**Taxonomy and Identification of Principal Foliar Nematode Species (***Aphelenchoides* **and** *Litylenchus***)**

### **Zafar Handoo \*, Mihail Kantor and Lynn Carta**

Mycology and Nematology Genetic Diversity and Biology Laboratory, USDA, ARS, Northeast Area, Beltsville, MD 20705, USA; mihail.kantor@usda.gov (M.K.); lynn.carta@usda.gov (L.C.)

**\*** Correspondence: zafar.handoo@ars.usda.gov

Received: 25 September 2020; Accepted: 2 November 2020; Published: 4 November 2020

**Abstract:** Nematodes are Earth's most numerous multicellular animals and include species that feed on bacteria, fungi, plants, insects, and animals. Foliar nematodes are mostly pathogens of ornamental crops in greenhouses, nurseries, forest trees, and field crops. Nematode identification has traditionally relied on morphological and anatomical characters using light microscopy and, in some cases, scanning electron microscopy (SEM). This review focuses on morphometrical and brief molecular details and key characteristics of some of the most widely distributed and economically important foliar nematodes that can aid in their identification. *Aphelenchoides* genus includes some of the most widely distributed nematodes that can cause crop damages and losses to agricultural, horticultural, and forestry crops. Morphological details of the most common species of *Aphelenchoides* (*A. besseyi*, *A. bicaudatus*, *A. fragariae*, *A. ritzemabosi*) are given with brief molecular details, including distribution, identification, conclusion, and future directions, as well as an updated list of the nominal species with its synonyms. *Litylenchus* is a relatively new genus described in 2011 and includes two species and one subspecies. Species included in the *Litylenchus* are important emerging foliar pathogens parasitizing trees and bushes, especially beech trees in the United States of America. Brief morphological details of all *Litylenchus* species are provided.

**Keywords:** foliar nematodes; taxonomy; *Aphelenchoides*; *Litylenchus*

#### **1. Introduction**

Foliar nematodes are mostly pathogens of ornamental crops in greenhouses, nurseries, and forest trees, as well as field crops [1]. Foliar nematodes include several nematode genera among which *Aphelenchoides*, *Anguina*, *Ditylenchus,* and *Litylenchus*. Foliar nematodes have been documented as associated with more than 1100 different species of plants, belonging to 126 botanical families, to include dicots, monocots, gymnosperms and angiosperms, ferns and mosses [2]. *Aphelenchoides,* as well as nematodes of genus *Litylenchus*, are phytoparasites known to infect leaves, stems, and buds [3]. The damage caused by the foliar nematodes can cause marketability problems in ornamentals because they interfere with the appearance of the plant or they can reduce yield in food crops [2].

#### **2. General Techniques**

For morphological observation, adult specimens of foliar nematodes can be extracted from fresh leaves. The best method for extracting nematodes from fresh leaves is by using the Baermann Funnel method. Another simple extraction method of nematodes from rice seeds was described by Hoshino and Togashi [4]. They cut the rice seeds longitudinally in two, then transferred the pieces into single plastic pipette tips, which were placed upright in glass vials with water. The pipet tips are transferred to new vials 2, 4, 8, and 24 h later the rate of nematode extraction can be observed. Nematodes are transferred to Syracuse watch glasses and counted. The final step includes dissection of seeds and counting the remaining nematodes after additional 24 h.

For light microscopic observation, fresh specimens are fixed using different methods, such as the ones described by Golden [5] and Hooper [6]. Another method of fixing nematodes was described by Ryss et al. [7] in which nematodes are placed into cold 4% formalin and 1% glutaraldehyde in 0.01 M phosphate buffer at pH 7.3, and then stored at 48 ◦C for light and scanning electron microscopy (SEM).

For SEM observations, nematode specimens can be fixed in phosphate-buffered aldehyde and transferred to special chambers [8], rinsed for 15 min in distilled water, transferred for 2 h in 1% aqueous osmium tetroxide, rinsed again in distilled water and dehydrated in increasing concentrations of ethanol (10% to 100%) in 10% increments for 30 min each, followed by three changes of 100% ethanol. Alcohol is removed using a critical point dryer and the dried specimens stored under vacuum over silica gel. Dried specimens can be mounted on double-sided adhesive tape placed on SEM stubs, sputter-coated with 30 nm of gold, and photographed [7]. To observe the nematode stylets, one individual (alive) specimen is placed in a 1 µL drop of 45% lactic acid on a 12-mm-round, glass cover slip. A small sliver of a broken cover slip, approximately 1 × 3 mm, is placed over the specimen and pressure is applied to it with a needle until the nematode ruptures and the stylet and guiding apparatus extrude. After 24 h, small triangles of filter paper are applied to the edge of the broken sliver to remove the lactic acid, which is exchanged with 2% formalin, followed by three changes of 50% ethanol. The sliver is then floated by adding 50% ethanol and removed with forceps. Stylets adhering to the glass cover slips are air-dried and prepared for SEM as described previously.

Other methods to prepare nematodes for low temperature SEM observations were described by Kantor et al. [9] and Carta et al. [10]. Nematodes can be placed in 1.5 Eppendorf tubes filled with a fixative composed of 2% Paraformaldehyde, 2.5% Glutaraldehyde, 0.05 M Na Cacodylate, and 0.005 M CaCl<sup>2</sup> for at least 12 h. After 12 h, specimens are rinsed in distilled water and individual nematodes placed onto ultra-smooth, round (12 mm diameter), carbon adhesive tabs (Electron Microscopy Sciences, Inc., Hatfield, PA, USA) secured to 15 mm × 30 mm copper plates. The nematode specimens are frozen conductively, in a Styrofoam box, by placing the plates on the surface of a pre-cooled (−196 ◦C) brass bar whose lower half is submerged in liquid nitrogen. After 20–30 s, the brass plate containing the frozen sample is transferred to the Quorum PP2000 cryo transfer system (Quorum Technologies, East Sussex, UK), attached in this case to an S-4700 field emission scanning electron microscope (Hitachi High Technologies America, Inc., Dallas, TX, USA). The specimens are freeze- etched inside the cryotransfer system to remove any surface contamination (condensed water vapor) by raising the temperature of the stage to −90 ◦C for 10–15 min. Following etching, the temperature inside the chamber is lowered below −130 ◦C, and the specimens coated with a 10 nm layer of platinum using a magnetron sputter head equipped with a platinum target. The specimens are transferred to a pre-cooled (−130 ◦C) cryostage in the SEM for observation. An accelerating voltage of 5 kV is used to view the specimens.

DNA extraction from live specimens can be performed using the freeze-thaw lysis with a single live nematode in a 0.2 mL PCR tube containing 25 µL of extraction buffer (10 mM Tris pH 8.2, 2.5 mM MgCl2, 50 mM KCl, 0.45% TWEEN 20 and 0.05% gelatin). Next, the PCR tube is submerged in liquid nitrogen for 10 to 15 s and then placed at 95 ◦C for 2 min in a thermal cycler. The tube is submerged one more time in liquid nitrogen for 10 to 15 sec and then slow-thawed at room temperature. After thawing, the sample is lysed with 1 µL of proteinase K (800 U/mL, Sigma-Aldrich, St. Louis, MO, USA) at 60 ◦C for 60 min, followed by 95 ◦C for 15 min to deactivate the proteinase K. It is recommended to use at least three single nematodes for the individual DNA extraction. The lysates can be stored at −20 ◦C until needed [11,12]. After extraction, the DNA fragments can be amplified using SSU rDNA (18S), D2D3 (28S) expansion region of the LSU rDNA and cytochrome oxidase subunit 1 of the mitochondrial DNA (mtCOI) markers [13]. The primers used for the 18S fragment amplification are 1813F (CTGCGTGAGAGGTGAAAT) and 2646R (GCTACCTTGTTACGACTTTT) and were first published by Holterman et al. [14]. Primers used for the amplification of the 28S region are D2A (ACAAGTACCGTGAGGGAAAGTTG) and D3B (TCCTCGGAAGGAACCAGCTACTA) [15]. The mtCOI fragment can be amplified using COI-F1(CCTACTATGATTGGTGGTTTTGGTAA TTG) and COI-R2 (GTAGCAGCAGTAAA ATAAGCACG) primers [16].

### **3. Genus** *Aphelenchoides* **Fischer, 1894**

After Goodey [17] the genus *Aphelenchoides* Fischer, 1894 is characterized as follows:


A polytomous key was developed and tested on 14 populations by Hockland [18] and the primary key characters were identified as:


A more detailed characterization of the genus was first given by Allen [19]:


Female and male tail never elongate filiform. A recent characterization of *Aphelenchoides* was given by Wheeler and Crow [20]:


According to Hunt [21] members of the *Aphelenchoides* genus can be diagnosed by the following morphological characteristics:


### **4. Genus** *Aphelenchoides* **Fischer, 1894**

### Emended Diagnosis [22]

These nematodes are small and slender, averaging around one millimeter in length and a width less than 20 microns. One characteristic of thee Aphelenchidae nematodes family is that they have a larger median bulb as compared to other plant parasitic nematodes in the order Tylenchida. The dorsal esophageal gland orifice connects to the esophageal lumen at the base of the stylet in most plant-parasitic nematodes, but in Aphelenchida, this duct empties into the esophageal lumen within the median bulb. In *Aphelenchoides*, males are more common, and they reproduce primarily by amphimixis. In most species, the vulva of the female is located near 2/3 the body length from the anterior. Females have a single, prodelphic (anteriorly outstretched) ovary and a post-uterine sac, while males have prominent, thorn-shaped spicules (paired, cuticularized copulatory structures). There is a considerable variation in the shape of the tail terminus within populations of species of the genus *Aphelenchoides.* The tail terminus can be used to divide *Aphelenchoides* species into four groups [22]. The four groups are:


### **5. Systematic Position**

The number of valid nominal species in the Aphelenchoidea is still debatable. However, modern molecular technology may help solve this problem soon. According to Hunt [23] there are 453 'valid' species in Aphelenchoidea, of which 33 belong to the Aphelenchidae and 420 to the Aphelenchoididae. From Aphelenchoididae family, *Aphelenchoides* genera has the most species, namely 153 [23]. A more recent 2015 study conducted by Sánchez-Monge et al. [2] assigned approximately 200 species to the genus. However, after conducting a through literature review, the authors have identified 182 valid nominal species assigned to the *Aphelenchoides* genus.

#### **6. Diagnostic Characters**

Some diagnostic characteristics of *Aphelenchoides* are presented below [3]:


### **7. Genus Synonyms**

Emended list of *Aphelenchoides* species and synonyms:

### **Type species:**

1. *A. kuehnii* Fischer, 1894 = *A.* (*Aphelenchoides*) *kuehnii* Fischer, 1894 (Filipjev, 1934)

### **Other species:**


#### **8. Principal Species**

The following four species have been selected for further discussion because of their commonality, economic importance, and/or worldwide distribution:


Each species is illustrated below (Figures 1–11). Data were obtained from various sources, including Allen [19]; Christie [24] De Jesus et al. [25], 2016; Xu et al. [26]; Siddiqi [27–30]; Shahina [22]; Siddiqui and Taylor [31]; Jen et al. [32]; Khan et al. [33]; Chizhov et al. [34]; Zhao et al. [35]; Khan et al. [36]; Hunt [21], Kanzaki et al. [37] 2019 and Carta et al. [11], and original descriptions and/or re-descriptions.

Because *Aphelenchoides besseyi* Christie, 1942, *Aphelenchoides fragariae* (Ritzema Bos, 1891) Christie, 1932, *Aphelenchoides ritzemabosi* (Schwartz, 1911) Steiner and Buhrer*, Aphelenchoides bicaudatus*(Imamura, 1931) Filipjev and Schuurmans Stekhoven, 1941 are of major economic importance and widely distributed all over the world, they will be discussed in detail.

#### **9. Rice White-Tip Nematode (***Aphelenchoides besseyi* **Christie, 1942)**

*Aphelenchoides besseyi* (Figure 1) is an economically important pathogen of rice and has been reported from many countries. However, it is not commonly found in ornamentals [38,39], with the exception of some reports on tuberose [36], begonia [40], gerbera [41], hydrangea [27], tuberose [42], and even on bird nest fern [43]. *A. besseyi* distribution is mostly in warmer climates, whereas *A. ritzemabosi* and *A. fragariae* are more commonly associated with temperate climates, while found in both tropical and temperate localities [1].

