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Nematode-free agricultural system of a fungus-growing termite

Abstract

Fungus-growing termites forage dead plant materials from the field to cultivate symbiotic Termitomyces fungi in the nest. Termite foraging behavior and the entry of symbiotic arthropod inquilines may transfer nematodes into a nest and adversely affect fungus production. To test whether nematodes were transferred to fungus gardens by termites and inquilines, we examined the occurrence of nematodes in fungus gardens, five termite castes, and nine species of inquilines of a fungus-growing termite, Odontotermes formosanus. Our results revealed that nematodes were commonly carried by foraging termites and beetle inquilines. Numerous nematodes were found under the beetle elytra. No nematodes were found on termite larvae, eggs, and wingless inquilines. In addition, nematodes rarely occurred in the fungus garden. By observing the response of nematodes to three species of Termitomyces spp. and the fungus gardens, we confirmed that the fungus and fungus gardens are not actually toxic to nematodes. We suggest that nematodes were suppressed through grooming behavior and gut antimicrobial activity in termites, rather than through the antimicrobial activity of the fungus.

Introduction

Insect agricultural systems

Agricultural systems are not only reported in humans but also in insects1,2. For example, attine ants (Hymenoptera: Formicidae) cultivate a wide range of fungal species3,4. Fungus-growing termites (Blattodea: Termitidae) cultivate fungal species of the genus Termitomyces (Agaricales: Lyophyllaceae)5,6. In addition, ambrosia beetles are wood-boring weevils that cultivate symbiont fungi in their tunnels as sources of nutrition, and such farming behavior has independently evolved more than eleven times in weevils7,8. Symbiosis between fungi and farming insects is obligate, and farming insects have evolved specific agricultural behaviors that are comparable to those of humans, such as seeding, cultivation, and harvesting of fungal products for food1. For instance, workers of fungus-growing termites forage outside the nest and bring plant materials into the nest. Incoming materials are chewed up by young workers, ingested, passed rapidly through their gut with little digestion, and then deposited as feces on the top rim of the fungus garden. At the same time, vegetative fungus spores are ingested from conidiophores (=vegetative fruiting bodies) produced on slightly older fungus garden, to inoculate the newly-deposited materials2,9,10. The termites then harvest aged substrates and fungal nodules growing on the surfaces of fungus gardens for food11. The symbiont fungi are a major source of nutrition for farming insects since the fungi hold nutrients critical for their growth, including essential vitamins, amino acids, and sterols2,12,13,14.

Nematodes as potential pests in agricultural systems of termites

Nematodes are found in various terrestrial environments15 and are widely regarded as agricultural pests. For example, plant-parasitic nematodes cause severe losses to crop production and are vectors of plant pathogens16,17. Nematodes are also considered as pests in both human18,19 and insect20 fungicultural systems. Termites have commonly been reported to carry nematodes, mostly microbe-feeders in the phoretic stage21,22,23,24,25,26,27,28,29. In addition, nematode species associated with termites differ from those randomly sampled from soil or epiphytic nematodes, which implies that termite-associated nematodes are transferred by termites and potentially other soil invertebrates as well, and inhabit the carriers’ habitats and/or nests30,31. Invasion of termite nests by nematodes is possible and may lead to production losses through the consumption of fungi and fungus gardens. Fungus gardens of termites are rich in carbohydrates, proteins, and lipids10,32. Fungus gardens are not only media for growing Termitomyces, the symbiont fungus, but also media for growing other fungal species, such as Xylaria fungi (Xylariales: Xylariaceae)33, and are potential media for the growth of bacteria and nematodes.

Transmission and management of pests in insect agricultural systems

Microbes that are pathogenic to fungi are potential pests in agricultural systems and are potentially transferred to fungus gardens when (1) insect foragers carry materials with microbes from extranidal environments and contaminate the cultivating substrates and fungus; (2) numerous “guests” (inquilines), such as myrmecophiles, associated with ants, and termitophiles, associated with termites34,35,36,37 enter nests of social insects through chemical mimicry or chemical insignificance38,39, and may carry pathogenic microbes into the agricultural system; and (3) pathogens are vertically transmitted through colonizers, such as the alates of ants and termites, carrying pathogens from parental colonies1 to newly founded colonies.

Pest management strategies have also been reported in agricultural systems of insects. For example, in fungus gardens of leaf-cutter ants, the parasitic microfungus Escovopsis (Hypocreales: Hypocreaceae) are suppressed by antimicrobial chemicals produced by actinomycete bacteria, a symbiont of leaf-cutter ants40,41,42. Um, et al.43 also observed that a strain of Bacillus in fungus gardens of termites suppressed non-Termitomyces microbes. Allogrooming behavior among nest mates, which has been reported in termites and ants, may remove pathogens from body surfaces2,44,45. In termites, partitioning of foraging and nest-caring tasks among individuals has been suggested to inhibit the transfer of pathogens in colony46. Fungus-growing termites were also hypothesized to suppress non-Termitomyces microbes by passing fungus garden materials through their guts, which generally have high levels of antimicrobial activity47.

Purpose of this study

We aimed to understand the transmission and management of nematodes in an agricultural system of a fungus-growing termite, Odontotermes formosanus (Shiraki). We investigated three transmission pathways of nematodes in the agricultural system: (1) vertical transmission via alate, (2) horizontal transmission via foragers, and (3) horizontal transmission via inquilines. In addition, we assessed the potential of fungus gardens as media for nematodes. We also examined factors that may suppress nematode populations in termite nests.

