaetoglobosin A (1) and 19-O-acetylchaetoglobosin A (2).
HMBC (arrows) and COSY (bold bonds) correlations.
S1 Text. Structure determination of chaetoglobosins.
S2 Text. Spectroscopic data for chaetoglobosin A (1) and 19-O-acetylchaetoglobosin A (2).
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.
- Conceptualization: SA AAD WM.
- Data curation: SA SH.
- Formal analysis: SA SH HJS MS WM.
- Funding acquisition: AAD WM.
- Investigation: SA HJS SH WM.
- Methodology: SA SH HJS MS WM.
- Project administration: AAD WM.
- Resources: AAD MS WM.
- Supervision: HJS MS WM.
- Visualization: SA KRP.
- Writing – original draft: SA HJS SH WM.
- Writing – review & editing: HJS SA MS WM.
- 1. Kühn J. 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. Z Ver Rübenzucker-Ind. 1877;27:452–7.
- 2. Stirling GR. Biological control of plant-parasitic nematodes: soil ecosystem management in sustainable agriculture. Second Edition ed. Stirling GR, editor. Wallingford, UK: CABI; 2014. xxiv + 510 p.
- 3. Goffart H. 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. 1932;20:1–28.
- 4. Kerry BR. A fungus associated with young females of the cereal cyst-nematode, Heterodera avenae. Nematologica. 1974;20(2):259–61.
- 5. Kerry BR, Crump DH. Two fungi parasitic on females of cystnematodes (Heterodera spp.). Transactions of the British Mycological Society. 1980;74(1):119–25.
- 6. Kerry BR. Fungi and the decrease of cereal cyst-nematode populations in cereal monoculture. EPPO Bulletin. 1975;5(4):353–61.
- 7. Kerry BR, Crump DH, Mullen LA. Natural control of the cereal cyst nematode, Heterodera avenae Woll., by soil fungi at three sites. Crop Protection. 1982;1(1):99–109.
- 8. Dackman C, Nordbring-Hertz B. Fungal parasites of the cereal cyst nematode Heterodera avenae in southern Sweden. Journal of Nematology. 1985;17(1):50–5. PMC2618414. pmid:19294057
- 9. Rodríguez-Kábana R, Morgan-Jones G. Potential for nematode control by mycofloras endemic in the Tropics. Journal of Nematology. 1988;20(2):191–203. PMC2618814. pmid:19290202
- 10. Khan A, Williams KL, Nevalainen HKM. Control of plant-parasitic nematodes by Paecilomyces lilacinus and Monacrosporium lysipagum in pot trials. BioControl. 2006;51(5):643–58.
- 11. Stadler M, Mayer A, Anke H, Sterner O. Fatty acids and other compounds with nematicidal activity from cultures of Basidiomycetes. Planta Med. 1994;60(2):128–32. pmid:8202563.
- 12. Li G, Zhang K, Xu J, Dong J, Liu Y. Nematicidal substances from fungi. Recent Pat Biotechnol. 2007;1(3):212–33. pmid:19075843.
- 13. Anke H, Stadler M, Mayer A, Sterner O. Secondary Metabolites with Nematocidal and Antimicrobial Activity from Nematophagous Fungi and Ascomycetes. Can J Bot. 1995;73:S932–S9.
- 14. Anke H. Insecticidal and nematicidal metabolites from fungi. In: Hofrichter M, editor. Industrial Applications. Berlin, Heidelberg: Springer Berlin Heidelberg; 2011. p. 151–63.
- 15. Sekita S, Yoshihira K, Natori S, Kuwano H. Structures of chaetoglobosins C, D, E, and F, cytotoxic indol-3-yl-cytochalasans from Chaetomium globosum. Tetrahedron Lett. 1976;17(17):1351–4. http://dx.doi.org/10.1016/S0040-4039(00)78062-2.
- 16. Sekita S, Yoshihira K, Natori S, Kuwano H. Structures of chaetoglobosin A and B, cytotoxic metabolites of Chaetomium globosum. Tetrahedron Lett. 1973;14(23):2109–12. http://dx.doi.org/10.1016/S0040-4039(01)86820-9.