**Figure 1.** *Aphelenchoides besseyi* Christie (**A**) female; (**B**) female head end; (**C**) female en face view; (**D**) Lateral field; (**E**,**F**) variation in female esophageal bulb and position of excretory pore with respect to nerve ring; (**G**) male anterior end; (**H**) female tail termini showing variation in shape mucro; (**I**–**K**) male tail ends; (**L**–**N**) variation in post-vulval uterine sac (**B** and **D** original, the rest after Fortuner, **Figure 1.** *Aphelenchoides besseyi* Christie (**A**) female; (**B**) female head end; (**C**) female en face view; (**D**) Lateral field; (**E**,**F**) variation in female esophageal bulb and position of excretory pore with respect to nerve ring; (**G**) male anterior end; (**H**) female tail termini showing variation in shape mucro; (**I**–**K**) male tail ends; (**L**–**N**) variation in post-vulval uterine sac (**B** and **D** original, the rest after Fortuner, 1970) after Franklin and Siddiqi [27]. Courtesy of *Commonwealth Institute of Helminthology*.

1970) after Franklin and Siddiqi [27]. Courtesy of *Commonwealth Institute of Helminthology*. Measurements

After Christie [24].

Females (*n* = 10): length = 0.66–0.75 mm; a = 32–42 (width = 17–22); b = 10.2–11.4 (esophagus = 64–68 µm); c = 17–21 (tail = 36–42 µm); V = 68–70%.

Males (*n* = 10): length = 0.54–0.62 mm; a = 36–39 (width = 14–17 µm); b = 8.6–8.8 (esophagus = 63–66 µm); c = 15–17 (tail = 34–37 µm); T = 44–61%

After Allen [19].

Females: length = 0.62–0.88 mm; a = 38–58; b = 9–12; c = 15–20; V = 66–72

Males: length = 0.44–0.72 mm; a = 36–47; b = 9–11; c = 14–19; T = 50–65%.

After De Jesus et al. [20]

Females: length = 0.65–0.75 mm; a = 42.8–49; c = 15.6–17.5; c' = 4.0–4.5.

Males: length = 0.65–0.75 mm; a = 42.8–49; c = 15.6–17.5; c' = 4.0–4.5; spicule = 14.1–18.3 µm. After Xu et al. [44]

Body length (*n* = 11) = 0. 656 ± 18.5 (0.546–0.729) mm; body width = 14.4 ± 0.32 (12.4–15.9) µm; pharynx = 124 ± 2.53 (111.0–137.8) µm; stylet = 12.5 ± 0.21 (10.6–13.3) µm; median bulb end to anterior end 69.7 ± 1.07 (65.7–75.3) µm; tail length 36.9 ± 0.38 (35.3–38.9) µm; anus/cloacal width 9.27 ± 0.47(7.5–12.1) µm.

#### Description

Female: female specimens share a slender body, slightly arcuate ventrally when relaxed, anteriorly tapering from the level of esophageal glands to the head, which is one half of the body width. Four lateral lines (occasionally six noted) are present in the lateral field (Figure 2). In en face view, the pore-like amphids are on outer margins of lateral lips; four papillae, one on each submedian lip (Figure 2). Lip region is non-striated and set off from body by a constriction as wide as or slightly wider than adjacent body; labial framework weakly developed; cheilorhabdions well sclerotized. Basal knobs of spear distinct, 2 µm across. Procorpus cylindrical; median esophageal bulb one and a half times to twice as long as wide, with refractive valvular apparatus slightly posterior to center. Esophageal glands extending over intestine 5 to 8 body widths. Excretory pore at 58 to 83 µm from anterior end, level with or slightly anterior to nerve ring. Hemizonid distinct in specimens from rice seeds (but not from cultured specimens), 11 to 15 µm behind excretory pore; hemizonion 20 to 30 µm behind hemizonid, usually difficult to see. Tail straight, slender, regularly tapering to a narrowly rounded end, 3–5 to 5 anal body diameters long; mucro with 3 to 4 processes. Ovary not extending to esophageal glands; oocytes in 2 to 4 rows; spermatheca very conspicuous, elongate oval, full of rounded sperms showing a central nucleolus usually surrounded by a circle of black dots of unknown nature. Post-vulval uterine sac short, slender and extending up to one fourth of the distance from vulva to anus (2.5 to 3 body diameters) often found empty and collapsed but more conspicuous and rounded in nematodes from cultured specimens. Vulval lips slightly protruding after Fortuner [45].

Male: tail end usually curved by 90◦ (a greater curvature has also been found) in specimens killed in 3% formaldehyde; mucro of diverse shape, with 2 to 4 processes. Spicule length between 17 to 21 µm along dorsal limb. Different morphometric characters, such as the shape of the head, the position of the excretory pore in relation to the nerve ring and the shape and length of the post-vulval uterine sac were found to be variable between populations [45].

#### Distribution

According to Devran et al. [46], *A. besseyi* was on the quarantine lists of nine countries in 1982 and up to 70 countries in 2002. Centre for Agriculture and Biosciences International (CABI), Invasive Species Compendium [47] lists *A. besseyi* being present in 75 countries around the world. The quarantine pests lists *A. besseyi* as the second most prevalent nematode after *Globodera rostochiensis* [39].

*Aphelenchoides bicaudatus* (Imamura, 1931) Filipjev and Schuurmans Stekhoven, 1941 *Aphelenchoides bicaudatus* (Imamura, 1931) Filipjev and Schuurmans Stekhoven, 1941 was originally described from a paddy field in Japan and previously considered a primarily mycophagous species. Since then, it has been reported to parasite more than 200 plant species [31,48].

*Plants* **2020**, *9*, x FOR PEER REVIEW 12 of 34

**Figure 2.** SEM photomicrographs of *Aphelenchoides besseyi* female (**A**) head end; (**B**–**C**) lateral fields; (**D**) tail end, after Khan et al. [36]. Courtesy of *Journal of Nematology*. **Figure 2.** SEM photomicrographs of *Aphelenchoides besseyi* female (**A**) head end; (**B**,**C**) lateral fields; (**D**) tail end, after Khan et al. [36]. Courtesy of *Journal of Nematology*.

Distribution Measurements

> According to Devran et al. [46], *A. besseyi* was on the quarantine lists of nine countries in 1982 After Imamura [49].

and up to 70 countries in 2002. Centre for Agriculture and Biosciences International (CABI), Invasive Species Compendium [47] lists *A. besseyi* being present in 75 countries around the world. The quarantine pests lists *A. besseyi* as the second most prevalent nematode after *Globodera rostochiensis*  Female (*n* = 18): L = 0.38–0.47 (0.43) mm; a = 31.3–31.7 (31.5); b = 6.8–8.4 (7.4); c = 9.4–12.6 (10.6); V%= 61.7–90.2 (0.4).

[39]. After Siddiqui and Taylor [31].

*Aphelenchoides bicaudatus* (Imamura, 1931) Filipjev and Schuurmans Stekhoven, 1941 *Aphelenchoides bicaudatus* (Imamura, 1931) Filipjev and Schuurmans Stekhoven, 1941 was originally Female (*n* = 50): L = 0.41–0.55 (0.46) mm; a = 25–31 (28.0); b = 7.3–9.6 (8.2); c = 9.8–13.7 (11.4); V% = 65–70 (67.5); stylet = 10–12 (11.2) µm.

described from a paddy field in Japan and previously considered a primarily mycophagous species. Since then, it has been reported to parasite more than 200 plant species [31,48]. Male: L = 0.385 mm; a = 22.6; b = 7.5; c = 11.4; stylet = 10 µm. After Jen et al. [32].

Measurements After Imamura [49]. Female (*n* = 18): L = 0.38–0.47 (0.43) mm; a = 31.3–31.7 (31.5); b = 6.8–8.4 (7.4); c = 9.4–12.6 (10.6); V%= 61.7–90.2 (0.4). After Siddiqui and Taylor [31]. Female (*n* = 50): L = 499.12 ± 67.95 (0.376–0.637) mm; maximum body width = 15.24 ± 2.69 (11–22) µm; a = 33.03 ± 2.42 (27.00–38.64); b = 9.0 ± 0.7 (7.5–10.0); b' = 5.13 ± 0.76 (3.61–7.94); c = 11.94 ± 0.93 (10.16–14.80); c' = 5.41 ± 0.56 (4.13–7.14); V% = 68.53 ± 1.20 (64.90–71.83); stylet = 10.38 ± 0.63 (9–12) µm; length of post-uterine sac expressed as % of length from vulva to anus = 18.98 ± 4.54 (9.23–33.80) µm.

Female (*n* = 50): L = 0.41–0.55 (0.46) mm; a = 25–31 (28.0); b = 7.3–9.6 (8.2); c = 9.8–13.7 (11.4); V% After Israr et al. [50].

= 65–70 (67.5); stylet = 10–12 (11.2) µm. Male: L = 0.385 mm; a = 22.6; b = 7.5; c = 11.4; stylet = 10 µm. After Jen et al. [32]. Female (*n* = 50): L = 499.12 ± 67.95 (0.376–0.637) mm; maximum body width = 15.24 ± 2.69 (11– 22) µm; a = 33.03 ± 2.42 (27.00–38.64); b = 9.0 ± 0.7 (7.5–10.0); b' = 5.13 ± 0.76 (3.61–7.94); c = 11.94 ± 0.93 (10.16–14.80); c' = 5.41 ± 0.56 (4.13–7.14); V% = 68.53 ± 1.20 (64.90–71.83); stylet = 10.38 ± 0.63 (9–12) µm; length of post-uterine sac expressed as % of length from vulva to anus = 18.98 ± 4.54 (9.23–33.80) µm. After Israr et al. [50]. Female (*n* = 2): L = 0.36 mm; a = 30.1, 32.7; b = 8.8,7.2; b' = 5.6, 5.8; c = 11.3, 12; c' = 2.9, 3.7; V% = 66.8–67.2; G1% = 25, 26.2; body diameter 12, 12,5; stylet = 10, 11 µm, median bulb length 10, 10 µm; median bulb width 7, 8 µm; median bulb length/ width 1.4, 1.3; distance anterior end to distal end of median bulb 51, 52 µm; anterior end to excretory pore 50, 51 µm; anterior end to nerve ring 55, 56 µm; anterior end to vulva 242, 248 µm; ovary length 95, 84 µm; distance from vulva to anus 85, 84 µm; post uterine sac length 24, 22 µm; post uterine sac length/vulva anus distance% 22.4, 24; esophageal length 90, 92 µm; esophageal intestinal junction 62, 64 µm; tail length 31, 30 µm; anal body width 31, 30 µm, anal body width 11, 8.

Male (*n* = 1): L = 0.40 mm; a = 30.7; b = 4.3; b' = 6.2; c = 10; c' = 3.9; T% = 52; body diameter 13; stylet = 10 µm, median bulb length 12 µm; median bulb width 9 µm; median bulb length/width 1.3; distance anterior end to distal end of median bulb 54 µm; anterior end to excretory pore 62 µm; anterior end to nerve ring 60 µm.

#### Description

Female: have a slender body, attenuated slightly anteriorly, and more prominently toward posterior end (Figure 3). When relaxed by gentle heat the position of the body is straight and only the tail region is slightly curved. Cuticle is finely striated, with annuli measuring between 0.47–0.58 µm wide and 0.39–0.51 µm thick. Lateral field has two lateral lines. Head distinctly set off from body. Lip region rounded, offset with no annules. Stylet weak, with small basal swellings. Metacorpus rounded, occupying approximately 73% of body width. Nerve ring is located about 1/2 body width behind metacorpus. Excretory pore opposite anterior margin of nerve ring. Vulva a transverse slit and slightly protruding, about 66% of body length from anterior end. Post-vulvar uterine sac extending for one-fifth of distance from vulva to end of tail. Rectum prominent, straight, near ventral body wall, and in length approximately three-fourths of anal body width. Tail gradually tapering to terminus, which is unevenly bifurcated with one prong longer than the other. *Plants* **2020**, *9*, x FOR PEER REVIEW 14 of 34

**Figure 3.** *Aphelenchoides bicaudatus* (Imamura) Filip. and Sch. Stek (**A**–**D**) Larvae, first of fourth stages; (**E**) egg; (**F**) cross section of female at mid-body; (**G**) lateral field; (**H**) face view; (**I**) framework around oral opening; (**J**) esophageal region in dorsal view; (**K**) whole female; (**L**) male tail after Siddiqi [28]. **Figure 3.** *Aphelenchoides bicaudatus* (Imamura) Filip. and Sch. Stek (**A**–**D**) Larvae, first of fourth stages; (**E**) egg; (**F**) cross section of female at mid-body; (**G**) lateral field; (**H**) face view; (**I**) framework around oral opening; (**J**) esophageal region in dorsal view; (**K**) whole female; (**L**) male tail after Siddiqi [28]. Courtesy of *Commonwealth Institute of Helminthology*.

