Phenotypic plasticity is one of the most important strategies used by organisms with low mobility to survive in fluctuating environments. Phenotypic plasticity plays a vital role in nematodes because they have small bodies and lack wings or legs and thus, cannot move far by themselves. Bursaphelenchus xylophilus, the pathogenic nematode species that causes pine wilt disease, experiences fluctuating conditions throughout their life history; i.e., in both the phytophagous and mycetophagous phases. However, whether the functional morphology changes between the life phases of B. xylophilus remains unknown. Our study revealed differences in the ultrastructure of B. xylophilus between the two phases. Well-developed lateral alae and atrophied intestinal microvilli were observed in the phytophagous phase compared with the mycetophagous phase. The ultrastructure in the phytophagous phase was morphologically similar to that at the dauer stage, which enables the larvae to survive in harsh environments. It suggests that the living tree represents a harsh environment for B. xylophilus, and ultrastructural phenotypic plasticity is a key strategy for B. xylophilus to survive in a living tree. In addition, ultrastructural observations of obligate plant-parasitic species closely related to B. xylophilus revealed that B. xylophilus may be in the process of adapting to feed on plant cells.
Many environments are heterogeneous and change continuously. Most highly mobile organisms are able to move around to find optimal or better habitats. For example, winged insects or birds can fly away from environments that become unfavorable on a seasonal basis. Mobile animals can migrate into a new environment for feeding and/or reproduction. On the other hand, low mobility or sessile organisms cannot escape from their habitat when it becomes unfavorable and often alter their phenotype to adapt to their changing habitat. This ability of a single individual to develop more than one phenotype is termed “phenotypic plasticity.” Phenotypic plasticity plays a key role in the survival and propagation of certain organisms1,2 in nature where large environmental fluctuations occur and could be one driver of evolution through the initiation of adaptive divergence, i.e., “plasticity-first” evolution3.
Phenotypic plasticity is well-studied in plants. Plants are sessile and cannot move even if the environment becomes unfavorable, making plasticity very important for their survival4; plants can recognize changes in their environment and alter their forms to adapt without moving. As an example, plant leaves are particularly plastic and exhibit great diversity in shape, size, and color in nature. Environmental factors, such as temperature, light quality and intensity, and humidity, all affect leaf morphology5,6,7. Phenotypic plasticity has also been reported in many animals. Many clones of Daphnia pulex develop thorns as antipredator devices in the presence of chemical signals from predators such as the insect Chaoborus americanus, and clones of D. pulex have been shown to develop thorns, known as neckteeth, until the end of the fourth juvenile instar, when they were exposed to chemical cues emitted from their predator, the Chaoborus flavicans larva8,9,10.
Nematoda is one of the most abundant and diverse groups of organisms on earth11,12. Nematodes have no wings or legs, and their body size is relatively small, resulting in a low dispersal capacity. Phenotypic plasticity is essential in nematodes because of their reduced mobility. The phenotypic plasticity of nematodes has been reported previously; for example, Caenorhabditis elegans enters a stress-resistant dauer stage in response to harsh environmental conditions13. Under optimum growth conditions, second-stage larvae (L2) develop into third-stage larvae, and then into adults. Under unfavorable growth conditions, L2 become dauer larvae, which enable survival and dispersal, and in this stage they are resistant to environmental stresses14.
The stomatal phenotypic plasticity of Diplogastridae nematodes has been well studied. Pristionchus pacificus has two feeding structures: a “wide-mouthed” eurystomatous morph with two large teeth and a “narrow-mouthed” stenostomatous morph with one small tooth. Eurystomatous animals can feed on other nematode species15, whereas stenostomatous animals have difficulty feeding on other nematodes, but are specialized as bacteria feeders when food bacteria are sufficient16.
Recently, stomatal phenotypic plasticity has been reported in another nematode clade, the genus Bursaphelenchus17. Most Bursaphelenchus feed on fungi, but a few species parasitize plants. Bursaphelenchus spp. commonly have a syringe-like feeding structure referred to as a “stylet” to pierce fungal and plant cells for nutrient uptake. However, Bursaphelenchus sinensis, which inhabits in dead pine trees, has a predatory form that has a stylet with a wider lumen than the mycetophagous form, and feeds on other nematodes.
