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Potato farmers conquer a devastating worm—with paper made from bananas

Low-tech approach can quintuple yield and slash need for soil pesticide

Female Golden Nematode (Globodera rostochiensis)
These yellow cysts, attached to potato roots, each contain several hundred eggs that hatch into microscopic worms.USDA/SCIENCE SOURCE

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Potato cyst nematodes are a clever pest. These microscopic worms wriggle through the soil, homing in the roots of young potato plants and cutting harvests by up to 70%. They are challenging to get rid of, too: The eggs are protected inside the mother’s body, which toughens after death into a cyst that can survive in the soil for years.

Now, researchers have shown a simple pouch made of paper created from banana tree fibers disrupts the hatching of cyst nematodes and prevents them from finding the potato roots. The new technique has boosted yields five-fold in trials with small-scale farmers in Kenya, where the pest has recently invaded, and could dramatically reduce the need for pesticides. The strategy may benefit other crops as well.

“It’s an important piece of work,” says Graham Thiele, a research director at the International Potato Center who was not involved with the study. But, “There’s still quite a lot of work to take it from a nice finding to a real-life solution for farmers in East Africa,” he cautions.

Soil nematodes are a problem for many kinds of crops. For potatoes, the golden cyst nematode (Globodera rostochiensis) is a worldwide threat. Plants with infected, damaged roots have yellowish, wilting leaves. Their potatoes are smaller and often covered with lesions, so they can’t be sold. In temperate countries, worms can be controlled by alternating potatoes with other crops, spraying the soil with pesticides, and planting varieties bred to resist infection.

These approaches aren’t yet feasible in many developing countries, in part because pesticides are expensive and resistant varieties of potatoes aren’t available for tropical climates. In addition, small-scale farmers, who can make decent money selling potatoes, are often reluctant to rotate their planting with less valuable crops.

In Kenya, the potato cyst nematode has expanded its range and thrived. “The nematode densities are just so astonishingly high,” says Danny Coyne, a nematode expert at the International Institute of Tropical Agriculture. This is leading to an additional problem of biodiversity loss: Potato farmers are cutting down forests to create new fields free of the nematodes.

The idea that banana paper could help farmers rid their soil of nematodes was hatched more than 10 years ago. Researchers at North Carolina State University (NC State) were looking for a way to help farmers in developing countries safely deliver small doses of pesticides. They experimented with various materials. What works best, they found, is paper made from banana plants. Their tubular, porous fibers slowly release pesticides in the soil for several weeks before breaking down. By then, the plant has developed enough so that even if it does get infected, it already has a healthy root system.

In a field trial, researchers added abamectin, a pesticide that kills nematodes, to the paper. They also planted potatoes in banana paper without abamectin as a control. To their surprise, those plants grew nearly as well as the ones with pesticides. Coyne mentioned this puzzling result to a colleague, a chemical ecologist named Baldwyn Torto who studies the interactions between pests and plants at the International Centre of Insect Physiology and Ecology. “This is fascinating indeed,” Torto recalls thinking.

Together with Juliet Ochola, now a graduate student at NC State, Torto devised several experiments to figure out what was going on. The duo discovered the banana paper holds onto key compounds released from the roots of young potato plants, some of which attract soil microbes that benefit the plant. Nematodes have also evolved to notice these compounds. Some, such as alpha-chaconine, are a signal for nematode eggs to hatch. “If a lot of them hatch at the same time, they’re able to bust open the cysts,” Ochola says. After hatching, the young nematodes sense the compounds and use them to seek out the tender potato roots.

Banana fibers absorbed 94% of the compounds, Ochola and colleagues found. When they exposed nematode eggs to the exudates using the paper, the hatching rate decreased by 85% compared with not using the paper, the team reports today in Nature Sustainability. Other experiments suggested the nematodes that do hatch are far less likely to be able to find potato roots enclosed in the paper.

In nematode-infested fields in Kenya, Coyne and colleagues showed planting potatoes wrapped in plain banana paper tripled the harvest compared with planting without the paper. A tiny dose of abamectin in the paper—just five-thousandths of what would normally be sprayed on the soil—boosted the harvest by another 50%. Presumably, any nematodes that happened to come across the potatoes were then killed by the abamectin. “We’ve got a win here,” Coyne says.

Now, researchers are figuring out how to bring the wrap-and-plant paper to potato farmers in East Africa. Banana plantations in Kenya and nearby countries could supply the fibers, which are now discarded as a waste product. Paper manufacturers could then make the pouches. The biggest challenge, Coyne suspects, will be convincing farmers to buy the paper for the first time.

Once the farmers try the pouches, they’ll find them easy to use, the researchers say. “It’s just wrap and plant,” Ochola says. Simple, yes, but wrapping a lot of potatoes will still be laborious, notes Isabel Conceição, a nematode expert at the University of Coimbra. If a machine is developed to wrap the potatoes, she says, it’s possible the approach might also be feasible on larger farms that use mechanical planters.

Meanwhile, Coyne and his colleagues say they have encouraging results from trials with other tuber crops, such as yam and sweet potato. He also hopes many kinds of vegetables, planted as seeds or seedlings, could be protected from soil pests and pathogens with small pots or trays made from banana fiber, impregnated with various pesticides or biocontrol agents.

The appeal is natural: Banana paper is a biodegradable product, recycled from waste, and it could help protect both farmers and the environment. “We are reducing the amount of pesticides by so much,” Ochola says. “To me, I feel like that’s amazing.”


doi: 10.1126/science.ada1727

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PLANTS & ANIMALS

ABOUT THE AUTHOR

Erik Stokstad

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Erik is a reporter at Science, covering environmental issues. 

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LSU awarded $5 million to look into invasive species harmful to sweet potatoes

A team of LSU AgCenter researchers, collaborating with scientists from four other universities, have been awarded a USDA National Institute of Food and Agriculture grant of more than $5 million, aiding them in developing sweet potato varieties resistant to the invasive guava root-knot nematode.

The AgCenter team is spearheaded by nematologist Tristan Watson. It has also received a sub-grant for $990,000 to support research on sweet potato breeding and characterization of resistance mechanisms and associated genes as well as extension of research findings to regional and national stakeholders.

Watson: “Root-knot nematodes are particularly damaging to the sweet potato. The overall goal of this project is to provide Louisiana sweet potato growers effective tools for the management of established and emerging root-knot nematode species.”

Source: lsuagcenter.com

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Dear Colleagues and Friends,

I want to announce that the handbook I developed together with to other editors and 70 chapter authors on: “Integrated Nematode Management: state-of-the-art and visions for the future”

is now officially available, gratis, in open-access format on the CABI website.

The book can be download for direct viewing on your computer or smart phone or both or saved as a pdf for future use.

Please forward this link to anyone you know who might be interested in what we feel is an important hand book on applied plant pathology and nematology.

The link to the e-Book is available on the CABI website at:

https://urldefense.com/v3/__https://www.cabi.org/bookshop/book/9781789247541/__;!!PvXuogZ4sRB2p-tU!WR2OAeEqidoosMzpHs-bVXo8jPYGPd3SSlo9hVPa2C09W-DLj0A5szz2OSyyOczDLg$

The editors put together the 500 pages of science in 65 chapters with over 250+ figures in the 12 month window we set. Good collaborators.

I look forward to any comments you have and hope the vast majority are positive! Of course nothing is perfect.

All the best over the holiday season whether Thanksgiving, Christmas, New Years or other special occasion.

Sincerely yours

Richard Sikora

rsikora@uni-bonn.de

——————————————————————————

Richard A. Sikora, Prof. em.

Institute for Crop Science & Resource Conservation Consultant Plant Health Management University of Bonn, Germany

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Beech leaf disease is ravaging North American trees

Two new studies gauge impact and cause of forest blight

A view underneath North American beech trees
The fast-spreading beech leaf disease is starting to kill the widespread, majestic American beech, which can rise to about 40 meters tall and live about 400 years.MIRCEA COSTINA/ALAMY STOCK PHOTO

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A tree disease first spotted 9 years ago in Ohio is now a leading threat to one of eastern North America’s most important trees. The poorly understood malady, called beech leaf disease, is spreading rapidly and killing both mature American beeches and saplings, new research shows.

“This study documents how rapidly [the disease] has spread since its first observation in 2012,” says Robert Marra, a forest pathologist at the Connecticut Agricultural Experiment Station who was not involved with the work.

American beeches (Fagus grandifolia) are found across the eastern United States and Canada. The trees, which can grow nearly 40 meters tall and live up to 400 years, are a major player in many forests. Beeches constitute more than 25% of forests in Vermont, for example.

Historically, a blight called beech bark disease has been the primary threat to the species. But now, beech leaf disease appears to pose a bigger danger. First spotted in northeastern Ohio, it causes parts of leaves to turn leathery and branches to wither. The blight can kill a mature tree within 6 to 10 years. It has now been documented in eight U.S. states and in Canada.

In Rhode Island, observers first spotted beech leaf disease in 2020, confined to a small area, says Heather Faubert of the University of Rhode Island’s Plant Protection Clinic who was not involved with the study. But, “This year, it’s everywhere.”

Beech leaf disease symptoms of dark banding between the leaf veins seen on beech tree leaves
Beech leaf disease causes some leaves to emerge with leathery, dark green parts. As the season continues, those parts may turn yellow or brown.MARY PITTS/ HOLDEN FORESTS AND GARDENS

To track the disease, Constance Hausman, an ecologist for a network of parks called the Cleveland Metroparks, and colleagues surveyed 64 0.04-hectare forest plots within 224 kilometers of Lake Erie in Ohio, Pennsylvania, New York, and Canada’s Ontario province. An analysis of 894 beeches in the plots found nearly half had the leaf disease, whereas just 34 had bark disease. Earlier surveys elsewhere had found the disease mostly attacked saplings, but the new work finds it is attacking mature trees, too, the team reported last month in Forest Ecology and Management. In forests near Lake Erie, beech leaf disease has now “become pervasive,” the group says.

