Nematode+ID+Workshop +Bari+Italy+2005+Manual

NEMATODE

WORKSHOP:

Diagnostics and Phylogenetics

of Major Groups of Nematodes Considered in Biological

Control

Diagnostics and Phylogenetics of Major Groups of Nematodes Considered in Biological Control Prepared by Dr. S. Patricia Stock - Locorotondo, Bari, Italy, June 12-13, 2005.

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NEMATODE WORKSHOP: Diagnostics and Phylogenetics of Major

Groups of Nematodes Considered in Biological Control

June 12-13, 2005 Locorotondo, Bari, Italy,

Sponsored by

10th European Meeting of the IOBC/WPRS Working Group

"Insect Pathogens and Insect Parasitic Nematodes"

in cooperation with

COST ACTIONS 842 and 850

“Entomophthorales” & “Biocontrol Symbiosis”

WORKSHOP MANUAL Prepared by:

Dr. S. Patricia Stock Division of Plant Pathology and Microbiology

Department of Plant Sciences – Department of Entomology University of Arizona, Tucson, AZ, USA

This manual is to be used only for scientific teaching purposes. It does not constitute a scientific publication, and may not be used for any commercial activity. Unauthorized reproduction may violate copyrights held by the author or by the copyright holders of the illustrations reproduced in this manual.


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TABLE OF CONTENTS

A. Classification B. Diagnosis of major groups C. Tables D. Polytomous keys E. Preparation of Nematodes for Identification F. Atlas of main morphology and terminology G. Molecular Diagnostics / Phylogenetics H. References


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A. Classification _____________________________________________________________

More than 30 nematode families are known to host taxa that parasitize or are associated with insects (Maggenti, 1981; Nickle, 1972; Poinar, 1975, 1983, 1990; Kaya and Stock, 1997). However, because of their biological control potential, research has concentrated on seven families: Mermithidae, Allantonematidae, Neotylenchidae, Sphaerularidae, Rhabditidae, Steinernematidae and Heterorhabditidae, the latter two families currently receiving the most attention as control agents of soil insect pests (Lacey et al., 2001).

The biological control potential of nematodes is not restricted to insects. Phasmarhabditis hermaphrodita (Schneider), a member of the family Rhabditidae, is known to suppress several slug species, and has recently been developed as a biological molluscicide. Several predatory nematodes such as mononchids, dorylaimids, nygolaimids, diplogasterids have also potential for management of plant-parasitic nematodes. Moreover, the fungal-feeding aphelenchid, Aphelenchus avenae Bastian, has also been considered as potential biological control agent of soilborne fungal pathogens of plants.

I have adopted the new classification scheme suggested by De Ley and Blaxter (2002) to list those groups with biological control potential. This classification is rooted on a phylogenetic interpretation of a preliminary evolutionary tree based on 18S ribosomal DNA proposed by Blaxter et al. (1998). This molecular framework recognizes the presence of three basal clades: 1) dorylaimids,2) enoplids and 3) chromadorids. Relationships between these clades have not been fully resolved yet, but available data support sister taxon status of dorylaims and enoplids (De Ley and Blaxter, 2002). In this new taxonomic scheme, dorylaims and enoplids are grouped within the class Enoplea Inglis, 1983. The Chromadorea Inglis, 1983 comprise the majority of taxa within

Nematoda, including all the former Secernentea.

In this classification system, seven out of eleven nematode families currently considered in biological control are grouped within the Chromadorea, the remaining families, the Mononchidae, Mermithidae, Dorylaimidae and Nygolaimidae are members of the Enoplea (Table 1).


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B. Diagnosis of major groups ____________________________________________________________________________________

CLASS CHROMADOREA

Family Steinernematidae Chitwood and Chitwood, 1937

Diagnostics: Adults with truncated to slightly round head. Lips fused six, but tips distinct, and each with one labial papilla. Four cephalic papillae present. Amphids small. Stoma reduced, short and wide, with inconspicuous sclerotised walls. Oesophagus rhabditioid, set off from intestine. Nerve-ring usually surrounding isthmus or anterior part of basal bulb. Excretory pore opening distinct. Female with paired opposed ovaries. Vagina short, muscular. Vulva located near middle of body, with or without protruding lips. Epiptygma present or absent. Male with single reflexed testis. Spicules paired, symmetrical. Gubernaculum present. One single midventral and 10-14 pairs of genital papillae present of which 7-10 pairs precloacal. Tail rounded, digitated or mucronated. Third-stage infective juvenile with collapsed stoma. Cuticle annulated, lateral field with 6-8 ridges in middle of body. Oesophagus and intestine collapsed. Specialized bacterial pouch located at beginning of intestine and of variable shape. Excretory pore distinct, anterior to nerve ring. Tail conoid or filiform, with variable hyaline portion. Phasmids present, prominent or inconspicuous.

The Steinernematidae currently comprise 2 genera, Steinernema Travassos, 1927 with more than 30 species and Neosteinernema Nguyen and Smart, 1994 with only one species N. longicurvicauda (Table 2)

Bionomic Notes: Steinernematids are obligate pathogens in nature and are characterized by their mutualistic association with bacteria of the genus Xenorhabdus. Of all nematodes studied for biological control of insects, the Steinernematidae together with the

Heterorhabditidae, have received the most attention because they possess many of the attributes of effective biological control agents.

Figure 1. Steinernema monticolum (From Stock et al, 1998). A. Male in toto, B. Male tail, C. Female, stomatal region, D. Female in toto, E. Infective juvenile, anterior end, F. Infective juvenile, tail.

Family Aphelenchidae Fuchs, 1937 Diagnostics: Labial cap distinct and often set off by a constriction. Hollow axial protrusible spear with slight basal thickenings. Oesophagus with a large metacorpus (median bulb). Dorsal

A B

C

D

E

F


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oesophageal gland opening into metacorpus. Oesophageal glands either forming a lobe or adjoined intestine. Male bursa supported by four pairs of caudal papillae (rays). Spicules ventrally arcuate and slender. Gubernaculum present.

Figure 2. Aphelenchus avenae Bastian. (After Stock and Hunt, 2005). A: Female; B: Male tail, lateral view. Bionomic Notes: Aphelenchids are fungal-feeding nematodes. They can be found in decaying plant tissues feeding on various fungal hyphae. Until now, Aphelenchus avenae has been studied as a biological control alternative to suppress fungal pathogens of plants. Family Allantonematidae Pereira 1931

Diagnostics: Preparasitic female and free-living male with small stylet (less than 15 µm long) with or without knobs. Oesophageal glands elongated, lobe-like; subventral glands extending past dorsal lobe. Tail conoid or subcylindrical. Preparasitic female with small vulva and short vagina. Postvulval sac short or absent. Uterus elongated. Parasitic female obese, sac-like, elongate or spindle-shaped. Reproductive organs filling body cavity. Uterus not everted. Vulva a small transverse slit or indistinct. Male with outstretched testis. Spicules arcuate, pointed, usually less than 25 µm long. Gubernaculum usually present. Bursa present or absent.

Figure 3. Thripinema rineraoi. A, F: Anterior and posterior region of partially free-living impregnated female; B: Male; C, D: Anterior and posterior regions of male; E: Entomoparasitic female from haemocoel of Megaluriothrips sp. (After Siddiqi, 1986, courtesy CAB International). Bionomic Notes: Allantonematids have a single heterorsexual life cycle. Females are obligate parasites of the haemocoel of mites and insects. Within this family, members of Thripinema Siddiqi, 1986 are known to parasitize thrips (Thysanoptera: Thripidae). A free-living stage occurs in flowers, buds and leaf galls of plants that attack thrips.

A

B


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Family Neotylenchidae Thorne, 1941

Diagnostics: (modified from Siddiqi, 2000). Free-living stages with smooth or finely striated cuticle. Stylet well developed, less than 20 µm long, basal knobs may be bifid. Oesophagus fusiform, basal bulb absent. Oesophageal glands free in body cavity, extending over intestine. Orifice of dorsal gland close to stylet base. Nerve ring generally circumintestinal, posterior to, or at level of, oesophago-intestinal junction. Excretory pore anterior or posterior to nerve ring. Female monodelphic prodelphic. Vulva in posterior region, postvulval sac present or absent. Tail conoid, subcylindroid or cylindroid. Male monorchic, testis outstretched. Bursa present or absent. Spicules paired, small, cephalated or arcuate, distally pointed. Gubernaculum present or absent. Preadult female (free-living) with hypertrophied stylet and oesophagus. Ovary immature. Uterus long. Mature parasitic female, obese, sausage-shaped or elongate tuboid. Stylet and oesophagus non-functional. Uterus hypertrophied but not everted.