The plant-parasitic nematode, Bursaphelenchus xylophilus, the causal pathogen of pine wilt disease, is thought to inhabit more fluctuating environments than B. sinensis. B. xylophilus mostly experiences two different environmental conditions, living pine trees and dead, moldy pine trees, whereas B. sinensis experiences only dead pine trees. Although no stomatal plasticity was observed in B. xylophilus17, B. xylophilus likely has environmentally sensitive traits. B. xylophilus uses epithelial cells and resin canal parenchyma cells as food sources in a living pine tree18. In this study, we refer to the phase in a living pine tree as the “phytophagous phase.” In many cases, infection by B. xylophilus causes pine wilt disease and results in the death of the infected tree. Various bacteria and fungi grow on the dead pine tree, and B. xylophilus feeds on the fungi and propagates rapidly19. Herein, we refer to this phase as the “mycetophagous phase.” In the fluctuating environment, B. xylophilus adapts its phenotype to survive. Thus, we compared the phenotypes between the phytophagous and mycetophagous phases, as this was the best method to understand the survival strategy of B. xylophilus in relation to its phenotypic plasticity. Tsai et al.20 showed that the expression of the collagen gene family differed significantly between these two phases, which indicated that the nematode exhibited morphological phenotypic plasticity. Although collagens are essential in structural formation and modification in nematodes21, compound microscopy revealed that the only morphological difference between these two phases was in the tail shape of adult females.
In this study, we used transmission electron microscopy (TEM) to investigate differences in the ultrastructure of the cuticle and intestines of B. xylophilus as these structures consist mainly of collagen22,23. Differences were examined between the phytophagous and mycetophagous phases, and the functional significance of any structural change is discussed. Furthermore, to understand the evolutionary adaptation of B. xylophilus to its host plant, we examined the internal ultrastructure and intestines of an obligate plant-parasitic species (phytophagous phase), Schistonchus sp., which is closely related to B. xylophilus, belongs to the family Aphelenchoididae, and evolved from soil-inhabiting mycetophagous species24. Findings were compared with those observed from the phytophagous phase of B. xylophilus.
Nematodes of B. xylophilus in the phytophagous phase were recovered from inoculated pine trees at 2, 3, and 4 weeks after inoculation. Throughout the sampling time, initial visible symptoms of disease onset were noted. The tips of the pine needles turned yellow, but the other needle parts remained green. No differences were observed in the ultrastructure of B. xylophilus between the three different time points (2, 3, and 4 weeks). Therefore, all the morphological information recorded during the phytophagous phase described hereafter was obtained from B. xylophilus recovered from inoculated pine trees at 2 weeks post-inoculation.
Observations of ultrastructure using TEM
We observed the cuticle, lateral alae, and intestinal ultrastructure of B. xylophilus grown on live pines, dead pines inoculated with fungus, and on fungal cultures.
Tsai et al. showed that the expression of the collagen gene family differed significantly between the mycetophagous phase and phytophagous phases20, which suggests that the cuticle, a structure consisting mainly of collagen, changes between the two phases. However, we observed no qualitative differences in the cuticular structure of B. xylophilus between sexes and phases (Fig. 1). The structure of the cuticle was morphologically the same as that previously reported25. It consisted of three parts: an epicuticle (EPI), cortical and median zones (CZ and MZ), and a basal zone (BZ). The EPI consisted of three layers: an electron-dense outermost layer (surface coat) and two inner layers. However, the triple layer was sometimes indistinct and appeared as a single or double layer. The CZ and MZ were electron-lucent and not distinguishable from each other, forming an amorphous zone. In comparison, the BZ was distinguishable from the MZ due to its radial striation. The thickness of the cuticle was significantly different between the two different phases, i.e., the phytophagous and mycetophagous phases cultured on agar plates for both females and males (P < 0.05).
Observing the cuticle structure, we found that the structures of the lateral alae differed markedly between the mycetophagous and phytophagous phases in males. The lateral alae were not conspicuous, i.e., were relatively flattened with a smooth surface, in the mycetophagous phase (Fig. 2D, E) in males and in both phases in females (Fig. 2A–C). On the other hand, the lateral alae were very well developed, with each band expanded with a mushroom-like outline in cross section in the phytophagous phase (Fig. 2F). To evaluate the degree of protrusion from cuticle, the protruding area of lateral alae was measured (Table 1). There were significant differences in the protrusion area between the phytophagous and mycetophagous phases in males cultured on agar plates (P < 0.05) and on pine stems (P < 0.01), whereas no qualitative or quantitative differences were observed in the lateral alae of females between phases.