The disease is “attacking the life cycle of beech trees in both directions,” Hausman says. The number of trees could fall so much in some forests that the species no longer serves key ecological functions, she warns, such as providing food and shelter for birds and other animals.

Another recent study by a different team examines an ongoing mystery: What exactly causes the disease? Earlier work raised suspicions that a tiny, previously unknown nematode worm that feeds on beech buds and leaves, dubbed Litylenchus crenatae mccannii, plays a role in spreading the blight.

Now, researchers report in Phytobiomes that when they examined diseased beech leaves, the tissues contained a fungus and four bacterial species also carried by the nematode. That suggests both the nematode and a pathogen it carries are contributing to the disease, says study co-author Pierluigi “Enrico” Bonello, an ecologist at Ohio State University, Columbus.

Marra is skeptical, however. He says one of the study’s suspects, Wolbachia, is known only to help its hosts. So he thinks its role in beech leaf disease, if any, might just be to strengthen the nematode’s attack.

So far, researchers haven’t identified a practical, cost-effective treatment for the disease, although some beeches appear to be resistant. But using those trees to breed new resistant strains could take decades, researchers say.

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Bacterial endosymbionts protect beneficial soil fungus from nematode attack

 View ORCID ProfileHannah Büttner,  View ORCID ProfileSarah P. Niehs,  View ORCID ProfileKoen Vandelannoote,  View ORCID ProfileZoltán Cseresnyés, Benjamin Dose,  View ORCID ProfileIngrid Richter,  View ORCID ProfileRuman Gerst,  View ORCID ProfileMarc Thilo Figge,  View ORCID ProfileTimothy P. Stinear,  View ORCID ProfileSacha J. Pidot, and  View ORCID ProfileChristian Hertweck

 See all authors and affiliationsPNAS September 14, 2021 118 (37) e2110669118; https://doi.org/10.1073/pnas.2110669118

  1. Edited by Nancy A. Moran, The University of Texas at Austin, Austin, TX, and approved August 5, 2021 (received for review June 9, 2021)

Significance

Soil is a complex and competitive environment, forcing its inhabitants to develop strategies against competitors, predators, and pathogens. Identifying and understanding the molecular mechanisms has translational value for medicine, ecology, and agriculture. In this study, we show that a member of important soil-dwelling fungi (Mortierella) forms a tight alliance with toxin-producing bacteria (Mycoavidus) that live within the fungal hyphae and protect their host from nematode attack. This discovery is relevant since Mortierella species correlate with healthy soils and are used as plant growth–promoting fungi in agriculture. Unraveling an ecological role for fungal endosymbionts in Mortierella, our results contribute to the understanding of a mainspring in fungal–endobacterial symbioses and open the possibility for the development of new biocontrol agents.

Abstract

Fungi of the genus Mortierella occur ubiquitously in soils where they play pivotal roles in carbon cycling, xenobiont degradation, and promoting plant growth. These important fungi are, however, threatened by micropredators such as fungivorous nematodes, and yet little is known about their protective tactics. We report that Mortierella verticillata NRRL 6337 harbors a bacterial endosymbiont that efficiently shields its host from nematode attacks with anthelmintic metabolites. Microscopic investigation and 16S ribosomal DNA analysis revealed that a previously overlooked bacterial symbiont belonging to the genus Mycoavidus dwells in M. verticillata hyphae. Metabolic profiling of the wild-type fungus and a symbiont-free strain obtained by antibiotic treatment as well as genome analyses revealed that highly cytotoxic macrolactones (CJ-12,950 and CJ-13,357, syn. necroxime C and D), initially thought to be metabolites of the soil-inhabiting fungus, are actually biosynthesized by the endosymbiont. According to comparative genomics, the symbiont belongs to a new species (Candidatus Mycoavidus necroximicus) with 12% of its 2.2 Mb genome dedicated to natural product biosynthesis, including the modular polyketide-nonribosomal peptide synthetase for necroxime assembly. Using Caenorhabditis elegans and the fungivorous nematode Aphelenchus avenae as test strains, we show that necroximes exert highly potent anthelmintic activities. Effective host protection was demonstrated in cocultures of nematodes with symbiotic and chemically complemented aposymbiotic fungal strains. Image analysis and mathematical quantification of nematode movement enabled evaluation of the potency. Our work describes a relevant role for endofungal bacteria in protecting fungi against mycophagous nematodes.

A healthy soil nourishes plants and animals, purifies water and air, and promotes sustainable agriculture. Characteristic for highly complex and competitive soil ecosystems are the frequent and direct interactions between all soil-dwelling microorganisms, animals, and plants (12), all of which need to be provided with minerals and carbon sources. Thus, carbon cycling, mainly promoted by fungal saprophytes and decomposers that release nutrients from decaying matter, plays a pivotal role for soil health (34). Fungi belonging to the genus Mortierella are the most common soil-dwelling fungi, ubiquitously distributed in all parts of the world, inhabiting highly diverse niches including the rhizosphere and plant tissues (59). Owing to their ability to degrade biopolymers as well as xenobiotics, they not only deliver energy-rich carbon sources but also clear the environment from pollutants (1011). Typically associated with healthy soils, Mortierella species are recognized as valuable plant growth–promoting fungi in agriculture (912).

Even so, all fungi, including Mortierella species, are threatened by micropredators such as nematodes (1315). In order to oppose these predators, fungi have developed a diverse set of defense strategies. These include the production of toxic proteins and nematocidal natural products, hyphal piercing, trapping, egg parasitism, and endoparasitism (1316). Information on defense strategies employed by Mortierella species against nematodes is, however, scarce. It is known that Mortierella globalpina traps nematodes by means of its hyphae and penetrates the nematode’s cuticula. In this way, M. globalpina may protect its host plants from plant-parasitic nematodes (e.g., Meloidogyne chitwoodi) (17). Antinematode activities have been implicated for some Mortierella species (1819), including Mortierella alpina [against Meloidogyne javanica or Heterodera sp. (2021)], but it is not a general trait of Mortierella (2122). Apart from the hyphal trapping strategy, insight into the molecular basis of the antinematode activities of Mortierella is missing. Furthermore, on a more general note, it is remarkable that thus far no Mortierella secondary metabolites have been associated with potential protective roles against nematodes.

Here, we report a so far unknown strategy of a Mortierella species to protect itself from nematode attack. We provide evidence that cytotoxic benzolactones initially isolated from fungal cultures are in fact produced by bacterial endosymbionts that have been overlooked thus far. We also show that the bacteria dwelling in the fungal hyphae protect their host from predatory nematodes.

Results and Discussion

Mortierella Fungus Harbors Bacterial Endosymbionts Producing Toxic Macrolactones.

We reasoned that benzolactones CJ-12,950 and CJ-13,357 (Fig. 1A) (23) from cultures of Mortierella verticillata [synonym Podila verticillata (24)] could play a role as nematode defense metabolites. Although the initial report on CJ-12,950 and CJ-13,357 only stated that these compounds enhance the expression of the low-density lipoprotein receptor in human hepatocytes (23), they share the benzolactone enamide architecture with structurally related vATPase inhibitors (2526). Moreover, the architectures of CJ-12,950 and CJ-13,357 specifically resemble those of Burkholderia sp. strain B8 produced necroximes A to D (1 to 4), which proved to be cytotoxic (27). Since only the two-dimensional structures of CJ-12,950 and CJ-13,357 had been reported (23), we assigned their absolute configurations by examining the structural relationships with necroximes C and D. Optical rotation comparison, high-performance liquid chromatography (HPLC)–based coelution experiments and comparison of tandem mass spectrometry (MS/MS) fragmentation indicated that necroxime D (4) is identical to CJ-12,950, and necroxime C (3) is identical to CJ-13,357 (Fig. 1B and SI Appendix, Table S4). These assignments were corroborated by comparison of the NMR spectra of purified metabolites (SI Appendix, Table S9).

Fig. 1.

Fig. 1.

Bacterial origin of cytotoxic benzolactones from M. verticillata cultures. (A) Cytotoxic lactone compounds assigned to endofungal symbionts from the fungus R. microsporus (14), M. verticillata (34), Pseudomonas sp. (5), and a tunicate and the bacterium Gynuella sunshinyii (6). (B) Metabolic profiles of extracts from Burkholderia sp. strain B8 and M. verticillata NRRL 6337 as symbiont or cured strain as total ion chromatograms in the negative mode. (C) Fluorescence micrograph depicting endosymbionts living in the fungal hyphae; staining with Calcofluor White and Syto9 Green. (D) Phylogenetic relationships of Mortierella symbionts, Burkholderia sp. strain B8, and other bacteria based on 16S rDNA. BRE, Burkholderia-related endosymbiont of Mortierella spp. (E) Metabolic profiles of extracts from M. verticillata NRRL 6337 and other necroxime-negative M. verticillata strains analyzed for endosymbionts in this study as total ion chromatograms in the negative mode. M, medium component. (F) Growth of symbiotic M. verticillata NRRL 6337 in comparison to the cured strain.

Given the bacterial origin of the necroximes (27) and related benzolactones (252830), we questioned the biosynthetic capability of M. verticillata and sought to identify the true producer. Since several Mortierella spp. have been reported to live in symbiosis with bacteria (3132), we suspected an endosymbiont to be the true source of 3 and 4. Yet, a 2018 report investigating the prevalence of Burkholderiaceae-related bacteria within Mortierella spp. stated that strain NRRL 6337 was devoid of endosymbionts (32). Nonetheless, we re-examined the same strain for endosymbionts by staining fungal hyphae with the chitin-binding Calcofluor White dye, and tentative endobacteria with the nucleic acid dye Syto9 Green (Fig. 1C). Fluorescence microscopy revealed the presence of endosymbiotic organisms in M. verticillata NRRL 6337 (SI Appendix, Fig. S1).