- 17. Maruyama K, Oosawa M, Tashiro A, Suzuki T, Tanikawa M, Kikuchi M, et al. Effects of chaetoglobosin J on the G-F transformation of actin. Biochimica et biophysica acta. 1986;874(2):137–43. http://dx.doi.org/10.1016/0167-4838(86)90110-X. pmid:3778915.
- 18. Cutler HG, Crumley FG, Cox RH, Cole RJ, Dorner JW, Springer JP, et al. Chaetoglobosin K: a new plant growth inhibitor and toxin from Diplodia macrospora. J Agric Food Chem. 1980;28(1):139–42. pmid:7358926.
- 19. Meyer SLF, Huettel RN, Liu XZ, Humber RA, Juba J, Nitao JK. Activity of fungal culture filtrates against soybean cyst nematode and root-knot nematode egg hatch and juvenile motility. Nematology. 2004;6(1):23–32.
- 20. Xue M, Zhang Q, Gao JM, Li H, Tian JM, Pescitelli G. Chaetoglobosin Vb from endophytic Chaetomium globosum: absolute configuration of chaetoglobosins. Chirality. 2012;24(8):668–74. pmid:22593034.
- 21. Zheng QC, Kong MZ, Zhao Q, Chen GD, Tian HY, Li XX, et al. Chaetoglobosin Y, a new cytochalasan from Chaetomium globosum. Fitoterapia. 2014;93:126–31. pmid:24418656.
- 22. Stadler M, Anke H, Arendholz W-R, Hansske F, Anders UWE, Sterner O, et al. Lachnumon and lachnumol A, new metabolites with nematicidal and antimicrobial activities from the ascomycete Lachnum papyraceum (Karst.) Karst. I. Producing organism, fermentation, isolation and biological activities. The Journal of Antibiotics. 1993;46(6):961–7. pmid:8344878
- 23. Niu XM, Wang YL, Chu YS, Xue HX, Li N, Wei LX, et al. Nematodetoxic aurovertin-type metabolites from a root-knot nematode parasitic fungus Pochonia chlamydosporia. J Agric Food Chem. 2010;58(2):828–34. Epub 2009/12/17. pmid:20000774.
- 24. Ghisalberti EL. Secondary metabolites with antinematodal activity. In: Atta ur R, editor. Studies in Natural Products Chemistry. Volume 26, Part G: Elsevier; 2002. p. 425–506.
- 25. Kundu A, Saha S, Walia S, Dutta TK. Anti-nemic secondary metabolites produced by Fusarium oxysporum f. sp. ciceris. Journal of Asia-Pacific Entomology. 2016;19(3):631–6. http://dx.doi.org/10.1016/j.aspen.2016.06.003.
- 26. Hu Y, Zhang W, Zhang P, Ruan W, Zhu X. Nematicidal activity of chaetoglobosin A Poduced by Chaetomium globosum NK102 against Meloidogyne incognita. J Agr Food Chem. 2013;61(1):41–6. pmid:23214998
- 27. Coyne DL, Nicol JM, Claudius-Cole B. Practical plant nematology: a field and laboratory guide: SP-IPM Secretariat, International Institute of Tropical Agriculture (IITA), Cotonou, Benin; 2007. 82 p.
- 28. Nirenberg H. Untersuchungen über die morphologische und biologische Differenzierung in der Fusarium-Section Liseola. Mitteilungen der Biologischen Bundesanstalt für Land- und Forstwirtschaft. 1976;169:1–117.
- 29. Stadler M, Wollweber H, Muhlbauer A, Henkel T, Asakawa Y, Hashimoto T, et al. Secondary metabolite profiles, genetic fingerprints and taxonomy of Daldinia and allies. Mycotaxon. 2001;77:379–429.
- 30. Kornerup A, Wanscher JH. Methuen Handbook of colour. London: Methuen and Co Ltd; 1967.