All individuals had some microvilli on the internal surface of the intestines (Fig. 3). The microvilli were connected to the basement membrane and supported by an axial core of actin filaments. However, morphological and quantitative characteristics differed between the mycetophagous and phytophagous phases. Microvilli on the internal surface of the intestines were longer and more numerous in the mycetophagous phase (cultured on either agar plates or pine stems) than in the phytophagous phase (Fig. 3).
The intestinal structure of Schistonchus sp., an obligate plant parasite, was also observed in this study for comparisons with the structure of the facultative plant parasite B. xylophilus. The intestinal structure of Schistonchus sp. was similar to that of B. xylophilus in the mycetophagous phase; i.e., the microvilli were long and numerous (Fig. 4).
We studied the phenotypic plasticity of B. xylophilus to investigate its adaptation strategy in response to the changing environment in pine trees. We focused on its ultrastructure and made comparisons between the mycetophagous and phytophagous phases using TEM. As a result, we revealed that nematodes in the phytophagous phase had more developed lateral alae and more atrophic intestinal microvilli than those in the mycetophagous phase.
The lateral alae of nematodes in the phytophagous phase were mushroom-shaped, whereas those of nematodes in the mycetophagous phase were smooth. Shinya et al.26 observed the surface structure in B. xylophilus in the mycetophagous and phytophagous phases using scanning electron microscopy. Although they did not refer to the various forms of the lateral alae, Fig. 5 in their paper showed that nematodes in the phytophagous phase had well-developed lateral alae compared to nematodes raised on fungus. Alae are thickenings or projections of the cuticle that occur in the lateral or sublateral region. Lateral alae occur in both sexes, two per individual, and run longitudinally along the length of the nematode body27. They enable the nematode to change shape and the cuticle to flex during dorsoventral contractions. The exact function of the lateral alae remains unknown; however, their structure varies considerably, not only among different taxa but also across developmental stages within a species27. Therefore, we hypothesize that the lateral alae are one of the most functionally important structures of the nematode surface features, formed by elaborations of the cuticle27. The alae have a complex structure, which differs from that of the general body cuticle, and they may provide a degree of longitudinal stiffening. Furthermore, because nematodes lie and move on their sides28, the alae are in contact with the substrate, rather like the tread of a car tire27, where they probably assist in locomotion by increasing traction and preventing slipping, although their absence in some forms does not appear to inhibit movement. For example, the lateral alae in the dauer stage in Caenorhabditis elegans are different from that in other developmental stages and are mushroom-shaped29. Dauer larvae display a specific behavior known as “nictation,” in which the nematodes lift and wave the anterior part of their bodies13, enabling them to attach to vectors such as insects30. The lateral alae are considered to play an important role in this specific behavior. The alae run longitudinally along the length of the nematode body and are also thought to support the body. It is likely that the alae enable nematodes to undertake three-dimensional activities, such as nictation as well as crawling.
Phytophagous individuals of B. xylophilus have well-developed and complex lateral alae. This suggests that B. xylophilus moves more actively in a living pine tree than on a dead pine tree and fungal mat. Nematodes were collected from fungal culture plates or pine trees, and phytophagous individuals exhibited three-dimensional activity in water, whereas mycetophagous individuals did not. From a pathological viewpoint, migration ability in a living tree is an indispensable trait for B. xylophilus to cause pine wilt disease31. Bursaphelenchus xylophilus nematodes migrate to the xylem tissues, spreading throughout the infected pine tree, and parenchymatous cell death and cavitation occur following the nematode migration. Finally, the infected pine tree dies due to a lack of water. Ichihara et al.31 compared the migration between virulent and avirulent strains of B. xylophilus in the tissues of living pine trees. They reported that the avirulent strain barely invaded the xylem resin canals and cortical tissue, whereas the virulent strain did. They concluded that the migration of B. xylophilus is an important factor that increases the severity of disease symptoms in infected pine trees. Considering these results, we hypothesize that well-developed lateral alae are necessary for B. xylophilus to actively move through tree tissues and cause disease.