To identify the observed bacterial endosymbionts, we cut a small piece of fungal mycelium and extracted holobiont DNA, followed by PCR amplification of the 16S ribosomal DNA (rDNA) region using universal primers. Sequencing of the 16S rDNA region (SI Appendix, Table S1) and BLAST analysis indicated that the symbiont of M. verticillata NRRL 6337 is a Mycoavidus species. Notably, members of this genus have been reported as symbionts of soil-dwelling fungi (32). So far, the full genomes of only three Mycoavidus cysteinexigens strains from Mortierella elongata and Mortierella parvispora have been sequenced (313335). PCR-amplified bacterial 16S rDNA sequences from other Mortierella fungi, however, revealed further Mycoavidus endosymbionts with three phylogenetically distant clades (Mortierella-associated Burkholderia-related endosymbiont [MorBRE] groups A to C) (32). Through phylogenetic analysis, we found that the Mycoavidus symbiont of M. verticillata NRRL 6337 falls into MorBRE group A (Fig. 1D and SI Appendix, Fig. S3) comprising symbionts of Mortierella humilisMortierella gamsiiMortierella basiparvispora, and M. elongata (M. cysteinexigens). To better understand the occurrence of Mycoavidus endosymbionts in M. verticillata strains, we investigated five additional M. verticillata strains for the presence of endosymbionts. Amplification of the 16S rDNA regions from gDNA of symbionts of these strains revealed a conserved occurrence of Mycoavidus endosymbionts in M. verticillata strains. Interestingly, these additional endosymbionts all fall into another phylogenetic group together with Burkholderia sp. strain B8. Furthermore, analysis of the metabolic profiles of the respective fungi did not show any production of necroximes (Fig. 1 D and E). This finding shows that endosymbionts may frequently occur in Mortierella and other species of the order Mucorales, but they can be phylogenetically different.

To clarify whether bacterial endosymbionts are the true producers of 3 and 4, we aimed at curing M. verticillata NRRL 6337 of its symbiont through the addition of antibiotics (36). Over the course of several months, we subcultivated the fungal strain on agar plates containing kanamycin, ciprofloxacin, or chloramphenicol. During treatment, changes of the fungal growth were noticeable (Fig. 1F). Finally, we confirmed the absence of the symbionts by fluorescence staining, microscopic inspection, and PCR analysis (SI Appendix, Figs. S2 and S4). The metabolic profiling of the symbiont-free fungal strain by liquid chromatography (LC) combined with high-resolution electrospray ionization revealed the complete absence of 3 and 4 (Fig. 1B). These findings indicate that Candidatus Mycoavidus necroximicus is the true producer of the benzolactones.

Ca. M. necroximicus Dedicates 12% of Its Genome to Secondary Metabolism.

To gain insight into the symbiont’s biosynthetic potential, with particular focus on the molecular basis of necroxime biosynthesis, we aimed at sequencing the genome of the endosymbiont. Attempts to isolate and cultivate the endosymbiont in the absence of the fungal host, however, proved to be futile. Methods previously used to axenically cultivate similar fungal endobacteria did not enable growth of the endosymbionts (3337), indicating a strong dependence of the bacterial symbiont on the host environment. Thus, we sought to enrich the symbiotic bacteria for DNA isolation. Initially, physical disruption of the host’s mycelium resulted in high levels of contamination with fungal DNA, which complicated the assembly of the endosymbiont’s genome. Eventually, we succeeded in retrieving a bacterial cell pellet by filtration and centrifugation of the turbid supernatant of shaking cultures in baffled flasks and isolated the genomic DNA from resuspended bacteria.

The genome of the bacterial endosymbiont was sequenced using a combination of Oxford Nanopore MinION and Illumina NextSeq sequencing, and both data sets were used to generate a hybrid genome assembly. Of the 118 contigs, a single 2.4 Mb contig of putative bacterial origin was identified through homology searches using the Mycoavidus-like 16S rDNA sequence previously amplified from M. verticillata NRRL 6337. Following trimming of overlapping ends (suggesting a circular chromosome) the final 2.2 Mb contig was found to contain 1,768 CDS, 6 rRNAs, 42 tRNAs, and a GC content of 50.6% (genome accession number: PRJNA733818). The 16S rDNA sequence of the new strain has 98.82% nucleotide identity to M. cysteinexigens B1-EBT (33). Even so, genomic comparisons showed an average nucleotide identity of only 81.85% across the two genomes. By current standards for molecular species discrimination, the newly identified Mortierella endosymbiont should be considered a new species (Ca. M. necroximicus) (3839).

By comparative genomic analyses, we noted that the genomes of the two endofungal strains AG77 and B1-EBT isolated from M. elongata (3335) are 400 to 500 kb larger than the genome of Ca. M. necroximicus. Only the genome of strain B2-EB isolated from M. parvispora (34) is smaller (∼500 kb) than the genome of Ca. M. necroximicus (2.2 Mb). When investigating shared protein orthologs, we noted that a core genome encoding 1,164 proteins exists among the four genomes at the 70% identity level (Fig. 2A). However, a further all-versus-all comparison showed B1-EBT and AG77 to be the most closely related as they share ∼75% of their deduced proteome. The B2-EB and Ca. M. necroximicus strains are more distantly related to B1-EBT and AG77, as well as each other, with only a small number of proteins shared exclusively with either B1-EBT (20 and 17 proteins, respectively) or AG77 (17 and 22 proteins, respectively) (Fig. 2B).

Fig. 2.

Fig. 2.

Comparative genomic analyses of Mycoavidus spp. (A) Number of orthologous proteins among the four Mycoavidus strains at 70% identity. (B) Circos plot of shared protein orthologs, and secondary metabolite loci (detected by antiSMASH v5) in Mycoavidus genomes. Outer blocks (orange, brown, yellow, green) represent genome sizes, while the inner blocks represent genomic positions of secondary metabolite loci. Lines linking the three genomes show position of genes whose proteins are orthologous at 70% identity. Depicted are the genome sequences of M. cysteinexigens strains AG77, B1-EB, B2-EB, and Ca. M. necroximicus (Ca. M. nec.). (C) Number of gene clusters putatively coding for natural products in Mycoavidus spp. detected by antiSMASH and by manual assignment. (D) BGCs and their encoded assembly lines identified from the endofungal Ca. M. necroximicus are displayed. A, adenylation; AT, acyltransferase; C, condensation; DH, dehydratase; E, epimerization; Gnat, GCN5-related N-acetyltransferase; KR, ketoreductase; KS, ketosynthase; MT, methyltransferase; OX, oxygenase; TE, thioesterase domains. Acyl carrier (light blue) and peptidyl carrier proteins (dark blue) are shown as circles without designators. (E) Homologous benzolactone BGCs in the genome of Burkholderia strain B8 and Ca. M. necroximicus.

Whereas biosynthetic gene clusters (BGCs) are present in the genomes of all four studied Mortierella symbionts, antiSMASH analysis (40) revealed that the biosynthetic potential for secondary metabolites is by far the greatest in Ca. M. necroximicus (Fig. 2C). Despite the relatively small genome for Mycoavidus standards, ∼12% of its protein-encoding capacity is dedicated to natural product biosynthesis. We identified nine nonribosomal peptide synthetase (NRPS) gene clusters, two polyketide synthase (PKS) gene clusters, two hybrid PKS/NRPS gene clusters, and five other BGCs (Fig. 2D). Notably, several large PKS and NRPS gene loci present in the Ca. M. necroximicus genome are absent in the genomes of strains B1-EBT, B2-EB, and AG77 (Fig. 2B). This BGC list includes a cryptic BGC (Mcyst_0009–0017) encoding a PKS/NRPS hybrid that shows high similarity to the necroxime assembly line from Burkholderia sp. strain B8 (97% coverage, ∼70% amino acid identity), which has been unequivocally linked to necroxime biosynthesis by targeted gene knockouts (Fig. 2E) (27). The only major difference between the two BGCs is the NRPS gene necA, which is missing in the genome of Ca. M. necroximicus. This finding is in full agreement with the current biosynthetic model, since NecA is responsible for the attachment of the peptide side chain in 1 (Fig. 1A) (27), which is absent in 3 and 4. Furthermore, the architecture of the encoded PKS/NRPS modules is perfectly in line with the biosynthesis of the benzolactone enamide backbone of 3 and 4. Based on these in silico predictions, we inferred that this PKS/NRPS hybrid gene cluster codes for the biosynthesis of 3 and 4 (SI Appendix, Figs. S6 and S8). Together with the metabolic profiling of the cured fungal strain, these data indicate that the bacterial endosymbionts, not the fungus, are the true producers of the benzolactones 3 and 4. CJ-12,950 and CJ-13,357 are thus important additions to the small group of natural products that were believed to be fungal metabolites but are actually produced by bacterial endosymbionts; rhizoxins (41) and rhizonins (42) from symbionts of Rhizopus microsporus (43), and endolides from Stachylidium bicolor (44). From an ecological viewpoint, it is remarkabe that endosymbiotic bacteria were identified as the true producers of the virulence factor of the rice-seedling blight fungus R. microsporus (363745). Given the different ecological context of Mortierella, however, we assumed that the necroximes may have another function in microbial interactions.

Necroximes Protect the Fungal Host from Nematode Attacks.