Materials and Methods

Hosts of nematodes

Five nests of Odontotermes formosanus from different localities in Taiwan were located by searching for fruiting bodies (mushrooms) of Termitomyces on the ground and were excavated to investigate their potential as hosts of nematodes in nests, including fungus gardens, termites, and inquilines (Figs 1 and 2; Supplementary Information, Table S1); 3–7 fungus gardens were collected from each nest. Since the compositions of microbes may vary between the upper and the lower parts of fungus gardens47, 6 g from the upper and lower parts were collected separately (Supplementary Information, Table S2) to investigate the presence of nematodes. The upper parts of fungus gardens are dark-colored fresh substrates, comprising partially-digested plant material. By contrast, the lower parts are whitish aged substrates, comprising dense fungal hyphae and highly decomposed plant materials (Fig. 1e)48. Between the dark-colored fresh substrates and the whitish old substrates is the active zone in which the Termitomyces conidiophores are mainly produced. The upper and lower parts were sampled separately by dividing the active zone at the middle, and therefore the relatively fresh and aged active zone will be sampled and included in the upper and lower parts, respectively.

Figure 1
figure1

Major components of Odontotermes formosanus agriculture. (a) Swarming alates; (b) minor and major workers; (c) termite larvae and egg mass; (d) a fungus comb in situ in the soil, part of a large nest of O. formosanus; (e) an O. formosanus fungus garden (note that the dark upper layer is comprised of fresh plant material and the pale lower layer is aged and decomposed material).

Figure 2
figure2

Inquilines of Odontotermes formosanus. (a) Cycloxenus sp. (Coleoptera: Cerylonidae), adult in dorsal view; (b) Cycloxenus sp., adult in ventral view; (c) larva of Cycloxenus sp.; (d) Ziaelas formosanus (Coleoptera: Tenebrionidae); (e) Clitelloxenia audreyae (Diptera: Phoridae); (f) Clitelloxenia formosana (Diptera: Phoridae); (g) Selenophora shimaidai (Diptera: Phoridae); (h) Pseudotermitoxenia nitobei (Diptera: Phoridae); (i) Horologiphora sinensis (Diptera: Phoridae); (j) Platystylea sp. (Thysanura: Nicoletiidae); (k) millipede (Diplopoda: Polydesmida: Pyrgodesmidae).

Termites and inquilines in each nest were sought and collected exhaustively. To investigate whether the foraging termites carry nematodes, foraging termites on feeding sites (logs, leaves, tree bark) found within 20 m of nests were collected. Feeding sites of O. formosanus were distinguished by the presence of soil-sheeting, which is specifically built by foragers of O. formosanus. Inquilines in O. formosanus nests were identified based on the morphological descriptions of termitophilous insects in Taiwan49 (Fig. 2). To examine whether inquilines carry nematodes in the dispersal stage, dispersing adults of a termitophilous beetle, Ziaelas formosanus Hozawa (Coleoptera: Tenebrionidae) (Fig. 2d) were collected using a light trap at Xiaping Tropical Botanical Garden, the Experimental Forest, National Taiwan University, Nantou, Taiwan (23.77°N, 120.67°E). Ziaelas formosanus is commonly observed in fungus gardens of O. formosanus. Dispersing termite alates (Fig. 1a) were collected from four localities (Supplementary Information, Table S3) to examine whether nematodes were transferred from parental colonies. Voucher specimens of nematodes, termites, and inquilines were deposited at National Chung Hsing University (NCHU).

Isolation of nematodes

Fungus gardens, eggs, larvae, major workers, minor workers, and alates of termites were deposited on 2% agar plates (ø = 90 mm)29, squashed with tweezers, and observed using a Leica® M205 C stereomicroscope or Leica® DM750 microscope (Leica, Wetzlar, Germany) to inspect the presence of nematodes. To clearly observe nematodes on agar plates, termite eggs were evenly spread on agar plates. Head capsules and digestive systems of termite larvae, workers, and alates were further dissected. To avoid high densities of body parts interfering with the observation of nematodes, the number of larvae and major and minor workers on each agar plate was limited to 50 individuals, and that of termite alates was limited to 40 individuals (20 males and 20 females). Nematodes of inquilines were examined using the same methodology applied to termites. The number of inquilines on each agar plate was approximately 30 individuals. All inquilines were examined on a single agar plate if the nest contained less than 30 individuals. The fungus gardens, eggs, dissected bodies of termites, and inquilines were kept on agar plates for one month at room temperature to allow nematodes to propagate for inspection. The percentage of nematodes present in fungus gardens, termite castes, and inquilines was calculated by dividing the number of plates with nematodes with the total number of plates examined and multiplying by 100.

Morphotyping and molecular characterization of nematodes

When nematode propagation was confirmed on agar plates, the nematodes were removed manually, their morphologies assessed, and were transferred to nematode digestion buffer50,51 for molecular identification. However, if the number of individuals was sufficient to establish cultures, they were transferred to artificial media (i.e., nematode growth medium inoculated with OP50 Escherichia coli strain for bacteria feeders or 2% malt extract agar inoculated with Botrytis cinerea Pers. for fungal feeders). Cultured nematodes were kept as laboratory strains and the detailed taxonomic studies are presented elsewhere.