Figure 4. Beddingia (=Deladenus) siricidicola Bedding (From Stock and Hunt, 2005). A: Oesophageal region of fungus feeding female; B: Oesophageal region of entomoparasitic preadult female; C: Male tail region; D: Posterior region of fungus feeding female. (After Siddiqi, 2000, courtesy CAB International).

Bionomic Notes: Neotylenchids are characterized by having a free-living generation alternating with an insect parasitic generation. Beddingia Thorne, 1941 currently comprises 17 nominal species with B. siricidicola Bedding, 1968, a parasite of the wood wasp Sirex noctilio, being the only taxon currently used in biological control

Family Rhabditidae Örley, 1880

Diagnostic Characters: Stoma commonly cylindrical without distinct separation of cheilo-, gymno- and stegostom. Stoma two or more times as long as wide. Usually with six distinct lips, each with one cephalic papilla. Amphids porelike. Oesophagus clearly divided into corpus (procorpus and metacorpus) and postcorpus (isthmus and valvated muscular portion). Male spicules separate or fused distally. Gubernaculum present. Bursa mostly well developed, peloderan or leptoderan, occasionally small or rudimentary. Nine or ten pairs of genital papillae (bursal rays). Female with one or two ovaries.

Figure 5. Phasmarhabditis hermaphrodita (Schneider). A-B. Female, stoma dorsal view (A) and subventral (B); C. Oesophagus; D-E. Female tail, lateral view (D) and ventral (E); F. dauer juvenile,


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anterior regionl, G. Female gonad, anterior end, L. Male tail, lateral view. (From Hooper et al., 1999)

Bionomic Notes: Most members of this family are free-living bacterivores, but, two species of Phasmarhabditis, P. hermaphrodita (Schneider, 1859) and P. neopapillosa (Schneider, 1866) have parasitic associations with terrestrial slugs and snails. Phasmarhabditis hermaphrodita is capable of killing several slug and snail pests and is the only species currently used as a biological control agent and is mass produced and commercialized as a molluscicide (See Wilson and Grewal, Chapter 23 and Glenn and Ester, Chapter 24).

Family Heterorhabditidae Poinar, 1976

Diagnostics: Adults with six distinct protruding pointed lips surrounding oral aperture. Each lip bearing one labial papilla. Stoma short and wide. Oesophagus rhabditoid. Corpus cylindrical, metacorpus not differentiated. Isthmus short. Basal bulb pyriform with reduced valve. Excretory pore usually located at level of basal bulb. Hermaphrodite (first generation) with an ovotestis. Vulva located near midbody. Postanal swelling present or absent. Tail terminus blunt, with or without a mucro. Female (second generation) amphidelphic, ovaries with reflexed portions often extending past vulva opening. Vulva located near middle of body, with or without protruding lips. Tail conoid; postanal swelling present or absent. Male (second generation) monorchic. Spicules paired, symmetrical, straight or arcuate, with pointed tips. Gubernaculum slender, about half length of spicules. Bursa open, peloderan, attended by a complement of nine pairs of bursal rays (papillae). Third-stage infective juvenile ensheathed in cuticle of second-stage juvenile. Cuticle of J2 with longitudinal ridges throughout most of body length, and a tessellate pattern in anterior-most region. Lateral field with two ridges. Prominent cuticular dorsal tooth present. Excretory pore

located posterior to basal bulb. Tail short, conoid, tapering to a small spike-like tip. Heterorhabditidae consists of one genus, Heterorhabditis Poinar, 1976, with H. bacteriophora as the type and 10 other species described (Table 3).

Figure 6. Heterorhabditis downesi Stock et al. (from Stock et al. 2003) . A. Hermaphrodite; B. Spicules; C. Gubernaculum; D. Hermaphrodite,vuvla; E. Male tail, ventral view; F. Male, anterior end; G. Female tail. Bionomic Notes: Heterorhabditids have a similar life cycle to steinernematids, but adults resulting from infective juveniles are hermaphroditic. Eggs laid by the hermaphrodites produce juveniles that develop into males and females or infective juveniles. The males and females mate and produce eggs that develop to infective juveniles.


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Family Diplogasteridae Micoletzky, 1922

Diagnostics: Lip region, never set off by a constriction, usually composed of six distinct lips or six fused lips. Amphids pore-like. Stylet absent. Stoma variable, usually broad and short with stegostom containing denticles, warts or teeth. Oesophagus with a median valvated bulb and a basal valveless bulb. Female gonad usually paired. Male with paired spicules and gubernaculum. Bursa usually small or absent. Male tail often with nine pairs of genital papillae and a pair of phasmids. Three pairs of genital papillae located preanal.

Figure 7. Aduncospiculum halicti Giblin y Kaya A. Adult, anterior end; B-C. Male tail, ventral view (B), and lateral (C); Female tail, lateral view; E. Female, middle-body region. (From Giblin and Kaya) Bionomic Notes: Most Diplogasteridae members are predators or omnivores but a few taxa are also bacterial feeders. Of all of them, only a few genera (i.e. Butlerius, Fictor and Mononchoides) have been studied as biological control agents of plant parasitic nematodes

CLASS ENOPLEA

Family Mononchidae Chitwood, 1937 Diagnostics: Generally large, stout nematodes. Cuticle usually appearing non-striated and smooth. Lateral field usually not differentiated. Head not distinctly offset, composed of 6 or fewer confluent lips, each carrying at least two papillae. Amphids small, cup-shaped. Stylet absent. Stoma forming a small to large barrel-shaped cuticularized chamber bearing an immovable dorsal tooth. Subventral teeth and/or rows of denticles or ridges may also be present. Oesophagus stout, muscular, glandular and almost cylindrical with some posterior swelling. Oesophago-intestinal junction tuberculate or non-tuberculate. Excretory pore usually absent. Females usually with paired ovaries, opposite and reflexed. Males with paired opposed testes leading to a common vas deferens. Spicules paired. Gubernaculum present. Lateral guiding piece often present. Midventral row of precloacal papillae always present on males. Tail variable in form. Bursa absent

Figure 8. Mononchidae. A-B Mononchus sp. A. anterior region; B: female tail region. C-D Mylonchulus minor. C. Anterior region; D: Female tail region. Modified from Stock and Hunt, 2005. Bionomic Notes: Mononchids are predominantly predaceous nematodes feeding on small invertebrates (including other nematodes) in soil and fresh water. Many genera have been proposed, but only Mylonchulus (Cobb, 1916), Mononchus

A

B

C

D


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Bastian and Iotonchus (Cobb, 1916) have been explored as biocontol agents. A few taxa have been used against plant-parasitic nematode species Family Mermithidae Braun, 1883 Diagnostics: Long slender nematodes sometimes reaching a length of 50 cm, but usually between 1 and 10 cm. Cuticle smooth or with criss-cross fibers. Anterior end containing two, four or six cephalic papillae and rarely a pair of lateral mouth papillae. Amphids tube-like or modified pouch like. Oesophagus modified into a slender tube surrounded posteriorly by stichosomal tissue. Intestine modified into a trophosome or food storage organ forming a blind sac soon after the nematodes enter a host. Preparasitic juveniles (J2) with a functional stylet and a pair of penetration glands that degenerate after host invasion. Ovaries paired; muscular vagina straight or curved. Males with a single fused or paired spicules. Gubernaculum and bursa absent. Several rows of genital papillae usually present

Figure 9. Mermithidae 1.Mermis nigrescens Dujardin. A. lateral view of female head, B. Postparasitic juvenile tail, C. Preparasitic larva, D. Vagina, lateral view, E. egg with byssi, F. Male tail, lateral view, G. female tail, lateral view, H. Parasitized grasshopper (From Nickle, 1972). Bionomic Notes: There are numerous described genera, many of which are poorly characterised by contemporary standards. The group is in urgent need of revision before a workable key can be

constructed. All known species are obligate parasites of terrestrial and aquatic arthropods and other invertebrates. Mermithids parasitize many different insect groups, including Orthoptera, Dermaptera, Hemiptera, Lepidoptera, Diptera, Coleoptera, and Hymenoptera. Mermithids with significant biological control potential include Romanomermis culicivorax, a parasite of mosquito larvae (Petersen, 1984, 1985), Oesophagomermis (=Filipjevimermis) leipsandra, a parasite of larval banded cucumber beetle Diabrotica balteata (Creighton and Fassuliotis, 1983), Mermis nigrescens, a parasite of grasshoppers (Webster and Thong, 1984) and Agamermis unka, a parasite of white and brown planthoppers (Choo et al., 1989, 1994) Figure 10. Romamnomermis culicvorax. A. Female head, B. postparasitic juvenile tail, C. Male papillae pattern, D. Female vagina, E. egg, F. Male tail, G. female tail, H. preparasitic larva, I. Postparasitic juvenile stoma showing tooth, J. parasitized mosquito. (From Nickle, 1972).