Courtesy of *Commonwealth Institute of Helminthology*.

*Plants* **2020**, *9*, 1490

Females of *A. bicaudatus* (Figure 4) can be differentiated from other members of the genus by having an unevenly bifurcated tail tip with prongs of different lengths [51].

Male: extremely rare. *Plants* **2020**, *9*, x FOR PEER REVIEW 15 of 34

**Figure 4.** *Aphelenchoides bicaudatus* (Imamura, 1931) Filipjev and Schuurmans Stekhoven, 1941. (**A**) Entire female; (**B**) neck region; (**C**) female reproductive system; (**D**) female posterior region. Scale bars: (**A**) = 50 µm, (**B**,**D**) = 10 µm, (**C**) = 20 µm after Kim et al. [52]. Courtesy of *Animal Systematics Evolution and Diversity Journal*. **Figure 4.** *Aphelenchoides bicaudatus*(Imamura, 1931) Filipjev and Schuurmans Stekhoven, 1941. (**A**) Entire female; (**B**) neck region; (**C**) female reproductive system; (**D**) female posterior region. Scale bars: (**A**) = 50 µm, (**B**,**D**) = 10 µm, (**C**) = 20 µm after Kim et al. [52]. Courtesy of *Animal Systematics Evolution and Diversity Journal*.

#### Distribution *A. bicaudatus* was recorded in most of the tropical and subtropical regions of the world as well Distribution

as some warmer temperate areas [21]. More specifically, it was reported in the following countries: Australia, Brunei, France, Japan, USA, Russia, Venezuela [28], South Korea [52], Taiwan [32]. *A. bicaudatus* was recorded in most of the tropical and subtropical regions of the world as well as some warmer temperate areas [21]. More specifically, it was reported in the following countries: Australia, Brunei, France, Japan, USA, Russia, Venezuela [28], South Korea [52], Taiwan [32].

#### **10. Strawberry Crimp Nematode (***Aphelenchoides fragariae* **(Ritzema Bos, 1891) Christie, 1932)**

*Aphelenchoides fragariae* was originally described by Ritzema Bos (1891) in specimens recovered from strawberry plants sent to him from England (Figure 5). When compared to all the *Aphelenchoides* species mentioned previously, it has the widest distribution as well as hosts range (more than 600 species), to include ferns, herbaceous perennials and bedding plants [2,33,53]. *A. fragariae* is an ectoand endo-parasite of the above ground parts of a plant, but it can also be mycetophagous [2,21,33]. The nematodes enter the plant leaves through stomata or wounds [1,47]. In the leaves, nematodes feed on mesophyll cells which causes characteristic vein delimited lesions [1,47]. *A. fragariae* survives overwinter in soil, dormant buds, dry leaves, but not in roots [18,47]. Research showed that *A. fragariae* nematodes can tolerate temperature as high as 40 ◦C and as low as −80 ◦C once in leaf tissues [18].

#### Measurements

After Allen [19].

Females: length = 0.45–0.80 mm; a = 45–60; b = 8–15; c = 12–20; V%= 64–71. Males: length = 0.48–0.65 mm; a = 46–63; b = 9–11; c = 16–19; T% = 44–61.

After Franklin [54].

Females: length = 0.552–0.886 (0.796) mm; a = 36–63 (53); body width = 12–17 (15) µm.

Males: length = 0.573–0.864 mm; a = 40–63; body width= 12–17 (14) µm.

After Khan et al. [33].

Females (*n* = 7): length = 0.620–0.895 mm; a = 46.2–64.5; b = 9.0–13.2; c = 13.4–20.3, V% = 66.5–72.2; stylet = 10.0–11.5 µm.

Males (*n* = 7): length = 0.480–0.623 mm; a = 45.7–61.7; b = 9.3–10.8; c = 15.7–18.5, T% = 45.6–60; stylet = 10.0–11.2 µm; spicules = 16.9–19.0 µm.

After Chizhov et al. [34].

Females (*n* = 25): length = 0.525–0.685 (0.579 ± 0.043) mm; a = 37.1–59.8 (48.7 ± 4.8); b = 7.6–9.1 (8.1 ± 0.3); c = 15.2–20.6 (17.0 ± 1.2), c' = 3.6–5.7 (4.7 ± 0.3); V = 65.0–74.0% (69.0 ± 2.0); stylet = 8.0–11.0 (9.0) µm; head region width = 4.0–5.0 µm; head region high = 3.0 µm; distance from anterior end to: medial bulb base = 52.0–64.0 (58.0) µm, nerve ring = 63.0–78.0 (72.0) µm, excretory pore = 68.0–85.0 (76.0) µm and esophageal gland base = 100.0–150.0 (128.0) µm; post uterine sac length = 58.0–98.0 (77.0) µm; tail length = 28.0–40.0 (34.0) µm; body width at vulva level = 10.0–16.0 (12.0) µm and anus level = 6.0–8.0 (7.0) µm.

Males (*n* = 24): length = 0.435–0.562 (0.493 ± 0.037) mm; a = 41.2–54.8 (46.8 ± 3.1); b = 6.5–8.1 (7.2 ± 0.4); c = 15.9–24.1 (18.5 ± 1.8); stylet = 8.0–10.0 (9.0) µm; head region width = 4.0–5.0 µm; head region height = 3.0 µm; spicule length = 10.0–13.0 (12.0) µm; distance from anterior end to: medial bulb base = 52.0–62.0 (57.0) µm and esophageal gland base = 100.0–135.0 (118.0) µm; nerve ring = 68.0–77.0 (71.0) µm; excretory pore = 70.0–82.0 (76.0) µm; testis length = 204.0–289.0 (250.0) µm; maximal body width = 10.0–13.0 (11.0) µm; tail length = 21.0–33.0 (27.0) µm.

#### Description

Body very slender (a = 45–63 µm), straight or arcuate when relaxed. Cuticle marked by fine transverse striae about 0.9 µm apart; lateral field with two incisures, 1/7th of body-width. Cephalic region, smooth, anteriorly flattened with straight to curved side margins, almost continuous with neck contour. Lips without annulation. Stylet slender, approximately 10 µm long, with small but distinct basal knob. Median esophageal bulb well developed, oval. Nerve ring about one body width behind median bulb. Excretory pore level at or close behind nerve ring. Esophageal glands stretched five body widths behind the medium bulb, joining esophagus immediately behind the medium bulb. Tail elongate-conoid, terminus bearing a terminal peg which is simple, spike-like.

Female: vulva a transverse slit, at approximately 64–71% of body. Spermatheca elongate-oval. Posterior uterine sac more than half the vulva-anus distance, often containing sperm. Ovary single, with oocytes in a single row. Tail terminus with a single mucronate points point enlarged at the base.

Male: abundant. Male tail curved to about 45–90 degrees. Three pairs of ventro-submedian copulatory papillae (1st slightly post-anal, 2nd midway, and 3rd near the end). Testis single, outstretched; sperm large-sized, rounded, in a row. Spicules large and prominent, ventrally curved, rose-thorn-shaped, with moderately developed dorsal and ventral processes (apex and rostrum) at proximal end; dorsal limb 14–17 µm long. Male: abundant. Male tail curved to about 45–90 degrees. Three pairs of ventro-submedian copulatory papillae (1st slightly post-anal, 2nd midway, and 3rd near the end). Testis single, outstretched; sperm large-sized, rounded, in a row. Spicules large and prominent, ventrally curved, rose-thorn-shaped, with moderately developed dorsal and ventral processes (apex and rostrum) at proximal end; dorsal limb 14–17 µm long.

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**Figure 5.** *Aphelenchoides fragariae*. (**A**) Female head end. (**B**) Male head end; (**C**) a, female; b male of *A. olesistus* Ritzema Bos, 1893 (= *A. fragariae*); (**D**) a, male; b, posterior portion of female, of *Aphelenchus fragariae* Ritzema Bos, 1891; (**E**) male; (**F**) female; (**G**) female tail; (**H**) lateral field; (**I**) female tail tip (**J**,**K**) male tails. (**L**) Spicules a, drawn from paratypes of Allen (1952); b, from specimens ex *Cornus canadensis* from Surrey, England after Siddiqi [29]. Courtesy of *Commonwealth Institute of Helminthology*.

#### Distribution

*A. fragariae* has a widespread distribution in Europe, Russia, Japan and North America [21]. According to the CABI Invasive Species Compendium [47], *A. fragariae* is currently reported to be present in 37 countries.

#### **11. Chrysanthemum Nematode (***Aphelenchoides ritzemabosi* **(Schwartz, 1911) Steiner and Buhrer)**

*Aphelenchoides ritzemabosi* (Schwartz, 1911) Steiner and Buhrer, also known as the Chrysanthemum foliar nematode, is a common plant-parasite infecting more than 300 plant species, second only to *A. fragariae* [2] in the *Aphelenchoides* genus based on the number plants they parasitize.

#### Measurements

After Allen [13].

Females: length = 0.77–1.2 mm; a = 40–54; b = 10–13; c = 18–24; V% = 66–75. Males: length = 0.70–0.93 mm; a = 31–50; b = 10–14; c = 16–30; T% = 35–64. After Chizhov et al. [29].

Females (*n* = 15): length = 0.768–1.027 (0.916 ± 0.067) mm; a = 43.4–60.5 (51.2 ± 3.7); b = 8.1–9.5 (9.1 ± 0.3); c = 16.8–21.2 (19.3 ± 1.1); c' = 4.0–5.1 (4.6 ± 0.2); V% = 68–71 (69 ± 0.2); stylet = 9.0–11.0 (10.0) µm; head region width = 6.0–7.0 µm; head region height = 3.0 µm; distance from anterior end to: medial bulb base = 71.0–77.0 (74.0) µm; nerve ring= 95.0–108.0 (100.0) µm; excretory pore= 108.0–130.0 (121.0) µm and esophageal gland base = 145–185 (170) µm; postuterine sac length= 105.0–160.0 (134) µm; tail length= 41.0–54.0 (48.0) µm; body width at vulva level = 16.0–23.0 (18.0) µm and anus level= 8.0–12.0 (10.0) µm.

Males (*n* = 15): length = 0.625–0.852 (0.721 ± 0.053) mm; a = 36.9–53.3 (46.3 ± 3.3); b = 6.5–9.4 (7.9 ± 0.6); c = 17.3–22.4 (19.9 ± 1.1); stylet = 9.0–11.0 (10.0) µm; head region width = 6.0–7.0 µm; head region height= 3.0 µm; spicule = 15–18 (16) µm; distance from anterior end to: medial bulb base = 67.0–72.0 (69.0) µm, nerve ring = 85.0–108.0 (93.0) µm; excretory pore = 92.0–118.0 (105.0) µm and esophageal gland base = 156.0–180.0 (169.0) µm; testis length = 353.0–512.0 (442.0) µm; tail length = 34.0–39.0 (36.0) µm.

#### Description (Figure 6)

Female: nematodes with slender body, with fine transverse striae on the cuticle. Four lines present in the lateral field. Lip region set off, wider than neck at base of lips with no annulations. Hexaradiate framework weakly sclerotized. Stylet approximately 12 µm long, with small but well-developed basal knobs. Median esophageal bulb well developed, oval in shape. Nerve ring 1.5 body widths behind median bulb. Excretory pore located behind nerve ring, approximately 0.5–2 body widths posterior to nerve ring. Esophageal glands extending 4 body widths over the intestine, joining esophagus immediately behind median bulb. Oocytes in multiple rows, several in a cross-section at middle of ovary. Posterior uterine branch extending for more than half the vulva-anus distance, usually containing sperms. Tail elongated-conoid. Terminus peg-like armed with two-four small mucronate points pointing posteriorly.

Male: males are common, having a tail curvature at about 180 degrees when relaxed. Testis single. Three pairs of ventro-submedian papillae. First pair adanal, second midway on tail, third near end. Spicules smoothly ventrally curved, the ventral piece without a ventral process at the distal end; dorsal limb 20–22 µm long. Terminus peg-like armed with two-four small mucronate points.

#### Distribution

*Aphelenchoides ritzemabosi* is a major pest of chrysanthemum in Europe, Russia, North America, South Africa, New Zealand, Australia, and Brazil [25]. According to the CABI Invasive Species Compendium [55], *A. ritzemabosi* is currently reported to be present in 35 countries around the world.

