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Citation Information : Journal of Nematology. Volume 50, Issue 4, Pages 459-472, DOI: https://doi.org/10.21307/jofnem-2018-042
License : (PUBLISHER)
Published Online: 03-December-2018
In this study juvenile stages of
The nematode species Bursaphelenchus crenati Rühm, 1956 belonging to Sexdentati group was originally described from the galleries of Hylesinus crenati Fabricius in bark of ash Fraxinus excelsior L. in Bavaria, Germany and later was re-described by Gu et al. (2017) using the populations from Poland. This species was recently detected in several localities of Russia and Belarus (Ryss et al., unpublished). Transmissive dauer juveniles of B. crenati were found in senior larvae, pupae, and female imagoes of the beetle species. From these dauer juveniles, adult nematodes were grown in cultures of the spore-less strain of the fungus Botrytis cinerea on the 2% potato dextrose agar (PDA). All records revealed associations of B. crenati with wilt symptoms of the ash Fraxinus excelsior. In the same beetle vector, the fungi belonging to the family Ophiostomataceae were extracted and cultured in 2% PDA and 2% MEA (malt agar). Nutrition relations between the nematodes and fungi were established, it has been shown that the nematode population grown in slow rate (5 wk) comparing to that culture with Botrytis cinerea (7 d at room temperature). To study a functional role of different juvenile stages of B. crenati in the adaptations to vector and plant and fungus host, as well as in survival of winter seasonal stress, the identification of the life cycle stages would be necessary.
Special attention was given to observe the genital primordium, because according to previous studies this organ is most useful in identification of stage and sex of juveniles (Hirschmann, 1962, 1971; Hirschmann and Triantaphyllou, 1968; Ryss, 1981, 1988; Ryss and Chernetskaya, 2009, 2010; Ryss and Polyanina, 2017). The purpose of this research is to characterize the juvenile stages of the species.
The bark and wood samples with the beetle bark tunnels of Hylesinus crenatus Fabricius, in the inner bark layer and sapwood in wilted ash tree, Fraxinus excelsior L. were collected by A.V. Petrov in the forestry plantation “Tellermanovskoye Opytnoye Lesnichestvo ILAN RAS,” Tellermanovsky settlement, Gribanovsky District, Voronezh Oblast, Russia, June 2017, in a location with the GPS. 51°21′46.8″N 42°02′40.3″E. Samples were sent to Zoological Institute RAS. In the laboratory, the nematode Bursaphelenchus crenati dauers were collected from beetle imagoes, where they formed semi-dry spot-like assemblages (“nematangia”) on the inner surface of elytra.
Nematodes were multiplied from dauers in the sterile laboratory cultures on the lawn of spore-less strain of the fungus Botrytis cinerea, inoculated in the 2% potato dextrose agar media in petri dish (Ryss, 2015). Over 2 wk after 20 nematode specimen inoculation, nematodes occupied all the petri dish space (diam. 6 cm), devastating the fungus lawn, showing active oviposition and population growth. To maintain the nematode isolate, approximately 100 nematode specimens were transferred to the fresh culture of B. cinerea, multiplying population occupied all the petri dish space in 5 to 7 d at 20 to 22°C.
To study the morphology, nematodes were washed out from petri dish with 1 ml of distilled water and fixed in hot TAF as described by Ryss (2017b). Over 2 d after fixation, nematodes were stained with acetic orcein or methylene blue. A modification of the staining technique (Ryss, 1988; Ryss and Chernetskaya, 2009; Ryss and Polyanina, 2017) was used to stain nematodes with acetic orcein. A 1.5 ml Eppendorf tube with nematodes in TAF was placed in upright position for 10 min to settle nematodes down, and the upper layer of fixative was removed with a syringe up to the two fold level of the nematode suspension height (usually 100 µL). Then 200 µL of 70% acetic acid was added into the tube. After shaking the tube, two drops of saturated acetic orcein solution in 65% acetic acid were added, which is usually used for chromosome staining. The mixture was kept overnight for staining procedure and then a drop of staining suspension was placed into the glycerin in a cavity slide. The stained nematodes picked out by needle and transferred to 5 µL drop of glycerin.
To stain with methylene blue, the nematodes were fixed in TAF and processed to glycerin according to Ryss (2017a). Then, nematodes were picked out and transferred from glycerin suspension into a small 1 µL drop of glycerin on a glass slide, and a 5 µL drop of saturated methylene blue water solution was added from above. The drop was placed for 12 hr in open air at room temperature for water evaporation and staining. Then nematodes were transferred in another small glycerin drop on glass slide, heated softly and quickly for stain contrasting and differentiation.
