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Nematoda and Nematomorpha

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The phylum Nematoda comprises a wide range of roundworms that can be grouped either in nutritional (microbotrophic, predaceous, or parasitic in plants, invertebrates, or vertebrates) or ecological (soil, marine, freshwater, ectoparasitic, endoparasitic, or free-living) categories. Nematode representatives of essentially all the nutritional categories mentioned above occur in freshwater during one or more life stages. The chapter describes those nematodes in which all or a large part of life cycle occurs in freshwater; and briefly discusses the nematode parasites of vertebrates that live in or frequent freshwater usually occur in the freshwater habitat only as eggs or within intermediate hosts. The phylum Nematomorpha comprises the freshwater (Gordiaceae) and marine (Nectonematoidea) hairworms. All hairworms have parasitic lifestyles. Although freshwater hairworms usually develop in large terrestrial invertebrates (mostly insects), they all enter freshwater to mate, oviposit, and produce infective larvae, which in turn encyst in various aquatic invertebrates and amphibians. This chapter offers information on the phylogeny, morphology, physiology, developmental history, life history, and ecology of Nematoda and Nematomorpha.
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Ecology and Classification of North American Freshwater Invertebrates
Copyright © 2010, Elsevier Inc. All rights reserved.2010
Chapter 9
237
Nematoda and Nematomorpha
I. NEMATODA
A. Introduction
The phylum Nematoda comprises a wide range of round-
worms that can be grouped either in nutritional (microbo-
trophic, predaceous, or parasitic in plants, invertebrates, or
vertebrates) or ecological (soil, marine, freshwater, ectopar-
asitic, endoparasitic, or free-living) categories[63]. Relatively
little attention has been devoted to this important group of
invertebrates by freshwater biologists, probably because
of difficulties associated with sampling, extraction, and
identification.
Nematode representatives of essentially all the nutritional
categories mentioned above occur in freshwater during one
or more life stages. However nematodes are treated here only
if all or a large part of their life cycle occurs in freshwater
habitats (these habitats are discussed elsewhere). Nematode
parasites of vertebrates that live in or frequent freshwa-
ter usually occur in the freshwater habitat only as eggs or
within intermediate hosts (Fig. 9.12) and are only briefly
mentioned in this chapter. Mermithid parasites of aquatic
insects (Fig. 9.17) are included since the eggs, infective
stages, postparasitic juveniles, and adults occur in freshwa-
ter habitats. Also, because of their large size, they are easily
spotted by collectors.
The keys include select nematode genera reported from
North America that are considered obligate aquatic species
(those which live nowhere else) as well as a few that also
occur in wet soil or even brackish waters. It is often dif-
ficult to place ecological tags on nematode genera since
each species may live in a different ecozone. The system-
atic arrangement of these genera is listed in Appendix 9.1.
The reader should note that undescribed species or species
in genera other than those included here may be encoun-
tered in routine sampling.
B. Phylogeny and Fossil Record
The origins of the Nematoda are shrouded in mystery. Even
the phylogenetic relationships of the various clades are not
completely known. Molecular studies are being used to
show relationships within the phylum, and the previous sys-
tem of classification is now crumbling and being replaced
by one supported by genomic studies. The classification
system used in the present chapter differs considerably from
that of the earlier editions and is based on the molecular
approach used in a recently published book on freshwater
nematodes[23]. This system will certainly be modified in
forthcoming years as more is learned about morphological
and molecular convergence. A complete phylogenetic sys-
tem can only be ascertained when the full genome of each
species is determined. In the meantime, readers should refer
George O. Poinar Jr.
Department of Zoology, Oregon State
University, Corvallis, Oregon
I. Nematoda
A. Introduction
B. Phylogeny and Fossil Record
C. Morphology and Physiology
D. Development and Life History
E. Ecology
F. Collecting and Rearing Techniques
G. Identification of Selected Nematoda
II. Nematomorpha
A. Introduction
B. Phylogeny and Fossil Record
C. Morphology and Physiology
D. Development and Life History
E. Sampling
F. Identification of Nematomorpha Genera
III. Selected References
Ecology and Classification of North American Freshwater Invertebrates
238
to earlier works[12] summarizing ideas on nematode phylog-
eny and morphological evidence suggesting that Nematoda
arose from a basal clade of Gastrotricha[46].