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**Figure 6.** *Aphelenchoides ritzemabosi.* (**A**) Female head; (**B**) female; (**C**) female tail ends; (**D**) male tail ends; (**E**) female tail; (**G**) spicules; (**H**) lateral field; (**I**) male tail region. (**A**, **E**, and **F** syntypes; **B**, **C**, and **H** Specimens from chrysanthemum, Stockholm; I Specimen from *Buddleia* leaf, Sussex, England) after Siddiqi [25]. Courtesy of *Commonwealth Institute of Helminthology*. **Figure 6.** *Aphelenchoides ritzemabosi.* (**A**) Female head; (**B**) female; (**C**) female tail ends; (**D**) male tail ends; (**E**) female tail; (**G**) spicules; (**H**) lateral field; (**I**) male tail region. (**A**, **E**, and **F** syntypes; **B**, **C**, and **H** Specimens from chrysanthemum, Stockholm; I Specimen from *Buddleia* leaf, Sussex, England) after Siddiqi [25]. Courtesy of *Commonwealth Institute of Helminthology*.

#### Distribution Identification

Accurate identification of foliar nematodes (*Aphelenchoides* spp.) is crucial for effective disease control. Major efforts should be geared towards rapid and accurate classification of the pathogens so that appropriate control measures could be taken. In addition, timely and accurate diagnosis is also needed to make sound decisions regarding quarantine of imported and exported plant material and commodities. Nevertheless, the identification of foliar nematodes to species level remains a challenging endeavor. The diagnosis and/or relationship between conserved morphology, variable morphometrics, host effects, intraspecific variation, existence of cryptic species, and the ever-increasing number of described species, still vary significantly. To add to the confusion, there is verification of mixed populations and/or detection of rare species which require(s) identification techniques, including morphology of adult females; male, and labial region shape, and stylet morphology; V% age, body length, and shape of tail and tail terminus, and, in some cases, biochemical or molecular methodologies. Because of an increasing number of described species, the value of many of these characters often show large intraspecific variation. Isozyme electrophoresis has discriminated a number of these otherwise cryptic species. Currently used PCR-based molecular methodologies offer hope for a future relying on bigger genebanks that could be used by scientists for a more accurate specie identification. Integrated morphology and molecular approaches are essential to future improved identification of Anguinata nematodes. Detailed diagnostic characters differentiating various species of foliar nematodes have been given by authors such as Allen [19], Hunt [21], Shahina [22].

#### **12. Genus** *Litylenchus* **Zhao, Davies, Alexander and Riley, 2011**

Genus *Litylenchus* Zhao, Davies, Alexander and Riley, 2011 is a new genus with much smaller number of species when compared to *Aphelenchoides* genus. *Litylenchus crenatae* Kanzaki, 2019*, Litylenchus crenatae mccannii* Carta 2020, are emerging foliar pathogens of major economic importance. Nematodes from this genus parasitize trees (*Fagus grandifolia*) and bushes (*Coprosma repens*). *Litylenchus crenatae mccannii* described by Carta et al. [11] seems to be a very aggressive subspecies with devastating effects on beech trees (*Fagus grandifolia*). Even though *Litylenchus crenatae mccannii* was initially found infesting beech trees in Ohio [11], it was also reported in several other states and provinces, to include Pennsylvania, New York, Ontario, Canada [56], Connecticut [57], New Jersey, Rhode Island, and West Virginia (unpublished data).

After Zhao [35] the genus *Litylenchus* Zhao, Davies, Alexander and Riley, 2011 is characterized as follows:


#### **13. Systematic Position**

Based on phylogenetic analyses, *Litylenchus* genus [35] is close to *Subanguina.* However, the two genera have many morphological differences as highlighted below:

• *Litylenchus* genus. does not induce typical galls like *Anguina* and *Nothanguina*;


List of *Litylenchus* species and synonyms:

Type species:

1. *Litylenchus coprosma*

Other species


#### **14.** *Litylenchus coprosma* **Zhao, Davies, Alexander and Riley, 2011**

#### Measurements

After Zhao et al. [35].

Slender female (*n* = 13): L = 743 ± 50 (649–816) µm; a = 55.2 ± 4.0 (51.5–63.3); b = 4.4 ± 0.6 (3.9–5.8); c = 18.7 ± 1.3 (16.3–21.3); V %= 81.5 ± 2.4 (76.5–85.3); stylet = 10.8 ± 0.9 (8.9–11.7) µm.

Obese female (*n* = 15): L = 856 ± 72 (710–940) µm; a = 32.8 ± 3.7 (24.9–37.7); b = 5.1 ± 0.6 (4.2–6.8); c = 19.4 ± 2.5 (15.4–25.0); V%= 82.2 ± 1.6 (78.8–84.7); stylet = 10.9 ± 0.3 (10.2–11.4) µm.

Male (*n* = 11): L = 899 ± 66 (768–994) µm; a = 52.0 ± 4.4 (44.5–60.2); b = 5.4 ± 0.4 (4.8–6.2); c = 21.1 ± 1.9 (18.2–24.1); stylet = 10.5 ± 0.5 (9.7–11.3) µm; spicule= 16.2 ± 0.7 (14.9–17.0) µm.

#### Description

*Litylenchus coprosma* has adult females with two distinct forms, one described as semi-obese (a = 20–40) and the other slender (a = 45–65) (Figures 7 and 8).

Semi-obese female: when killed by heat body is almost straight, semi-obese. Maximum body width is at mid-body. Body cuticle finely striated, almost smooth. Four lines can be observed in lateral field extending almost to tail terminus. Head offset, cephalic framework, and stylet as described for male. Excretory pore located ca 3–3.5 body diameter from anterior, opening near anterior end of terminal bulb, duct with obvious cuticular lining. Hemizonid, pharynx, pharyngeal glands, and pharyngo-intestinal junction as described for male. Nerve ring is located approximately 100 µm from anterior extremity. Deirids and phasmids not seen. Gonads are monodelphic, prodelphic, outstretched, crustaformeria forming a quadricolumella. Oocytes arranged in single row. Oviduct with several cells forming a valve just anterior to elongate, sac-like spermatheca. Vulva located 7–11 anal body diameter anterior to anus (80–85% of body length). Vulval slit occupying almost half body diameter when viewed laterally, vagina almost perpendicular to body wall. Post-uterine sac extending 20–70% of distance from vulva to anus, approximately 2.7 anal body long, sometimes with sperms, lacking cellular relicts of posterior ovary. Rectum difficult to see, anus pore-like, opening in a cuticular depression. Tail approximately 4–5 anal body diameter long, conoid, straight, with a variable tail terminus, may be bluntly rounded, more or less bifurcate, or appear bilobed. Mucro not observed.

Slender female: very similar to the semi-obese females, but slender. Head capsule is a little bit bigger, 59–77% of body diameter at level of stylet knobs compared to the semi-obese females, where the head capsule is between 48–62%. Quadricolumella cells are smaller than in semi-obese female.

Male: when killed, the nematodes assume a smoothly ventrally arcuate shape, body cylindrical, narrowing to a bluntly rounded conoid tail. Body cuticle smooth with three incisures in the lateral field

visible in the region of procorpus increasing to four incisures at mid-body and extending almost to tail tip. Head is set off from the body, smooth, and not annulated. Lightly sclerotized cephalic framework with six sectors. *Plants* **2020**, *9*, x FOR PEER REVIEW 23 of 34

**Figure 7.** *Litylenchus coprosma*. All males in lateral view, except E which is ventral. (**A**) Anterior region; (**B**) pharynx showing median bulb; (**C**) tail with bursa; (**D**) tail showing spicules and variation in shape of tail tip; (**E**) spicules. (Scale bars = 10 µm) after Zhao et al. [35]. Courtesy of *Nematology*. **Figure 7.** *Litylenchus coprosma*. All males in lateral view, except E which is ventral. (**A**) Anterior region; (**B**) pharynx showing median bulb; (**C**) tail with bursa; (**D**) tail showing spicules and variation in shape of tail tip; (**E**) spicules. (Scale bars = 10 µm) after Zhao et al. [35]. Courtesy of *Nematology*.

*robusta* [26].

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**Figure 8.** *Litylenchus coprosma*. All in lateral view, except C, D en face. (**A**) Head of mature, semi-obese female; (**B**) terminal pharyngeal bulb; (**C**) sub-terminal head showing amphidial apertures; (**D**) apical view of head; (**E**) lateral fields at mid-body showing four incisures; (**F**) lateral fields at pharyngeal region showing three incisures; (**G**) vulva and post-uterine sac; (**H**) second-stage juvenile within egg; (**I**) quadricolumella; (**J**) female tail. (Scale bars = 10 µm) after Zhao et al. [35]. Courtesy of *Nematology*. **Figure 8.** *Litylenchus coprosma*. All in lateral view, except C, D en face. (**A**) Head of mature, semi-obese female; (**B**) terminal pharyngeal bulb; (**C**) sub-terminal head showing amphidial apertures; (**D**) apical view of head; (**E**) lateral fields at mid-body showing four incisures; (**F**) lateral fields at pharyngeal region showing three incisures; (**G**) vulva and post-uterine sac; (**H**) second-stage juvenile within egg; (**I**) quadricolumella; (**J**) female tail. (Scale bars = 10 µm) after Zhao et al. [35]. Courtesy of *Nematology*.

Distribution *Litylenchus coprosma* was reported in New Zealand from *Coprosma repens* [35] and from *Coprosma*  En-face view shows amphidial apertures appearing as small lateral slits. Stylet robust, with welldeveloped rounded knobs, conus comprising ca 40% of stylet length, diameter narrowing sharply to be distinctly less than that of shaft. The opening of dorsal esophageal gland is located just posterior to stylet

knobs. Nerve ring is located 70–110 µm from anterior extremity, surrounding isthmus, *ca* one body diameter long. Excretory pore is located ca 5–6 body diameter from anterior end, opening posterior to nerve ring. Hemizonid located immediately anterior to excretory pore. Procorpus cylindrical, fusiform, non-muscular median bulb which is approximately one body diameter long and narrowing sharply to isthmus which is slender, cylindroid, marked off from terminal bulb, pharyngeal glands enclosed in a pyriform terminal bulb containing three large nuclei. Esophago-intestinal junction is immediately posterior to terminal bulb and covered by it in some specimens, valve present, without hyaline cells. Deirids and phasmids were not observed. Testis outstretched, reflexed in some specimens, reaching to nerve ring in some specimens, with spermatocytes arranged in a single row. Spicule paired, similar, arcuate, 2–3 µm wide at anterior end, gradually narrowing towards tip. Capitulum absent. Gubernaculum simple and arcuate. Tail conoid with a variable in shape tail terminus, usually bluntly rounded, but may have terminal process; no mucron observed. Bursa membranous, crenate in some, arising ca 1–2 cloacal body diameter anterior to cloacal aperture, extending nearly (90–95% of tail length) to tail tip.

#### Distribution

*Litylenchus coprosma* was reported in New Zealand from *Coprosma repens* [35] and from *Coprosma robusta* [26].

#### **15.** *Litylenchus crenatae* **Kanzaki, Ichihara, Aikawa, Ekino, and Masuya, 2019**

#### Measurements

After Kanzaki et al. [37].

Mature female (*n* = 10): L = 816 ± 32 (758–870) µm; a = 35.9 ± 3.4 (30.2–41.1); b = 6.6 ± 0.4 (6.1–7.6); c = 24.5 ± 1.9 (21.8–28.1); V%= 81.5 ± 1.0 (79.4–83.2); stylet = 10.6 ± 0.5 (9.9–11.3) µm.

Immature female (*n* = 10): L = 868 ± 33 (837–915) µm; a = 67.5 ± 5.8 (60.7–74.4); b = 4.3 ± 0.3 (3.9–4.8); c = 15.7 ± 0.7 (14.4–16.7); V% = 77.4 ± 0.5 (76.6–78.3); stylet = 8.0 ± 0.4 (7.4–8.5) µm.

Mature male (*n* = 9): L = 805 ± 21 (766–840) µm; a = 41.0 ± 2.4 (37.4–44.4); b = 6.4 ± 0.4 (5.9–7.3); c = 24.8 ± 2.5 (21.4–30.3); stylet = 10.5 ± 0.4 (9.9–11.3) µm; spicule = 18.3 ± 1.0 (16.7–20.2) µm; gubernaculum = 8 ± 0.4 (7.1–8.5) µm.

Immature male (*n* = 8): L = 707 ± 41 (642–773) µm; a = 57.2 ± 4.7 (48.9–61.9); b = 5.3 ± 0.6 (4.5–6.3); c = 21.1 ± 2.0 (18.5–25.1); stylet = 10.2 ± 0.4 (9.9–11.0) µm; spicule = 15.6 ± 1.2 (14.2–17.7) µm; gubernaculum = 6.5 ± 0.4 (6.0–7.1) µm.