To learn more about the potential role of necroximes (3 and 4) in the ecological context of the MortierellaMycoavidus symbiosis, we investigated whether these toxins could impair the growth of, or even kill, competitors. Therefore, we considered that the common natural habitat of Mortierella species, including M. verticillata NRRL 6337, is soil, and that microbial survival in the soil environment is not only determined by the capacity to grow under harsh conditions but also by the ability to defend oneself from (micro)predators (46). Among the most abundant fungal predators are nematodes, which share the same soil habitat as Mortierella (47).

To determine if 3 and 4 or any other endobacteria-derived substance have anthelmintic activity, we first performed a viability assay against the model organism, Caenorhabditis elegans (48). We cultivated both cured (Mycoavidus-free) and symbiotic M. verticillata NRRL 6337 on potato dextrose agar (PDA agar). Cultures were extracted, and each extract was fractionated by preparative HPLC. The individual fractions (F1 to F9) were subsequently tested against C. elegans. Anthelmintic activity in this assay was determined by the ability of C. elegans to feed on a supplied Escherichia coli food source in the presence of the different fractions. Consumption of bacteria indicates unimpeded nematodes, whereas growth of E. coli indicates that the nematodes are negatively affected by the added substances (Fig. 3A). Notably, all fractions of the extract obtained from the cured strain culture were found to be inactive in the C. elegans assay. In contrast, we observed a marked nematocidal activity of fraction 6 from the extract of the symbiotic fungus. By LC/MS measurements we confirmed the presence of 3 and 4 in the active fraction. In order to determine the anthelmintic potency of the major metabolite 4, we performed the viability assay against C. elegans using increasing concentrations of the pure substance and determined an inhibitory concentration at 50% (IC50) value of 11.3 µg ⋅ mL−1 (24.66 µM) (Fig. 3B). Interestingly, the amount of isolated necroximes from fungal cultures grown on agar plates is ∼11 µg ⋅ mL−1. Assuming that the actual concentration in fungal hyphae is slightly higher due to an uneven diffusion into the agar and some loss during the purification steps, we conclude that the concentrations inside and around the fungal mycelium are sufficiently high to fully protect it from mycophagous nematodes.

Fig. 3.

Fig. 3.

Nematocidal activity of symbiont-derived toxins. (A) Viability assay of C. elegans in presence of extract fractions of symbiotic and cured M. verticillata NRRL 6337. HPLC profiles of extracts are shown with corresponding effect on nematodes, measured as effect on the E. coli optical density (OD). When nematode growth is impaired by the fraction, E. coli cells are not consumed, and thus the OD600 is not altered (error bars represent mean of three biological replicates). The red asterisk represents 4. (B) Toxicity screening of 4 against C. elegans. The red line marks IC50 at 11.3 µg ⋅ mL−1 (24.66 µM; 95% CI, 21.45 to 28.37 µM; error bars as mean of five biological replicates). (C) Nematode counts from propagation assay of M. verticillata and A. avenae cocultures. Bars represent relative nematode numbers compared to the mean of the nematode count from cured M. verticillata NRRL 6337 cultures. cur., cured; sym., symbiotic. *P < 0.02; ***P < 0.001; ****P < 0.0001. Data represent three biological replicates with three technical replicates each. (D) Workflow of image analysis and mathematical evaluation of A. avenae mobility in fungal–nematodal coincubations. Processing of time series is demonstrated by one time frame. Exemplary images of nematodes from two time frames (frame 1 and 26) are shown to illustrate differences in motility. Results of calculated mobility ratios (MR) were used for live or paralyzed/dead categorization. (E) Results of image analysis and mathematical quantification of nematode movement. Bars show ratio between moving/living nematodes and paralyzed/dead nematodes, which were harvested from cocultures of A. avenae with symbiotic M. verticillata NRRL 6337 cultures, cured NRRL 6337 cultures, or CSB 225.35 cultures. Numbers and error bars were calculated from minimal 176 worms from three biological replicates. (F) Stereomicroscopic images and schematic picture of chemical complementation assay with a magnitude of 25×. Sample of nematodes harvested from plates containing symbiotic, cured, or cured and with 4 chemically complemented M. verticillata NRRL 6337 cultures. (G) Schematic summary of tripartite interaction between fungal host, bacterial endosymbiont, and mycophagous nematodes.

In order to corroborate a potential host-protective role of the symbiont-derived toxin, we next focused on a fungivorous nematode. Therefore, we selected Aphelenchus avenae, a predator using a stylet to feed on fungi, which pierces the fungal cell wall and allows the fungivore to ingest the fungal cytoplasm (4950). Sharing the same soil habitat, A. avenae represents a realistic predator of Mortierella spp. (51). To investigate the effects of symbiotic and cured M. verticillata strains on the feeding behavior and survival of A. avenae, we determined the number of animals that were harvested from fungal–nematodal cocultures. In addition, we compared the mobility ratios of the nematodes in correlation to the presence or absence of the bacterial symbiont. As a control, we employed the symbiont-bearing, but necroxime-negative, M. verticillata strain CSB 225.35 (Fig. 1E), thus ruling out an influence solely based on the presence of bacterial symbionts.

To determine nematodal propagation rates, we inoculated plate cultures of symbiotic and cured fungi with A. avenae and cocultivated both organisms for 17 to 24 d (three biological triplicates). Subsequently, nematodes were isolated from the cocultivation plates by Baermann funneling (52), transferred onto water-agar plates, and counted by stereomicroscopic visualization. We found that significantly fewer nematodes are able to grow in the presence of the necroxime-producing endosymbionts (Fig. 3C and SI Appendix, Fig. S9 and Tables S5 and S6).

To scrutinize the effect of the toxin on the fitness of the fungivorous nematodes, we harvested the animals from cocultures and determined their movement—and thus the mobility ratios—by image analysis and mathematical quantification (Fig. 3D). Using stereoscopic time series to track their movement, we compared the area covered by each moving nematode during the time series to the area covered solely by its body without movement, allowing us to differentiate active (living) from inactive (dead or paralyzed) animals. Analyzing a minimum of 176 nematodes from three independent experiments, we observed a significant decrease in the mobility ratio of nematodes grown on symbiotic M. verticillata NRRL 6337 compared to cured NRRL 6337 cultures and to necroxime-negative CSB 225.35 cultures (Fig. 3E and SI Appendix, Fig. S10 and Tables S7 and S8).

HPLC analyses of the plate extracts detected necroximes only in symbiotic cultures of NRRL 6337 but not in cured strains or CBS 225.35, correlating once again the toxins with reduced numbers and lower fitness of the nematodes. To unambiguously assign the nematocidal activity in the propagation assay to the necroximes, we repeated the A. avenae assay with the cured Mortierella strain and chemically complemented the major toxin. Specifically, we overlaid the cured strain NRRL 6337 with solutions of 4 in increasing concentrations (25 µM [IC50], 50 µM, 109 µM, and 219 µM). We then compared A. avenae propagation in necroxime-complemented cultures to untreated cured as well as symbiotic fungi by microscopic examination after two weeks of coincubation. For cultures supplemented with 25 µM or 50 µM of 4, we noted a moderate reduction of nematode propagation, whereas in cultures supplemented with 109 µM or 219 µM of 4 the presence of nematodes in the fungus was abolished (Fig. 3F and SI Appendix, Figs. S11 and S12). The elevated concentrations compared to the IC50 value can be explained due to an uneven distribution of 4 into deeper layers of the hydrophobic fungal colony and the ongoing growth of fungal hyphae, which were not wetted with toxin solution. Nonetheless, these experiments unambiguously verified that the chemical complementation restores the anthelmintic effect. Thus, we uncovered an important role of a natural product in the complex tripartite interplay of symbiont, host, and (micro)predator (Fig. 3G).

Conclusions

In this study, we uncovered a previously overlooked bacterial endosymbiont that protects the important soil-dwelling fungus M. verticillata from a fungivorous nematode. Comparative genomics indicate that the yet unculturable bacterial symbionts belong to a new species that is endowed with a high biosynthetic potential. Through metabolic profiling of the symbiotic wild type and cured aposymbiotic fungi, we provide evidence that the endofungal bacteria are the true producers of highly toxic macrolides that were previously believed to be fungal metabolites. Importantly, these compounds (necroximes) efficiently protect the host from nematode attack, as demonstrated by coculture experiments, chemical complementation, and image analyses. Thus, this work not only reveals an ecological role of endofungal bacteria but also introduces a strategy to ward off micropredators. Consequently, the bacterial biosynthesis of necroximes provides an advantage of the fungal–bacterial alliance over other aposymbiotic or necroxime-negative symbiotic M. verticillata strains in the soil niche. Beyond inspiring the discovery of related tactics in symbioses, our findings may set the basis for new biocontrol agents, with the prospect of shielding plant hosts from plant-pathogenic nematodes.

Materials and Methods

Isolation of Natural Products.

For 3 and 4 isolation, M. verticillata NRRL 6337 was cultivated on PDA plates (Bacto, BD) at 26 °C. The culture was extracted twice with 1:1 volume of ethyl acetate overnight. The organic phase was concentrated under reduced pressure and the residue was dissolved in methanol. The extracts were prefractionated on an open Sephadex LH-20-column with methanol as eluent. Necroxime-containing fractions were further purified with a preparative HPLC under following conditions: A, H2O + 0.01% TFA; B, methanol; and 15 to 100% B in 35 min, 15 mL ⋅ min−1 [Phenomenex, Luna, 10 µm, C18(2), 100 Å, 250 × 21.2 mm]. NMR analysis was carried out on a 600 MHz Avance III Ultra Shield (Bruker), and signals were referenced to the residual solvent signal (DMSO-d6).

Identification of Endosymbionts in M. verticillata.

For the preparation of cured fungal strains, fungi were continuously subcultivated at 24 °C on PDA plates containing 40 µg ⋅ mL−1 ciprofloxacin or 50 µg ⋅ mL−1 kanamycin for several months. After phenotypic changes were observed by eye, an agar plate of each fungal culture was extracted with 20 mL ethyl acetate and controlled for the absence of 3 and 4 by LC/MS. Final verification of the cured fungal strains was performed by fluorescence staining (Calcofluor White Stain [Sigma] and SYTO 9 Green [Invitrogen]).