The discovery of Palaeonema shows that certain clades
of the Enoplida had formed associations with early land
plants some 400 mya (million years ago)[80] (Fig. 9.1). It is
difficult to determine whether the ancestors of Palaeonema
occurred in a marine or freshwater habitat since mem-
bers of that group today occur in moist soils as well as in
both the marine and freshwater habitats (but not as plant
endoparasites). It is possible that its presence inside plant
tissues was associated with osmotic requirements found in
its previous habitat. The ecology of the Devonian habitat
appears to be one of geyser activity with hot springs peri-
odically flooding the region with silica-rich waters.
After Palaeonema, the next oldest fossil freshwa-
ter nematode is Cretacimermis libani Poinar, Acra,
and Acra[79] parasitizing an adult true midge (Diptera:
Chironomidae) in 120–135-million-year-old amber from
Lebanon. Almost as old is Cretacimermis burmitis emerg-
ing from an adult biting midge (Diptera: Ceratopogonidae)
in 100-million-year-old amber from Burma (Fig. 9.2)[67].
Some possible fossil tracks of aquatic nematodes occur
in the Upper Precambrian of Australia and Europe[32]. Tracks
similar to these have been reported in the Green River sedi-
ments of Lake Uinta (Middle Eocene) (Fig. 9.3)[54]. These
tracks were made during the second lacustrine phase of the
Green River sediments when Lake Uinta was a large fresh-
water lake occupying the Uinta and Piceance Creek Basins
in Utah and Colorado. Some Precambrian tracks could well
be nematodes since it is highly probable that the phylum
dates back to that period.
C. Morphology and Physiology
Nematodes are nonsegmented, wormlike invertebrates
lacking jointed appendages but possessing a body cavity
and a complete alimentary tract (Fig. 9.4). While lacking
specialized respiratory and circulatory systems, nematodes
possess a well-developed nervous system, an excretory
system, and a set of longitudinal muscles.
With the exception of the family Mermithidae, most
freshwater nematodes are under 1 cm in length. Although
FIGURE 9.1 The oldest known nematode, Palaeonema phyticus, occurred
in the stomatal chambers of the Early Devonian (396 mya) land plant,
Aglaophyton major. Tentatively assigned to the order Enoplia, this fossil
has characters of freshwater nematodes; however, it appeared to have been
a plant endoparasite.
FIGURE 9.2 The Early Cretaceous Cretacimermis protus leaving its
biting midge host (Diptera: Ceratopogonidae) in Burmese amber. The
nematode probably lived in a freshwater source some 100 mya.
FIGURE 9.3 Fossil tracts thought to be made by freshwater nematodes.
From the 50-million-year-old (Middle Eocene) Green River sediments of
Lake Uinta in the Soldier summit area near Provo, Utah.
Chapter | 9 Nematoda and Nematomorpha 239
the basic body shape and anatomic plan of all nematodes are
similar, some characteristics of most freshwater nematodes
are lacking in many terrestrial forms. One of these is the
presence of three unicellular hypodermal glands com-
monly located in the upper part of the tail, immediately
behind the anus (Figs. 9.4, 9.5b, 9.7d). These caudal glands
produce a secretion that is carried by a canal to the tip of
the tail. At this location, the canal is attached to a valve
(spinneret), which passes through a pore in the tail termi-
nus (Fig. 9.5b). The secretion produces an adhesive deposit
by which the nematodes can attach themselves to surfaces
of submerged rocks, plants, invertebrates, and debris.
Mucus may also be produced by pharyngeal glands of
aquatic nematodes. The deposits from the caudal and pha-
ryngeal glands can be used to form slimy traces over the
substratum. Microorganisms becoming entrapped in these
deposits serve as food when the nematodes ingest the
mucus together with the attached microorganisms[85].
1.  Cuticle
The exterior body covering on all nematodes is a noncellu-
lar, flexible, multilayered structure called the cuticle (Fig.
9.6). It is secreted by a layer of underlying hypodermal
cells and covers the entire external surface of the nematode,
with portions entering various external openings such as
the stoma, rectum, vagina, amphids (lateral sense organs),
and sometimes the excretory pore. The cuticle is composed
of an outer cortex, a middle matrix, and an inner fiber or
basal zone. All these regions are constructed of sublayers,
which vary in thickness according to the species and stage.
The outermost lipoidal layer of the cortex is important
because it serves as a semipermeable membrane, allowing
water and solutes to pass in and out of the body cavity.