#### Description

Female (Figure 9a): when killed, the nematodes assume a smoothly ventrally arcuate shape, body cylindrical, vermiform to semi-obese. Anterior part and cuticular morphology similar to mature male. Female gonad single, anteriorly outstretched reaching to level of pharyngeal glands. Oocytes are arranged in single row in entire ovary. Oviduct is short and spermatheca is elongated oval filled with large sperm, posteriorly connected to crustaformeria, which consists of four rows of four large and rounded cells, i.e., forming a quadricolumella, posteriorly connected to uterus by a cluster of small cells. Uterus, a thick-walled tube, sometimes containing an egg. Vagina at right angles to body axis or slightly inclined anteriorly. Vulva, a horizontal slit. Post uterine sac present, well-developed, with a thin wall and a short appendage comprising several rounded cells at distal end. Rectum is about less than one anal body diameter in length, with muscular constriction at intestine-rectal junction. Tail is short and broad, abruptly narrowing at the end with a conoid and bluntly pointed terminus, sometimes appearing like a conical blunt mucron.

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**Figure 9.** Males and Females of *Litylenchus crenatae.* (**a**) Female reproductive system and tail of *Litylenchus crenatae;* (A,B) posterior part of gonad of mature female in different focal planes; (C,D) posterior part of gonad of immature female in different focal planes; (E,F) tail of mature female; (G) tail of immature female. Ovary (ov), oviduct (od), spermatheca (sp), crustaformeria (cr), uterus (ut), and post-uterine sac (pus) are shown in (A–D), and anal opening is indicated by arrowheads in (E– G). (**b**) Male reproductive system of *Litylenchus crenatae* (A–K) are mature individuals, (L) is an immature individual. (A) Anterior end of testis; (B) middle part of mature testis; (C) posterior part of testis; (D) posterior end of testis and *vas deferens;* (E,G) ventral view of tail in different focal planes; (H,K) right lateral view of tail in different focal planes; (L) left lateral view of tail of immature individual. After Kanzaki et al. [32]. Courtesy of Nematology. **Figure 9.** Males and Females of *Litylenchus crenatae.* (**a**) Female reproductive system and tail of *Litylenchus crenatae;* (**A**,**B**) posterior part of gonad of mature female in different focal planes; (**C**,**D**) posterior part of gonad of immature female in different focal planes; (**E**,**F**) tail of mature female; (**G**) tail of immature female. Ovary (ov), oviduct (od), spermatheca (sp), crustaformeria (cr), uterus (ut), and post-uterine sac (pus) are shown in (A–D), and anal opening is indicated by arrowheads in (**E**–**G**). (**b**) Male reproductive system of *Litylenchus crenatae* (**A**–**K**) are mature individuals, (**L**) is an immature individual. (**A**) Anterior end of testis; (**B**) middle part of mature testis; (**C**) posterior part of testis; (**D**) posterior end of testis and *vas deferens;* (**E**,**G**) ventral view of tail in different focal planes; (**H**,**K**) right lateral view of tail in different focal planes; (**L**) left lateral view of tail of immature individual. After Kanzaki et al. [32]. Courtesy of Nematology.

Male (Figure 9b): when killed, the nematodes assume a smoothly ventrally arcuate shape, body cylindrical, not clearly obese or semi-obese. Body cuticle annulated with six incisures in the lateral field at the anterior part of body, 6–8 incisures around mid-body, and posteriorly connected to bursa. Deirids present in middle of lateral field slightly posterior to hemizonid and excretory pore. Lip region slightly offset from body, with a truncated shape, separated by a very shallow constriction.

**Figure 10.** Anterior region of mature adults of *Litylenchus crenatae*; (**A**) anterior end to pharyngointestinal junction; (**B**) lip region; (**C**) metacorpus (median bulb); (**D**–**G**) pharyngeal gland region in Stylet with narrow lumen and a shaft with prominent rounded basal knobs (3.6 µm in diameter). Dorsal esophageal gland is located posterior to stylet knobs. Procorpus is cylindrical. Median esophageal bulb is weakly developed, with small metacarpal valve at mid-bulb length. Isthmus is cylindrical, but narrower than the procorpus, enveloped by the nerve ring in its mid-length. Broad and glandular gland lobe with three large nuclei were observed (Figure 10). Hemizonid found at level the beginning of expansion of pharynx. Excretory pore located slightly posterior to hemizonid, with clear secretory-excretory duct. Nuclei of the esophageal overlap observed between hemizonid and pharyngo-intestinal junction, two being just anterior to the third, and latter located slightly anterior to junction. Gonad single, anteriorly outstretched reaching to level of pharyngeal glands. Testis outstretched with spermatocytes arranged in single row from anterior to middle part of testis and in multiple rows in posterior section. *Vas deferens* is visible, consisting of rounded cells, sometimes containing well-developed sperm. Spicules paired, smoothly arcuate ventrally, forming a smoothly curved horn-like blade with bluntly pointed distal end in lateral view (V-shaped). Gubernaculum simple, crescent or bow-shaped in lateral view. Bursa peloderan,

different focal planes. Nerve ring (nr), excretory pore (ep), pharyngeal gland nuclei (n), hemizonid

well developed arising three cloacal body diameter anterior to cloacal opening and terminating near tail tip. Tail is conoid, bluntly pointed in lateral view. testis; (D) posterior end of testis and *vas deferens;* (E,G) ventral view of tail in different focal planes; (H,K) right lateral view of tail in different focal planes; (L) left lateral view of tail of immature individual. After Kanzaki et al. [32]. Courtesy of Nematology.

immature individual. (A) Anterior end of testis; (B) middle part of mature testis; (C) posterior part of

**Figure 9.** Males and Females of *Litylenchus crenatae.* (**a**) Female reproductive system and tail of *Litylenchus crenatae;* (A,B) posterior part of gonad of mature female in different focal planes; (C,D) posterior part of gonad of immature female in different focal planes; (E,F) tail of mature female; (G) tail of immature female. Ovary (ov), oviduct (od), spermatheca (sp), crustaformeria (cr), uterus (ut), and post-uterine sac (pus) are shown in (A–D), and anal opening is indicated by arrowheads in (E–

*Plants* **2020**, *9*, x FOR PEER REVIEW 26 of 34

**Figure 10.** Anterior region of mature adults of *Litylenchus crenatae*; (**A**) anterior end to pharyngointestinal junction; (**B**) lip region; (**C**) metacorpus (median bulb); (**D**–**G**) pharyngeal gland region in different focal planes. Nerve ring (nr), excretory pore (ep), pharyngeal gland nuclei (n), hemizonid **Figure 10.** Anterior region of mature adults of *Litylenchus crenatae*; (**A**) anterior end to pharyngointestinal junction; (**B**) lip region; (**C**) metacorpus (median bulb); (**D**–**G**) pharyngeal gland region in different focal planes. Nerve ring (nr), excretory pore (ep), pharyngeal gland nuclei (n), hemizonid (h), pharyngo-intestinal junction (pij) and deirid (d) are indicated in (**D**–**G**) after Kanzaki et al. [37]. Courtesy of *Nematology*.

#### Distribution

*Litylenchus crenatae* was reported so far from Japan from *Fagus crenata* [37].

The phylogenetic relationships among anguinid nematodes inferred from three ribosomal RNA loci were provided by Kanzaki et al. [37]. The marker sequences derived from *Litylenchus crenatae* specimens, LC383723 (SSU), LC383725 (D2-D3 LSU), and LC383724 (ITS) were deposited to GenBank.

#### **16.** *Litylenchus crenatae* **Kanzaki et al., 2019** *mccannii* **ssp. Carta, Handoo, Li, Kantor, Bauchan, McCann, Gabriel, Yu, Reed, Koch, Martin, Burke 2020**

#### Measurements

After Carta et al. [11].

Immature female (*n* = 10): L = 823 ± 61 (750–947) µm; a = 72.9 ± 3. (61.0–86.0); b = 5.4 ± 0.7 (4.5–6.6); c =17.4 ± 3.3 (13.0–25.0); V%= 76.9 ± 1.2 (75.0–79.0); stylet = 9.7 ± 0.9 (8.5–11.2) µm.

Mature male (*n* = 4): L = 548 ±16.7 (534.5–566.7) µm; a = 36.1 ± 5.4 (33.4–44.1); b = 4.8 ± 0.2 (4.6–4.9); c = 15.5 ± 0.2 (15.3–15.9); stylet = 11.1 ± 0.5 (10.5–11.4) µm; spicule= 16.3 ± 1.4 (14.9–17.6) µm; gubernaculum = 5.3 ± 0.8 (4.3–6.1) µm.

#### Description

Females have long and slender bodies, a lip region slightly offset with 5 annules. Stylet measures 9.7 ± 0.9 µm in young females with 5% of the pharynx length, and 7–10% of the pharynx length in males. Median bulb is weak without an obvious valve. The vulval region is kinked and irregular and the anterior gonad is relatively long, nearly five times the length of the post uterine sac. The post uterine sac is about three times the vulval body width and one quarter of the vulval anal distance. The rectum is approximately one quarter of the tail length and the anus is pore-like and obscure in most specimens. Tail is conical, slender and asymmetrically pointed, with a gradually tapering and the

tail tip often with mucronate extension (Figure 11). There is a shape variation in tails of immature and mature females. *Plants* **2020**, *9*, x FOR PEER REVIEW 28 of 34

**Figure 11.** Males and Females of *Litylenchus crenatae mccannii*. (**A**) Mature Female; (**B**) male; (**C**) LT-SEM of young Female. Courtesy of Gary Bauchan and Shiguang Li of Electron and Confocal Microscopy and Mycology and Nematology Genetic Diversity and Biology Laboratory (MNGDBL), USDA, ARS, Beltsville, MD, respectively. **Figure 11.** Males and Females of *Litylenchus crenatae mccannii*. (**A**) Mature Female; (**B**) male; (**C**) LT-SEM of young Female. Courtesy of Gary Bauchan and Shiguang Li of Electron and Confocal Microscopy and Mycology and Nematology Genetic Diversity and Biology Laboratory (MNGDBL), USDA, ARS, Beltsville, MD, respectively.

Molecularly, *Litylenchus crenatae mccannii* from Ohio, Pennsylvania, and the neighboring province of Ontario, Canada, showed some differences in morphometric averages among females when compared to the Japanese population described by Kanzaki et al. [32]. Ribosomal DNA marker Female: *Litylenchus crenatae mccannii* ssp. n. young female population from North America can be differentiated from the *Litylenchus crenatae* described from Japan by:


Male: males of *Litylenchus crenatae mccannii* ssp. n. are very similar to *Litylenchus crenatae* males described from Japan. Carta et al. [11] noted some differences between the North America and the Japan population such as:


Molecularly, *Litylenchus crenatae mccannii* from Ohio, Pennsylvania, and the neighboring province of Ontario, Canada, showed some differences in morphometric averages among females when compared to the Japanese population described by Kanzaki et al. [32]. Ribosomal DNA marker sequences were nearly identical to the population from Japan [11]. The 18S rDNA and internal transcribed spacer (ITS) rDNA sequences for *Litylenchus crenatae* from Japan are 99.9% and 99.7% similar, respectively, to *Litylenchus crenatae mccannii* from North America. A sequence for the COI marker was also generated, although it was not available in the Japanese population [11]. The marker sequences derived from *Litylenchus crenatae mccannii* specimens, 104H78 and 104H82 were deposited to GenBank with accession numbers for rDNA (MK292137, MK292138) and COI (MN524968, and MN524969).

Phylogenetic trees for 18S rDNA of *Aphelenchoides* and *Litylenchus* are shown in Figures 12 and 13. *Plants* **2020**, *9*, x FOR PEER REVIEW 29 of 34

**Figure 12.** Phylogenetic Bayesian tree of 18S rDNA sequences for *Aphelenchoides* and related genera from multiple sequence alignment made with Clustal Omega (EMBL-EBI, https://www.ebi.ac.uk/Tools/msa/clustalo/); tree processed from 1,100,000 iterations in MrBayes version 3.2.6 [58] within Geneious Prime Version 2020.2.4 (Biomatters, Ltd., Auckland, NZ). Pathogenic species are indicated by arrows. **Figure 12.** Phylogenetic Bayesian tree of 18S rDNA sequences for *Aphelenchoides* and related genera from multiple sequence alignment made with Clustal Omega (EMBL-EBI, https://www.ebi.ac.uk/Tools/ msa/clustalo/); tree processed from 1,100,000 iterations in MrBayes version 3.2.6 [58] within Geneious Prime Version 2020.2.4 (Biomatters, Ltd., Auckland, NZ). Pathogenic species are indicated by arrows.