Since all nematodes collected during dissection were dauer (dispersal) juveniles that do not have genus/species-specific morphological characters, they were transferred to nematode digestion buffer and molecularly characterized based on ribosomal RNA sequences, that is, near full-length 18S (SSU) and/or D2-D3 expansion segments of 28S (D2-D3 LSU) regions. Molecular sequences were determined through PCR-based direct sequencing according to Kanzaki and Futai52 and Ye, et al.53. Generated sequences were compared with those deposited in the GenBank database (http://www.genome.jp/dbget-bin/www_bfind?genbank-today) using the BLAST homology search program (http://blast.ddbj.nig.ac.jp/blastn?lang=en).

Fungus toxicity bioassay

To confirm whether the fungal symbiont Termitomyces spp. suppressed nematode populations, three Termitomyces spp. (Supplementary Information, Table S4) were isolated and cultured on Potato Dextrose Agar (PDA) to examine their nematocidal activity against four nematode genera isolated from termites, fungus gardens, or inquilines. Termitomyces spp. strains and their identification based on the internal transcribed spacer (ITS) region were provided by Mycology Laboratory of NCHU (Supplementary Information). Fungus cultures were inoculated with five individuals of one species from each nematode genus. To test whether nematocidal activity was present on fresh and aged substrates of fungus gardens, three fungus gardens were collected from NCHU and separated into fresh and aged parts. Six grams of fresh or aged substrates were vortexed for homogenization and each of the 0.3-g substrates placed on different 2% agar plates (ø = 50 mm). Each plate was inoculated with 10 individuals of a single nematode species. Locomotion and pharyngeal pumping in the nematodes were examined at 1, 24, or 48 hours after exposure to fungal hyphae to observe if the fungal culture would exhibit any toxicity to the nematodes.

Results

Nematodes isolated

Seven genotypes of nematodes were recognized from the culture plates, namely, a fungal feeder, Aphelenchoides sp. and 6 bacteria feeders, comprising three types of Acrostichus sp. (Type A, B, and C), 2 types of Halicephalobus spp. (Type A, B), and a Diplogastrellus sp. The molecular sequences, BLAST results, and notes on each genotype determined in the present study are presented in Supplementary Information.

No nematodes were observed when dissecting 21 fresh and 21 aged substrates of fungus gardens (Table S2). A few Halicephalobus sp. (type B) were observed in only one plate 3 weeks after inoculation of the fresh substrate from Xiaping (1 plate detected/21 plates cultured, ~4% occurrence) (Table S2). No nematode was observed in the plates hosting the aged substrates (0/21 plates, 0%) (Table S2). The results indicate that termite fungiculture systems were almost nematode free.

No nematodes were isolated from the 160 swarming alates (80 males and 80 females) of O. formosanus from the four localities that were examined (Tables S3 and S5) (0/4 plates, 0%); the results indicated that nematodes were not transferred from parental to incipient colonies. Therefore, within-colony transmission of nematodes via alates was unlikely.

Nematodes were not found when dissecting the termite workers collected in or outside the nests. After culturing on plates, four nematode genotypes were identified from foraging major workers (Supplementary Information, Table S5): an Aphelenchoides sp., a Diplogastrellus sp., and two genotypes of Halicephalobus spp. (type A and B) (3/6 plates, 50%) (Table S5). The major workers collected in the nest hosted three nematode genotypes, namely Diplogastrellus sp., Halicephalobus spp. (type A and B) (4/7 plates, 57%) (Table S5). Minor workers in nests hosted two nematode genotypes (Aphelenchoides sp. and Diplogastrellus sp.) (1/5 plates, 25%) (Table S5). No nematodes were detected in termite eggs (0/6 plates, 0%) or larvae (0/13 plates, 0%) (Table S5). The results suggest that nematodes are largely transferred to nests by major workers, which forage and bring plant materials to nests. Therefore, it is likely that horizontal transmission of nematodes via foragers occurred frequently.

Nematodes were examined in nine inquiline species previously described by Liang and Li49: two coleopteran inquilines, Cycloxenus sp. (Cerylonidae) (Fig. 2a–c) and Ziaelas formosanus (Tenebrionidae) (Fig. 2d); five dipteran inquilines of the family Phoridae, Clitelloxenia audreyae (Fig. 2e), C. formosana (Fig. 2f), Selenophora shimaidai (Fig. 2g), Pseudotermitoxenia nitobei (Fig. 2h), and Horologiphora sinensis (Fig. 2i); one thysanuran inquiline, Platystylea sp. (Thysanura: Nicoletiidae) (Fig. 2j), and a newly recorded species of termitophilous millipede (Diplopoda: Polydesmida) (Fig. 2k) (Table S6). The sample sizes of each of the species or developmental stages of inquilines that were investigated are listed in Table S6. Three nematode genotypes (bacteriophagous Acrostichus sp. type A, B, and C) were isolated from adults of Z. formosanus and Cycloxenus sp. (Table S6), and no nematodes were observed in plates hosting larvae of Cycloxenus sp. and those of the other inquilines (Table S6). Most nematodes isolated from Z. formosanus adults were dauer juveniles, inhabiting the section beneath the elytra (Fig. 3). Based on the results, nematodes were transferred by coleopteran inquilines of termites. However, nematodes cultured from coleopteran inquilines and from termites were different.

Figure 3
figure3

Distribution of nematodes on the body of Ziaelas formosanus. (a) The habitus of Ziaelas formosanus; (b) nematodes on the ventral side of the beetle’s elytra; (c) nematodes on the beetle’s hindwing.

General information on presence of nematodes on termites, inquilines, and fungus gardens is summarized in Fig. 4. The results revealed that although nematodes were transferred to fungus gardens, the populations were suppressed before or after they entered fungus gardens.