Genome Assembly for Ca. M. necroximicus.

M. verticillata NRRL 6337 was grown in MM9 medium (53) and orbitally shaken at 160 rpm and 26 °C. The turbid supernatant, containing bacteria from disrupted hyphae, was twice filtered through a membrane (pore diameter, 40 µm) and centrifuged (12,000 × g, 25 °C, 10 min) until a stabile pellet occurred. The genomic DNA was extracted with the MasterPure DNA Purification Kit (Epicentre). For long-read sequencing on the MinION platform, DNA quality was evaluated by pulsed-field gel electrophoresis and prepared for sequencing according to the protocol of the Ligation Sequencing kit (Oxford Nanopore). DNA was loaded onto a single MinION flow cell, and data were collected over a 72-h period. DNA was prepared for sequencing on the Illumina NextSeq platform using the Nextera XT DNA preparation kit (Illumina) with ×150 bp paired end chemistry and with a targeted sequencing depth of >50×. Combined MINion and Illumina sequencing data were assembled using the Unicycler hybrid assembler (54) to form a single contig 2.2 Mb containing a 98.82% match to the M. cysteinexigens rDNA gene. The evaluation of secondary metabolite loci was performed with antiSMASH version 5 (55).

Nematode Assays.

Liquid assays for active-fraction determination and potency assessment against C. elegans were conducted as previously described (48). For A. avenae coincubation assay, an aliquot of hyphae of each tested Mortierella strain was transferred to a PDA plate and incubated at 24 °C overnight. Nematodes were sterilized and starved. After one washing step with K-medium, nematodes were resuspended in 300 µL K-medium and aliquots of 50 µL were distributed onto the fresh fungal cultures. Plates were dried and controlled for living nematodes before they were incubated for 17 to 24 d at 20 °C. For the evaluation, nematodes were harvested via Baermann funneling (56). Funneled A. avenae were transferred on 1.5% water-agar plates containing 200 mM geneticin and 50 µg ⋅ mL−1 kanamycin overnight and subsequently monitored with a Zeiss Axio Zoom.V16 Stereomicroscope for worm count and bioinformatics (https://www.jipipe.org/). Remaining plates were extracted with ethyl acetate to control the metabolite production and processed as described before. For the A. avenae chemical complementation assay, an aliquot of hyphae of the respective fungus was transferred into 12-well plates filled with 1 mL PDA and incubated overnight. The 4 dissolved in 200 µL 50% MeOH was applied and evaporated at room temperature. Nematode suspensions of 50 µL were distributed onto the fungi, dried, and coincubated for 14 d at 20 °C. For evaluation, the coculture was removed from the well and washed in 5 mL K-medium overnight. The mixture was filtered through miracloth (Merck) to avoid agar carryover and left at 4 °C for 1 h. The remaining worms were transferred onto 6-well plates containing 5 mL 1.5% water-agar with 200 mM geneticin and 50 µg ⋅ mL−1 kanamycin. After the plates were dried, the worm count from each plate was assessed with a Zeiss Axio Zoom.V16 Stereomicroscope.

Data Availability

Genome sequence data have been deposited in GenBank (PRJNA733818). The 16S rDNA sequences of the Mortierella endosymbionts were deposited at the NCBI database (BRE_MvertCBS_346.66: MZ330684; BRE_MvertCBS_220.58: MZ330685; BRE_MvertCBS_225.35: MZ330686; BRE_MvertCBS_315.52: MZ330687; BRE_MvertCBS_100561: MZ330688).

Acknowledgments

Mortierella strains were supplied by ARS Culture Collection (NRRL) and the Jena Microbial Resource Collection. C. elegans was provided by the Caenorhabditis Genetics Center, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440). A. avenae was received as a kind gift from Prof. Dr. M. Künzler (ETH Zürich). We thank E. Bratovanov (HKI) for helpful discussions. Assistance by K. Martin and S. Linde (HKI) is gratefully acknowledged. H.B. and S.P.N. were funded by the Deutsche Forschungsgemeinschaft (DFG; German Research Foundation) Project No. 239748522 SFB 1127, the Cluster of Excellence “Balance of the Microverse,” and also the Leibniz Award (to C.H.). Z.C. and M.T.F. were funded by the DFG Project No. 316213987 SFB 1278 (Z01). I.R. acknowledges financial support from the European Union Horizon 2020 Research and Innovation Program under the Marie Sklodowska-Curie grant agreement No. 794343. R.G. was funded by the International Leibniz Research School for Microbial and Biomolecular Interactions Jena.

Footnotes

  • Accepted August 5, 2021.
  • Author contributions: H.B., S.P.N., and C.H. designed research; H.B., S.P.N., K.V., Z.C., B.D., I.R., R.G., and S.J.P. performed research; H.B., S.P.N., K.V., Z.C., B.D., I.R., R.G., M.T.F., T.P.S., and S.J.P. analyzed data; and H.B., S.P.N., S.J.P., and C.H. wrote the paper.
  • The authors declare no competing interest.
  • This article is a PNAS Direct Submission.
  • This article contains supporting information online at https://www.pnas.org/lookup/suppl/doi:10.1073/pnas.2110669118/-/DCSupplemental.
  • Copyright © 2021 the Author(s). Published by PNAS.

This open access article is distributed under Creative Commons Attribution-NonCommercial-NoDerivatives License 4.0 (CC BY-NC-ND).