*Plants* **2020**, *9*, x FOR PEER REVIEW 30 of 34

**Figure 13.** Phylogenetic Bayesian tree from 1,100,000 iterations created in MrBayes version 3.2.6 [59] from multiple sequence alignment made with Clustal Omega (EMBL-EBI, https://www.ebi.ac.uk/Tools/msa/clustalo/) within Geneious Prime Version 2020.2.4 (Biomatters, Ltd., Auckland, NZ). **Figure 13.** Phylogenetic Bayesian tree from 1,100,000 iterations created in MrBayes version 3.2.6 [59] from multiple sequence alignment made with Clustal Omega (EMBL-EBI, https://www.ebi.ac.uk/Tools/ msa/clustalo/) within Geneious Prime Version 2020.2.4 (Biomatters, Ltd., Auckland, NZ).

#### **17. Conclusions and Future Prospects**

**17. Conclusions and Future Prospects**  Until recently, morphology used to be the only way to differentiate nematodes. With recent developments of molecular approaches in taxonomy gaining more widespread use, molecular identification has the potential to become an indispensable tool in the near future. As the GenBank continues to expand, molecular identification can become a reliable resource for nematode identification. Classical morphology continues to play a very important role in nematode identification, being reliable, cheap and quick. Molecular approaches can complement classical morphology and are crucial for species with similar morphological characters. A blend of both morphological (including SEM), morphometric, and molecular data is essential for future new foliar nematode species. The prospects in foliar nematode taxonomy and diagnostics are dependent on molecular-based methodologies that will discriminate not only species but also at the level of host races and pathotypes. This finer discrimination provides opportunities for more focused management strategies. These techniques can provide rapid diagnostics and help resolve the present problems associated with morphologically conservative organisms. When widely employed, these characterization techniques will allow differentiation between nominal species, also enhancing our Until recently, morphology used to be the only way to differentiate nematodes. With recent developments of molecular approaches in taxonomy gaining more widespread use, molecular identification has the potential to become an indispensable tool in the near future. As the GenBank continues to expand, molecular identification can become a reliable resource for nematode identification. Classical morphology continues to play a very important role in nematode identification, being reliable, cheap and quick. Molecular approaches can complement classical morphology and are crucial for species with similar morphological characters. A blend of both morphological (including SEM), morphometric, and molecular data is essential for future new foliar nematode species. The prospects in foliar nematode taxonomy and diagnostics are dependent on molecular-based methodologies that will discriminate not only species but also at the level of host races and pathotypes. This finer discrimination provides opportunities for more focused management strategies. These techniques can provide rapid diagnostics and help resolve the present problems associated with morphologically conservative organisms. When widely employed, these characterization techniques will allow differentiation between nominal species, also enhancing our understanding of the phylogeny of the genus and its relationship with other plant-parasitic nematodes.

**Author Contributions:** Conceptualization, Z.H. and M.K.; methodology, Z.H., M.K., L.C.; software, Z.H., M.K., L.C.; validation, Z.H., M.K. and L.C.; formal analysis, Z.H., M.K.; investigation, M.K., Z.H.; resources, Z.H. M.K.; data curation, Z.H., M.K., L.C.; writing—original draft preparation, M.K.; writing—review and editing, Z.H., M.K., L.C.; visualization, Z.H., M.K., L.C.; supervision, Z.H.; project administration, Z.H.; funding acquisition, Z.H. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research received no external funding.

**Acknowledgments:** Mihail Kantor was supported in part by an appointment to the Research Participation Program at the Mycology and Nematology Genetic Diversity and Biology Laboratory USDA, ARS, Northeast Area, Beltsville, MD, administered by the Oak Ridge Institute for Science and Education (ORISE) through an interagency agreement between the U.S. Department of Energy and USDA-ARS. This research was funded by ORISE ARS Research Participation Program Outgoing Interagency Agreement number (60-8042-0-057). Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U.S. Department of Agriculture. USDA is an equal opportunity provider and employer. We would also like to thank Brill Publishers for allowing us to use some of the images published in this review.

**Conflicts of Interest:** The authors declare no conflict of interest.

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## *Review* **Nematode Identification Techniques and Recent Advances**

### **Mesfin Bogale, Anil Baniya and Peter DiGennaro \***

Department of Entomology and Nematology, University of Florida, Gainesville, FL 32611, USA; mazene@ufl.edu (M.B.); anilbaniya1@ufl.edu (A.B.)

**\*** Correspondence: pdigennaro@ufl.edu; Tel.: +1-352-273-3959

Received: 28 July 2020; Accepted: 22 September 2020; Published: 24 September 2020

**Abstract:** Nematodes are among the most diverse but least studied organisms. The classic morphology-based identification has proved insufficient to the study of nematode identification and diversity, mainly for lack of sufficient morphological variations among closely related taxa. Different molecular methods have been used to supplement morphology-based methods and/or circumvent these problems with various degrees of success. These methods range from fingerprint to sequence analyses of DNA- and/or protein-based information. Image analyses techniques have also contributed towards this success. In this review, we highlight what each of these methods entail and provide examples where more recent advances of these techniques have been employed in nematode identification. Wherever possible, emphasis has been given to nematodes of agricultural significance. We show that these alternative methods have aided nematode identification and raised our understanding of nematode diversity and phylogeny. We discuss the pros and cons of these methods and conclude that no one method by itself provides all the answers; the choice of method depends on the question at hand, the nature of the samples, and the availability of resources.

**Keywords:** nematode identification; morphology-based methods; DNA-based methods; protein-based methods; image analysis

#### **1. Introduction**

Comprising over a million species [1], nematodes are likely the most diverse and numerous metazoans in soil and aquatic sediments. Despite this, nematodes are among the least studied organisms with less than 0.01% of their species diversity described to date [2]. Among some 26,000 described species, about 4100 are plant parasitic, which cause drastic economic losses to all crops [3]. Nematodes are also of significant medical and veterinary importance [4], and free-living nematodes are crucial to nutrient recycling in the environment. Therefore, accurate identification is of paramount significance to understand nematode diversity and design efficient control and management strategies. Traditionally, identification is based on characteristics such as body length, morphology of sexual organs, mouth and tail parts, and other physical characters. This morphology-based classification can prove inadequate due to lack of clear variation among closely related taxa and the need for highly skilled taxonomists, whose number is on the decline [5]. Morphology-based identification is also a demanding endeavor, especially when large numbers of samples are involved. Various sub-organismal (protein- and DNA-based) methods have been employed to supplement or circumvent the limitations associated with morphology-based classification of nematodes. The highly influential work of Blaxter et al. [6] employed sequencing of nematode ribosomal DNA (rDNA) and led to improved understanding of nematode evolutionary relationships and identification. We will not spend time discussing the evolution of nematodes and phylogenetic relationships, but it is important to understand the significance of correct nematode identification and, more to the point, how we define a nematode species. As pointed

out by Adams [7] there is a trade-off between an operational species definition and that with a strong philosophical integrity. While there is a justified need to place species within the correct evolutionary lineage, more often, nematode identification techniques are driven by an operational definition of species to assess potential threats to animal and plant health. Here, we review current methods and their progenitors in nematode taxonomic techniques and suggest potential advances.

#### **2. Morphological and Image-Based Analyses**

#### *2.1. Classical Morphological Identification*

Classic identification of nematodes is based on morphological and anatomical differences using microscopic image analysis. Morphological identification is among the cheaper identification methods and helps relate morphology with possible function [5]. While most effective for nematodes that have distinct differences, nematodes that share subtle morphological and morphometric differences like body length, presence, and shape of a stylet, the shape of the tail, etc., are difficult to distinguish morphologically. For example, root-knot nematodes (RKN; *Meloidogyne* spp.) were previously diagnosed based on adult female perineal patterns [8,9], i.e., posterior region comprising the vulva-anus area (perineum), tail terminus, phasmids, lateral lines and surrounding cuticular striae; a set of characters that was originally proposed to distinguish among *Meloidogyne incognita*, *M. javanica, M. arenaria* and *M. hapla* [10]. With the discovery of new species, however, perineal patterns became inadequate because perineal patterns (and other morphometric characters; [11]) overlapped between species [12,13]. Currently, RKN species are identified using a combination of morphological and molecular characteristics (e.g., [14,15]).

Another example is in cyst nematodes (*Heterodera* spp. and *Globodera* spp.), which are among the major pathogenic plant parasitic nematodes with worldwide distribution [16]. *Heterodera* and *Globodera* can be distinguished from each other by the morphology of their cysts: lemon shaped in the former and round in the latter [17]. Species identification within *Heterodera* is based on few morphological traits including vulval cone [18], cone top [19], vaginal [20] and lip [21,22] structures. Taxonomic distinction within *Globodera* is mainly based on morphology of cyst and second stage juveniles [23]. Host plant association may also be indicative of the cyst nematode species, though this may be misleading at times as is the case with the cereal cyst nematode group of *Heterodera* [17]. Morphological identification of cyst nematodes requires taxonomic expertise and can be challenging if samples contain mixed species. Moreover, both genera include species complexes whose members are difficult to distinguish based on morphology alone [24,25].

Important morphological identification characters in nematodes include shape of head, number of annules, body length, length of stylet, shape of stylet knob, structure of lateral fields, presence/absence and shape of spermatheca, shape of female tail terminus, shape and length of spicule and gubernaculum [26]. Measurements of these characteristics and processing of samples for this purpose requires skilled taxonomists, whose number is on the decline [5]. Morphology may also be altered due to variation in geographic location, host plant, nutrition, and other environmental factors as is observed among some free-living and plant parasitic nematodes. Concisely, it can be difficult for non-specialists to identify a nematode species with a high level of confidence based on morphology alone [27], and an integration of sub-organismal data such as DNA sequence can be required for accurate identification. However, recent advances in high performance computing may augment human image analyses.

#### *2.2. Machine Learning*

Advances in machine learning, also referred to as deep learning or artificial intelligence (AI), have opened a new avenue for nematode identification and quantification based on image analysis. The technique is especially suitable for handling large numbers of samples as well as detecting rare and microscopic objects, such as nematode eggs in complex backgrounds.

Machine learning for automated detection of phenotypes takes place in multiple stages. First, a large number of images (of nematodes, their eggs, or cysts) is taken and independently annotated (labeled) by a group of experts to reduce subjectivity. These are then used to build an algorithm that learns (captures) the salient features of the objects from the images in a layer-wise hierarchy while masking (rejecting) the noise in the background. The pattern of interest in the in-put images is then reconstructed using a network model with a supervised learning scheme. Using this technique, Akintayo et al. [28] designed a novel end-to-end Convolutional Selective Autoencoder (CSAE) to identify soybean cyst nematode (SCN) eggs in different backgrounds to cover for variations in background noise across samples from different sources. The authors trained the CSAE to identify SCN eggs using many labeled image segments (patches) that were smaller than the entire image. Information from multiple overlapping local patches was then combined to reconstruct a complete image and determine the existence of an egg in a particular patch. The model correlates pixel intensity values to reconstructed images to show the degree of confidence in predicting the object in the image is indeed an SCN egg. Tests done using two sets of samples collected from regions with different soil properties showed that egg counts done by trained personnel and using this AI technique were comparable at the 95% confidence level.

Another AI technique developed by Hakim et al. [29] using *Caenorhabditis elegans* combines the capabilities of different image processing programs for a fully automated and simultaneous processing of informative phenotypic features in a single platform called WorMachine. The image processor in this platform binarizes, identifies and crops individual worms from still in-put images taken using the bright-field with or without overlapping fluorescent acquisitions. Morphological and fluorescent features are then extracted from the cropped worm masks and analyzed individually by the feature extractor, which also allows labeling of different worms. Based on the features and labels obtained, the machine learner algorithm then conducts a binary classification or scoring of complex phenotypes using principal component analysis (PCA) and *t*-distributed stochastic neighbor embedding (*t*-SNE). The authors distinguished between males (XO), hermaphrodites (XX) and a range of phenotypes in between using fluorescent reporters for sex-specific expression patterns in mutant *C. elegans*. To demonstrate that WorMachine can be used to quantify continuous morphological phenotypes, they used strain CB5362 that is mutated in the sex-determination genes, and quantified intersex phenotypes in worms grown at different temperatures. For each worm, they determined the degree of masculinization from measurements of tail shape, gonad width (larger mid-width in egg-bearing worms), body length and area (males being smaller), brightness of head and tail (darker tails in males in bright-field), analyzed using PCA and *t*-SNE. They reported that the results agreed with those from previous studies, which showed increased masculinity at higher temperatures.