The intestinal structure of B. xylophilus differed between the mycetophagous and the phytophagous phases. Although the number of microvilli could not be counted for technical reasons, the properties of the microvilli were observed. Both females and males had more developed microvilli in the mycetophagous phase than in the phytophagous phase. The microvilli in the mycetophagous nematodes were thicker and longer than those in the phytophagous nematodes. Generally, animals absorb nutrients through their intestines from their food. Previous reports showed that the intestinal and microvilli structure on the internal surface of the intestine changes according to nutritional conditions in many animal species, e.g., chicks32, pigs33, and pythons34. In terms of nematodes, the intestinal lumen in the dauer larvae of Caenorhabditis elegans and B. xylophilus was small, and the brush border was so compact that individual microvilli were difficult to discern25,35.
In the phytophagous phase, B. xylophilus most likely uses epithelial cells and the resin canal parenchyma cells as food sources in living trees. Given that the intestinal microvilli of B. xylophilus are shrunken in the phytophagous phase, epithelial cells of the living pine tree are not considered suitable food for B. xylophilus. This coincides with the fact that the reproductive rate of B. xylophilus on fungal mats (e.g., Botrytis cinerea) is higher than in pine seedlings19,36.
To understand the intestinal adaptation of B. xylophilus to the host plant in a phylogenetic context, we observed the intestinal ultrastructure of an undescribed Schistonchus sp. This species is an obligate plant-parasitic species and closely related to B. xylophilus, i.e., both species belong to the family Aphelenchoididae24. Interestingly, Schistonchus sp. feeding on plant (fig) tissues had well-developed microvilli on the surface of the intestine. It was also reported that the L2 and L3 of the obligate plant-parasitic nematode Heterodera glycines had a moderate proliferation of microvilli37. These results suggest that plant-parasitic species, including Schistonchus sp., can ingest sufficient nutrients from plant tissues. Schistonchus share an ancestor with B. xylophilus and is physiologically adapted as a plant parasite, indicating that nematodes are genetically plastic. Considering that the ancestral species of B. xylophilus is a mycetophagous species, it appears that B. xylophilus is in the process of adapting to being able to feed on plant cells.
In this study, we showed that B. xylophilus was able to alter its ultrastructure according to the environment. This is the first report of phenotypic plasticity in B. xylophilus at the ultrastructural level. The lateral alae were more developed in the phytophagous phase than in the mycetophagous phase. On the other hand, the intestine was less developed in the phytophagous phase than in the mycetophagous phase. The ultrastructure in the phytophagous phase was similar to that at the dauer stage, enabling the nematode to survive harsh environments. This suggests that ultrastructural phenotypic plasticity in B. xylophilus is a strategy for surviving harsh environments such as those encountered while actively migrating within the living pine tree instead of feeding. This rapid dispersion, regulated by the phenotypic plasticity, could cause the rapid development of symptoms observed in infected pine trees. To date, little research has focused on phenotypic plasticity in B. xylophilus20,38. However, investigating phenotypic plasticity is essential to aid our understanding of the survival strategy of B. xylophilus, and eventually, the survival strategy of Nematoda.
Materials and methods
Mycetophagous and phytophagous phases were prepared. To understand the effects of growth medium, the mycetophagous phase was cultured on two grown media, i.e., agar plates and dead pine stems.
Mycetophagous phase cultured on agar plates
The B. xylophilus Ka4 C1 strain was cultured with Botrytis cinerea on malt extract agar plates (1.5% malt extract and 4.0% agar) at 25 °C. Nematodes were collected using the Baermann funnel technique overnight. Nematodes were rinsed three times with ion-exchange water (IEW), and females and males were randomly harvested.
Mycetophagous phase cultured on pine stems
Mycetophagous phase growing on pine twigs was prepared following Kanzaki et al.17. The B. xylophilus Ka4 C1 strain was cultured as described above. Ten stems (ca 7 mm in diam. and 4 cm long) of 3-year-old Japanese black pine (Pinus thunbergii) were autoclaved and individually transferred to 15-mL sterile plastic centrifuge tubes. B. cinerea was inoculated onto the stems. One week after inoculation, 500 nematodes were inoculated into the tubes. At 2 weeks after inoculation, the nematodes were extracted from the wood using the Baermann funnel technique overnight. The extracted nematodes were lumped together, transferred to a 15 mL sterile plastic centrifuge tube, and rinsed three times with IEW. Then, females and males were randomly harvested.