References

    1. N. Fierer
    , Embracing the unknown: Disentangling the complexities of the soil microbiome. Nat. Rev. Microbiol. 15, 579–590 (2017).CrossRefPubMedGoogle Scholar
    1. R. D. Bardgett, 
    2. W. H. van der Putten
    , Belowground biodiversity and ecosystem functioning. Nature 515, 505–511 (2014).CrossRefPubMedGoogle Scholar
    1. L. Tedersoo et al
    ., Fungal biogeography. Global diversity and geography of soil fungi. Science 346, 1256688 (2014).Abstract/FREE Full TextGoogle Scholar
    1. E. Ozimek et al
    ., Synthesis of indoleacetic acid, gibberellic acid and ACC-deaminase by Mortierella strains promote winter wheat seedlings growth under different conditions. Int. J. Mol. Sci. 19, 3218 (2018).CrossRefGoogle Scholar
    1. D. Zhou et al
    ., Deciphering microbial diversity associated with Fusarium wilt-diseased and disease-free banana rhizosphere soil. BMC Microbiol. 19, 161 (2019).Google Scholar
    1. J. Yuan et al
    ., Predicting disease occurrence with high accuracy based on soil macroecological patterns of Fusarium wilt. ISME J. 14, 2936–2950 (2020).Google Scholar
    1. D. Liu, 
    2. H. Sun, 
    3. H. Ma
    , Deciphering microbiome related to rusty roots of Panax ginseng and evaluation of antagonists against pathogenic Ilyonectria. Front. Microbiol. 10, 1350 (2019).Google Scholar
    1. S. Edgington, 
    2. E. Thompson, 
    3. D. Moore, 
    4. K. A. Hughes, 
    5. P. Bridge
    , Investigating the insecticidal potential of Geomyces (Myxotrichaceae: Helotiales) and Mortierella (Mortierellacea: Mortierellales) isolated from Antarctica. Springerplus 3, 289 (2014).Google Scholar
    1. E. Ozimek, 
    2. A. Hanaka
    Mortierella species as the plant growth-promoting fungi present in the agricultural soils. Agriculture 11, 7 (2021).Google Scholar
    1. L. Ellegaard-Jensen, 
    2. J. Aamand, 
    3. B. B. Kragelund, 
    4. A. H. Johnsen, 
    5. S. Rosendahl
    , Strains of the soil fungus Mortierella show different degradation potentials for the phenylurea herbicide diuron. Biodegradation 24, 765–774 (2013).Google Scholar
    1. J. Zeng et al
    ., Lignocellulosic biomass as a carbohydrate source for lipid production by Mortierella isabellina. Bioresour. Technol. 128, 385–391 (2013).Google Scholar
    1. F. Li et al
    ., Mortierella elongata‘s roles in organic agriculture and crop growth promotion in a mineral soil. Land Degrad. Dev. 29, 1642–1651 (2018).Google Scholar
    1. M. Künzler
    , How fungi defend themselves against microbial competitors and animal predators. PLoS Pathog. 14, e1007184 (2018).CrossRefGoogle Scholar
    1. J. H. J. Leveau, 
    2. G. M. Preston
    , Bacterial mycophagy: Definition and diagnosis of a unique bacterial-fungal interaction. New Phytol. 177, 859–876 (2008).CrossRefPubMedGoogle Scholar
    1. S. Zhang, 
    2. R. Mukherji, 
    3. S. Chowdhury, 
    4. L. Reimer, 
    5. P. Stallforth
    , Lipopeptide-mediated bacterial interaction enables cooperative predator defense. Proc. Natl. Acad. Sci. U.S.A. 118, e2013759118 (2021).Abstract/FREE Full TextGoogle Scholar
    1. T. Degenkolb, 
    2. A. Vilcinskas
    , Metabolites from nematophagous fungi and nematicidal natural products from fungi as an alternative for biological control. Part I: Metabolites from nematophagous ascomycetes. Appl. Microbiol. Biotechnol. 100, 3799–3812 (2016).CrossRefGoogle Scholar
    1. M. J. DiLegge, 
    2. D. K. Manter, 
    3. J. M. Vivanco
    , A novel approach to determine generalist nematophagous microbes reveals Mortierella globalpina as a new biocontrol agent against Meloidogyne spp. nematodes. Sci. Rep. 9, 7521 (2019).CrossRefGoogle Scholar
    1. O. Topalović, 
    2. M. Hussain, 
    3. H. Heuer
    , Plants and associated soil microbiota cooperatively suppress plant-parasitic nematodes. Front. Microbiol. 11, 313 (2020).CrossRefGoogle Scholar
    1. W. Qiu et al
    ., Organic fertilization assembles fungal communities of wheat rhizosphere soil and suppresses the population growth of Heterodera avenae in the field. Front. Plant Sci. 11, 1225 (2020).Google Scholar
    1. T. A. Al-Shammari, 
    2. A. H. Bahkali, 
    3. A. M. Elgorban, 
    4. M. T. El-Kahky, 
    5. B. A. Al-Sum
    , The use of Trichoderma longibrachiatum and Mortierella alpina against root-knot nematode, Meloidogyne javanica on tomato. J. Pure Appl. Microbiol. 7, 199–207 (2013).Google Scholar
    1. S. Meyer et al
    ., Activity of fungal culture filtrates against soybean cyst nematode and root-knot nematode egg hatch and juvenile motility. Nematology 6, 23–32 (2004).CrossRefGoogle Scholar
    1. M. K. Hasna, 
    2. V. Insunza, 
    3. J. Lagerlöf, 
    4. B. Rämert
    , Food attraction and population growth of fungivorous nematodes with different fungi. Ann. Appl. Biol. 151, 175–182 (2007).Google Scholar
    1. K. A. Dekker et al
    ., Novel lactone compounds from Mortierella verticillata that induce the human low density lipoprotein receptor gene: Fermentation, isolation, structural elucidation and biological activities. J. Antibiot. (Tokyo) 51, 14–20 (1998).PubMedGoogle Scholar
    1. N. Vandepol et al
    ., Resolving the Mortierellaceae phylogeny through synthesis of multi-gene phylogenetics and phylogenomics. Fungal Divers. 104, 267–289 (2020).Google Scholar
    1. M. R. Boyd et al
    ., Discovery of a novel antitumor benzolactone enamide class that selectively inhibits mammalian vacuolar-type (H+)-atpases. J. Pharmacol. Exp. Ther. 297, 114–120 (2001).Abstract/FREE Full TextGoogle Scholar
    1. M. Pérez-Sayáns, 
    2. J. M. Somoza-Martín, 
    3. F. Barros-Angueira, 
    4. J. M. Rey, 
    5. A. García-García
    , V-ATPase inhibitors and implication in cancer treatment. Cancer Treat. Rev. 35, 707–713 (2009).CrossRefPubMedGoogle Scholar
    1. S. P. Niehs et al
    ., Mining symbionts of a spider-transmitted fungus illuminates uncharted biosynthetic pathways to cytotoxic benzolactones. Angew. Chem. Int. Ed. Engl. 59, 7766–7771 (2020).Google Scholar
    1. Y. Hayakawa et al
    ., Oximidine III, a new antitumor antibiotic against transformed cells from Pseudomonas sp. II. Structure elucidation. J. Antibiot. (Tokyo) 56, 905–908 (2003).PubMedGoogle Scholar
    1. D. L. Galinis, 
    2. T. C. McKee, 
    3. L. K. Pannell, 
    4. J. H. Cardellina, 
    5. M. R. Boyd
    , Lobatamides A and B, novel cytotoxic macrolides from the tunicate Aplidium lobatum. J. Org. Chem. 62, 8968–8969 (1997).CrossRefGoogle Scholar
    1. R. Ueoka et al
    ., Genome mining of oxidation modules in trans-acyltransferase polyketide synthases reveals a culturable source for lobatamides. Angew. Chem. Int. Ed. Engl. 59, 7761–7765 (2020).Google Scholar
    1. Y. Sato et al
    ., Detection of betaproteobacteria inside the mycelium of the fungus Mortierella elongata. Microbes Environ. 25, 321–324 (2010).CrossRefPubMedGoogle Scholar
    1. Y. Takashima et al
    ., Prevalence and intra-family phylogenetic divergence of Burkholderiaceae-related endobacteria associated with species of Mortierella. Microbes Environ. 33, 417–427 (2018).Google Scholar
    1. S. Ohshima et al
    ., Mycoavidus cysteinexigens gen. nov., sp. nov., an endohyphal bacterium isolated from a soil isolate of the fungus Mortierella elongata. Int. J. Syst. Evol. Microbiol. 66, 2052–2057 (2016).CrossRefGoogle Scholar
    1. Y. Guo et al
    ., Mycoavidus sp. Strain B2-EB: Comparative genomics reveals minimal genomic features required by a cultivable Burkholderiaceae-related endofungal bacterium. Appl. Environ. Microbiol. 86, e01018-20 (2020).Abstract/FREE Full TextGoogle Scholar
    1. J. Uehling et al
    ., Comparative genomics of Mortierella elongata and its bacterial endosymbiont Mycoavidus cysteinexigens. Environ. Microbiol. 19, 2964–2983 (2017).CrossRefGoogle Scholar
    1. L. P. Partida-Martinez, 
    2. C. Hertweck
    , Pathogenic fungus harbours endosymbiotic bacteria for toxin production. Nature 437, 884–888 (2005).CrossRefPubMedGoogle Scholar
    1. G. Lackner, 
    2. N. Moebius, 
    3. C. Hertweck
    , Endofungal bacterium controls its host by an hrp type III secretion system. ISME J. 5, 252–261 (2011).CrossRefPubMedGoogle Scholar
    1. C. Jain, 
    2. L. M. Rodriguez-R, 
    3. A. M. Phillippy, 
    4. K. T. Konstantinidis, 
    5. S. Aluru
    , High throughput ANI analysis of 90K prokaryotic genomes reveals clear species boundaries. Nat. Commun. 9, 5114 (2018).CrossRefPubMedGoogle Scholar
    1. P. Yarza et al
    ., Uniting the classification of cultured and uncultured bacteria and archaea using 16S rRNA gene sequences. Nat. Rev. Microbiol. 12, 635–645 (2014).CrossRefPubMedGoogle Scholar
    1. K. Blin et al
    ., antiSMASH 5.0: Updates to the secondary metabolite genome mining pipeline. Nucleic Acids Res. 47 (W1), W81–W87 (2019).CrossRefPubMedGoogle Scholar
    1. K. Scherlach, 
    2. L. P. Partida-Martinez, 
    3. H. M. Dahse, 
    4. C. Hertweck
    , Antimitotic rhizoxin derivatives from a cultured bacterial endosymbiont of the rice pathogenic fungus Rhizopus microsporus. J. Am. Chem. Soc. 128, 11529–11536 (2006).CrossRefPubMedGoogle Scholar
    1. L. P. Partida-Martinez et al
    ., Rhizonin, the first mycotoxin isolated from the zygomycota, is not a fungal metabolite but is produced by bacterial endosymbionts. Appl. Environ. Microbiol. 73, 793–797 (2007).Abstract/FREE Full TextGoogle Scholar
    1. G. Lackner, 
    2. L. P. Partida-Martinez, 
    3. C. Hertweck
    , Endofungal bacteria as producers of mycotoxins. Trends Microbiol. 17, 570–576 (2009).CrossRefPubMedGoogle Scholar
    1. C. Almeida et al
    ., Unveiling concealed functions of endosymbiotic bacteria harbored in the ascomycete stachylidium bicolor. Appl. Environ. Microbiol. 84, e00660-18 (2018).Abstract/FREE Full TextGoogle Scholar
    1. K. Scherlach, 
    2. B. Busch, 
    3. G. Lackner, 
    4. U. Paszkowski, 
    5. C. Hertweck
    , Symbiotic cooperation in the biosynthesis of a phytotoxin. Angew. Chem. Int. Ed. Engl. 51, 9615–9618 (2012).CrossRefPubMedGoogle Scholar
    1. S. Geisen et al
    ., The soil food web revisited: Diverse and widespread mycophagous soil protists. Soil Biol. Biochem. 94, 10–18 (2016).Google Scholar
    1. J. van den Hoogen et al
    ., A global database of soil nematode abundance and functional group composition. Sci. Data 7, 103 (2020).Google Scholar
    1. M. P. Smith et al
    ., A liquid-based method for the assessment of bacterial pathogenicity using the nematode Caenorhabditis elegans. FEMS Microbiol. Lett. 210, 181–185 (2002).CrossRefPubMedGoogle Scholar
    1. E. J. Ragsdale, 
    2. J. Crum, 
    3. M. H. Ellisman, 
    4. J. G. Baldwin
    , Three-dimensional reconstruction of the stomatostylet and anterior epidermis in the nematode Aphelenchus avenae (Nematoda: Aphelenchidae) with implications for the evolution of plant parasitism. J. Morphol. 269, 1181–1196 (2008).CrossRefPubMedGoogle Scholar
    1. S. S. Schmieder et al
    ., Bidirectional propagation of signals and nutrients in fungal networks via specialized hyphae. Curr. Biol. 29, 217–228.e4 (2019).CrossRefGoogle Scholar
    1. G. W. Yeates, 
    2. T. Bongers, 
    3. R. G. De Goede, 
    4. D. W. Freckman, 
    5. S. S. Georgieva
    , Feeding habits in soil nematode families and genera-an outline for soil ecologists. J. Nematol. 25, 315–331 (1993).PubMedGoogle Scholar
    1. A. Tayyrov, 
    2. S. S. Schmieder, 
    3. S. Bleuler-Martinez, 
    4. D. F. Plaza, 
    5. M. Künzler
    , Toxicity of potential fungal defense proteins towards the fungivorous nematodes Aphelenchus avenae and Bursaphelenchus okinawaensis. Appl. Environ. Microbiol. 84, e02051-18 (2018).Abstract/FREE Full TextGoogle Scholar
    1. R. Hermenau et al
    ., Gramibactin is a bacterial siderophore with a diazeniumdiolate ligand system. Nat. Chem. Biol. 14, 841–843 (2018).CrossRefGoogle Scholar
    1. R. R. Wick, 
    2. L. M. Judd, 
    3. C. L. Gorrie, 
    4. K. E. Holt
    , Completing bacterial genome assemblies with multiplex MinION sequencing. Microb. Genom. 3, e000132 (2017).Google Scholar
    1. S. Blanton et al
    ., A web-based carepartner-integrated rehabilitation program for persons with stroke: Study protocol for a pilot randomized controlled trial. Pilot Feasibility Stud. 5, 58 (2019).Google Scholar
    1. S. Bleuler-Martínez et al
    ., A lectin-mediated resistance of higher fungi against predators and parasites. Mol. Ecol. 20, 3056–3070 (2011).CrossRefPubMedGoogle Scholar