The cuticle of freshwater nematodes may bear longitu-
dinal striations, punctations, bristles, setae, or somatic alae
(Figs. 9.7b, 9.8a,d, 9.18b,c). When present, the bursa is
attached to the male tail and is composed of flattened flaps
of cuticle which guide and hold the male in place during
mating (Fig. 9.7c). These cuticular structures are useful
characters for identifying nematode groups or species. The
cuticle is normally shed four times during development. At
each molt (ecdysis), the old cuticle splits and is discarded.
In a few forms, the old cuticle is retained, producing a
double cuticle or sheath around the nematode.
2.  Hypodermis
The hypodermis is a layer of tissue that is located beneath the
cuticle which is responsible for the formation of the cuticle.
FIGURE. 9.4 Adult female of Plectus (Plectidae). Center insert shows tail of male. M, mouth; S, stoma; P, pharynx; N, nerve ring; E, excretory pore;
B, valvated basal bulb of pharynx; I, intestine; R, rectum; A, anus; C, caudal glands; T, spinneret; V, vulva; O, ovary; F, egg; D, spicule; G, gubernacu-
lum; L, genital papillae. Drawing by A. Maggenti, University of California, Davis; labeling by G. Poinar.
Ecology and Classification of North American Freshwater Invertebrates
240
3.  Muscular System
Nematode movement is controlled by longitudinal muscles
because these organisms do not possess circular muscles.
These somatic muscles are composed of many adjacent
cells which are innervated by nerves lodged in the ventral
and dorsal hypodermal cords. Most nematode movement
superficially resembles that of many other elongate, stiff-
bodied, appendage-less vertebrates (e.g., snakes) and inver-
tebrates (e.g., larvae of biting midges). In locomotion, the
side-to-side bending of the body is produced by alternat-
ing contractions and relaxations of opposite muscle sec-
tors, which are controlled by the central nervous system.
When placed on flat surfaces in the laboratory, nematodes
normally crawl on their sides. Some aquatic nematodes can
swim by rapid vibratory or thrashing, side-to-side move-
ments of the body (Fig. 9.5c).
4.  Digestive System
All freshwater nematodes except postparasitic and adult
Mermithidae contain a continuous alimentary tract com-
posed of a stoma, pharynx, and intestine (Fig. 9.4). The
stoma (mouth) is the most anterior part of the digestive tract
and is usually lined with cuticle. Its walls are formed by
a fusion of various smaller segments or rings (rhabdions),
which may be separated as is found in the Cephalobidae,
or fused, as in the Rhabditidae (Fig. 9.25a). The structure
of the stoma generally reflects the food selection of the
FIGURE 9.5 (A) Gradually tapering pharynx of a freshwater Dorylaimus;
note the absence of distinct valve in basal portion of pharynx; arrow shows
the nerve ring. (B) Tail of a freshwater nematode showing three caudal
glands (CG) and the caudal gland tube (T). (C) Juvenile freshwater nema-
tode capable of swimming by rapid vibratory body movements.
FIGURE 9.6 As shown here, the nematode cuticle is composed of many
layers. The number of layers depends on the nematode species and stage.
It often expands into the coelom to form longitudinal cords
between the muscle fields. Most nematodes possess lateral,
dorsal, and somewhat ventral cords that contain the nuclei
and other cytoplasmic inclusions of the hypodermis. The
hypodermal cords originate as distinct cells but later may
lose their identity and form a syncytium.
Chapter | 9 Nematoda and Nematomorpha 241
bearer. For example, predaceous nematodes tend to have a
wide stoma armed with teeth (Fig. 9.8b), a stylet, or a spear
(Figs. 9.8a, 9.26b), whereas microbotrophic forms gener-
ally have a small tubular stoma (Fig. 9.25a) and omnivores
a funnel-shaped mouth cavity (Fig. 9.18a). Plant parasites
have protrusible stylets which can be inserted into plant tis-
sues (Fig. 9.26a), and the infective stages of invertebrate
parasites often have a small stylet used in penetrating the
cuticle or intestinal wall of the host (Fig. 9.11c).
The pharynx (or esophagus) lies between the stoma and
intestine and passes food from the mouth into the intestine
(Fig. 9.4). It is usually composed of radial muscles, which
upon dilation allow the pharynx to function as a pump or
vacuum. The pharynx also contains glands that produce
FIGURE 9.7 (A) Midbody region of a
female Pellioditis (Rhabditidae) showing
the vagina (arrow) and paired opposite
uteri filled with developing eggs; note
copulation deposit surrounding the vulva
opening. (B) Longitudinal striations lining
the cuticle of Dorylaimus (Dorylaimidae).