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Family Dorylaimidae De Man, 1876

Diagnostics: Generally large and robust nematodes. Stoma with an axial odontstyle, the aperture of which is located dorsally. Oesophagus cylindrical and divided into two parts: anterior portion usually slender, sometimes with small muscular swellings, followed by an expanded posterior portion. Excretory pore rudimentary or absent. Females with one or two ovaries. Males with two testes. Spicules robust and separated. Gubernaculum usually absent, but lateral guiding pieces present. Bursa absent. Setae and caudal glands absent.

Figure 11. Dorylaimidae. A-C, Mesodorylaimus. A: Head region; B: Female tail; C: Male tail. D-F, Allodorylaimus. D: Head region; E: Female tail; F: Male tail. G-J, Eudorylaimus. G: Head region; H: Vulval region; I: Male tail, J: Female tail. K, L, Discolaimus. K: Pharyngeal region; L: Head region. M-O, Labronema. M: Head region; N. Male tail region; O: Female tail. P-R, Pungentus. P: Head region; Q: Female tail; R: Male tail region. (After Jairajpuri & Ahmad, Dorylaimida. Free-living, Predaceous and Plant-parasitic Nematodes (1992) Bionomic Notes: The feeding habits of many members are not known, although some are acknowledged as being predaceous on other nematodes and invertebrates.

Family Nygolaimidae Thorne, 1935 Diagnostics: Stoma armed with mural tooth of variable shape. Dorylaimoid oesophagus with posterior portion enclosed in a sheath. Three large cardiac glands at oesophag-intestinal junction. Ovaries paired, opposed and reflexed. Spicules arcuate. Gubernaculum and lateral guiding pieces present in some males. Bionomic Notes: Likewise dorylaimids, nygolaimids are predaceous, a few taxa considered for biological control of plant-parasitic nematodes


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C. TABLES ____________________________________


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Table 1. Major groups in Nematoda with biological control potential. (Classification based on De Ley and Blaxter, 2002) CLASS SUBCLASS ORDER SUBORDER INFRAORDER SUPERFAMILY FAMILY

Mononchida Jairajpuri, 1969

Mononchina Kirjanova y Krall, 1969

Mononchoidea Chitwood, 1937

Mononchidae Chitwood, 1937

Dorylaimida Pearse, 1942

Nygolaimia Thorne, 1935

Nygolaimoidea Thorne, 1935

Nygolaimidae Thorne, 1935

ENOPLEA Inglis 1983

DORYLAIMIA Inglis, 1983

Mermithida Hyman, 1951

Mermithina Andrássy, 1974

Mermithoidea Braun, 1883

Mermithidae Braun, 1883

Panagrolaimomorpha De Ley y Blaxter, 2002

Strongyloidoidea Chitwood y McIntosh, 1934

Steinernematidae Travassos, 1927

Aphelenchoidea Fuchs, 1937

Aphelenchidae Fuchs, 1937 Allantonematidae Pereira, 1931

Tylenchina Thorne, 1949

Tylenchomorpha De Ley y Blaxter, 2002

Sphaerularoidea Lubbock, 1861

Neotylenchidae Thorne, 1941

Rhabditoidea Örley, 1880

Rhabditidae Örley, 1880

Rhabditomorpha De Ley y Blaxter, 2002 Strongyloidea

Baird, 1853 Heterorhabditidae Poinar, 1975

CHROMADOREA Inglis, 1983

CHROMADORIA Pearse, 1942

RHABDITIDA Chitwood, 1933

Rhabditina Chitwood, 1933

Diplogasteromorpha De Ley y Blaxter, 2002

Diplogasteroidea Micoletzky, 1922

Diplogasteridae Micoletzky, 1922

* Families within Sphaerularoidea are listed based on the classification proposed by Siddiqi (2000) which recognizes three families within the Sphaerularoidea: Sphaerulariidae, Lubbock, 1861; Allantonematidae, Pereira, 1931 and Neotylenchidae Thorne, 1941


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Table 2. Taxonomic summary of the family Steinernematidae (Modified from Stock and Hunt, 2005). Family Steinernematidae Chitwood and Chitwood, 1937 Syn. Neoaplectanidae Sobolev, 1953 Taxa Geographic Origin* Type genus: Steinernema Travassos, 1927 Type species: Steinernema kraussei (Steiner, 1923) Travassos, 1927 Europe (Germany), North America Other species: S. abbasi Elawad, Ahmad and Reid, 1997 Asia (Oman) S. affine (Bovien, 1937) Wouts, Mrácek, Gerdin and Bedding, 1982 Europe (Denmark) S. anatoliense Hazir, Stock and Keskin, 2003 Asia (Turkey) S. apuliae Triggiani, Mracek and Reid, 2004 Europe (Italy) S. arenarium (Artyukhovsky, 1967) Wouts, Mrácek, Gerdin and Bedding, 1982 Asia (Central Russia) S. asiaticum Anis, Shahina, Reid and Rowe, 2002 Asia (Pakistan) S. bicornutum Tallosi, Peters and Ehlers, 1995 Europe (Yugoslavia) S. carpocapsae (Weiser, 1955) Wouts, Mrácek, Gerdin and Bedding, 1982 Asia, Europe (Czechoslovakia), North

America, South America S. caudatum Xu, Wang and Li, 1991 Asia (China) S. ceratophorum Jian, Reid and Hunt, 1997 Asia (China) S. cubanum Mrácek, Hernandez and Boemare, 1994 Central America (Cuba) S. diaprepesi Nguyen and Duncan, 2002 North America (USA) S. feltiae (Filipjev, 1934) Wouts, Mrácek, Gerdin and Bedding, 1982 Europe (Denmark), North America,

South America S. glaseri (Steiner, 1929) Wouts, Mrácek, Gerdin and Bedding, 1982 Asia, Europe, North America (USA),

South America S. hermaphroditum Stock, Griffin and Chaenari, 2004 Asia (Indonesia) S. intermedium (Poinar, 1985) Mamiya, 1988 North America (USA), Europe S. karii Waturu, Hunt and Reid, 1997 Africa (Kenya) S. kushidai Mamiya, 1988 Asia (Japan) S. loci Phan, Nguyen and Moens, 2001 Asia (Vietnam) S. longicaudum Shen and Wang, 1992 Asia (China), North America S. monticolum Stock, Choo and Kaya, 1997 Asia (Korea) S. neocurtillae Nguyen and Smart, 1992 North America (USA) S. oregonense Liu and Berry, 1996 North America (USA) S. pakistanense Shahina, Anis, Reid, Rowe and Maqbool, 2001 Asia (Pakistan)


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S. puertoricense Román and Figueroa, 1994 Central America (Puerto Rico) S. rarum (de Doucet, 1986) Mamiya, 1988 South America (Argentina), North

America S. riobrave Cabanillas, Poinar and Raulston, 1994 North America (USA) S. ritteri de Doucet and Doucet, 1990 South America (Argentina) S. sangi Phan, Nguyen and Moens, 2001 Asia (Vietnam) S. scapterisci Nguyen and Smart, 1990 South America (Uruguay) S. scarabaei Stock and Koppenhöfer, 2003 North America (USA) S. serratum Liu, 1992† Asia (China) S. siamkayai Stock, Somsook and Kaya, 1998 Asia (Thailand) S. tami Van Luc, Nguyen, Spiridonov and Reid, 2000 Asia (Vietnam) S. thanhi Phan, Nguyen and Moens, 2001 Asia (Vietnam) S. thermophilum Ganguly and Singh, 2000 Asia (India) S. websteri Cutler and Stock, 2003 Asia (China) S. weiseri Mrácek, Sturhan and Reid, 2003 Europe (Germany) Genus: Neosteinernema Nguyen and Smart, 1994 Type and only species: Neosteinernema longicurvicauda Nguyen and Smart, 1994 North America (USA) * Country of original isolation in parenthesis † species inquirenda