Figure 4
figure4

Occurrence of nematodes in Odontotermes formosanus termite society and its inquilines. Encircled abbreviations: H, nematode genus Halicephalobus; D, nematode genus Diplogastrellus; Ap, nematode genus Aphelenchoides; Ac, nematode genus Acrostichus. Yellow circles mean nematodes are present; white circles represent absence of nematodes. Numbers of nematode species and their occurrence rates are denoted for each component of the termite society. Vertical transmission through reproductive castes (4a) to egg or larvae (4e) was not observed. Different nematode species found on termites (4bcd) and inquilines (4hij) indicates that nematodes could enter termite nest through two exclusive horizontal transmission pathways. Extremely low occurrence of nematodes on vulnerable fungus gardens (4fg) indicates termite hygienic behavior plays a critical role in nematode control in the nest.

Fungus toxicity bioassays

Five isolated nematode genotypes, including Acrostichus sp., Aphelenchoides sp., Diplogastrellus sp., and Halicepholobus spp., were successfully cultured, (Table S7). No nematodes died or exhibited repellent responses to Termitomyces or fungus gardens within 3 hours of inoculation (0/24 plates, 0%). In addition, a fungus-feeding nematode, Aphelenchoides sp., was repeatedly observed boring into the fungus garden (Supp. Video 1. https://youtu.be/BIobuqNaw60). The results suggested Termitomyces and fungus gardens are not likely to suppress nematodes by themselves.

Discussion

Transmission and management of nematodes

The results of the present study revealed that nematodes commonly enter termite nests via two sources: (1) termite foraging workers and (2) coleopteran inquilines. However, being a potential pest, nematodes were not transferred to other castes after entering nests, considering the absence of nematodes on swarming alates, larvae, and eggs. In addition, although nematodes were not repelled by fungus and fungus gardens, they rarely occurred on fungus gardens, which indicated that nematodes were likely suppressed before entering fungus gardens. We suggest that the low quantities of nematodes in agricultural systems of fungus-growing termites were due to integrated management by termites through three strategies: (1) prevention of transmission of nematodes by partitioning of tasks and diets; (2) suppression of nematodes in fungus gardens by passing fresh substrates through their guts; and (3) decontaminating the vectored nematodes through allogrooming behavior.

In fungus-growing termites, major workers are generally foragers while minor workers are mostly nest-keepers. For example, in a fungus-growing termite, O. distans Holmgren and Holmgren, 94.8% of the individuals collected from feeding sites were major workers, and 95.1% of the individuals collected from queen chambers were minor workers54. Similarly, in another fungus-growing termite, Macrotermes subhyalinus Rambur, 88.8% of foragers were major workers, and 56.1% of individuals in the nest were minor workers55. Such partitioning of tasks predictably decreases the level of interactions between major workers and queens, larvae, or eggs, and may lower the probability of transferring nematodes. In addition to task partitioning, age polyethism on diets was also observed in M. subhyalinus. Young major workers that molted less than 30 days before stay in the nest, consume the plant material collected by foragers, and construct fungus gardens, while the older major workers that molted more than 30 days before are more likely to forage for plant materials in the field and largely consume aged substrates of fungus gardens in their diets55. The diet partitioning in fungus-growing termites separates the constructors of fungus gardens and the foragers, which likely decreases the probability of foragers contaminating fresh substrates.

Thomas47 reported that an average of 229.8 and 8.6 fungal isolates were found in a gram of plant material collected and in fresh substrate of fungus gardens constructed by the fungus-growing termite, M. bellicosus (Smeathman), respectively, which supported the claim that microbial populations were suppressed after passing the gut of major workers. We propose that O. formosanus controlled the populations of nematodes inhabiting the plant materials collected by integrating the task and diet partitioning, and suppressing the microbial populations in fresh substrates of fungus gardens.

Allogrooming behavior, which cleans microbes growing on body surfaces56,57,58,59, was reported in a fungus-growing termite, Macrotermes michaelseni (Sjöstedt)60. Suppression of microbial populations by allogrooming behavior has been confirmed in multiple groups of termites, such as genus Reticulitermes (Blattodea: Rhinotermitidae)61 and genus Zootermopsis (Blattodea: Archotermopsidae)59. In the present study, no nematodes were observed on the body surfaces of alates, workers, larvae, eggs, and most of the inquilines. Even in the millipedes, which commonly harbor internal parasites62,63,64 and external phoretic nematodes65, no nematodes were isolated. For inquilines, nematodes were only found beneath the elytron of coleopteran inquilines, which is a site that is not likely to be cleaned by allogrooming behavior in termites. In the laboratory, we observed that O. formosanus performed allogrooming behavior on all inquilines (unpublished data). We propose that allogrooming behavior in the fungus-growing termites not only managed the populations of nematodes carried by termites, but also the populations carried by inquilines.

Diversity of nematodes of termites and inquilines

Three nematode genera were isolated from O. formosanus: Halicephalobus, Diplogastrellus, and Aphelenchoides. Halicephalobus is one of the most ubiquitous nematode genus, inhabiting various environments66,67,68,69. The genus Diplogastrellus is commonly found in rotten plants70 and insects associated with rotten plants71, as well as in humid soils that are rich in organic matter72. Similarly, the genus Aphelenchoides sp. is associated with humid soils that are rich in organic matter. The nematode species composition in O. formosanus is likely associated to soil environment, similar to in termites foraging or nesting in soil. For example, Halicephalobus was isolated from the subterranean termite Reticulitermes lucifagus in Corsica23, from dry/dampwood termites in both Florida and Taiwan27,73, from Coptotermes formosanus in Florida74, and from many different termite species in Central America30. Aphelenchoides was isolated from two termite species: Reticulitermes lucifagus in Corsica23 and Hodotermopsis sjostedti in Japan75. We suggest that the nematodes isolated from termite workers were from their foraging environments, e.g., soils and/or substrates that the termites foraged.