The B. xylophilus Ka4 C1 strain was cultured as described above. Nematodes were washed three times with IEW and adjusted to 10,000 individuals per 500 µL of IEW. Two-year-old Japanese black pine trees were each inoculated with 10,000 nematodes on April 25, 2019. Four pine trees were inoculated in all. At 2, 3, and 4 weeks after inoculation, nematodes were extracted from two, one, and one pine tree, respectively, using the Baermann funnel technique overnight. The extracted nematodes were lumped together, and transferred to a separate 15-mL sterile plastic centrifuge tube for each week, rinsed three times with IEW, and females and males were randomly harvested.
Obligate plant-parasitic species closely-related to Bursaphelenchus xylophilus
Fruits of Ficus superba were collected in June 2017 from Ishigaki Island, Okinawa, Japan. A collection permit was not necessary for F. superba because it was not obtained inside a protected area. The fig fruits were identified visually and dissected under a light microscope (S8 Apo; Leica). Parasitic females of an undescribed Schistonchus sp. were identified under the light microscope (Eclipse 80i; Nikon; 200× or 400×) and used for the ultrastructural observations.
Observation of ultrastructure using TEM
Samples were prepared for TEM as described by Ekino et al.39 Adult nematodes were fixed in 1.25% glutaraldehyde and 1.5% picric acid (only 1% glutaraldehyde was used for fixing Schistonchus sp.) in 0.1 mol/L phosphate buffer (pH 7.4) for more than 24 h. Then, the heads and tails were excised from the fixed adults, and the mid-body regions were used; almost all positions were of equal thickness. The nematodes were arranged in a parallel array on a 2% agar pad prepared on a microscope slide40. Molten 2% agarose was dripped onto the pad containing three to five nematodes. After the agarose had solidified, we trimmed it into a cube and dripped molten 2% agarose to cover the surfaces of the agarose cube. It was then trimmed again to form a larger cube. Several cubes were prepared for each nematode species. The cubes were rinsed six times with phosphate buffer (10 min for each rinse) and post-fixed in 1% osmium tetroxide in IEW for 90 min. Then, the samples were dehydrated in a graded series of ethanol baths (one bath each of 50%, 70%, 80%, and 90% ethanol, and three baths of 99.5% ethanol), and cleaned three times with propylene oxide for 5 min. The samples were infiltrated overnight with a mixture of 50% Eponate resin and 50% propylene oxide. On the following night, they were infiltrated with undiluted resin. Finally, the samples were embedded in Epon resin.
We used a diamond knife fitted to an ultramicrotome to section the mid-body region of the nematodes. Sections were only used when the nematodes were cut vertically along the long axis of the body. The sections were collected on Formvar-coated copper grids for electron microscopy. The grid was stained with EM Stainer (Nissin EM, Tokyo, Japan) for 30 min, followed by lead citrate. Grid-mounted sections were photographed at 100 kV using a JEOL JEM-2010 electron microscope (Tokyo, Japan). One section in which the structures were observed clearly in each nematode was used for measurement. The thickness of the total cuticle and the total cross section area were measured from these cross-sections using ImageJ Software version 1.52v (National Institutes of Health, Bethesda, Maryland, USA)41. The cuticle was measured once where the cuticle structure was observed clearly and outside the annuli. The body radius (r) was calculated from the total cross-section area.
To evaluate the degree of protrusion of lateral alae from the cuticle, the protrusion area (S; areas indicated by asterisks in Fig. 5) was calculated. First, the area of the lateral alae (S′; area indicated by dots in Fig. 5) was measured with a polygon-section tool in ImageJ, and their width of (x; arrow in Fig. 5) was measured with the straight-tool in ImageJ. S was calculated using the following formula:
Significant differences in cuticle thickness and protrusion area of lateral alae between the phytophagous and mycetophagous phases (cultured on media plates and on pine stems) were identified using Welch’s t-test. The significance level was adjusted uisng the Bonferroni method. The procedure was performed using Excel 2013 for Windows (Microsoft Corporation, Redmond, WA, USA). A value of P < 0.05 was considered to indicate statistical significance.