These studies show that AI can play a big role in the detection, quantification as well as classification of nematodes. As such, it will help address some of the limitations associated with the traditional morphology-based classification including the dwindling number of taxonomists, subjective decision making, and provide fast and accurate identification. Ironically, however, generating sufficient training data may present a bottleneck in developing AI due to the declining number of taxonomists. Limitations arising from shared morphological features between taxa would likely remain, but there is the possibility that machine learning will be able to elucidate unique characters discriminating nematodes that have been undetected even by the trained human eye.

#### *2.3. Autoflorescence*

A potential supplement to traditional light microscopy is the utilization of natural autofluorescence of microorganisms. Bhatta et al. [30] demonstrated that the emission and excitation spectra of the bacterial genera *Lactobacillus* and *Saccharomyces* were distinct. They also reported on the potential of these spectroscopic fingerprints to discriminate between different fungal species within the genus *Saccharomyces* without the need for fluorescent staining. Qazi et al. [31] built on this and demonstrated that eggs of different helminths revealed characteristic florescence when illuminated at different wavelengths ranging from white light to infrared. They also showed that differences in florescence lifetime values (decay in florescence intensity) were diagnostic of the species considered, *Ascaris lumbricoides* and *A. suum*. Qazi et al. [31] concluded that spectroscopic features and lifetime value measurements of autofluorescence in nematodes are promising tools in the taxonomy of these organisms.

#### **3. DNA-Based Methods**

Many forms of DNA-based methods have been developed for the identification of nematodes (e.g., [32–37]). These can be broadly categorized into fingerprint- and nucleotide-based methods. Fingerprint-based methods may include Restriction Fragment Length Polymorphism (RFLP), Amplified Fragment Length Polymorphism (AFLP), Random Amplification of Polymorphic DNA (RAPD) and the use of species-specific primers, which relies on the presence/absence of a PCR amplification product. Except for RFLP, where PCR may not be needed, all fingerprint-based methods involve PCR followed by electrophoresis. The resulting DNA fingerprint, i.e., the pattern of resolution of the DNA fragments, is used for identification and/or phylogenetic analyses of the nematode taxa considered. On the other hand, nucleotide-based methods involve PCR amplification, specific probe hybridizations and sequencing of a region(s) of the DNA, which is then used in phylogenetic analyses. Each of these methods has its own advantages and/or disadvantages compared to other nematode identification methods, DNA-based or otherwise. However, it is notable that nematode sequences have greatly altered our understanding of the evolutionary relationships between taxa [6].

#### *3.1. Fingerprint-Based Methods*

RFLP analyses can be made using fingerprints generated from genomic DNA (gDNA) digested with one or more endonucleases. Alternatively, fingerprints may be generated from PCR-amplicons (PCR-RFLPs) (e.g., [37–39]). gDNA-RFLPs tend to be complex, but potentially reveal more polymorphisms owing to the size of the gDNA template. Also, gDNA-RFLPs do not require knowledge of sequence information a priori, which is not the case with PCR-RFLPs. In both cases, however, care must be taken to let restriction digestions go to completion since incomplete digestions may lead to non-reproducible fingerprints.

The AFLP technique improves upon gDNA-RFLP by selectively amplifying fewer restriction products and producing less-complex fingerprints (e.g., [32,40]). gDNA is digested with two restriction enzymes that produce sticky ends, to which are ligated adaptors. A subset of these adaptor-ligated fragments is then selectively amplified using primer sets that recognize sequences of the adaptors, the sticky ends, and one to three nucleotides inside the restriction sites. As with gDNA-RFLPs, AFLPs do not require prior knowledge of sequence information, and completion of restriction digestions is crucial for reproducible fingerprints.

RAPD involves PCR amplification of gDNA fragments using short (usually 10 bp) primers of arbitrary sequences (e.g., [34,41]). The primers bind to several regions on the DNA, and amplification results if two primers bind on opposite strands of the DNA with their 30 -ends facing each other at a distance that can be traversed by the polymerase. Consequently, fragments of various sizes may be generated, with sizes of the larger fragments dependent on efficiency of the polymerase used. The use of large, intact gDNA template is important for this reason. Because RAPDs are done at lower temperatures, which create lower stringency for primer annealing, reproducibility especially between laboratories also poses a limitation. One advantage of this method is that it does not require prior knowledge of sequence information about the template DNA.

The use of primer sets that amplify a PCR product only in a taxon of interest is commonplace nowadays (e.g., [42–44]). Such primer sets can be designed based on fragments that uniquely identify the taxon in fingerprint analyses or based on taxon-dependent nucleotide sequence differences in aligned sequence data. In either case, care must be taken to include as much of the genetic variation within the taxon of interest as well as that of its close phylogenetic relations to ensure specificity of the

primer sets. The diagnostic value of species-specific primers is based on amplification of a product only in the species for which they are designed. Therefore, it is necessary to have an internal control for a successful PCR and avoid false negatives by multiplexing the reaction with a second set of primers that amplify a product nonspecifically; after electrophoresis, two bands would be diagnostic of the species of interest while single bands corresponding to the internal control indicate otherwise.

#### *3.2. Microarrays and Probe-Based Methods*

DNA microarray is a collection of pico-moles of microscopic DNA fragments fixed at defined positions on a solid surface such as a glass slide. For nematode identification, these DNA fragments can be generated from sequence characterized amplified regions (SCARs) and are used as probes to which test samples containing florescent-labeled PCR products or gDNA are made to hybridize in high-throughput diagnostics. Data from hybridized slides are acquired using an array scanner at the emission wavelengths of the florescent dyes used. François et al. [45] investigated the suitability of DNA microarray technique for identification of nematodes using *M. chitwoodi*-specific oligonucleotides as probes. The probes were designed based on nucleotide sequences internal to binding sites of the primer sets used to amplify SCAR and satellite DNA fragments in *M. chitwoodi,* but not in *M. arenaria, M. javanica, M. fallax* and *M. hapla*. In agreement with the specificity of the primer sets in standard PCRs, both SCAR- and satellite DNA-based probes detected *M. chitwoodi* irrespective of the geographical origin of the nematode. However, cross-hybridization with *M. chitwoodi* targets was observed when satellite DNA-based probes designed from the pMfFd satellite DNA family of *M. fallax,* a closely related species, was used. This shows that careful selection of probes is important. This is the only study that we came across where DNA microarray technology was used in nematode diagnostics.

TaqMan qPCR also employs labelled DNA probe(s) for the detection and quantification of nematodes. At the start of TaqMan qPCR, the labelled probe binds to the template DNA within the site circumscribed by the primers. As the reaction progresses and the polymerase reaches the probe, its endogenous 50 nuclease activity cleaves the probe, separating the dye from the quencher at the 3 0 -end of the probe. With each PCR cycle, more dye molecules are released, resulting in an increase in fluorescence intensity proportional to the amount of amplicon synthesized. The inclusion of probe(s) makes the technique more specific than standard PCRs and the amount of florescence detected can be used to quantify the number of nematodes in the sample. Primers and probes may be designed from aligned sequence data (e.g., [46]) as described above for species-specific primers, or from SCARs (e.g., [47]). Using this technique, Sapkota et al. [46], for example, developed a real-time PCR assay for the detection of M. hapla in soil and in root galls. They were able to differentiate *M. hapla* DNA from among those of 14 other *Meloidogyne* spp. included in their study except for *M. minor*. Based on aligned sequences from the 14 species, the authors concluded that the *M. minor* DNA must have been contaminated with that of *M. hapla* for amplification to result using these primer sets and probe. The authors reported *M. hapla* DNA extracted from 250 mg of soil (containing the equivalent of a third of an egg) could be detected by this technique. Similar studies have been carried out for other nematode taxa as well, which reported on the suitability of TaqMan qPCR for detection and quantification of nematode taxa (e.g., [47,48]).

#### *3.3. Sequence-Based Methods*

Sequence-based methods may involve analyses of nucleotide sequence information from specific segment(s) of the nuclear DNA, mitochondrial DNA (mtDNA), or the whole genome (for examples of gene regions and the corresponding primer sets, see: [42,49–54]). The rDNA and mitochondrial cytochrome c oxidase subunit I (*COX1*) genes are preferred by most studies (e.g., [54–58]) for diagnostic purposes because they have variable regions circumscribed by conserved ones. The higher level of sequence diversity in the variable region makes *COX1* preferable for resolution at lower taxonomic levels such as species and subspecies groups (e.g., [59]), while the higher level of sequence conservation in the flanking regions, which allows for 'universal' primers to be designed [56], has made the rDNA more suitable for use in wider taxonomic levels. The bulk of the sequence variability in the rDNA is harbored in the internal transcribed spacer (ITS), which is interrupted by the 5.8S coding region in the rDNA cistron into ITS1 and ITS2 [60], making the ITS useful in molecular systematics of closely related nematode species (e.g., [61–63]). ITS2 alone has been used for species diagnosis in *Caenorhabditis* [64] involving genetic crosses of newly collected isolates with known biological species, though the authors do not advocate for the use of ITS2 as an absolute criterion for species diagnosis because of the potential that distinct species may share identical ITS2 sequences. An added advantage of *COX1* and rDNA is that both genes occur in multiple copies in nematode genomes enabling PCR amplifications form small amounts of DNA templates such as that can be obtained from single nematodes. Sequence information generated is then used in character-based or phylogenetic analyses to resolve and/or identify the taxa involved; the latter analysis allows for evolutionary inferences.

The rDNA encompasses conserved coding regions (28S, 18S, and 5.8S subunits) and variable non-coding regions (ITS and ETS; the external-transcribed region) organized as tandem repeats, with intergenic spacers separating the repeating units [60]. As mentioned above, the rDNA provides phylogenetic resolution at a wide range of taxonomic levels and allows 'universal' primers to be designed for use in these taxa. This has led researchers to propose different regions of the rDNA for use as DNA barcode in different organisms; unique nucleotide sequences that can potentially be used to identify each species. Proposed DNA regions include ITS for fungi [65], 16S for bacteria [66], and 18S for nematodes [67,68]. The barcode region used in animals is the *COX1* region [69]. As such, DNA barcodes use sequence information from defined regions of the DNA to identify species using primers that are applicable for the broadest possible taxonomic group. Intraspecific variations should be smaller than interspecific variations in the barcode region.

Floyd et al. [67] used sequence information from the 18S (small subunit; SSU) to group soil nematodes into molecular operational taxonomic units (MOTUs). Each of these MOTUs was comprised of a cluster of sequences that differed from one another by less than three bases over aligned sequence data. The aligned data contained 349 to 396 nucleotides after removal of gaps, ambiguous characters and unresolved base calls from 450–500 nucleotide-long raw sequences generated using primer SSU94 [6]. MOTU content was predicted from neighbor joining trees generated using absolute character differences as a measure of distance. The authors reported that MOTUs largely corresponded with morphologically defined species or genera. Powers et al. [70] also studied a region of the 18S as a potential barcode for nematodes of suborder *Criconematina*. This region does not overlap with that used by Floyd et al. [67] and lies closer to ITS1. The authors used both phylogenetic and character state differences to define MOTUs. Among the 132 polymorphic sites in the aligned dataset, 56 were singletons and defined 56 MOTUs, each consisting of identical sequences. Most clades did not have strong statistical support, and morphologically identified species did not correspond with phylogenetically supported clades except for Clade B. Apart from a single MOTU, Clade B exclusively consisted of *Discocriconemella limitanea*, represented by 11 MOTUs, which may be cryptic species according to the authors. Conversely, some individual MOTUs identified a complex of species. For example, MOTU 76 corresponded to *Ogma* spp. that have scales arranged singularly in longitudinal rows along the length of the body, or arranged in rows consisting of clusters of 4–6 scales, or with scales densely packed on the annules forming a continuous elongated fringe.

The value of a barcode is directly related to the taxonomic rank it can effectively be applied to. The regions of SSU tested for their potential as barcodes by Powers et al. [70] and Floyd et al. [67] resolved the respective soil nematodes into named taxa and/or MOTUs. However, it is evident that MOTUs cannot be compared between the two studies because they were established based on incongruent sequence information; a phylum-wide barcode would be more powerful, but possible only if taxa representing the whole phylum were analyzed for the same DNA region. The sequence heterogeneity in individual nematodes that was reported by Powers et al. [70] is also suggestive of sequence variation among different copies of the SSU in the rDNA tandem repeat. Though, Dorris et al. [71] and Floyd et al. [67] stated that there is no evidence in nematodes of one species carrying more than

one very distinct sequence variant. Bik et al. [72], however, have demonstrated that there exists intragenomic rRNA polymorphism and copy number variation in nematodes, and that the existence of minor variant gene copies in the rRNA repeats presents substantial challenges for biodiversity estimates and the analysis of marker-based datasets. Care must be taken to exclude such variable sites during analyses if the variation is greater than the cut-off value (see below). Another issue that needs to be addressed is how to interpret the barcode. DeSalle et al. [73] contend that a non-tree-based population aggregation analysis (PAA; [74]) is the most appropriate approach because tree-building approaches are flawed for many reasons. Firstly, morphology-based methods are character-based rendering the union of classical methods and distance-based DNA barcoding difficult. Secondly, tree-building methods are hierarchical while the underlying system consisting of individuals and populations is not. Thirdly, cut-off values are rather subjective; there is no objective set of criteria to delineate taxa when using distances. DeSalle et al. [73] emphasize that the best approach is to look for diagnostic characters in the aligned sequences themselves.