- 31. Gams W, Hoekstra ES, Aptroot A. CBS course of mycology. Fourth edition ed: Baarn Centraalbureau voor Schimmelcultures; 1998.
- 32. Saghai-Maroof MA, Soliman KM, Jorgensen RA, Allard RW. Ribosomal DNA spacer-length polymorphisms in barley: mendelian inheritance, chromosomal location, and population dynamics. Proceedings of the National Academy of Sciences of the United States of America. 1984;81(24):8014–8. PMC392284. pmid:6096873
- 33. de Hoog GS, Gerrits van den Ende AH. Molecular diagnostics of clinical strains of filamentous Basidiomycetes. Mycoses. 1998;41(5–6):183–9. pmid:9715630.
- 34. Vilgalys R, Hester M. Rapid genetic identification and mapping of enzymatically amplified ribosomal DNA from several Cryptococcus species. J Bacteriol. 1990;172(8):4238–46. PMC213247. pmid:2376561
- 35. Gardes M, Bruns TD. ITS primers with enhanced specificity for basidiomycetes—application to the identification of mycorrhizae and rusts. Mol Ecol. 1993;2(2):113–8. pmid:8180733.
- 36. White TJ, Bruns T, Lee S, Taylor J. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: Innis M, Gelfand D, Sninsky J, White T, editors. PCR Protocols: a guide to methods and applications. San Diego: Academic Press; 1990. p. 315–22.
- 37. Rehner SA, Samuels GJ. Taxonomy and Phylogeny of Gliocladium Analyzed from Nuclear Large Subunit Ribosomal DNA-Sequences. Mycol Res. 1994;98(6):625–34.
- 38. Castlebury LA, Rossman AY, Sung GH, Hyten AS, Spatafora JW. Multigene phylogeny reveals new lineage for Stachybotrys chartarum, the indoor air fungus. Mycological research. 2004;108(Pt 8):864–72. http://dx.doi.org/10.1017/S0953756204000607. pmid:15449591.
- 39. Samuels GJ, Dodd SL, Lu B-S, Petrini O, Schroers H-J, Druzhinina IS. The Trichoderma koningii aggregate species. Studies in mycology. 2006;56:67–133. http://dx.doi.org/10.3114/sim.2006.56.03. pmid:18490990
- 40. O’Donnell K, Cigelnik E. Two divergent intragenomic rDNA ITS2 types within a monophyletic lineage of the fungus Fusarium are nonorthologous. Molecular Phylogenetics and Evolution. 1997;7(1):103–16. pmid:9007025
- 41. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ. Basic Local Alignment Search Tool. J Mol Biol. 1990;215(3):403–10. pmid:2231712
- 42. Hirooka Y, Kobayashi T, Ono T, Rossman AY, Chaverri P. Verrucostoma, a new genus in the bionectriaceae from the Bonin Islands, Japan. Mycologia. 2010;102(2):418–29. pmid:20361508.
- 43. Jaklitsch WM, Voglmayr H. Stromatonectria gen. nov. and notes on Myrmaeciella. Mycologia. 2011;103(2):431–40. pmid:21139029; PubMed Central PMCID: PMC3076889.
- 44. Katoh K, Standley DM. MAFFT Multiple Sequence Alignment Software Version 7: Improvements in Performance and Usability. Mol Biol Evol. 2013;30(4):772–80. pmid:23329690
- 45. Rambaut A. Se-Al: Sequence Alignment Editor, 2.0a11. Available: http://evolve.zoo.ox.ac.uk and http://tree.bio.ed.ac.uk/software/seal/. 1996.
- 46. Ronquist F, Huelsenbeck JP. MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics. 2003;19(12):1572–4. Epub 2003/08/13. pmid:12912839
- 47. Huelsenbeck JP, Ronquist F. MRBAYES: Bayesian inference of phylogenetic trees. Bioinformatics. 2001;17(8):754–5. pmid:11524383
- 48. Nylander J. MrModeltest v2. Program distributed by the author. Evolutionary Biology Centre, Uppsala University. 2004.
- 49. Silvestro D, Michalak I. raxmlGUI: a graphical front-end for RAxML. Organisms Diversity & Evolution. 2012;12(4):335–7.