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Table 3. Taxonomic summary of described Heterorhabditis species (Modified from Stock and Hunt, 2005). Taxa Biogeography*

Type and only genus: Heterorhabditis Poinar, 1976 Syn. Chromonema Khan, Brooks and HIrschmann, 1976

Type species: Heterorhabditis bacteriophora Poinar, 1976 Syn. Chromonema heliothidis Khan, Brooks and Hirschmann, 1976 H. heliothidis (Khan, Brooks and Hirschmann, 1976) Poinar, Thomas and Hess, 1977 H. argentinensis Stock, 1993§

Africa, Asia, Australia, Central America, Europe, North America (USA) , South America

Other species:

H. baujardi Phan, Subbotin, Nguyen and Moens, 2003 Asia (Vietnam)

H. brevicaudis Liu, 1994 Asia (China)

H. downesi Stock, Burnell and Griffin, 2002 Europe (Ireland)

H. indica Poinar, Karunakar and David, 1992 Syn. H. hawaiiensis Gardner, Stock and Kaya, 1994§

Asia (India), Central America, North America

H. marelata Liu and Berry, 1996 Syn. H. hepialius Stock, Strong and Gardner, 1996

North America (USA)

H. megidis Poinar, Jackson and Klein, 1987 North America (USA), Europe

H. mexicana Nguyen , Shapiro-Ilan, Stuart, McCoy, James and Adams, 2004 North America (Mexico)

H. poinari Kakulia and Mikaia, 1997** Europe (Georgia)

H. taysearae Shamseldean, Abou El-Sooud, Abd-Elgawad and Saleh, 1996 Asia (Egypt)

H. zealandica Poinar, 1990 Australia (New Zealand)

* Country of original isolation in parenthesis; ** Species inquirenda. § As proposed by Stock (in press).


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D. Polytomous keys


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Key 1. Polytomous key for Steinernematidae. All data from original descriptions unless otherwise specified. Morphometrics are given in microns (From Stock and Hunt, 2005)

Neosteinernema Key Diagnostic Features: Adults and third-stage infective juvenile with very conspicuous amphids. Male with ventrally arcuate spicules with a very prominent

manubrium. Third-stage juvenile with very long (as long as oesophagus length) and filiform tail. First Generation Adults Third Stage Infective Juvenile Male Female

Species L W EP T D% E% LF SPL GL SW D% M EPI VL longicurvicauda 920

789-1084 24

20-31 68

61-76 167

141-190 41

38-46 41

37-48 8 61

52-67 59

52-66 1.03

0.8-1.15 NA A A V

Steinernema Key Diagnostic Features: Adults and third-stage infective juvenile with phasmids not visible. Shape of spicules variable but not with a manubrium shape as in

Neostreinernema. Third-stage infective juvenile with conoid tail variable in size. First Generation Adults Third Stage Infective Juvenile Male Female