Only a single nematode genus, Acrostichus, was isolated from inquilines, which was not found in termites. The result indicated that nematodes were not transferred between termites and inquilines, and the sources of nematodes in inquilines were different from the sources of nematodes in termites. Nevertheless, the life cycle and ecology of Acrostichus spp. is unknown.

Conclusion

Vertical transmission of nematodes through termite dispersal alates to eggs and larvae was not observed in the fungus-growing termite, O. formosanus. Nematodes were transferred into termite nests through two horizontal transmission pathways, including via termite foragers and inquilines. Nematode species observed on termite foragers are associated with the soil environment, but inquilines carried specific nematode species, which indicates the two horizontal transmission pathways are exclusive. Laboratory experiments revealed neither fungus garden substrates nor Termitomyces fungi were toxic to nematodes, but nematodes were barely present in fungus gardens in the field, which indicates termite hygienic behavior plays a vital role in nematode management in the nests.

Data Availability

The sequences of nematodes and GenBank accession number of Termitomyces spp. are available as Supplementary Information to this paper.

References

  1. 1.

    Mueller, U. G., Gerardo, N. M., Aanen, D. K., Six, D. L. & Schultz, T. R. The evolution of agriculture in insects. Annu. Rev. Ecol. Evol. Syst. 36, 563–595 (2005).

  2. 2.

    Nobre, T., Rouland-Lefèvre, C. & Aanen, D. K. Comparative biology of fungus cultivation in termites and ants in Biology of termites: a modern synthesis (eds Bignell, D. E., Roisin, Y., & Lo, N.) 193–210 (Springer, Heidelberg, 2011).

  3. 3.

    Chapela, I. H., Rehner, S. A., Schultz, T. R. & Mueller, U. G. Evolutionary history of the symbiosis between fungus-growing ants and their fungi. Science 266, 1691–1694 (1994).

  4. 4.

    Schultz, T. R. & Brady, S. G. Major evolutionary transitions in ant agriculture. Proc. Natl. Acad. Sci. USA 105, 5435–5440 (2008).

  5. 5.

    Darlington, J. P. E. C. Nutrition and evolution in fungus-growing termites in Nourishment and evolution in insect societies (eds Hunt, J. H. & Nalepa, C. A.) 105–130 (Westview press, Boulder, CO, 1994).

  6. 6.

    Aanen, D. K. et al. The evolution of fungus-growing termites and their mutualistic fungal symbionts. Proc. Natl. Acad. Sci. USA 99, 14887–14892 (2002).

  7. 7.

    Jordal, B. H. & Cognato, A. I. Molecular phylogeny of bark and ambrosia beetles reveals multiple origins of fungus farming during periods of global warming. BMC Evol. Biol. 12, 133 (2012).

  8. 8.

    Hulcr, J. & Stelinski, L. L. The ambrosia symbiosis: from evolutionary ecology to practical management. Annu. Rev. Entomol. 62, 285–303 (2017).

  9. 9.

    Johnson, R. A. Colony development and establishment of the fungus comb in Microtermes sp. nr. usambaricus (Sjöstedt) (Isoptera: Macrotermitinae) from Nigeria. Insectes. Soc. 28, 3–12 (1981).

  10. 10.

    Li, H. et al. Lignocellulose pretreatment in a fungus-cultivating termite. Proc. Natl. Acad. Sci. USA 114, 4709–4714 (2017).

  11. 11.

    Rouland, C. & Bignell, D. E. Cultivation of symbiotic fungi by termites of the subfamily Macrotermitinae in Symbiosis (ed. Seckbach, J.) 733–756 (Kluwer Academic Publishers, the Netherlands, 2001).

  12. 12.

    Kok, L. T., Norris, D. M. & Chu, H. M. Sterol metabolism as a basis for a mutualistic symbiosis. Nature 225, 661–662 (1970).

  13. 13.

    Beaver, R. A. Insect-fungus relationships in the bark and ambrosia beetles in Insect-fungus interactions (eds Wilding, N., Collins, N., Hammond, P., & Webber, J.) 121–143 (14th Symposium of the Royal Entomological Society of London, Academic Press, London, 1989).

  14. 14.

    Rouland-Lefèvre, C., Inoue, T. & Johjima, T. Termitomyces/termite interactions in Intestinal microorganisms of termites and other invertebrates (eds König, H. & Varma, A.) 335–350 (Springer, Heidelberg, 2006).

  15. 15.

    Poinar, G. O. Jr. The natural history of nematodes. 323 pp. (Prentice-Hall, Englewood Cliffs, New Jersey, USA, 1983).

  16. 16.

    Hunt, D. J. Aphelenchida, Longidoridae and Trichodoridae: their systematics and bionomics. 352 pp. (CABI, Wallingford, UK, 1993).

  17. 17.

    Siddiqi, M. R. Tylenchida: parasites of plants and insects, 2nd edition. (CABI, Wallingford, UK, 2000).

  18. 18.

    Tsuda, K., Futai, K. & Kosaka, H. The tripartite relationship in gill-knot disease of the oyster mushroom, Pleurotus ostreatus (Jacq.: Fr.) Kummer. Can. J. Zool. 74, 1402–1408 (1996).