- 50. Stamatakis A. RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics. 2014;30(9):1312–3. PMC3998144. pmid:24451623
- 51. Saitou N, Nei M. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol. 1987;4(4):406–25. pmid:3447015
- 52. Swofford DL. PAUP*. Phylogenetic Analysis Using Parsimony (*and Other Methods). Version 4. Sinauer Associates, Sunderland, Massachusetts. 2002.
- 53. Kimura M. A simple method for estimating evolutionary rates of base substitutions through comparative studies of nucleotide sequences. J Mol Evol. 1980;16(2):111–20. Epub 1980/12/01. pmid:7463489.
- 54. Felsenstein J. Confidence limits on phylogenies: an approach using the Bootstrap. Evolution. 1985;39(4):783–91. pmid:28561359
- 55. Spatafora JW, Sung GH, Sung JM, Hywel-Jones NL, White JF. Phylogenetic evidence for an animal pathogen origin of ergot and the grass endophytes. Molecular Ecology. 2007;16(8):1701–11. pmid:17402984
- 56. Rossman AY, McKemy JM, Pardo-Schultheiss RA, Schroers H- J. Molecular studies of the Bionectriaceae using large subunit rDNA sequences. Mycologia. 2001;93(1):100–10.
- 57. Schroers HJ. A monograph of Bionectria (Ascomycota, Hypocreales, Bionectriaceae) and its Clonostachys anamorphs. Studies in mycology. 2001;(46):1–214.
- 58. Luo J, Zhuang WY. Bionectria vesiculosa sp. nov. from Yunnan, China. Mycotaxon. 2010;113:243–9.
- 59. Stenroos S, Laukka T, Huhtinen S, Döbbeler P, Myllys L, Syrjänen K, et al. Multiple origins of symbioses between ascomycetes and bryophytes suggested by a five-gene phylogeny. Cladistics. 2010;26(3):281–300.
- 60. Hirooka Y, Rossman AY, Chaverri P. A morphological and phylogenetic revision of the Nectria cinnabarina species complex. Studies in mycology. 2011;68:35–56. PMC3065984. pmid:21523188
- 61. Hirooka Y, Rossman AY, Samuels GJ, Lechat C, Chaverri P. A monograph of Allantonectria, Nectria, and Pleonectria (Nectriaceae, Hypocreales, Ascomycota) and their pycnidial, sporodochial, and synnematous anamorphs. Studies in mycology. 2012;71:1–210. http://dx.doi.org/10.3114/sim0001. pmid:22685364
- 62. Chaverri P, Salgado C, Hirooka Y, Rossman AY, Samuels GJ. Delimitation of Neonectria and Cylindrocarpon (Nectriaceae, Hypocreales, Ascomycota) and related genera with Cylindrocarpon-like anamorphs. Studies in mycology. 2011;68:57–78. PMC3065985. pmid:21523189
- 63. Herrera CS, Rossman AY, Samuels GJ, Chaverri P. Pseudocosmospora, a new genus to accommodate Cosmospora vilior and related species. Mycologia. 2013;105(5):1287–305. pmid:23921243
- 64. Summerbell RC, Gueidan C, Schroers HJ, de Hoog GS, Starink M, Rosete YA, et al. Acremonium phylogenetic overview and revision of Gliomastix, Sarocladium, and Trichothecium. Studies in mycology. 2011;68:139–62. http://dx.doi.org/10.3114/sim.2011.68.06. pmid:21523192
- 65. Kiyuna T, An KD, Kigawa R, Sano C, Miura S, Sugiyama J. Molecular assessment of fungi in ''black spots'' that deface murals in the Takamatsuzuka and Kitora Tumuli in Japan: Acremonium sect. Gliomastix including Acremonium tumulicola sp nov and Acremonium felinum comb. nov. Mycoscience. 2011;52(1):1–17.