Species TBL MBW EP TL D% E% LF SPL GUL SW D% M EPI VL carpocapsae-group

(IJ average size < 600 µm) asiaticum 425*

360-450 23

20-25 32

28-34 NA 32*

30-36 78*

60-90 6 68*

61-74 53*

46-62 * P44

35-57 P P SP

siamkayai 446 398-495

21 18-24

35 29-38

36 31-41

37 31-43

96 95-112

6-8 77.5 75-80

54 47-65

1.7 1.4-2.2

42 35-49

P P PR

ritteri 510 470-590

22 19-24

43 40-46

49 44-54

46 44-50

88 79-97

6 69 8-75

44 33-50

1.56 1.44-1.57

47 44-50

A A PR

rarum 511 443-573

23 18-26

38 32-40

51 4-56

35 30-39

72 63-80

6 47 42-52

34 23-38

0.94 0.91-1.05

50 44-51

P A PR

tami 530 400-600

23 19-29

36 34-41

50 42-57

31 28-34

73 67-86

6-8 77 71-84

48 38-55

2.0 1.4-3.0

44 30-60

P A NPR

abbasi 541 496-579

29 27-30

48 46-51

56 52-61

53 51-58

86 79-94

8 65 57-74

45 33-50

1.56 1.07-1.87

60 51-68

A P PR

anatoliense 545 507-580

24.5 21-28

37 36-39

52 46-58

35 31.5-39

72 68-81.5

6 74 68-84

47 42-59

1.75 1.6-1.9

48.5 46.5-55

P A SP

thermophilum 555 510-620

21 21-23

40 37-46

45 40-52

46 42-53

96 81-102

8 61 44-72

36 30-42

1.7 1.2-2.8

63 50-87

A P PR


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carpocapsae 558 438-650

25 20-30

38 30-60

53 46-61

26 23-28

60 54-66

6 66 58-77

47 39-55

1.72 1.40-2.00

41 27-55

P A PR

scapterisci 572 517-609

24 18-30

39 36-38

54 48-60

31 27-40

73 60-80

6 83 72-92

65 59-75

2.52 2.04-2.8

38 32-44

P P SP

websteri 584 553-631

21 17-25

36 29-40

47 37-56

31 24-34

77 62-102

6 68 64-72

49 42-56

1.8 1.6-2.1

40 30-50

P A NP

kushidai 589 524-662

26 22-31

46 42-50

50 44-59

41 38-44

92 84-95

8 63 NA

44 NA

1.5 51 NA

A A PR

intermedium-group IJ average size between 600-800 µm

riobrave 622 561-701

28 26-30

56 51-64

54 46-59

49 45-55

105 93-111

NA 67 62.5-75

51 47.5-56

1.14 71 60-80

A A SP

intermedium 671 608-800

29 25-32

65 59-69

66 53-74

51 48-58

96 89-108

6-8

91 84-100

64 56-75

1.24 NA

67 58-76

A A SP

pakistanense 683 649-716

27 24-29

54 49-58

58 53-62

47 42-53

91 87-102

8 68 62-73

41 36-45

1.8 1.0-2.2

60 50-60

P P SP

affine 693 608-880

30 28-34

62 51-69

66 64-74

49 43-53

94 74-108

8 70 67-86

46 37-56

1.17 NA

61 NA

P A PR

ceratophorum 706 591-800

27 23-34

55 47-70

66 56-74

45 40-56

84 74-96

6-8 71 54-90

40 25-45

1.4 1.0-2.0

51 33-65

A A SP

monticolum 706 612-821

37 32-46

58 54-62

77 71-95

47 44-50

76 63-86

8** 70 61-80

45 35-54

1.4 1.2-1.5

55 49-61

P A NP

weiseri 740 586-828

25 24-29

57 43-65

60 49-68

51 44-55

NA 8 68 62-72

53 46-57

1.8 1.5-2.4

49 39-60

A A SP

sangi 753 704-784

35 30-40

51 46-54

81 76-89

40 36-44

62 56-70

8 63 58-80

40 34-46

1.5 1.2-1.6

49 42-63

P A PR

bicornutum 769 648-873

29 25-33

61 53-65

72 63-78

50 40-60

84 80-100

8 62 53-70

48 38-50

2.22 2.18-2.26

52 50-60

A A NP

feltiae-group IJ average size between 800 - 1000 µm

feltiae 849 736-950

26 22-29

62 53-67

81 70-92

45 42-51

78 69-86

8 70 65-77

41 34-47

1.13 0.99-1.3

60 NA

P P PR

thanhi 851 720-960

31 27-39

75 68-84

63 52-72

58 52-67

119 101-138

8 72 67-78

49 40-56

1.8 1.5-2.1

73 64-82

A A PR

neocurtillae 885 741-988

34 28-42

18 14-22

80 64-97

12 10-15

23 18-30

6 58 52-64

52 44-59

1.43 1.18-1.64

19 13.26

P P V

scarabaei 918 890-959

31 25-37

77 72-81.5

76 71-80

60 50-75

100 90-110

8 75 67-83

44 36-50

1.7 1.5-2.0

66 53-77

P A SP

karii 932 33 74 74 57 96 8 83 57 NA 66 A P SP


20

876-982 31-35 68-80 64-80 NA NA 73-91 42-64 57-78 kraussei 957

797-1102 33

30-36 63

50-6 79

63-86 47 NA

80 NA

8 55 52-57

33 23-38

1.10 NA

53 NA

P A SP

oregonense 980 820-1110

34 28-38

66 60-72

70 64-78

50 40-60

100 90-110

6-8 71 65-73

56 52-59

1.51 NA

73 64-75

A A SP

loci 986 896-1072

37 30-45

80 71-86

75 66-83

57 52-63

107 94-120

8 71 60-80

46 40-52

1.9 1.7-2.1

73 61-80

A A PR

glaseri-group IJ average size > 1000 µm

longicaudum 1063 NA

40 NA

81 NA

95 NA

56 NA

85 NA

8 77 NA

48 NA

1.60 62 NA

A A PR

apuliae

1064 945-1212

37 34-41

95 86-102

71 63-80

66 63-70

130 120-150

8 72 64-80

50 46-54

1.6 1.5-1.8

81 75-92

A A SP-VP

caudatum 1106 933-1269

36 34-41

82 76-89

88 80-100

52 NA

94 87-100

8 75 NA

52 NA

2.22 71 NA

A A V

glaseri 1130 864-1448

43 31-50

102 87-110

78 62-87

65 58-71

131 122-138

8 77 64-90

55 44-59

2.1 1.6-2.4

70 60-80

A A PR

puertoricense 1171 1057-1238

51 47-54

95 90-102

94 88-107

66 62-74

101 88-108

8 78 71-88

40 36-45

1.52

77

A P PR

cubanum 1283 1149-1508

37 33-46

106 101-114

67 61-77

70 NA

160 NA

8 58 50-67

39 37-42

1.41

70

A A PR

References: E%= EP/TL x 100; EP= excretory pore; EPI= epiptygma; D%= EP/ oesophagus length x 100; GuL= gubernaculum length; LF= number of ridges of lateral field at midbody level; M= mucro; MBW= maximum body width; SpL= spicule length; SW= SpL/ cloacal body width; TBL= total body width; TL= tail length; VL= vulval lips. A= absent; NA= not available; P= present; PR= protruding, NP= not protruding * Morphometric values of type isolate have incongruent and/or erroneous data in tables and text in original publication. **After Stock, unpublished data.


21

Key 2. Key diagnostic features of Phasmarhabditis spp. All measurements are in microns (From Stock and Hunt, 2005).

FEMALE MALE Tail

Species TBL

Shape Length TBL Bursa shape SpL

hermaphrodita1

(Schneider,1859) Andrássy, 1983 1799

1509-2372 Elongate, conoid 3-4 anal body

widths long

Males are extraordinarily rare neopapillosa1

(Mengert in Osche, 1952) Andrássy, 1983

2227 1817-2449

Elongate, conoid 3-4 anal body widths long

1585 1432-1771

Well developed 1.5 times as long tail

nidrosiensis2

(Allgén, 1933) Andrássy, 1983 1000-1750 Cupola-shaped

w/pointed tip 1.5-2 anal body

widths long 900-1720 Small and narrow Twice as long as tail

papillosa2

(Schneider, 1866) Andrássy, 1976 1600-3400 Cupola-shaped


widths long 1200-2400 Well developed 1-1.5 times as long as tail

valida2

(Sudhaus, 1974) Andrássy, 1983 NA Cupola-shaped


widths long NA Well developed NA

References: * Type species; NA= not available; SpL= spicule length; TBL= total body length. 1 After Hooper et al., 1999; 2 after Andrássy, 1983.


22

Key 3. Polytomus key to Heterorhabditis spp. All data from original descriptions unless otherwise specified. Morphometrics are given in microns (From Stock and Hunt, 2005).

Adults Third Stage Infective Juvenile Hermaphrodite Male Female

Species

TBL MBW EP TL RF D% E% T shape SpL GuL TREF D% BR PAS indica-group

IJ average size < 550 µm

poinari NA 350-410

NA 18-22

NA NA NP NA NA conoid NA 43-55

NA 24-32

NA NA NA NA

taysaerae 418 332-499

19 17-23

90 74-113

55 44-70

NP 82 71-96

180 110-230

conoid 39 30-42

18 14-21

122 100-146

NA 7,8 do not reach the

bursal rim

P

indica 528 479-573

20 19-22

98 88-107

101 93-109

NP 84 79-90

94 83-103

conoid 43 35-48

21 18-23

106 78-132

122 NA

1 may be out of bursa 4, 7

outwards

V

bacteriophora-group IJ average size 550-700 µm

bacteriophora 588 512-671

23 18-31

103 87-110

98 83-112

NP 84 76-92

112 103-130

conoid 40 36-44

20 18-25

76 61-89

117 NA

4, 7 outwards

P

baujardi 551 497-595

20 18-22

97 91-103

90 83-97

NP 84 78-88

108 98-114

conoid 40 33-45

20 18-22

91 63-106

NA NA P

brevicaudis 572 528-632

22 20-24

111 104-116

76 68-80

NP 90 NA

147 NA

conoid 47 44-48

22 20-24

194 162-240

88 NA

NA P

zealandica 685 570-740

27 22-30

112 94-123

102 87-119

NP 80 70-84

108 103-109

conoid 51 48-55

22 19-25

132 88-173

118 NA

4, 7 outwards

V

marelata 654 588-700

28 24-32

102 81-113

107 99-117

NP 77 60-86

96 89-110

pipette-shaped 45 42-50

19 18-22

91 67-136

113 NA

4, 7 outwards

8 does not touch

bursal rim

P


23

mexicana 578 530-620

23 20-24

102 83-109

99 91-106

NP 81 72-86

104 87-111

conoid 41 30-47

23 18-32

96 65-130

129 114-149

Arrangement: 1,2,3,2

With 4, 7 outwards. Pairs 7-9 varaible

P

megidis-group

IJ average size >700 µm

megidis 768 736-800

29 27-32

131 123-142

119 112-128

NP 85 81-91

110 103-120

conoid 49 46-54

21 17-24

128 122 NA

4,7 outwards 2,3 fused

P

downesi 879 669-1066

39 33-55

97 64-107

33 28-42

P 83 77-92

169 129-216

blunt and mucronated

46 40-53

23 40-53

NA NA 4, 7 outwards 8 does not

touch bursal rim

P

References: BR= bursal rays; D%= EP/oesophagus length x 100; E%= EP/TL x 100; EP= excretory pore; MBW= maximum body width; NA= information not available; PAS= post-anal swelling; RF= tail refractile spine; T=tail; TBL= total body width; TL= tail length; TREF= testis reflexion; V= variable.


24

Key 4. Key diagnostic features of mononchid genera considered in biological control (From Stock and Hunt, 2005).

Mononchus Bastian, 1865

Mylonchulus Cobb, 1916

Iotonchus Cobb, 1916

Oesophago-intestinal junction

Non-tuberculate Non-tuberculate Tuberculate

Position/direction of dorsal tooth

Anterior half and forward Anterior half and forward Posterior half and forward

Subventral teeth or denticles

Absent Small pair of teeth usually opposite base of dorsal tooth. Walls with 2-13 transverse rows of

minute denticles.

Absent

Key 5. Key diagnostic features of mermithid genera considered in biological control (From Stock and Hunt, 2005).

Diagnostic Features

Agamermis Cobb, Steiner and

Christie, 1923

Mermis Dujardin, 1842

Oesophagomermis Artyukhovsky, 1969

Romanomermis Coman, 1961

Strelkovimermis Rubzov, 1969

Cephalic papillae

6 4 6 6 6

Labial papillae Absent Present (2) Absent Absent Absent Oral opening Terminal Absent Terminal or slightly shifted

to ventral side Terminal Terminal or slightly shifted to

ventral side Hypodermal cords

6 6 6 8 6

Vagina shape S-shaped S-shaped S-shaped Pear-shaped S-shaped Bursal sleeve Absent Absent Absent Absent May be present Parasitic and Post-parasitic tail

With crater-like terminus

With tail appendage With small tail appendage With tail appendage With tail appendage

Eggs Without byssi With byssi Without byssi Without byssi Without byssi


25

E. Preparation of Nematodes for Identification ________________________________________________________________________ 1. Obtaining nematodes