Stained nematodes were picked out with needle under a stereomicroscope and placed in a minute drop of glycerin on a microscopic slide. The drop was covered with 20 × 20 mm coverslip. The drop was spread under coverslip to press nematodes to obtain flattened worm bodies for detailed study of nematode genital primordia structures. After drop spreading, coverslip was sealed with nail polish along borders. All stained nematodes in slides were studied, measured, and photographed under the automated microscope system Leica DM5000 B with differential interference contrast (DIC) and with the Leica DFC320 (R2) digital camera with Leica DFC Twain Software for PC and Leica IM50 Image Manager for PC. All measurements were made using the software ImageJ 1.48 v (National Institute of Health, USA, http://imagej.nih.gov/ij).
Measures: L: body length; BW: body width; pharynx length till the pharyngo-intestinal valve; pharynx length till the end of the pharyngeal gland lobe; tail length; tail width at anus (ABW); GP: genital primordium length; GW: genital primordium width; spicule length along arc; hyaline part of tail in male juveniles (future bursal flap of male). Ratios: a: body length divided to body width; b: body length divided to pharynx length till the pharyngo-intestinal valve; b’: body length divided to pharynx length till the end of pharyngeal glands; c: body length divided to tail length; c’: tail length divided to tail width; V or (V): distance from anterior body end to vulva, or to the center of the genital primordium, divided to the body length, in %; GP/GW: genital primordium length divided to its width; GP/L: genital primordium length divided to body length, in %.
Because the bodies were flattened, the re-calculation of the body and tail diameter was used: the flattened width was considered as a half of the circle (πR or pi R), thus the diameter (2 R) was calculated as the flattened body width (BW) × (2/pi) = 0.64 × BW.
The stained nematodes and eggs were studied. The first molt took place inside the egg shell; thus during hatching the second stage juvenile (J2) broke the egg-shell and came out (Fig. 1E,F,G).
Other stages may be classified as J2, J3, J4, and adult males and females, separated by three molts. Molts were detected by cuticle separation from the body at body extremities, the head and tail tips. The sex of J3 and J4 may be recognized, whereas it cannot be determined in J2. The stage and sex of the juvenile may be determined in the genital primordium structure and its position in the body. The body size and the ratio between genital primordium length and the body are useful to identify the stage. Genital primordia in every stage and sex are described and illustrated below.
The genital primordium consists of four cells: two large germinal cells in the central part and two small somatic nuclei at extremities (Fig. 2A). Primordium is in the middle of the intestine part of the body (Fig. 4).
In the genital primordium, 3 to 5 germinal cells in the posterior part and 12 somatic nuclei in anterior part (Fig. 2G,H) are present. In addition, a single somatic apical nucleus at every extremity of the primordium is found. The primordium is situated in the middle of the intestine part of the body. Cloaca primordium is present as a dense agglomeration of somatic nuclei around the rectum (Fig. 2I,J).
In the genital primordium, 2 large germinal cells in the anterior part and 12 somatic nuclei in the posterior primordium part are present together with a single somatic apical nucleus on every extremity of the primordium (Fig. 2C,D). A circle of six somatic nuclei interrupt the ventral hypodermal cord chain of somatic nuclei. It is a primordium of vulva.
Differences of male juvenile of the third stage from female juvenile are manifested by the presence of cloaca primordium and the position of germinal cells in the posterior part of the primordium. In female juveniles, germinal cells are located in the anterior part of the primordium. Third stage juveniles differ from J2 in the number of cells of the primordium (more than 10 vs 4 in J2). From J4 juveniles, the third stage juveniles differ by the length of primordium occupying 6% of the body length or less vs 11% or more in J4 and adults.
Cloaca primordium is massive, with numerous somatic nuclei around rectum and a transparent cavity anterior to rectum with rudimentary spicules. The genital primordium is distinctly divided into an anterior germinal part of 30 to 60 large cells and a somatic part of two rows of somatic nuclei with 15 to 16 nuclei in every row. The somatic part is not divided into sections (Fig. 3G). Genital primordium occupies 11 to 25% of the body length (Fig. 4). Tail tip with narrowly conical 6 (4–9) µm hyaline zone is curved ventrally, corresponding to bursal flap of adult male.
Fourth stage female juvenile (Figs. 3,5; Table 1). Genital primordium is divided into the anterior germinal part, consisting of 20 to 30 large cells and the posterior somatic part consisting of more than 60 somatic nuclei. On the ventral side of body wall in the middle of somatic primordium part, the lens-like invagination surrounded by massive structure attached to the ventral body wall, is distinct. It is the vulva primordium. Genital primordium occupies 16 to 33% of the body length (Fig. 5).
Differences between male juveniles of the fourth stage from female juveniles are manifested by the presence of cloaca primordium. In female juveniles, the lens-like transparent vulva primordium at the level of uterus primordium part is present. In J4 male juveniles, the vulva primordium is absent. The fourth stage juveniles differ from J3 in the larger genital primordium (11% of body length or more vs 6% or less in J3) and in numerous cells of the primordium: more than 50 in J4 vs 20 or less in J3. From adult nematodes, J4s differ in the absence of copulative organs vs spicules or vulva is present in adult nematodes.