- 66. Grum-Grzhimaylo AA, Georgieva ML, Debets AJM, Bilanenko EN. Are alkalitolerant fungi of the Emericellopsis lineage (Bionectriaceae) of marine origin? IMA fungus. 2013;4(2):213–28. PMC3905940. pmid:24563834
- 67. Platas G, Rossman AY, Farr DF, N G. Hydropisphaera fungicola Rossman, Farr & Newcombe, sp. nov. Fungal Planet. 2008;24.
- 68. Lechat C, Fournier J. Four new species of Ijuhya (Bionectriaceae) from Belgium, metropolitan France and French Guiana. Ascomyceteorg. 2017;9(1):11–8.
- 69. Lechat C, Lesage-Meessen L, Favel A. A new species of Ijuhya, I. fournieri, from french Guiana. Ascomyceteorg. 2015;7(3):101–4.
- 70. Luo J, Zhuang W-Y. New species and new Chinese records of Bionectriaceae (Hypocreales, Ascomycota). Mycol Prog. 2009;9(1):17–25.
- 71. Lechat C, Fournier J. Two new species of Lasionectria (Bionectriaceae, Hypocreales) from Guadeloupe and Martinique (French West Indies). Mycotaxon. 2013;121(1):275–80.
- 72. Chaverri P, Liu M, Hodge KT. A monograph of the entomopathogenic genera Hypocrella, Moelleriella, and Samuelsia gen. nov. (Ascomycota, Hypocreales, Clavicipitaceae), and their aschersonia-like anamorphs in the Neotropics. Studies in mycology. 2008;60:1–66. PMC2275321. pmid:18490956
- 73. Jaklitsch WM, Voglmayr H. Persistent hamathecial threads in the Nectriaceae, Hypocreales: Thyronectria revisited and re-instated. Persoonia: Molecular Phylogeny and Evolution of Fungi. 2014;33:182–211. PMC4312933. pmid:25737600
- 74. Hirooka Y, Rossman AY, Zhuang W-Y, Salgado C, Chaverri P. Species delimitation for Neonectria coccinea group including the causal agents of beech bark disease (BBD) in Asia, Europe, and North America. Mycosystema. 2013;32:485–517.
- 75. Salgado-Salazar C, Rossman A, Samuels GJ, Capdet M, Chaverri P. Multigene phylogenetic analyses of the Thelonectria coronata and T. veuillotiana species complexes. Mycologia. 2012;104(6):1325–50. pmid:22778168
- 76. Crous PW, Wingfield MJ, Richardson DM, Le Roux JJ, Strasberg D, Edwards J, et al. Fungal Planet description sheets: 400–468. Persoonia. 2016;36:316–458. pmid:27616795
- 77. Sung GH, Hywel-Jones NL, Sung JM, Luangsa-Ard JJ, Shrestha B, Spatafora JW. Phylogenetic classification of Cordyceps and the clavicipitaceous fungi. Studies in mycology. 2007;57:5–59. Epub 2008/05/21. pmid:18490993; PubMed Central PMCID: PMCPmc2104736.
- 78. Kuhnert E, Surup F, Wiebach V, Bernecker S, Stadler M. Botryane, noreudesmane and abietane terpenoids from the ascomycete Hypoxylon rickii. Phytochemistry. 2015;117:116–22. http://dx.doi.org/10.1016/j.phytochem.2015.06.002. pmid:26071840
- 79. Chepkirui C, Richter C, Matasyoh JC, Stadler M. Monochlorinated calocerins A-D and 9-oxostrobilurin derivatives from the basidiomycete Favolaschia calocera. Phytochemistry. 2016;132:95–101. http://dx.doi.org/10.1016/j.phytochem.2016.10.001. pmid:27745908
- 80. Helaly SE, Richter C, Thongbai B, Hyde KD, Stadler M. Lentinulactam, a hirsutane sesquiterpene with an unprecedented lactam modification. Tetrahedron Lett. 2016;57(52):5911–3. http://dx.doi.org/10.1016/j.tetlet.2016.11.075.