  19. 19.

    Nagesh, M. & Reddy, P. P. Status of mushroom nematodes and their management in India. Integr. Pest Manag. Rev. 5, 213–224 (2000).

  20. 20.

    Mueller, U. G. & Gerardo, N. Fungus-farming insects: multiple origins and diverse evolutionary histories. Proc. Natl. Acad. Sci. USA 99, 15247–15249 (2002).

  21. 21.

    Sudhaus, W. & Koch, C. The new nematode species Poikilolaimus ernstmayri sp. n. associated with termites, with a discussion on the phylogeny of Poikilolaimus (Rhabditida). Russ. J. Nematol. 12, 143–156 (2004).

  22. 22.

    Carta, L. K. & Osbrink, W. Rhabditis rainai n. sp. (Nematoda: Rhabditida) associated with the Formosan subterranean termite, Coptotermes formosanus (Isoptera: Rhinotermitidae). Nematology 7, 863–879 (2005).

  23. 23.

    Fürst von Lieven, A. & Sudhaus, W. Description of Oigolaimella attenuata n. sp. (Diplogastridae) associated with termites (Reticulitermes) and remarks on life cycle, giant spermatozoa, gut-inhabiting flagellates and other associates. Nematology 10, 501–523 (2008).

  24. 24.

    Kanzaki, N., Giblin-Davis, R. M., Scheffrahn, R. H. & Center, B. J. Poikilolaimus floridensis n. sp. (Rhabditida: Rhabditidae) associated with termites (Kalotermitidae). Nematology 11, 203–216 (2009).

  25. 25.

    Kanzaki, N., Giblin-Davis, R. M., Scheffrahn, R. H., Center, B. J. & Davies, K. A. Pseudaphelenchus yukiae n. gen., n. sp. (Tylenchina: Aphelenchoididae) associated with Cylindrotermes macrognathus (Termitidae: Termitinae) in La Selva, Costa Rica. Nematology 11, 869–881 (2009).

  26. 26.

    Kanzaki, N., Giblin-Davis, R. M., Herre, E. A., Scheffrahn, R. H. & Center, B. J. Pseudaphelenchus vindai n. sp. (Tylenchomorpha: Aphelenchoididae) associated with termites (Termitidae) in Barro Colorado Island, Panama. Nematology 12, 905–914 (2010).

  27. 27.

    Kanzaki, N. et al. Poikilolaimus carsiops n. sp. (Rhabditida: Rhabditidae) associated with Neotermes koshunensis (Kalotermitidae) in Kenting National Park, Taiwan. Nematology 13, 155–164 (2011).

  28. 28.

    Kanzaki, N., Li, H.-F., Lan, Y.-C. & Giblin-Davis, R. M. Description of two Pseudaphelenchus species (Tylenchomorpha: Aphelenchoididae) associated with Asian termites and proposal of Tylaphelenchinae n. subfam. Nematology 16, 963–978 (2014).

  29. 29.

    Kanzaki, N., Ragsdale, E. J., Herrmann, M., Mayer, W. E. & Sommer, R. J. Description of three nematode species from Japan, Pristionchus exspectatus n. sp., P. arcanus n. sp., and P. japonicus n. sp., which form a cryptic species complex including the model organism P. pacificus (Diplogastridae). Zool. Sci. 29, 403–417 (2012).

  30. 30.

    Kanzaki, N. et al. Reverse taxonomy for tying diversity of termite-associated nematodes: a practical application of methodology. PLoS One 7, e43865 (2012).

  31. 31.

    Powers, T. O. et al. Tropical nematode diversity: vertical stratification of nematode communities in a Costa Rican humid lowland rainforest. Mol. Ecol. 18, 985–996 (2009).

  32. 32.

    Mishra, S. C. & Sen-Sarma, P. K. Nutritional significance of fungus comb and Termitimyces albuminosus (Berk.) Heim in Odontotermes obesus (Rambur) (Isoptera: Termitidae). Material und Organismen, 205–213 (1985).

  33. 33.

    Batra, L. R. & Batra, S. W. T. Termite-fungus mutualism in Insect-fungus symbiosis-nutrition, mutualism and commensalism (eds Batra, L. R.) 117–163 (Wiley, NY, 1979).

  34. 34.

    Kistner, D. H. The biology of termitophiles in Biology of termites Vol. 1 (eds Krishna, K. & Weesner, F. M.) 525–557 (Academic Press, New York, 1969).

  35. 35.

    Kistner, D. H. Social and evolutionary significance of social insect symbionts. Social Insects 1, 339–413 (1979).

  36. 36.

    Kistner, D. H. The social insects’ bestiary. Social Insects 3, 1–244 (1982).

  37. 37.

    Holldobler, B. & Wilson, E. O. The Ants. (The Belknap Press of Harvard University Press, Cambridge, Massachusetts, 1990).

  38. 38.

    Howard, R. W., McDaniel, C. & Blomquist, G. J. Chemical mimicry as an integrating mechanism: cuticular hydrocarbons of a termitophile and its host. Science 210, 431–433 (1980).

  39. 39.

    Lenoir, A., D’Ettorre, P., Errard, C. & Hefetz, A. Chemical ecology and social parasitism in ants. Annu. Rev. Entomol. 46, 573–599, https://doi.org/10.1146/annurev.ento.46.1.573 (2001).

  40. 40.

    Currie, C. R., Scott, J. A., Summerbell, R. C. & Malloch, D. Fungus-growing ants use antibiotic-producing bacteria to control garden parasites. Nature 398, 701–704 (1999).