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NEWS RELEASE 29-JUN-2021

DNA barcodes decode the world of soil nematodes

To understand soil ecosystems and contribute to advanced agriculture

TOYOHASHI UNIVERSITY OF TECHNOLOGY (TUT)

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IMAGE: SOIL SAMPLING SITES (TOP). CLASSIFICATION OF SOIL NEMATODE COMMUNITIES BY FEEDING GROUP (RESULTS FOR BARCODE REGION 4) (BOTTOM). view more CREDIT: COPYRIGHT (C) TOYOHASHI UNIVERSITY OF TECHNOLOGY. ALL RIGHTS RESERVED.

Overview

The research team of Professor Toshihiko Eki of the Department of Applied Chemistry and Life Science (and Research Center for Agrotechnology and Biotechnology), Toyohashi University of Technology used a next-generation sequencer to develop a highly efficient method to analyze soil nematodes by using the 18S ribosomal RNA gene regions as DNA barcodes. They successfully used this method to reveal characteristics of nematode communities that inhabit fields, copses, and home gardens. In the future, the target will be expanded to cover all soil-dwelling organisms in agricultural soils, etc., to allow investigations into a soil’s environment and bio-diversity. This is expected to contribute to advanced agriculture.

Details

Similar to when the UN declared 2015 to be the International Year of Soils, there have recently been many efforts worldwide to raise awareness of the importance of the soil that covers our Earth and its conservation. Diverse groups of organisms such as bacteria, fungi, protists, and small soil animals inhabit the soil, and together they form the soil ecosystem. Nematodes are a representative soil animal; they are a few millimeters long and have a shape resembling a worm. They play an important role in the cycling of soil materials. Many soil nematodes are bacteria feeders, but they have a wide variety of feeding habits, such as feeding on fungi, plant parasitism, or being omnivorous. In particular, plant parasitic nematodes often cause devastating damage to crops. Therefore, the classification and identification of nematodes is also important from an agricultural standpoint. However, nematodes are diverse, and there are over 30,000 species. Additionally, because nematodes resemble one another, morphological identification of nematodes is difficult for anyone but experts.

The research team focused on “DNA barcoding” to identify the species based on their unique nucleotide sequences of a barcode gene, and they established a method using a next-generation sequencer that can decode huge numbers of nucleotide sequences. They used this to analyze nematode communities from different soil environments. Initially, four DNA barcode regions were set for the 18S ribosomal RNA genes shared by eukaryotes. The soil nematodes used for analysis were isolated from an uncultivated field, a copse, and a home garden growing zucchini. The PCR was used to amplify the four gene fragments from the DNA of the nematodes and determine the nucleotide sequences. Additionally, the nematode-derived sequence variants (SVs) representing independent nematode species were identified, and after taxonomical classification and analysis of the SVs, it was revealed that plant parasitizing nematodes were abundant in the copse soil and bacteria feeders were abundant in the soil from the home garden. It was also determined that predatory nematodes and omnivorous nematodes were abundant in the uncultivated field, in addition to bacteria feeders.

This DNA barcoding method using a next-generation sequencer is widely used for the analysis of intestinal microbiota, etc., but analyses of eukaryotes such as nematodes are still in the research stage. This research provides an example of its usefulness for the taxonomic profiling of soil nematodes.

Development Background

Research team leader Toshihiko Eki stated, “Through genetic research, I have been working with nematodes (mainly C. elegans) for around 20 years. As a member of our university’s Research Center for Agrotechnology and Biotechnology, I came up with this theme while considering research that we could perform that is related to agriculture. As a test, we isolated nematodes from the university’s soybean field and unmanaged flowerbed and analyzed the DNA barcode for each nematode. Bacteria feeders were abundant in the soybean field, and that was used for comparison with the flowerbed, where weed-parasitizing nematodes and their predator nematodes were abundant. This discovery was the start of our research (Morise et al., PLoS ONE, 2012). If that method using one-by-one DNA sequencing was the first generation, the current method using the next-generation sequencer is the second generation, and we were able to clarify characteristics of nematode communities representing the three ecologically different soil environments according to expectations.”

Future Outlook

Currently, the research team is developing the third-generation DNA barcoding method which involves purifying DNA directly from the soil and analyzing the organisms in the whole soil instead of isolating and analyzing any particular soil-dwelling organisms. They are currently analyzing the soil biota of cabbage fields, etc. They are aiming to precisely analyze how communities of soil-dwelling organisms including microbes change with crop growth, clarify the effects that cultivated plants have on these organisms, and investigate biota closely related to plant diseases. If this research moves forward, crops can be cultivated and managed logically based on biological data in agricultural soils, and it can contribute to advancing smart agriculture in Japan, such as in the prominent Higashi-Mikawa agriculture region and beyond.

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This research was performed with the support of the Takahashi Industrial and Economic Research Foundation.

Reference

Harutaro Kenmotsu, Masahiro Ishikawa, Tomokazu Nitta, Yuu Hirose and Toshihiko Eki (2021). Distinct community structures of soil nematodes from three ecologically different sites revealed by high-throughput amplicon sequencing of four 18S ribosomal RNA gene regions.
PLoS ONE, 16(4): e0249571.

Disclaimer: AAAS and EurekAlert! are not responsible for the accuracy of news releases posted to EurekAlert! by contributing institutions or for the use of any information through the EurekAlert system.

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Nigeria can attain sustainable food security using biological pesticides – Don

ByNaija247news Media, New YorkJune 23, 2021 010 Share

ood Security

June 24, 2021 

June 24, 2021 Naija247news Media, New Yorkhttps://www.naija247news.com/Naija247news is an investigative news platform that tracks news on Nigerian Economy, Business, Politics, Financial and Africa and Global Economy.

By Akeem Abas
Ibadan, June 18, 2021 A Professor of Nematology, Prof. Timothy Olabiyi, says sustainable food security can be attained through the use of biological pesticides.
Olabiyi disclosed this on Friday while delivering the 45th inaugural lecture of Ladoke Akintola University of Technology (LAUTECH), Ogbomoso.
He said that the only sustainable food security measure is the use of non-synthetic chemicals.
He noted that the world population is increasing on daily basis and many have suffered ill-health as a result of food poison and toxin which are also increasing.
“Sustainable food security can only be attained through the use of biological pesticides that are ecologically friendly and bio-degradable with no chemical residues leading to safe-to-eat food.
“Mass production of biological pesticides and making it available to farmers is germane to disease management and sustainable food security in Nigeria,” he said.
The university don called on Federal Government to establish Biological Pesticide Production Industries that could produce adequate and required biological pesticides.

“Such industries will provide jobs for the youth aside from the fact that the teaming population will have the right to eat safe food,” he said.

He said that the presence of farmer’s hidden enemy is inevitable, adding that they are present everywhere and all year round.

Olabiyi said that sustainable management of these disease-causing micro-organisms is our best option.

According to him, “my target is to produce biological nematicide for farmers in Nigeria.

“I am almost at the point of patenting those products, so that farmers can get to the shop and buy it for their farms.
“At the moment, I have supplied so many farmers nationwide, even in the North East and West.

“They have used it effectively and have given me very good report that it is good and we can use it to replace the synthetic nematicide,” he said.

He called for support to set up an industry for the production of biological nematicide for farmers in Nigeria.
A nematicide is a type of chemical pesticide used to kill plant-parasitic nematodes.

The varsity don said that the issue of farmers-herders’ clashes was due to climate change, saying he could not blame either of the parties.

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Expert calls for healthy food cultivation in Nigeria

By Chidinma Ewunonu-Aluko Ibadan, Oct. 16, 2020 Dr Abayomi Olaniyan, Executive Director, National Horticultural Research Institute, Ibadan, says it is imperative for the country to increase agricultural productivity by cultivating healthy food that is diverse in nature. Olaniyan made the remark in an interview with newsmen on Friday in Ibadan…October 16, 2020

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NEMEDUSSA CONSORTIUM ADVANCING NEMATOLOGY EDUCATION IN SUB-SAHARA AFRICA

To develop the research and educational capacity in Sub-Sahara Africa in the field of nematology, or the study of roundworms, a joint Erasmus+ KA2 project was recently launched. The Erasmus+ project, Capacity Building in Higher Education (CBHE): Nematology Education in Sub-Sahara Africa (NEMEDUSSA), is a joint effort by a consortium of Universities from Sub-Sahara Africa and Europe.

This three-year project (2021-2023) is co-funded by the European Union (Erasmus+ KA2 CBHE) and VLIR-UOS, and is linked to the objectives of the Erasmus+ Programme. The aims are to encourage cooperation between the EU and Partner Countries and support eligible Partner Countries in addressing challenges in the management and governance of their higher education institutions.