- 81. Richter C, Helaly SE, Thongbai B, Hyde KD, Stadler M. Pyristriatins A and B: Pyridino-Cyathane Antibiotics from the Basidiomycete Cyathus cf. striatus. J Nat Prod. 2016;79(6):1684–8. pmid:27231731
- 82. Rossman AY, Samuels GJ, Rogerson CT, Lowen R. Genera of Bionectriaceae, Hypocreaceae and Nectriaceae (Hypocreales, Ascomycetes). Studies in mycology. 1999;(42):1–248.
- 83. Silverton JV, Akiyama T, Kabuto C, Sekita S, Yoshihira K, Natori S. X-ray analysis of chaetoglobosin A, an indol-3-yl-cytochalasan from Chaetomium globosum. Tetrahedron Lett. 1976;17(17):1349–50. http://dx.doi.org/10.1016/S0040-4039(00)78061-0.
- 84. Probst A, Tamm C. 19-O-Acetylchaetoglobosin B and 19-O-Acetylchaetoglobosin D, Two New Metabolites of Chaetomium globosum. Helvetica Chimica Acta. 1981;64(7):2056–64.
- 85. Sekita S, Yoshihira K, Natori S, Kuwano H. Chaetoglobosins, Cytotoxic 10-(Indol-3-yl)- cytochalasans from Chaetomium spp. III. Structures of chaetoglobosins C, E, F, G, and J. Chemical & pharmaceutical bulletin. 1982;30(5):1629–38.
- 86. Spöndlin C, Tamm C. Chaetoglobosin M, a new metabolite of a mutant of Diplodia macrospora, belonging to the family of (1H-indol-3-yl)-substituted 10,11-diethyl-10,11-dinorcytochalasans. Helvetica Chimica Acta. 1988;71(8):1881–4.
- 87. Samuels GJ. Perfect states of Acremonium the genera Nectria, Actiniopsis, Ijuhya, Neohenningsia, Ophiodictyon, and Peristomialis. New Zealand J Bot. 1976;14(3):231–60.
- 88. Samuels GJ. Fungicolous, lichenicolous and myxomyceticolous species of Hypocreopsis, Nectriopsis, Nectria, Peristiomialis and Trichonectria. Memoirs of the New York Botanical Gardens 1988;48:1–78.
- 89. Samuels GJ. Some species of Nectria having Cylindrocarponim perfect states. New Zealand J Bot. 1978;16(1):73–82.
- 90. Lechat C, Courtecuisse R. A new species of ljuhya, I. antillana, from the French West Indies. Mycotaxon. 2010;113:443–7.
- 91. Lechat C, Hairaud M. A new species of Ijuhya, I. oenanthicola. Mycotaxon. 2012;119:249–53.
- 92. Siddiqui ZA, Mahmood I. Biological control of plant parasitic nematodes by fungi: A review. Bioresource Technology. 1996;58(3):229–39.
- 93. Yang E, Xu L, Yang Y, Zhang X, Xiang M, Wang C, et al. Origin and evolution of carnivorism in the Ascomycota (fungi). Proceedings of the National Academy of Sciences of the United States of America. 2012;109(27):10960–5. PMC3390824. pmid:22715289
- 94. Chen SY, Dickson DW. Biological control of nematodes by fungal antagonists. In: Chen ZX, Chen SY, Dickson DW, editors. Nematology: advances and perspectives Volume 2: Nematode management and utilization. Tsinghua University Press, China: CABI; 2004. p. 979–1039.
- 95. Ahmed M, Laing MD, Nsahlai IV. Use of Clonostachys rosea against sheep nematodes developing in pastures. Biocontrol Science and Technology. 2014;24(4):389–98.
- 96. Zhang L, Yang J, Niu Q, Zhao X, Ye F, Liang L, et al. Investigation on the infection mechanism of the fungus Clonostachys rosea against nematodes using the green fluorescent protein. Appl Microbiol Biotechnol. 2008;78(6):983–90. pmid:18292995
- 97. Lopez-Llorca LV, Olivares-Bernabeu C, Salinas J, Jansson H-B, Kolattukudy PE. Pre-penetration events in fungal parasitism of nematode eggs. Mycol Res. 2002;106(4):499–506. http://dx.doi.org/10.1017/S0953756202005798.