  41. 41.

    Fernández-Marín, H., Zimmerman, J. K., Nash, D. R., Boomsma, J. J. & Wcislo, W. T. Reduced biological control and enhanced chemical pest management in the evolution of fungus farming in ants. Proc. R. Soc. B Biol. Sci. 276, 2263–2269 (2009).

  42. 42.

    Sen, R. et al. Generalized antifungal activity and 454-screening of Pseudonocardia and Amycolatopsis bacteria in nests of fungus-growing ants. Proc. Natl. Acad. Sci. USA 106, 17805–17810 (2009).

  43. 43.

    Um, S., Fraimout, A., Sapountzis, P., Oh, D.-C. & Poulsen, M. The fungus-growing termite Macrotermes natalensis harbors bacillaene-producing Bacillus sp. that inhibit potentially antagonistic fungi. Sci. Rep. 3, 3250 (2013).

  44. 44.

    Currie, C. R. & Stuart, A. E. Weeding and grooming of pathogens in agriculture by ants. Proc. R. Soc. B Biol. Sci. 268, 1033–1039 (2001).

  45. 45.

    Hughes, W. O., Eilenberg, J. & Boomsma, J. J. Trade-offs in group living: transmission and disease resistance in leaf-cutting ants. Proc. R. Soc. B Biol. Sci. 269, 1811–1819 (2002).

  46. 46.

    Naug, D. & Camazine, S. The role of colony organization on pathogen transmission in social insects. J. Theor. Biol. 215, 427–439 (2002).

  47. 47.

    Thomas, R. J. Distribution of Termitomyces Heim and other fungi in the nests and major workers of Macrotermes bellicosus (Smeathman) in Nigeria. Soil Biol. Biochem. 19, 329–333 (1987).

  48. 48.

    Rouland-Lefèvre, C. Symbiosis with fungi in Termites: evolution, sociality, symbioses, ecology (eds Abe, T., Bignell, D. E., & Higashi, M.) 289–306 (Springer, Dordrecht, 2000).

  49. 49.

    Liang, W.-R. & Li, H.-F. New discovery and review of termitophile fauna in Taiwan in Proceedings of the 11 th pacific rim termite research group conference (Kunming, China, 2016).

  50. 50.

    Kikuchi, T., Aikawa, T., Oeda, Y., Karim, N. & Kanzaki, N. A rapid and precise diagnostic method for detecting the pinewood nematode Bursaphelenchus xylophilus by loop-mediated isothermal amplification. Phytopathology 99, 1365–1369 (2009).

  51. 51.

    Tanaka, R., Kikuchi, T., Aikawa, T. & Kanzaki, N. Simple and quick methods for nematode DNA preparation. Appl. Entomol. Zool. 47, 291–294 (2012).

  52. 52.

    Kanzaki, N. & Futai, K. A. PCR primer set for determination of phylogenetic relationships of Bursaphelenchus species within the xylophilus group. Nematology 4, 35–41 (2002).

  53. 53.

    Ye, W., Giblin-Davis, R. M., Braasch, H., Morris, K. & Thomas, W. K. Phylogenetic relationships among Bursaphelenchus species (Nematoda: Parasitaphelenchidae) inferred from nuclear ribosomal and mitochondrial DNA sequence data. Mol. Phylogenet. Evol. 3, 1185–1197 (2007).

  54. 54.

    McMahan, E. A., Kumar, S. & Sen-Sarma, P. Male/female (size) polyethism in workers of Odontotermes distans Holmgren and Holmgren (Isoptera: Termitidae: Macrotermitinae). Ann. Entomol. Soc. Am. 77, 429–434 (1984).

  55. 55.

    Badertscher, S., Gerber, C. & Leuthold, R. Polyethism in food supply and processing in termite colonies of Macrotermes subhyalinus (Isoptera). Behav. Ecol. Sociobiol. 12, 115–119 (1983).

  56. 56.

    Chouvenc, T. & Su, N.-Y. Apparent synergy among defense mechanisms in subterranean termites (Rhinotermitidae) against epizootic events: Limits and potential for biological control. J. Econ. Entomol. 103, 1327–1337 (2010).

  57. 57.

    Mankowski, M. E., Kaya, H. K., Grace, K. J. & Sipes, B. Differential susceptibility of subterranean termite castes to entomopathogenic nematodes. Biocontrol Sci. Technol. 15, 367–377 (2005).

  58. 58.

    Wilson-Rich, N., Stuart, R. J. & Rosengaus, R. B. Susceptibility and behavioral responses of the dampwood termite Zootermopsis angusticollis to the entomopathogenic nematode Steinernema carpocapsae. J. Invertebr. Pathol. 95, 17–25 (2007).

  59. 59.

    Rosengaus, R. B., Maxmen, A. B., Coates, L. E. & Traniello, J. F. Disease resistance: a benefit of sociality in the dampwood termite Zootermopsis angusticollis (Isoptera: Termopsidae). Behav. Ecol. Sociobiol. 44, 125–134 (1998).

  60. 60.

    Sieber, R. & Leuthold, R. H. Behavioural elements and their meaning in incipient laboratory colonies of the fungus-growing termite Macrotermes michaelseni (Isoptera: Macrotermitinae). Insectes Soc. 28, 371–382 (1981).

  61. 61.