(C) Tail of male Pellioditis (Rhabditidae)
showing spicule (S), gubernaculum (G),
bursa (B); note cuticular rays supporting
the bursa. (D) Tail of an aquatic nematode
showing caudal gland mucous being emit-
ted from the terminus (arrow).
Ecology and Classification of North American Freshwater Invertebrates
242
FIGURE 9.8 (A) Anterior end of
an aquatic nematode showing a nar-
row tube-shaped stoma (arrow) and
cuticular bristles or setae. (B) Large
oval, cup-shaped stoma of a preda-
tory aquatic nematode; note the dor-
sal tooth (arrow) on stomal wall. (C)
Nematode bearing a medium-sized,
circular amphid. (D) Dorsal view of
a nematode with large cup-shaped
amphids (arrows).
secretions used in digestion, hatching, molting, and pen-
etration of host tissue. There are normally three portions
to the pharynx: (1) the corpus (sometimes enlarged to
form a distinct metacorpus or median bulb); (2) the isth-
mus (usually surrounded by the nerve ring); and (3) the
basal bulb (often containing valves to keep food particles
from reversing their direction of flow) (Fig. 9.25a). When
nematodes feed, material is sucked up into the mouth by
reduced pressure resulting from contraction of the radial
muscles of the pharynx. Food is passed along the lumen
by waves of alternating contractions and relaxations of the
pharyngeal muscles. The pharyngeal lumen opens behind
the food, thus creating suction pressure, and closes in front
of the food, forcing it through.
The intestine is a single-cell-thick cylindrical tube that
runs from the pharynx to the anal opening (cloacal opening
in males). It is composed of an anterior ventricular region,
a mid-intestine section or intestine proper, and a posterior
rectal region (Fig. 9.18d) (not always obvious). The inner
surfaces of the cells facing the lumen are lined with micro-
villae, which seem to be responsible for the major uptake
of nutrients. A pharyngeal-intestinal valve is usually
Chapter | 9 Nematoda and Nematomorpha 243
located in the anterior portion, behind the basal bulb of the
pharynx. In some freshwater forms, this valve is quite dis-
tinct and elongated. An intestinal-rectal valve is located at
the junction of the intestine and rectum. This unicellular
sphincter muscle controls the amount of water passing out
of the body. Many rhabditid-type nematodes have three
rectal glands, which empty into the rectum. These glands
are not to be confused with the caudal glands, which pro-
duce secretions from the tail terminus (Fig. 9.4). In adult
Mermithidae, the intestine becomes detached from the
remainder of the alimentary tract and serves as a food stor-
age organ (trophosome).
5.  Excretory System
Although variable in structure, the excretory system of
nematodes falls into two basic types. The first is frequently
found in freshwater forms and consists of a ventral excre-
tory gland (renette cell) connected by an excretory duct
to the excretory pore on the surface of the cuticle. The
ventral pore usually opens in the pharyngeal or anterior
intestinal region of the body. The second type is a tubu-
lar system consisting of a series of longitudinal excretory
canals; pooled contents of these canals pass into a joining
canal which connects with the excretory duct and pore.
Most freshwater nematodes excrete nitrogen in the form
of ammonia or urea. The excretory system also has an
osmoregulatory function of removing water that diffuses
into the pseudocoelom and so regulates the amount of tur-
gor pressure in the body cavity; some scientists believe
that this is the primary function of the “excretory” system.
6.  Respiration
Most nematodes are aerobic organisms, at least during
their developmental period. Many nematodes can survive
short periods of anaerobic conditions but only a few forms
can survive anoxia indefinitely. Strayer[97] listed represent-
atives of three nematode genera that he considered as ben-
thic species found under anoxic conditions in Mirror Lake.
A specialized respiratory system is lacking in nematodes.
Instead, they obtain and lose gases by simple diffusion.
Body cavity cells, when present, appear to play no role in
oxygen transport.
7.  Nervous System
The nervous system of nematodes centers around a central
anterior mass of ganglia or “brain,” which connects with
nerve cords extending anteriorly and posteriorly through the
body. The central ganglionic mass is closely associated with
the circumpharyngeal commissure or nerve ring (Figs. 9.4,
9.5a, 9.25a). Nerves extending anteriorly from the nerve
ring innervate the amphids and cephalic sense organs, while
those running posteriorly control the male tail papillae.