West-Eberhard, M. J. Phenotypic plasticity and the origins of diversity. Annu. Rev. Ecol. Syst. 20, 249–278 (1989).
Bateson, P. et al. Developmental plasticity and human health. Nature 430, 419–421 (2004).
Levis, N. A. & Pfennig, D. W. Evaluating ‘plasticity-first’ evolution in nature: Key criteria and empirical approaches. Trends Ecol. Evol. 31, 563–574 (2016).
Schlichting, C. D. The evolution of phenotypic plasticity in plants. Annu. Rev. Ecol. Syst. 17, 667–693 (1986).
Dengler, N. G. Comparative histological basis of sun and shade leaf dimorphism in Helianthus annuus. Can. J. Bot. 58, 717–730 (1980).
Sultan, S. E. Phenotypic plasticity for plant development, function and life history. Trends Plant Sci. 5, 537–542 (2000).
Rozendaal, D. M. A., Hurtado, V. H. & Poorter, L. Plasticity in leaf traits of 38 tropical tree species in response to light; relationships with light demand and adult stature. Funct. Ecol. 20, 207–216 (2006).
Dodson, S. I. The ecological role of chemical stimuli for the zooplankton: Predator-induced morphology in Daphnia. Oecologia 78, 361–367 (1989).
Schwartz, S. S. Predator-induced alterations in Daphnia morphology. J. Plankton Res. 13, 1151–1161 (1991).
Tollrian, R. Neckteeth formation in Daphnia pulex as an example of continuous phenotypic plasticity: Morphological effects of Chaoborus kairomone concentration and their quantification. J. Plankton Res. 15, 1309–1318 (1993).
Yeates, G. W. Soil nematodes in terrestrial ecosystems. J. Nematol. 11, 213–229 (1979).
Bongers, T. & Bongers, M. Functional diversity of nematodes. Appl. Soil Ecol. 10, 239–251 (1998).
Cassada, R. C. & Russell, R. L. The dauerlarva, a post-embryonic developmental variant of the nematode Caenorhabditis elegans. Dev. Biol. 46, 326–342 (1975).
Gutteling, E. W., Riksen, J. A. G., Bakker, J. & Kammenga, J. E. Mapping phenotypic plasticity and genotype–environment interactions affecting life-history traits in Caenorhabditis elegans. Heredity 98, 28–37 (2007).
Serobyan, V., Ragsdale, E. J. & Sommer, R. J. Adaptive value of a predatory mouth-form in a dimorphic nematode. Proc. R. Soc. B Biol. Sci. 281, 20141334 (2014).
Serobyan, V., Ragsdale, E. J., Müller, M. R. & Sommer, R. J. Feeding plasticity in the nematode Pristionchus pacificus is influenced by sex and social context and is linked to developmental speed. Evol. Dev. 15, 161–170 (2013).
Kanzaki, N., Ekino, T. & Giblin-Davis, R. M. Feeding dimorphism in a mycophagous nematode, Bursaphelenchus sinensis. Sci. Rep. 9, 13956 (2019).
Mamiya, Y. Pine wilting disease caused by the pine wood nematode, Bursaphelenchus lignicolus in Japan. Jpn. Agric. Res. Q. 10, 206–211 (1976).
Futai, K. Population dynamics of Bursaphelenchus lignicolus (Nematoda: Aphelenchoididae) and B. mucronatus in pine seedlings. Appl. Entomol. Zool. 15, 458–464 (1980).
Tsai, I. J. et al. Transcriptional and morphological changes in the transition from mycetophagous to phytophagous phase in the plant-parasitic nematode Bursaphelenchus xylophilus: How B. xylophilus adapts in the host environment. Mol. Plant Pathol. 17, 77–83 (2016).
Johnstone, I. L. Cuticle collagen genes. Trends Genet. 16, 21–27 (2000).
Cox, G. N., Kusch, M. & Edgar, R. S. Cuticle of Caenorhabditis elegans: Its isolation and partial characterization. J. Cell. Biol. 90, 7–17 (1981).
Graham, P. L. et al. Type IV collagen is detectable in most, but not all, basement membranes of Caenorhabditis elegans and assembles on tissues that do not expressed it. J. Cell. Biol. 137, 1171–1183 (1997).