    Rosengaus, R. B., Traniello, J. F. A. & Bulmer, M. S. Ecology, behavior and evolution of disease resistance in termites in Biology of termites: a modern synthesis (eds Bignell, D. E., Roisin, Y. & Lo, N.) 165–191 (Springer, Heidelberg, 2011).

  62. 62.

    Hunt, D. J. Four new species of the genus Rhigonema Cobb, 1898 (Nematoda: Rhigonematida: Rhigonematidae) parasitic in diplopods from Papua, New Guinea. Nematol. Mediterr. 23, 217–234 (1995).

  63. 63.

    Hunt, D. J. & Spiridonov, S. A new genus of Carnoyidae (Nematoda: Rhigonematida) with descriptions of Cattiena trachelomegali gen. n., sp. n. and C. trigoniuli gen. n., sp. n., parasites of Spirobolida (Diplopoda) from Vietnam. Nematology 3, 559–571 (2001).

  64. 64.

    García, N. & Morffe, J. A new genus and two new species of Xustrostomatidae Hunt, 2002 (Nematoda: Rhigonematomorpha) from the West Indies. Pap. Avulsos. Zool. (Sao Paulo) 55, 91–101 (2015).

  65. 65.

    Ragsdale, E. J., Kanzaki, N. & Herrmann, M. Taxonomy and natural history: the genus Pristionchus in Pristionchus pacificusc-a nematode model for comparative and evolutionary biology (ed. Sommer, R. J.) 77–120 (Brill Academic Pub, Leiden, Netherlands, 2015).

  66. 66.

    Kinde, H., Mathews, M., Ash, L. & St. Leger, J. Halicephalobus gingivalis (H. deletrix) infection in two horses in southern California. J. Vet. Diagn. Investig. 12, 162–165 (2000).

  67. 67.

    Nadler, S. A. et al. Molecular phylogenetics and diagnosis of soil and clinical isolates of Halicephalobus gingivalis (Nematoda: Cephalobina: Panagrolaimoidea), an opportunistic pathogen of horses. Int. J. Parasitol. 33, 1115–1125 (2003).

  68. 68.

    Borgonie, G. et al. Nematoda from the terrestrial deep subsurface of South Africa. Nature 474, 79–82 (2011).

  69. 69.

    Steel, H. et al. Survival and colonization of nematodes in a composting process. Invertebr. Biol 132, 108–119 (2013).

  70. 70.

    Kiontke, K. & Sudhaus, W. Diplogaster (Diplogastrellus) cerea sp. n. from saguaro cactus rot and a revision of the subgenus Diplogastrellus (Nematoda: Diplogastridae). Nematologica 42, 173–197 (1996).

  71. 71.

    Kanzaki, N., Giblin-Davis, R. M., Zeng, Y., Ye, W. & Center, B. J. Diplogastrellus metamasius n. sp. (Rhabditida: Diplogastridae) isolated from Metamasius hemipterus (L.) (Coleoptera: Dryophthoridae) in southern Florida and Costa Rica. Nematology 10, 853–868 (2008).

  72. 72.

    Khan, R., Bajaj, H. K., Sultana, R. & Tahseen, Q. Description of Diplogastrellus gracilis (Bütschli, 1876) Paramonov, 1952, D. sikorai sp. n. and Fictor composticola sp. n. (Nematoda: Diplogastrina) from India. Nematology 10, 153–166 (2008).

  73. 73.

    Kanzaki, N., Giblin-Davis, R. M., Scheffrahn, R. H. & Center, B. J. Dauer juveniles of Poikilolaimus floridensis (Rhabditida: Rhabditidae) isolated from drywood termites (Kalotermitidae). Nematropica 39, 305–310 (2009).

  74. 74.

    Foley, J. R., Chouvenc, T., Giblin-Davis, R. M., Su, N.-Y. & Kanzaki, N. Phoresy and within-colony transmission of nematodes associated with alates of subterranean termites (Isoptera: Rhinotermitidae). Environ. Entomol. 47, 1107–1116 (2018).

  75. 75.

    Kanzaki, N., Ohmura, W. & Giblin-Davis, R. M. Termite-associated nematode fauna in relation to their host/carrier termite habitat. XXV International Congress of Entomology 0951 (2016).

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Acknowledgements

We thank Kuan-Hung Chen and Kuan-Chih Kuan (Department of Entomology, NCHU), and Jie-Hao Ou (Department of Plant Pathology, NCHU) for locating and excavating termite nests. We thank Chih-Yu Kuo for drawing the Figure 4. We also thank Noriko Shimoda and Atsuko Matsumoto, FFPRI, for culturing nematodes and molecular sequencing. This study was supported by grants from the Ministry of Science and Technology, Taiwan (MOST 104-2311-B-005-002; MOST 105-2628-B-005-003-MY3).

Author information

N.K., W.R.L., C.I.C. and H.F.L. designed and performed the field experiments. W.R.L. collected and identified inquilines. N.K. and W.R.L. performed the nematode inspections on isolation plates. N.K. identified nematodes and performed molecular sequencing. N.K., C.T.Y. and Y.P.H. cultured nematodes for the nematocidal test. W.R.L., C.I.C., C.T.Y. and Y.P.H. performed the fungus toxicity bioassays. W.R.L. and H.F.L. designed the figures. N.K. and W.R.L. wrote the manuscript in consultation with C.I.C., C.T.Y., Y.P.H. and H.F.L. The project was supervised by H.F.L.

Correspondence to Hou-Feng Li.

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Supplementary information

Aphelenchoides sp. bored into termite fungus garden

Supplementary data of collection and identification of nematodes, fungi, inquilines

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