8.  Sense Organs
The size, shape, position, and spatial arrangement of sense
organs are all characteristic of nematode species or groups
and are therefore frequently used as taxonomic charac-
ters. The anterior sensory papillae are found on the nema-
tode head. The basic number is 16, but usually some have
become vestigial. An inner circle of six labial papillae is
normally present around the mouth, a second ring of six
labial papillae slightly behind the first set, and a third cir-
cle of four cephalic papillae still further back on the head.
In mermithids, the cephalic papillae may also include adja-
cent enlargements of the hypodermal tissue.
The amphids are paired, lateral sense organs that open
to the exterior on the nematode cuticle (Fig. 9.8c,d). They
may be located on the same circle as the labial or cephalic
papillae or further back in the neck region. The term
amphid generally refers to the exterior configuration of the
amphidial opening, although the amphid also consists of an
amphidial pouch, amphidial gland, and amphidial nerves.
The structure of the amphidial openings and their loca-
tion are very important taxonomic characters. They can
vary from small pores to large circles, pockets, or spirals.
Sexual dimorphism may be evident with either the male or
female possessing a considerably larger (usually the same
general shape) amphid. Amphids may be multifunctional
and evidence suggests that they serve as photoreceptors,
olfactors, or sensory glands. Some are sensitive to pH and
various ions.
Other innervated papillae on nematodes are deirids
(paired papillae located laterally near the nerve ring), post-
deirids (similar structures located at the midbody), and
genital papillae (tactile papillae on the ventral and ven-
tromedian surfaces around the cloacal opening of males).
When elongate, the genital papillae are usually attached
to thin membranous outgrowths of cuticle (bursa) (Fig.
9.7c). When the genital papillae possess organs of secre-
tion, which can be submerged, protrudint, echinate (with
projecting setae), they are called supplements. The pres-
ence or absence of a bursa and its structure are important
taxonomic characters.
A few freshwater nematodes (as well as marine spe-
cies) possess what are possibly light receptor organs, often
described as eyespots or pigment spots. These receptors nor-
mally consist of pigmented areas in the neck region, which
may or may not be associated with a cuticular-structured
lens. If a lensatic body is present, then the organ is some-
times referred to as an ocellus or pseudocellus.
In some of the Enoplida, threadlike structures occur
called metanemes located in the lateral epidermal chords
that are thought to serve as stretch receptor organs.
Members of the Enoplidae often have cephalic slits as an
extra pair of sensory organs located lateroventrally on the
tip of the head. Representatives of the Oncholaimidae pos-
sess a connection between the reproductive and digestive
Ecology and Classification of North American Freshwater Invertebrates
244
systems. Since these nematodes have traumatic semination
(sperm is injected into the body cavity), these tubes, which
are all part of the demanian system, play a role in conduct-
ing sperm to the ovary.
9.  Reproductive System
Most nematodes, including freshwater taxa, reproduce by
amphimixis (sperm and eggs come from separate individu-
als)[45]. However, some forms have developed uniparental
reproduction (i.e., autotoky), and this condition also occurs
in aquatic genera. In the latter, autotoky usually expresses
itself as parthenogenesis, where progeny arise from unfer-
tilized eggs. Hermaphroditism is apparently quite rare
amongst the freshwater nematodes[70], apparently occur-
ring in Chronogaster troglodytes[77] and some others.
Asexual reproduction does not occur in the Nematoda.
In amphimictic reproduction, the male nematode
always places sperm inside the female; external fertiliza-
tion has never been demonstrated. Some aquatic species
exhibit traumatic insemination when the male penetrates
the female cuticle with his spicules and releases sperm into
her body cavity.
Female nematodes contain one or two gonads, which
open to the exterior on the ventral side of the body at the
vulva (Fig. 9.4). The vulva is connected to a muscular
tube, the vagina (Fig. 9.7a), which in turn leads into the
uterus followed by the oviduct and ovary. A spermatheca
(sperm-collecting area) is usually present between the
uterus and oviduct. With double-ovary species (didelphic),
the gonads are usually opposite and join at a common
vagina (Fig. 9.4). Single-ovary forms (monodelphic) have
retained the anterior ovary, which is often reflexed into the
posterior part of the body.
The male may have a single testis (monorchic) or two
testes (diorchic), which lead to a common seminal vesi-
cle and vas deferens before entering the cloacal chamber,
a common opening for the reproductive and digestive
systems. Males of freshwater nematodes all possess one
or two sclerotized structures termed spicules (Figs. 9.4,
9.7c). These are inserted into the vagina of the female at
insemination (except in traumatic insemination when they
penetrate the cuticle of the female). Some males also have
another sclerotized structure, the gubernaculum.