Specifically, NEMEDUSSA aims to increase awareness of nematodes and expand educational and research capacities in higher education and other institutions in Sub-Sahara Africa in this field. Nematodes or roundworms cause significant damage and yield loss to a wide variety of crops often together with other pathogens. Unfortunately, nematodes are often overlooked or misdiagnosed, resulting in the unnecessary use of unhealthy agro-chemicals. Nematodes can also be used as bio-control agents against insect pests and/or as bio-control agents for environmental health and biodiversity.

Despite the profound adverse impact plant-parasitic nematodes have on productivity worldwide, it is striking how concealed the discipline of nematology has remained, particularly in Sub-Sahara Africa. This project aims to address the need for increased capacity and specialised training in handling these pathogens, so that plant-parasitic nematodes are managed correctly and beneficial nematodes can be implemented as biocontrol organisms.

To achieve this, the project focuses on 6 core activities:

  1. Developing Curricula. Develop curricula in nematology on BSc and MSc level for the integration into existing educational programmes in English and French, for both lecturers and students.
  2. Training Staff. Improve the nematological expertise of academic and technical staff to enhance teaching capacity.
  3. Upgrading lab facilities. Increase the number of student microscopes, lab and demonstration equipment to augment hands-on training.
  4. Nematology digital learning platform. Develop an open-access platform to share and disseminate nematological knowledge, develop curricular modules, knowledge clips, etc.
  5. Nematology Network. Enhance cooperation between nematologists in Sub-Sahara Africa by providing networking tools, workshops on relevant topics in nematology and sharing good practices in education, promoting collaboration with a focus on young nematologists.
  6. Creating awareness. Facilitate dissemination activities and involve a range of different stakeholders such as farmers, extension service workers, policy makers, students and private and public sector.

Ghent University (Belgium) coordinates NEMEDUSSA, in cooperation with:

  • University Abomey-Calavi, Benin
  • University of Parakou, Benin
  • Haramaya University, Ethiopia
  • Jimma University, Ethiopia
  • Kenyatta University, Kenya
  • Moi University, Kenya
  • Ahmadu-Bello University, Nigeria
  • University of Ibadan, Nigeria
  • North West University, South Africa
  • Stellenbosch University, South Africa
  • Makerere University, Uganda
  • Muni University, Uganda
  • University Côte d’Azur, France

The work of this project is further supported by 36 associated partners from the private and public sectors in Sub-Sahara Africa.

For more information about the NEMEDUSSA project, please see www.nemedussa.ugent.be or contact us at nemedussa@ugent.be.  

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fresh plaza logo

WSU researchers combat parasitic worm

Plants that fight back

So small it can’t be seen with the naked eye, a parasitic worm called the root-knot nematode causes mammoth problems for Northwest farmers. But potatoes, grapes and other crops could gain a new, nature-based way to fight back, thanks to Cynthia Gleason and Jennifer Watts, scientists at WSU.

Notorious thieves
Nematodes cause billions of dollars in crop losses nationwide every year. In Washington, they cause significant losses to crops such grapes, onions, garlic and the state’s $734 million potato industry.

“Root-knot nematodes are a huge problem for farmers,” said Gleason, plant pathologist with WSU’s College of Agricultural, Human and Natural Resource Sciences (CAHNRS). The soilborne parasites move into the roots of crops, “then just sit there and feed on the plant. They’re stealing nutrients and water.”

Nematodes don’t kill the plants, but they leave them stunted, wilted from lack of water, and more susceptible to other pathogens, ultimately reducing farmers’ yields.

Chemical-free weapon pursued
“Plants don’t have many natural resistances to root-knot nematodes, so we need a way to combat them,” Gleason said. Traditionally, farmers have used anti-nematode pesticides -nematicides- to eliminate the tiny worms.

“But there aren’t many chemical options left, and they’re very expensive,” she said. “I’m looking for new, chemical-free controls that help growers move on.”

Acid stops nematodes
To help, Gleason is using a new $47,400 Emerging Research Issues grant from the CAHNRS Office of Research to seek genetic defenses that help crops like potatoes and tomatoes fight back against the persistent pest. “I’m developing plants that are basically toxic to nematodes,” she said.

Jennifer Watts, researcher in the School of Molecular Biosciences, discovered that a dietary fatty acid stops parasites from multiplying. Partnering with Jennifer Watts, researcher in the College of Veterinary Medicine’s School of Molecular Biosciences, Gleason is adding genes that tell plants to secrete a specific fatty acid that stops the nematode reproductive cycle.

Watts and her team of student researchers discovered that a certain fatty acid, referred to as DGLA (20:3n-6), stops egg production in a cousin species of the root-knot nematode.

“These fatty acids aren’t normally produced in plant tissue,” says Watts. “My team and I are working with Cynthia to introduce genes into plants so they can make them. If it works, it could be a new, chemical-free method to control nematodes.”

While the fatty acid is not known to be toxic to people or animals at low levels, the researchers plan to only express it in cover crops and plant tissues that aren’t normally eaten.

Future pest fighters
Farmers could one day plant a seed, Gleason said, that grows into a cover or cash crop with its own natural pest control. As nematodes feed on the plants, their populations will fall — leading to healthier plants, bigger crops and an improved food supply.

“Usually, the study of nematodes and the challenges they bring is about new chemicals and pesticide controls,” said Gleason. “This is a new and different approach, one that’s chemical free.

“By working across colleges, mine and Jennifer’s teams are discovering and accomplishing much more than we could individually,” she added. “We can use that information to fight parasites, help Washington farmers, and grow more food. It’s a collaboration that benefits everyone.”

For more information:

Washington State University
Cynthia Gleason
Department of Plant Pathology
Tel.: +1 509-335-3742
Jennifer Watts
School of Molecular Biosciences
Tel.: +1 509-335-8554

 

Publication date: 6/21/2018

 

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CropLife

If you’re an ag retailer and you’re reading this report, I’d guess it’s fair to assume you’ve at least considered adding biopesticide products to your crop protection lineup.

My assumption is part idealism, part result of our 2017 CropLife® Biological Product Market Survey, which was sent to 29,000 ag retailers and other industry members nationwide. In the survey 67% of respondents said they plan to “increase the percentage of biological products” they sell/distribute in the future. Additionally, nearly half (49%) affirmed that their customers apply biologicals as “both seed treatments and topicals.”

Advanced Biological Marketing (ABM) is one such company finding success with seed-applied biological products. Dan Custis, CEO of the Van Wert, OH-based company, has been involved in the biologicals segment of the industry for almost 18 years now. He says that when the company first started marketing biologicals back in 2000 there was “very little adoption at all. Very little.”

“A lot of the types of products that we manufacture were referred to as kind of a bathtub mixture, or ‘Foo-foo Dust’,” Custis fondly recalls. “As we really got into it, we as a company put a lot of science and knowledge behind it.”

Ah yes, another aspect of biological products addressed in the survey. By far the top consensus among those surveyed was that biological products engender a “lack of trust around product performance” while a sizeable 72% of retailers responded that biopesticide products need “more research that demonstrates product effectiveness.”

At ABM, Custis says the company has research that shows about a seven bushel-per-acre yield increase over a five-year average on corn, and in soybeans that number is around two-and-a-half bushels per acre. Its top biopesticide, the seed-applied SabrEx (two strains of Trichoderma) is typically either applied downstream at the retailer, or on-farm by the grower. The company does work with some seed manufacturers as well, such as local Ohio seed company Rupp.

“We know that maybe we get six weeks of benefit at most from a chemical seed treatment depending on weather, unless it’s a systemic,” Custis says. “What biologicals bring to the table is the extension of that plant health beyond the six weeks. Biologicals are a living organism, they should be able to live on the root system of that plant up through flowering.”

ABM’s SabrEx is distributed via the traditional crop input retail channels, through well-known players such as Crop Production Services, WinField Uni­ted, Wil­bur-Ellis, and KOVA of Ohio. Production and formulation take place in Van Wert, while research & development is housed in the Finger Lakes region of Western New York in Geneva.

“Right now in R&D we’re taking a look at nematode control in soybeans and corn, that’s one of the products that we have committed to EPA for approval right now,” Custis shares. “That (product) would be a first, and we’ve certainly got other things in the pipeline that I’m not able to talk about at the moment.”

Where do others see the biopesticide industry headed in the next couple years? Again, we consult our survey responses, and with nearly three-fourths (72%) saying their customers prefer to apply biologicals not as one-off standalones, but actually in conjunction with conventional products. Well-known Iowa State University seed treatment expert Allison Robertson agrees.

“There has been quite a lot of work looking at biologicals, not as stand-alone treatments, but in partnership with treatments that address pathogens in the field,” she shared back in August. “In addition, nematicides have been developed recently to help fight off soybean nematodes.”

Which provides a perfect segue to discuss post-patent giant Albaugh and its intriguing BIOst system, which Director of Global Proprietary Products Chad Shelton describes as “the first complete biological seed treatment platform.”

“What’s really exciting for retailers,” he continues. “Is our BIOst 100 nematicide, which can be combined with synthetic chemistries to give both insect and nematode protection. This is the first biological nematicide registered for control of both soil dwelling pests, along with activity on nematodes. And when we combine that with a neonic seed treatment it’s giving the grower a better return-on-investment (ROI).”

That’s a trend Shelton is seeing play out more and more in the row crop biologicals space in the last couple years, shifting the deployment of biopesticides from one-off products to more integrated usage with conventional hard chemistries.

“It’s no longer about having one mode of action, or a specific agronomic response in the marketplace. To me that’s the biggest change,” he shares. “When you have biopesticides in combination with synthetics at a reduced rate you’re going to get enhanced performance plus ROI.”

Another area that Albaugh is focusing attention is developing products with what Shelton describes as “customization based on microclimate.”

“Our goal today is to customize seed treatment technologies based on micro climate and (regional) needs,” he adds.

 

 

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