Davies, K. A. et al. A review of the taxonomy, phylogeny, distribution and co-evolution of Schistonchus Cobb, 1927 with proposal of Ficophagus n. gen. and Martininema n. gen. (Nematoda: Aphelenchoididae). Nematology 17, 761–829 (2015).
Kondo, E. & Ishibashi, N. Ultrastructural differences between the propagative and dispersal forms in pine wood nematode, Bursaphelenchus lignicolus, with reference to the survival. Appl. Entomol. Zool. 13, 1–11 (1978).
Shinya, R., Morisaka, H., Takeuchi, Y., Ueda, M. & Futai, K. Comparison of the surface coat proteins of the pine wood nematode appeared during host pine infection and in vitro culture by a proteomic approach. Phytopathology 100, 1289–1297 (2010).
Bird, A. F. & Bird, J. The Structure of Nematodes (Academic Press, Cambridge, 1991).
Lee, D. L. Changes in adult Nippostrongylus brasiliensis during the development of immunity to this nematode in rats: 1. Changes in ultrastructure. Parasitology 59, 29–39 (1969).
White, J. The nematode Caenorhabditis elegans. Cold Spring Harb. Monogr. Ser. 17, 81–122 (1988).
Campbell, J. F. & Gaugler, R. Nictation behaviour and its ecological implications in the host search strategies of entomopathogenic nematodes (Heterorhabditidae and Steinernematidae). Behaviour 126, 155–169 (1993).
Ichihara, Y., Fukuda, K. & Suzuki, K. Early symptom development and histological changes associated with migration of Bursaphelenchus xylophilus in seedling tissues of Pinus thunbergii. Plant Dis. 84, 675–680 (2000).
Shamoto, K. & Yamauchi, K. Recovery responses of chick intestinal villus morphology to different refeeding procedures. Poult. Sci. 79, 718–723 (2000).
Li, D. F. et al. Effect of fat sources and combinations on starter pig performance, nutrient digestibility and intestinal morphology. J. Anim. Sci. 68, 3694–3704 (1990).
Lignot, J.-H., Helmstetter, C. & Secor, S. M. Postprandial morphological response of the intestinal epithelium of the Burmese python (Python molurus). Comp. Biochem. Physiol. A. Mol. Integr. Physiol. 141, 280–291 (2005).
Popham, J. D. & Webster, J. M. Aspects of the fine structure of the dauer larva of the nematode Caenorhabditis elegans. Can. J. Zool. 57, 794–800 (1979).
Tamura, H. & Mamiya, Y. Reproduction of Bursaphelenchus lignicolus on alfalfa callus tissues. Nematologica 21, 449–454 (1975).
Endo, B. Y. Infrastructure of the intestine of second and third juvenile stages of the soybean cyst nematode, Heterodera glycines. P. Helm. Sci. Wash. 55, 117–131 (1988).
Shinya, R. et al. Secretome analysis of the pine wood nematode Bursaphelenchus xylophilus reveals the tangled roots of parasitism and its potential for molecular mimicry. PLoS ONE 8, e67377 (2013).
Ekino, T., Yoshiga, T., Takeuchi-Kaneko, Y., Ichihara, Y. & Kanzaki, N. Sexual dimorphism of the cuticle and body-wall muscle in free-living mycophagous nematodes. Can. J. Zool. 97, 510–515 (2019).
Bargmann, C. I. & Avery, L. Laser killing of cells in Caenorhabditis elegans. Methods Cell Biol. 48, 225–250 (1995).
Abramoff, M. D., Magalhaes, P. J. & Ram, S. J. Image processing with ImageJ. Biophoton. Int. 11, 36–42 (2004) (This article is available online).
We sincerely thank Dr. Michio Sato, Meiji University, for his technical assistance with the TEM observations. This study was funded by grants from JSPS KAKENHI no. 19K23679 (to T.E.), JSPS Grant-in-Aid for Early-Career Scientists JP19K15853 (to R.S.), and JST PRESTO Grant no. JPMJPR17Q5 (to R.S.).
The authors declare no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
About this article
Cite this article
Ekino, T., Kirino, H., Kanzaki, N. et al. Ultrastructural plasticity in the plant-parasitic nematode, Bursaphelenchus xylophilus. Sci Rep 10, 11576 (2020). https://doi.org/10.1038/s41598-020-68503-3