A great advantage of sequence-based methods is that sequence information is stored in publicly available databases such as GenBank (ncbi.nlm.nih.gov) and NEMBASE (nematodes.org). This facilitates identification of nematodes based on sequence information through comparison with that available in these databases. Accuracy of identification, however, depends on the quality of sequences deposited in the databases and the authenticity of the taxa the sequences originated from.

Most journals require that sequences be submitted to open-access databases as part of the publication process. But there is no such requirement for alignments. Unavailability in these databases of aligned sequence datasets may affect identification, especially that based on character states. This is because though alignments are generated using software, they invariably need manual editing particularly when larger datasets containing ambiguous sites are involved, which may introduce variations in alignments.

While gene-specific sequence information is commonly employed at lower taxonomic levels, there is a growing effort to include whole mitochondrial or whole genome sequence information at all taxonomic levels now that sequencing has become more affordable. Comparative genomics enables retrieval of additional information such as synteny and gene order for the investigation of underlying evolutionary mechanisms like inversion, translocation, fusion, and so on, in addition to aiding a more advanced understanding of nematode biology. After the call to sequence 959 nematode genomes by Kumar et al. [75], progressively larger number of nematode genomes have been sequenced. It would be advantageous if whole genome sequencing projects involve morphological type specimens where possible as the availability of sequence information from type specimens in the databases would help improve the accuracy of sequence-based identification of nematode samples.

#### **4. Protein-Based Methods**

Like DNA-based methods, protein sequences, mass-to-charge ratios, and immunological techniques focus on using unique protein composition and structures to delineate nematode species. Proteins provide a reduced vocabulary compared to DNA due to redundancy of the genetic code; however, the alphabet used is vastly more complex, utilizing 20 plus characters compared to the four DNA bases. Additionally, protein structure and post-translational modifications increase the potential diversity available to define nematode species and facilitate identification. Nonetheless, the requisite specialization in protein-based techniques is often a significant deterrent.

#### *4.1. Isozyme Analyses*

Enzyme phenotypes were among the first non-morphology-based methods used for the identification of nematodes. Briefly, this technique involves the extraction of soluble proteins from whole nematodes in buffer solutions, resolving the resulting extracts by starch or polyacrylamide gel electrophoresis followed by staining for specific enzymes. This electrophoretic method, also known as Multi-locus Enzyme Electrophoresis (MEE), relies on the migration patterns of isozymes, owing to differences in electrical charge, molecular weight, and conformation stemming from slight variations in amino acid compositions. The most commonly utilized enzymes were esterases [76], though malate dehydorgenase, superoxide dismutase, and glutamate-oxaloacetate transaminase have also been employed to various degrees [76,77]. This technique supplemented morphological methods and shed light in the phylogenetic relationships, especially among the major species in the genus *Meloidogyne*. However, the method was still cumbersome and time consuming; and the need to include known samples for reference purposes are among its limitations [76].

#### *4.2. Two-Dimensional Gel Analyses*

Two-dimensional gel electrophoresis (2-DGE) has been employed in taxonomic studies of nematodes. The technique allows resolution of complex protein mixtures by charge using isoelectric focusing in one-dimension followed by mass-based resolution in a dimension perpendicular to the first. The resolution pattern is then compared among isolates to determine similarities/differences, which can be scored as presence/absence for phenetic and/or cladistic analyses of the resulting data matrix. Navas et al. [78] used 2-DGE to show proteomic variations among 18 root-knot nematodes representing four species. They demonstrated that some of these variations were species-specific, while other variations revealed evolutionary relationships among the different species.

The technique has a number of pros and cons as applied to nematode taxonomy. One of the pros of 2-DGE is that it allows evolutionary inferences to be made about the taxa considered. Species-specific polypeptides can also be excised and analyzed using mass spectrometry (see below) allowing inferences to be made about the encoding genes. The cons include that the number of polypeptides resolved, and the polymorphism observed depend on the procedure used and the number of samples analyzed. For example, the number of polypeptides Navas et al. [78] observed among the 18 isolates ranged from 73–203. The authors stated that scoring the spots was difficult at times because it was hard to assess if some of the observed differences were real or due to deformations in the gel. For this reason, they scored only 95 spots that were consistently expressed in the two replicates they used for each nematode. Thirty-seven of these spots were monomorphic and thus uninformative. Considering that two of the species in their study were represented by single isolates only, it can be concluded that both the total number as well as the number of informative spots would have been different from what they observed had they used larger number of isolates.

#### *4.3. Mass Spectral Analyses*

Matrix-assisted laser desorption/ionization (MALDI) is an ionization technique, which uses laser energy-absorbing matrix to generate gaseous ions from large molecules in solid state. Embedded in a suitable matrix, the sample is applied onto a plate and irradiated with pulsed laser resulting in vaporization of the sample and the matrix material. Molecules are ionized by loss/gain of proton(s) in the hot plume of ablated gases and accelerated into a mass spectrometer for detection. Time of flight mass spectrometer (ToF-MS) measures the time taken by these ions to reach the detector as determined by the mass/charge (*m*/*z*) values, with smaller and/or more charged ions travelling faster. Since MALDI results in minimum fragmentation, the ions generated are predominantly non-fragmented and single-charged, which makes it easy to determine parental ion masses from mass spectra [79].

The basis of taxonomic identification using MALDI-ToF-MS is the ability to detect protein/peptide ions or protein profiles that are diagnostic to the taxa being considered. Perera et al. [80] used intact second stage juveniles (J2s) and/or proteins extracted from these using various organic solvents and discriminated between *Anguina tritici, A. funesta* and *M. javanica* based on unique peaks in their spectra and/or the spectral profiles. However, the authors advised that care must be taken when selecting the solvent for protein extraction and the matrix material for the MALDI as reproducibility and quality of the spectra vary with the material used. Ahmad et al. [81] built on this study to show that single *M. incognita* nematodes (an adult female or a J2) washed or unwashed, crushed or intact, can be used for diagnostics using MALDI-ToF-MS. Their study revealed that protein profiles differed between

adults and J2s, each with its own diagnostic peaks; more masses and stronger peaks were also observed when washed and/or crushed samples were used. Both studies reported that careful optimization of instrument settings is also crucial.

Navas et al. [78] generated MALDI-ToF-MS spectral profiles for species-specific proteins obtained from excised 2-DGE gels to identify the proteins for use as biomarker molecules. Their attempt to identify the proteins using similarity matches, however, returned no hit for lack of sufficient information in the databases at the time. A similar study involving 2-DGE and MALDI-ToF-MS analyses of proteomes of two nematomorph species, *Paragordius tricuspidatus* and *Spinochordodes tellinii,* was carried out by Biron et al. [82]. Biron et al. [82] reported that while 36.2% of total protein spots on the 2-DGE analyses were shared between the two hairworm species, 38.0% were specific to *P. tricuspidatus* and 25.8% to *S. tellinii*; a genetic distance of 0.47 separated the two species confirming the distant relationship reported previously for these species. Unlike Navas et al. [78], Biron et al. [82] were successful in identifying MS fingerprints of proteins obtained from excised gel plugs using similarity searches in the databases.

These studies [78,80–82] have demonstrated that 2-DGE coupled with MALDI-ToF-MS provide a powerful tool in nematode taxonomy. The methods allow for inferences to be made regarding evolutionary relationships among taxa as well as for development of species-specific markers. Results, however, can be affected by a number of factors including the protein extraction method, the quality of 2-DGE runs, and setup of instrument. Protein expression profiles are also known to differ depending on the developmental stage of nematodes and the growth conditions.

#### *4.4. Serological Analyses*

Since Bird [83] first reported on the possibility of generating antisera against nematodes, the application of poly- and monoclonal antibodies (mAbs) has been explored by several researchers with mixed results (for a summary see [84]). For example, Lee [85] reported that antiserum raised against *M. incognita* did not produce the trademark arc-shaped precipitation band when paired with antigens from another species within the same genera, *M. hapla,* in the Ouchterlony double diffusion assay, indicating a lack of cross-reactivity. However, it was noted that the apparent specificity may be due to the small number of nematodes used in the assay. Further studies [86–88] also confirmed a lack of specificity in reactivity of antisera from *Meloidogyne* spp. Similar mixed results were also observed among cyst nematode *Heterodera* and *Globodera* species (summarized in [84]). Cross-reactivity of polyclonal antisera raised against whole macerated nematodes, including the associated microbiome and metabolites, in their bodies is to be expected.

The development of the hybridoma technique by Kohler and Milstein [89] raised the hope of the nematology community to develop mAbs for diagnostic purposes. The technique involved isolating mature B-cells from animals immunized with nematode antigens, fusing these B-cells with mouse lymphoid tumor cells to produce hybridomas that can be maintained indefinitely in vitro for continuous production of the antibodies. mAbs provide more specificity depending on the immunogen the antibodies were raised against. mAbs were raised against a variety of agriculturally important nematodes including *H. glycines* [90], *M. incognita* [91], *G. rostochiensis* and *G. pallida* [84] using the hybridoma technique. Schots et al. [84] reported that some mAbs differentiated between *G. rostochiensis* and *G. pallida* isolates. The authors also showed that these mAbs were so sensitive that protein equivalents of less than one egg were detected using immunoassays. The hybridoma technique becomes cumbersome with increasing number of nematode samples. The low proportion of successful fusions obtained between B- and tumor cells also presents a handicap. Next-generation sequencing technologies may prove to revitalize this line of nematode identification techniques as single B-cell receptor sequencing (scBCR-seq) can reconstruct antigen binding site sequences for comparative studies [92].

#### **5. Conclusions**

The purpose of taxonomy is to understand biodiversity, categorize organisms, and aid the communication of biological information. Scientific naming is a prerequisite for communication in taxonomy, and valid naming is only possible with type specimens and corresponding morphological information. However, this is not always possible, particularly when dealing with environmental samples (eDNA). Furthermore, it is now generally accepted that there are insufficient morphological features to describe biological diversity, and the use of molecular information to supplement and/or circumvent this limitation is commonplace. Nonetheless, a taxon is more meaningful if its members possess unique biological features, rather than the taxon only representing a group of individuals sharing similar morphological or molecular features.

Morphology-based classification forms the foundation of taxonomy. It has benefited from recent advances in image analysis. AI helps circumvent limitations associated with the scarcity of highly qualified taxonomists and enables objective decision making, coupled with fast and accurate identification. Spectroscopic features and lifetime value measurements of autofluorescence also provide additional traits that can be exploited for identification purposes.

The relative ease of molecular methods (Table 1) has led to the recognition of many new taxa; some, based on sequence information alone. These taxa would have been impossible to describe morphologically not only for lack of taxonomists and sufficient morphological differences, but also because members of these taxa are difficult to culture. Taxa identified using different molecular approaches, however, are not always congruent; for example, when sequence information from different regions of the DNA is used in different studies, or when sequence data generated from the same DNA region are analyzed differently between studies. Likewise, taxa based on morphological features do not always correspond with those based on molecular information and vice versa. Consequently, no single method by itself provides all the answers all the time; and the choice of method(s) depends on the question asked, the nature of the samples and the availability of resources.


**Table 1.** Comparison of different nematode identification methods.

If the question at hand is identification of a nematode sample, the most direct approach would be to examine the sample microscopically and assign the nematode to the lowest taxonomic rank possible. The source of the sample may also provide a clue in this regard. However, this may require some level of taxonomic expertise. Based on this information, a molecular technique may then be employed to identity the nematode to species or even subspecies level. If the question has to do with quarantine, molecular methods that are specific to the quarantined nematode species may be employed to ascertain whether the nematode at hand is quarantined. If the objective is assessment of diversity in a field population(s), any of the fingerprinting techniques and/or sequence analyses based on one or a few

genes may do. High-throughput sequencing using second or third generation technologies and the appropriate bioinformatic techniques are useful to study the diversity of nematodes in an environmental sample (eDNA).

**Author Contributions:** Drafting and writing of the manuscript, M.B.; drafting parts of the manuscript, A.B.; conceptualizing of the review and editing of the manuscript, P.D. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research received no external funding.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


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