- 98. Jansson H-B, Lopez-Llorca L. Biology of nematophagous fungi. In: Misra JK, Bruce WH, editors. Trichomycetes and other fungal groups Enfield (NH), USA: Science Publishers, Inc.; 2001. p. 145–73.
- 99. Dijksterhuis J, Harder W, Wyss U, Veenhuis M. Colonization and digestion of nematodes by the endoparasitic nematophagous fungus drechmeria coniospora. Mycol Res. 1991;95:873–8.
- 100. Leger RJS, Roberts DW, Staples RC. A model to explain differentiation of appressoria by germlings of Metarhizium anisopliae. J Invertebr Pathol. 1991;57(3):299–310. http://dx.doi.org/10.1016/0022-2011(91)90134-C.
- 101. Bird AF, McClure MA. The tylenchid (Nematoda) egg shell: structure, composition and permeability. Parasitology. 1976;72(1):19–28.
- 102. Perry RN, Trett MW. Ultrastructure of the eggshell of Heterodera schachtii and H. glycines (Nematoda:Tylenchida). Revue de Nematologie. 1986;9(4):399–403.
- 103. Morton O, Hirsch P, Kerry B. 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. 2004;6(2):161–70. doi:http://dx.doi.org/10.1163/1568541041218004.
- 104. Curtis RHC, Jones JT, Davies KG, Sharon E, Spiegel Y. Plant Nematode Surfaces. In: Davies K, Spiegel Y, editors. Biological Control of Plant-Parasitic Nematodes:: Building Coherence between Microbial Ecology and Molecular Mechanisms. Dordrecht: Springer Netherlands; 2011. p. 115–44.
- 105. Nordbring-Hertz B, Jansson H-B, Tunlid A. Nematophagous Fungi. Encyclopedia of life sciences: John Wiley & Sons, Ltd; 2006.
- 106. Jansson H-b, Friman E. Infection-related surface proteins on conidia of the nematophagous fungus Drechmeria coniospora. Mycol Res. 1999;103(2):249–56. http://dx.doi.org/10.1017/S0953756298007084.
- 107. Baral H-O, Weber E, Gams W, Hagedorn G, Liu B, Liu X, et al. Generic names in the Orbiliaceae (Orbiliomycetes) and recommendations on which names should be protected or suppressed. Mycol Progress. 2017.
- 108. Dijksterhuis J, Veenhuis M, Harder W, Nordbring-Hertz B. Nematophagous fungi: physiological aspects and structure-function relationships. Adv Microb Physiol. 1994;36:111–43. Epub 1994/01/01. pmid:7942313.
- 109. Kawahara T, Itoh M, Izumikawa M, Sakata N, Tsuchida T, Shin-ya K. New chaetoglobosin derivatives, MBJ-0038, MBJ-0039 and MBJ-0040, isolated from the fungus Chaetomium sp. f24230. J Antibiot (Tokyo). 2013;66(12):727–30. pmid:23881215.
- 110. Sekita S, Yoshihira K, Natori S, Kuwano H. Chaetoglobosins G and J, cytotoxic indol-3-yl-cytochalasans from Chaetomium globosum. Tetrahedron Lett. 1977;18(32):2771–4. http://dx.doi.org/10.1016/S0040-4039(01)83069-0.
- 111. Springer JP, Clardy J, Wells JM, Cole RJ, Kirksey JW, Macfarlane RD, et al. Isolation and structure determination of the mycotoxin chaetoglobosin C, a new  cytochalasin. Tetrahedron Lett. 1976;17(17):1355–8. http://dx.doi.org/10.1016/S0040-4039(00)78063-4.
- 112. Löw I, Jahn W, Wieland T, Sekita S, Yoshihira K, Natori S. Interaction between rabbit muscle actin and several chaetoglobosins or cytochalasins. Analytical Biochemistry. 1979;95(1):14–8. http://dx.doi.org/10.1016/0003-2697(79)90178-7. pmid:495949