A good stereomicroscope is essential for nematode identification and should have a range of magnification between 10 and 100X, a fairly flat field, and good resolution. Illumination by transmitted light should be as even as possible. Handling most nematodes in distilled water presents no problem; however, the use of a saline solution (i.e., Ringer's solution) is highly recommended to avoid osmotic shock. For living material, the nematodes can be mounted in Ringer's solution or water on a glass slide with supports and a cover-slide sealed with paraffin or nail polish. Perform dissections in a Petri dish with Ringer's solution and under stereomicroscope. Lift individual specimens out of the solution with the help of a handling needle of which there are many types: nylon toothbrush bristle, toothpick handling “L-shaped” needle, a dentist's probe, steel needle, a bamboo splinter, etc. For smaller nematodes, an eyebrow hair stuck on to the end of a mounted needle is very useful and does not damage the nematodes. Beginners will have some difficulty in picking up nematodes. It is recommended to use the lowest microscope magnification (to give good depth of focus and working distance) and to start with nematodes that are near the center of the dish. Place the needle underneath the nematode and lift up quickly so that the nematode is pulled through the meniscus. The time frame for doing the dissections may vary according to the different species/isolates. For steinernematids, dissections should be done 2-3 days after infection to recover first generation adults (males and females), and 5-6 days after infection for second generation adults. For heterorhabditids, dissections should be done 3-4 day after infection to recover first generation

hermaphrodites, and during the 6-9 day after infection for second generation adults (males and amphimictic females). 2. Killing and fixing Heat.-Nematodes may be killed by heating in a water bath at 60oC for 2 min. Once the specimens are killed, add the fixative (TAF) to an amount equal of Ringer's solution. The temperature of the fixative and the Ringer's solution should be approximately 60oC. Leave specimensl for 24h in this mixture of 1:1 Ringer-TAF. Then, replace solution with pure TAF. Nematodes fixed in TAF may remain in this solution for several months/years. If you desire to make permanent mount, nematodes should be dehydrated following the Seinhorst method (see section 3) An improved method that kills and fixes the nematodes in one process is that of Seinhorst (1966). The specimens are collected in a very small drop (ca. 1 ml) of water in a Syracuse watchglass or similar deep concave vessel. The fixative (i.e., TAF) is heated to 100oC and an excess (3-4 ml) is added to the nematodes. The nematodes remain in this solution for 12 hr, and then the solution is replaced with double-strength fixative. Cooling.- Samples can be stored in the refrigerator (ca. 4oC) or in water with ice to relax the nematodes by cooling. Once the nematodes are relaxed, add fixative heated previously at 65oC. Allow the fixative to act for at least 1 day. Transfer the nematodes to a Syracuse watchglass with as little fixative as possible. Ringer's solution (Woodring and Kaya, 1988) - 9 g NaCl - 0.4 g KCl


26

- 0.4 g CaCl2 - 0.2 g NaH2CO3 -1 liter distilled H2O M9 Buffer (Brenner, 1974) - 3 g KH2PO4 - 6 g Na2HPO4 - 5 g NaCl - 1 ml MgSO4 (1M) - 1 liter distilled H2O Fixatives (Seinhorst, 1959, 1966; Southey, 1986; Woodring and Kaya, 1988) Numerous fixatives have been recommended for studying different features of nematode anatomy. Double-strength fixatives are made up using half the amount of water indicated below. The commonly used fixatives are: TAF - 7 ml formalin (40% formaldehyde) - 2 ml triethanolamine - 91 ml distilled H2O F.A. 4:1 or F.A. 4:10 - 10 ml formalin - 1 or 10 ml glacial acetic acid - up to 100 ml distilled H2O 3. Preparing nematodes for permanent mounts The gonads and other structures of fixed nematodes may be obscured by the granular appearance of the intestine. Specimens can be cleared by processing to lactophenol or glycerin. Although both are good mounting media, nematodes will keep almost indefinitely if they are processed to glycerin. -Processing to glycerin Transfer fixed nematodes to a Syracuse watchglass containing 0.5 ml of Solution I (20 parts of 95% ethanol, 1 part of glycerin, 79 parts of distilled water) (Seinhorst, 1959). Place the Syracuse watchglass in a desiccator and add 95% ethanol to the desiccator so that the space

below the holding shelf is half full. Leave the watch-glass in this atmosphere for at least 12 hr in an oven at 35oC to allow slow evaporation of the ethanol from Solution I in the watchglass. Remove the watchglass from the desiccator. Then fill the watchglass with Solution II (5 parts of glycerin, 95 parts of 95% ethanol), place in a Petri dish which should be partially opened to allow slow evaporation of the ethanol, and maintain for 3 hr at 40oC. After processing the nematodes in pure glycerin, they are ready for mounting. -Mounting and labeling Aluminum double-coverslip slides (Cobb, 1917) are suitable for permanent mounts as they allow to view specimens from either side, are durable, and are less likely to crack during shipping or rough handling. If they are not available, good quality glass slides (76 x 25 x 1 mm) can be used. When using an aluminum slide, a square coverglass has to be slipped in the aluminum carrier between two pieces of cardboard. The edges of the carrier must then be pressed to hold the cardboard and the coverglass in place. Place a drop of glycerin in the center of the slide and place a round coverslip on top of the glycerine. Short pieces of glass fiber can be included to give support. Seal specimen slides with wax paraffin or a double ring of nail polish. A substance called Zut was previously recommended, but it is no longer commercially available. To seal specimen slides with paraffin, heat a 1.7 mm diameter metal tube with polished rim using a Bunsen burner or alcohol lamp. Press the tube into the paraffin and hold it against the glass to leave a ring of paraffin (the thickness of the ring should be in accordance with the size of the nematodes to be mounted). Place a drop of glycerin in the center of the ring and place a few nematodes in the bottom of the drop making sure that the nematodes are aligned and do not overlap. Place the slide on a hot plate and drop a coverglass on top as soon as the paraffin ring melts and immediately take the slide off the hot plate. Once the paraffin sets,


27

the slide is sealed and contains a small central area of glycerin and nematodes with a surrounding rim of paraffin. -Mounting cross sections The cephalic structures and the number of longitudinal chords are diagnostic characters for generic or specific determination of certain groups of nematodes (i.e., rhabditids and mermithids). These structures are best seen by making cross sections. Place the nematodes that have already been processed to glycerin in a drop of glycerin and cut into small sections (anterior end or midbody) with a razor blade or a small oculist’s scalpel. Using a needle, transfer the section to a drop of melted glycerin jelly in the center of a coverglass and orient in the proper position. Invert the coverglass and gently place on a slide with pieces of glass fiber to prevent the jelly from touching the slide. Seal the mount with paraffin or glycerin jelly.

Recipe for hard glycerin jelly (Southey, 1986): - Soak 20 g of gelatin in 40 ml of distilled water for 2 hr; - Add 50 ml of glycerin and 1 ml of phenol; - Place in water bath (70-80oC) for 10-15 min and stir until the mixture is homogenous.

4. Preparation for scanning electron microscopy Scanning electron microscope is a very useful tool for the visualization and interpretation of certain nematode features, that cannot be appreciated with a light microscope but are important for their taxonomic identification. To prepare nematodes for viewing, place them in a water bath at 60oC for 2 min to kill them. Rinse nematodes three times in Ringer’s solution (pH 7.3) or phosphate buffer (pH 7.4) (5 min each change) and prefix in 8% glutaraldehyde (glutaraldehyde 25% EM grade, diluted in Ringer’s solution). Leave nematodes overnight in this solution. The next day,

rinse nematodes three times in Ringer’s solution (5 min each change), and once in water (5 min). Post-fix in 1% osmium tetroxide (OsO4) for 2 hr. After post-fixation, rinse nematodes three times in water (5 min each) and dehydrate them using a series of ethanol washes (30, 50, 70, 90, 95 and 100%). The nematodes are then critical point dried with liquid CO2, mounted on SEM stubs, and coated with gold for 1 hr.

Plain agar plates for nematodes These plates are made at either 1% or 2% agar – 10g per liter for 1%, 20g per liter for 2%. 1. Combine agar and 1 L of double-distilled water in a large flask. 2. Autoclave for at least 30 minutes of heat. 3. Cool until comfortable to the touch. 4. Pour. 5. Allow to solidify and dry at room temperature overnight. 6. Re-bag and refrigerate.

Baby Food plates for nematodes (Stock et al., 2001) The baby food seems to have some heat resistant spores in it. To eliminate these, a double autoclave technique called Tindallization is used – autoclave agar, leave for 24 hours to allow bacteria to grow but not time enough for them to make resistant spores, then autoclave again. 1%: 10 g agar, 6 g Beechnut baby food (or any other 3 cereal baby mix) in 1 L distilled water. 1. Autoclave (at least 30 minutes of heat). 2. Leave at room temperature in the lab for 24 hours, then autoclave again. 3. Cool (water bath not necessary if swirled occasionally). 4. Pour 2%: 20 g agar, 0.5 g Beechnut baby food in 1 L distilled water. Autoclave (at least 30 minutes of heat), leave for 24 hours, then autoclave again, cool (water bath not necessary if swirled occasionally), and pour.