Staining did not reveal any new morphological features in addition to detailed description (Gu et al., 2017).
Molting individuals are characterized by the separation of molting cuticle at extremities: head and tail tip (Figs. 3C, 6B,D,E,H,J). The genital primordium structure of molting specimens is intermediate between structures of primordia described above for the juveniles of J2 to J4 stages.
In the center of primordium, 3 to 4 mitoses of somatic nuclei are distinct between germinal cells (Fig. 6A).
The genital primordium is initially straight, then transforming to loop-shaped (Fig. 6F,G), with the anterior apex and both somatic and germinal parts reversed posteriorly; the somatic part is set off by constriction from germinal part and its tip is curved hook-like (Fig. 6G). The cloaca primordium is enlarged with a transparent inner cavity anterior to the rectum (Fig. 6H). This molting phase is an indication of the orientation change of the somatic part, which in male juvenile J3 is located anteriorly and in J4 is shifted posteriorly, thus pulling a germinal part to the anterior direction (Fig. 6F,G).
The somatic part of the genital primordium in its posterior part possesses more than 20 somatic nuclei, the germinal part in the anterior part of the primordium with 5 to 10 large germinal cells.
The somatic part of the genital primordium reaches in some specimens the massive cloaca primordium, in which inner cavity transparent spicules outlines are distinct. The bursal flap is not formed, the tail is narrowly conical with 9 µm long hyaline zone.
Vulva primordium is a massive cellular agglomeration with an inner transversal slit, but without opening outside. All sections of the genital system are distinct: the ovary of 30 to 40 germinal cells, the oviduct, empty spermatheca, crustaformeria, and the uterus with inner empty cavity (Fig. 5).
Tabular key to juvenile stages of Bursaphelenchus crenati (Key is given in Table 3).
Individual growth is illustrated in diagrams (Fig. 7) with two main parameters: the body length and the ratio between the genital primordium length and the body length. The elongation of the body is faster after the third stage (Fig. 7A). At the same period, the maximum elongation and differentiation of the genital primordium takes place in both male and female development lines (Fig. 7B). Juvenile body increases between molts and within developmental stages, which is evident in the size ranges of every stage (Tables 1,2).
The dauer juveniles have the same genital primordium structure and size as J3. The differences are: the slender and straight body and straight long tail vs curved ventrally tail in the J3 of the propagative generation. Lateral field with two closely located bands of equal width (three incisures). Dauers differ in hemispherical continuous head with hyaline inner cap vs set off head and well developed cephalic circular framework. Stylet is very thin capillary tube, conus and basal thickenings not distinct. Median bulb elongated, its length twice as its width, central valve is weakly developed. Pharyngo-intestinal junction is one half of the median bulb length posterior to median bulb, surrounded by nerve ring with excretory pore at the posterior third of median bulb. Pharyngeal glands form very narrow band, gland nuclei indistinct.
In 3 hr after pouring beetle elytra with “nematangia” in water or 0.9% NaCl solution, the J3D juveniles started to molt to J4 juvenile with long genital primordium and well developed stylet and median bulb (Fig. 9).
The embryonic development with hatching of J2 after the first molt within egg shell is the ovoviviparity. The sex of juvenile is distinctly observed from the third stage. It is an indication of the possible physiological sex determination at the second stage of juvenile (Ryss and Polyanina, 2017). The fastest growth and genital primordium differentiation takes place after the J3 to J4 molt. Earlier study of the development of Bursaphelenchus mucronatus kolymensis (Ryss and Chernetskaya, 2009) and B. ulmophilus (Ryss and Polyanina, 2017) gave similar results: the first molt occurring in the egg shell and four stages outside of egg are divided by molts: J2 to J4 and adult worms. The only difference is the dauer juvenile stage. In B. mucronatus it is the fourth stage juvenile (Ryss, 2008), but in B. ulmophilus and B. crenati. dauers are J3D stage. Leaving the beetle surface, the B. crenati dauers start to molt to the fourth stage juvenile with elongated and differentiated genital primordium, just in 0.9% NaCl solution or in water. The same molting just after reviving in water was detected for dauers J3D of B. ulmophilus (Ryss et al., 2015). Difference between the Bursaphelenchus clades in the dauer juvenile stages (J3D or J4D) is the important character for the genus Bursaphelenchus phylogeny and intrageneric taxonomic diagnostics (Ryss and Subbotin, 2017). Dauers play the important role in the deciduous woody plant wilt spread, as the infective juveniles vectored by insects (Polyanina et al., 2016). The stage of dauers in nematode life cycles and their adaptations to insect vectors is of special interest.
The authors are grateful to the State Academic Programs FSR: АААА-А17-117030310322-3, АААА-А17-117080110040-3 and the grant RFBR 17-04-00360a. The authors are grateful to the forest entomologist Dr. Alexander V. Petrov (ILAN RAS) for wood and bark samples with tunnels of the bark beetle Hylesinus crenatus Fabricius.