28

F. Atlas _____________________________________________________________


29

General Morphology

From Maggenti, 1981 Transversal section

Stoma

Anterior region

Amphid

Oesophagus

Nerve Ring

Cuticle

Excretory cell

Digestive system

Tail

Reproductive system Egg

Excretory pore

Dorsal nerve

Intestinal Cavity

Muscles

Ventral nerve

Epidermis

Cuticle

Cuticle

General body cavity

Excretory canal

Glandular Cell


30

Excretory System

From Poinar, 1983 References: L: canal, G: excretory cell (gland), P: excretory pore; T: excretory canal A. H-shaped (e.g. Oxyuroids), B. I-shaped (e.g. Diplogasterids, Tylenchoids), C. Inverted U-shaped (e.g. Chromadoria), E. Single-cell (e.g. Chromadorids).


31

Digestive System

Types of Oesophagi

A. Crenate (e.g. Enoplids), B. Multibulbar (e.g. Enoplids), C. Stichosome (e.g. Mermithids), D. Two-part oesophagus (e.g. Dorylaimids), E-F. Three-part oesophagus (E. Rhabditis, F. Diplogasterid). From Maggenti, 1981 Types of corpus

A. Enoplida, B. Dorylaimida, C. Rhabditida, D. Tylenchida. From Maggenti, 1981


32

Nervous System

From Maggenti, 1981 Phasmid

From Maggenti, 1981

Nerve Ring

Nerve Ganglia

Nerve Canal


33

Male Reproductive System

From Dropkin, 1989

spicule

intestine

testis

spermatozoa

spermatocites

Vas deferens

gubernacule bursa


34

Female Reproductive System

From Dropkin, 1989


35

G. Molecular Diagnostics / Phylogenetics

PHENOL-CLOROPHORM EXTRACTION

Use: 1.7 ml Eppendorf tubes -Transfer nematodes to tubes and add the following: -500µl of TH pH 8.0 -15µl of 30% SDS -20µl of proteinase K (1ng/ µl) *note: if sample contains for example 100µl (nematode suspension) add ONLY 400µl of TE. This is adjust amount of TE added so final volume is 500µl. -Incubate at 50C in water bath (incubation time varies according to the sample). -Check periodically for digestion. If this is not completed at the end of the day, add 15µl more of proteinase K and leave overnight. -If digestion is completed place sample in freezer until next day. -The day after add 10µl of RNAase to each sample. -Vortex and incubate at 37C for 1 hour. -Centrifuge at lowest speed (4 in our centrifuge) for 2 minutes. -Transfer supernatant to a new 1.7 eppendorf tube. Be careful when pipetting, avoid collecting the debris. -Add equal (to the amount of volume that is already in the tube) volume of phenol (in the hood) to each tube -Vortex briefly and spin (full speed for 5min. -Pipet supernatant carefully, (do not touch the phenol interface) and place it in a new eppendorf tube. -Discard the phenol in the 'phenol disposable container" in the hood. -Add equal volume of 24:1 clorophorm/isoamyl alcohol. -Vortex briefly and spin at full speed for 5min. -remove upper interface carefully and transfer to a new eppendorf tube. -Discard the isoamyl alcohol in the 'phenol disposable container" in the hood.

-Centrifuge again the remaining clorophorm to be able to recover more upper interface (5min. at full speed) -Add Na acetate (to help precipitation) 10µl per 100µl. example if you have 400µl in the tube add 40µl of Na acetate (1M). -Add 100% ethanol to cover full volume of the tube and mix gently by hand. Place tubes in freezer and leave them overnight. -The following day spin in refrigerator's centrifuge at 4C for 10min at full speed. -Discard liquid and allow samples to dry in desiccator. -When samples are fully dried add 25µl of TE to each tube. If pellet is big, add 50µl of TE. Samples are now ready for spectrophotometry reading

CHELEX DNA EXTRACTION OF SINGLE NEMATODE

You will need: Chelex resin (Sigma sells this as "Chelex 100"), Proteinase K, Sterile water 1) make 5% chelex soln (e.g. for a 10mL solution) A) place stir bar in 50mL conical tube held upright in a beaker B) add 0.5g Chelex resin into conical C) fill to 10 mL with sterile water This solution can be stored in the refrigerator for up to 1 month 2) Use a sterile razor blade to cut the end off of a P1000 tip to make the opening bigger (Chelex beads are too big to fit otherwise) 3) With Stir Bar going, mix up chelex soln (make sure chelex beads are spinning in the water) and take 20uL and add to tubes(make sure chelex beads are in there ). 4) Add 1 uL of proteinase K solution to each tube (20mg/mL). 5) Add nematodes to tubes, cutting each in half with a clean razor blade. 6) Incubate at 56 C for 1 hour. 7) Boil at 100 C for 8 min (100 C program in thermocycler).


36

8) Cool down for a few seconds (if using thermocycler bring down to 40 C for about 30 sec). 9) Vortex for about 30 seconds. 10) Use 4 uL of this in your PCR reactions, making sure no beads get in there. DNAZOL DNA ISOLATION (protocol for individual nematodes) 1. Place each live nematode in 100 mL digestion solution in a 0.5 ml Phoenix tube 100 mM Tris HCL pH 7.6 200 mL 200 mM NaCl 200 mL 0.5 M EDTA pH 8.0 400 mL 10 % Sarkosyl 200 mL Proteinase K (10 mg/ml) 20 mL MP3A water 980 mL Note Proteinase K should be made fresh – usually 0.005 g/0.5 ml 2. Leave at room temperature for at least 0.5 hour to allow worm to digest solution 3. Digest overnight in 56 �C water bath. 4. Heat kill proteinase with 95 �C program in PCR machine (15 minutes) 5. Freeze and then thaw tubes 4 times 6. Centrifuge sample for 5 minutes at 10,000 rpm 7. Remove 95 mL of solution, leaving bottom 5 mL, and add it to 1 ml of DNAzol isolation reagent in a 1.7 ml tube 8. Add 4 mL Polyacryl Carrier solution, vortexing Carrier first 9. Mix tube by inverting 5 times 10 Add 0.5 ml 100 % ethanol and mix by inverting 10 times 11. Let sample sit at room temperature for 5 minutes 12. Pellet DNA by centrifuging at 7,000 rpm for 5 minutes 13. Pour off ethanol and then wash DNA twice with 800 mL 75% ethanol, spinning again if pellet breaks loose 14 Pour off final ethanol, then remove last of it with a pipetter 15. Allow visible ethanol to evaporate but do not allow pellet to dry 16 Resuspend in 6 mL lTE AGAROSE GEL 1. Make 1.3% gel: 40 ml of 1X or 0.5X TAE or TBE and 0.52g agarose Product: 7 mL water, 2 mL BPB dye, 1 mL vortexed product

Standard: 6 mL water, 2 mL BPB dye, 2 mL standard (Standard: 1 part Promega PhiX174/HaeIII marker, 2 parts lHind III marker) 2. Run 70-100 Volts for a couple of hours 3. Stain with 200 ml same running buffer and 35 mL Ethidium Bromide (1 mg/ml) on mixing machine for 15 minutes 4. Destain with 200 mL di water and 500 mL MgCl (0.1 M) on mixing machine for 20 minutes Notes: TBE resolves better than TAE but can precipitate and therefore stores less well. TAE runs slightly faster but disadvantage is slightly less resolution.

PRIMERS (dilutions)

PCR=10mM, Sequencing=3mM 1. Dilute with TE from stock for PCR primers (i.e. make 100 µL of 10 mM); degenerate primers use at 20 mM for PCR. 2. Dilute PCR primers to make Sequencing primers (usually make 50 µL:15 µL of 10 mM PCR primer and 35 µL lwater) – no TE here as it inhibits cycle sequencing. For new dry primers, add 250 µL lTE (Operon says pH 7), then determine concentration in mm based on pmoles given on oligo sheet. Sample calculation: DP585: Given 99126.16 pmoles on sheet and 250 µL lTE DNTPs Purchased at 100 mM, so each one is diluted to 10 mM for PCR and combined (all 4) in equal amounts. There are already 1.7 µL tubes with the diluted dNTPs in the same small blue box in freezer – just aliquote equal amounts of these into a smaller tube or dilute more into the 1.7 µL tubes if necessary.Enzymatic Treatment of PCR product


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Cloning Procedures1. PCR product Preparation:

1.Spin filtration

Use Ultrafree-MC spin filtration columns (Millipore 30,000 mwt filter) to clean up PCR product – see protocol in box for more details. a. Wash product (all remaining) with 300 µl of DEPC-treated water twice. Note that the filter must not be spun in excess of 5000 x g – do 5,000 rpms for 5 minutes on the Hermle centrifuge . b. Remove product from filter by rinsing surface of membrane with DEPC-treated water using pipettor (but do not touch membrane surface with pipet tip). c. Adjust volume to ~20 µl (or concentrate on speedvac if original gel band was light) 2. Ligation: 5 ul 2x ligation buffer (pre-aliquotted) 3 ul filtered PCR product (see step 1 above) 1 ul pGEMT vector – in blue freezer box 1 ul T4 ligase Incubate ligation overnight at ~4C in refrigerator 3. Transformations Before beginning make sure LB plates are dry and at room temperature – plates right from the refrigerator can be dried for 3 hours in the 37C incubator. a. Thaw competent cells on ice, mixing very gently b. Add 4 µl of ligation to a 2 ml eppendorf tube - freeze remaining 6 ul. c. Add 50 µl of thawed cells to eppendorf tube d. Gently mix (finger "vortexing") e. Place tube on ice for 90 minutes f. Heat-shock cells for 50 seconds at 42 C in water bath g. Place transformation on ice for 2 minutes h. Add 950 µl SOC medium (at RT) Continued on next page i. Incubate transformation at 37 C for 1 hr (not longer) using rotating wheel in incubator j. Spread 50 µl X-gal (20 mg/ml) onto room

temperature LB plates 1/2 hour before plating transformed bacteria. X-gal is kept in the –20 freezer in room 483. k. Plate 100 µl of cells per plate (plate with IPTG, amp, & X-gal). Minimum of 2 plates per tranformation. l. Place tubes with remaining transformed bacteria in autoclave waste under sink in room 582.

4. Boil preps a. Pick piece of colony from master plate to 50 µl of sterile water – vortex. b. Boil 10 minutes in dry block (at at least 100C) (put brick on top of tubes) c. Spin tubes for 3 minutes d. Use 1 ul of supernatant as DNA source for 20 µl PCR reaction – use red label pipettors for adding supernatant as PCR product has been amplified by the bacteria.

5. Storing clones a. Use cryotubes. Grow cells in 800 µl of LB with amp overnight (15 hr) on rotator wheel. b. Add 200 ul of 80% glycerol to grown cells. Mix by vortex. Freeze in –70C freezer. Transfer to liquid nitrogen tank and record placement on grid.

6. Broth culture/miniprep kit a. Put 5 ml LB with ampicillin broth in disposable culture tube. b. Inoculate with small piece of colony. c. Grow for 12 hours in shaker incubator at 37 C. d. Spin down tubes in large centrifuge for 5 minutes or until pelleted. e. Pour off broth, then pellet is ready to begin miniprep kit – see protocol in kit.

Media preparation LB plates 10g Tryptone 5 g Yeast Extract 5 g NaCl 1. Bring up to 1 L with MP3A water and pH to 7 with 5 N NaOH (about 200 µL should do it). 2. Add 15 g agar, autoclave (at least 30 minutes of heat), cool in 50 C water bath. 3. Once cooled, add necessary


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supplements: IPTG 100mM: add 5 µL to 1 L LB agar for final concentration of .5mM. This is kept in the refrigerator in room 483. Ampicillin - add 2 tubes (25 mg/µL) to 1 L of agar for final concentration of 75 mg/µL. Aliquote and kept in the freezer

4. Pour plates. Leave at room temperature several hours to solidify. 5. Place in 37C incubator overnight to dry

and check for contaminant growth. 6. Once plates are dry and contaminant-free, they can be re-bagged and placed in the refrigerator. Before use bring plates to room temperature and spread 50 µL of X-gal (20 mg/ml) at least 1/2 hour before spreading bacteria.


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References: 1. Generic and above taxonomic hierarchies (family, etc); 2. Species and/or infraspecies levels; 3. Species level; 4-5. Species and/or infraspecies levels; 6. Infraspecific level

12

3

54 6

Mitochondrial DNA

Ribosomal DNA

Schematic Representation of Genes considered for Nematode Phylogenetics and Diagnostics


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Origin of Nematode Parasitism in Insects and other Arthropods (Modified from Poinar, 1983)


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Phylogenetic Framework of the Phylum Nematoda based on SSU rDNA Sequence Analysis

(Modified from Blaxter et al. (1998)

References: * Parasitic and/or associated with insect pests


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H. References _____________________________________________________________________________ Brenner, S. (1974). The genetics of Caenorhabditis elegans. Genetics 77: 71-94. Choo, H.Y., Kaya, H.K. and Kim, J.B. (1989) Agamermis unka (Mermithidae) parasitism of Nilaparvata lugens in rice fields in Korea. Journal of Nematology 21, 254-259. Choo, H.Y. and Kaya, H.K. (1994) Biological control of the brown planthopper by a mermithid nematode. Korean Journal of Applied Entomology 33, 207-215. Cobb, N. A. (1917). Notes on nemas. Contribution Science. Nematology No. 5. pp 117-128. Creighton, C.S. and Fassuliotis, G. (1983) Infectivity and suppression of the banded cucumber beetle (Coleoptera: Chrysomelidae) by the mermithid nematode Filipjevimermis leipsandra (Mermithida: Mermithidae). Journal of Economic Entomology 76, 615-618. De Ley, P. and Blaxter, M. (2002) Systematic position and phylogeny. In: Lee, D. L. (ed.) The Biology of Nematodes. Taylor and Francis, New York, pp. 1-30. Dropkin, V.H.(1989). Introduction to Plant Nematology. John Willey & Sons, Inc. NY, 304 pp. Hooper, D. J., Wilson, M.J., Rowe, J.A., Glen, D.M., 1999: Some observations on the morphology and protein profiles of the slug-parasitic nematodes Phasmarhabditis hermaphrodita and P. neopapillosa (Nematoda: Rhabditidae). Nematology, 1, 173-182. Kaya, H.K. and Stock, S.P. (1997) Techniques in insect nematology. In: Lacey, L. A. (ed.) Manual of

techniques in insect pathology. Biological Techniques Series, Academic Press, San Diego, California, pp. 281-324. Lacey, L. A., Frutos, R., Kaya, H.K. and Vails, P. (2001) Insect pathogens as biological control agents: do they have a future? Biological Control 21, 230-248. Maggenti, A. R. (1981) General Nematology.Springer-Verlag, New York, pp. 372. Nickle, W.R. (1972) A contribution to our knowledge of the Mermithidae (Nematoda). Journal of Nematology 4, 113-146. Petersen, J.J. (1985) Nematode parasites. In: Chapman, H. C. (ed.) Biological control of mosquitoes. American Mosquito Control Association Bulletin 6, pp. 110-122. Poinar, G.O., Jr. (1975) Entomogenous Nematodes. E.J. Brill, Leiden, The Netherlands, 317pp. Poinar, G.O., Jr. (1983) The Natural History of Nematodes. Prentice-Hall, Inc., Englewood Cliffs, New Jersey, 323pp. Poinar, G.O., Jr. (1990) Entomopathogenic nematodes in biological control. In : Gaugler, R. and Kaya, K. H. (eds.) Taxonomy and biology of Steinernematidae and Heterorhabditidae. CRC Press, Inc., Boca Raton, Fl., pp. 23-74. Seinhorst, J. W. (1959). A rapid method for the transfer of nematodes from fixative to anhydrous glycerin. Nematologica 4: 117-128.

Seinhorst, J. W. (1966). Killing nematodes for taxonomic study with hot F. A. 4:1. Nematologica 12: 178.


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Southey, J. F. (1986). Laboratory methods for work with plant and soil nematodes. 6th Ed. Ministry of Agriculture, Fisheries and Food. Her Majesty's Stationary Office, London, UK. Stock, S. P. and Hunt, D. J. (2005). Nematode Morphology and Systematics. In: Grewal, P. S., Ehlers, R. U. and Shapiro-Ilan, D. I. (eds.) Nematodes as Biological Control Agents. CAB International (In Press).

Webster, J.M. and Thong, C.H.S. (1984) Nematode parasites of orthopterans. In: Nickle, W.R. (ed.) Plant and insect nematodes. Marcel Dekker, Inc., New York, USA, pp. 697-726. Woodring, J. L. and Kaya, H. K. (1988). Steinernematid and heterorhabditid nematodes: a handbook of techniques. Southern Cooperative Series Bulletin. 331. Arkansas Agricultural Experiment Station, Fayetteville, Arkansas. 30 pp.

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