The spicules are contained within the walls of the cloa-
cal chamber while the gubernaculum is attached to the
floor of the cloacal chamber. The latter structure supports
the spicules during their movement in and out of the cloa-
cal opening (Figs. 9.4, 9.7c).
Male and female nematodes locate each other with sex
attractants. Once together, the nematodes intertwine and
the posterior region of the male contacts the female vulva.
Sometimes an adhesive compound is produced to seal the
union. Spicules are then inserted and sperm is transferred
from the vas deferens of the male into the female’s vagina
and uterus. The pair may remain joined for only several
minutes or may be united for days. It is common to find
“balls” of mermithids in stream bottoms all coiled together
in continuous copulation. Individual sperm as well as sec-
ondary spermatocytes may be transferred during mating.
In many nematodes, sperm maturation is completed in the
female. Nematode sperm is variable in size and shape but is
always nonflagellated and usually amoeboid. An interesting
modification of sperm morphology occurs in the freshwater
mermithid Gastromermis anisotis, where the mature sperm
are elongate with membraneous organelles surrounding an
apical nuclear region while the mitochondria are arranged in
parallel rows and form an elongate basal sheath (Fig. 9.9a,c).
Pseudopodia are restricted to the main cell body and can
extend some distance from this region (Fig. 9.9b)[74].
Parthenogenetic forms can reproduce by mitotic or
meiotic parthenogenesis. In the former, the diploid somatic
number of chromosomes is retained; whereas in the lat-
ter, two maturation divisions occur which are similar to
oogenesis in amphimictic species. A diploid chromosome
number is established by fusion of the nonextruded polar
nucleus with the egg pronucleus or by doubling of the
chromosome before the first cleavage division occurs.
D. Development and Life History
The eggs of nematodes differ from all other stages in con-
taining chitin in their shells. During embryonic develop-
ment, nematodes show predeterminate cleavage, which
means that cells destined to form specific tissues appear
early in the embryo. Freshwater forms like the mermith-
ids may deposit eggs with fully formed juveniles ready
to hatch upon receiving the right stimuli. Other species
deposit eggs in the single-cell stage, with embryonic
development occurring in the environment (Fig. 9.10).
Postembryonic development (i.e., after hatching) is simi-
lar to the gradual type of metamorphosis in insects. Aside
from an increase in size, proportional changes of various
organs, and development of the gonads and second sexual
characteristics, few differences exist between juveniles and
adults. For this reason, the immature stages of nematodes
should be called juveniles and not larvae, because the lat-
ter usually implies complete metamorphosis where the
immature stages differ radically from the adults. Despite
these differences, however, the term larva is widely used
today in referring to the immature, postembryonic stages
of nematodes.
Hatching is influenced by temperature and external
stimuli. The eggs of the mermithid Pheromermis pachys-
oma hatch when ingested by aquatic insects. Eggs of other
freshwater nematodes hatch when the water reaches a cer-
tain temperature (Fig. 9.11c). Eggs that undergo anabiosis
as a result of desiccation will not hatch until the area is
flooded. This probably enhances survival for nematodes in
Chapter | 9 Nematoda and Nematomorpha 245
temporary water sources, although this point still requires
elucidation. In most cases, it is the first-stage juvenile that
emerges from the egg; however, in mermithids, juveniles
molt once in the egg and then emerge as second-stage
juveniles (Fig. 9.11c). If equipped with a spear or tooth,
the young nematode will use these to break out of the
shell, otherwise it simply employs pressure and pharyngeal
secretions to soften the shell wall.
All nematodes undergo four molts and have six stages
during their development: egg, first-stage juvenile, second-
stage juvenile, third-stage juvenile, fourth-stage juve-
nile, and adult. During each molt, the cuticle is shed
FIGURE 9.9 (A) Two sperm of a mermithid
nematode showing the bead-like mitochon-
dria at one end (small arrows) and the nar-
row elongate nuclei (large arrows). (B) Single
sperm that has produced pseudopodia along
both sides (arrows). (C) Electron micrograph
(TM) of a sperm showing the dark nuclear
areas (arrows) (same sperm as in (A) and (B)
(photograph by Roberta Poinar).
Ecology and Classification of North American Freshwater Invertebrates
246
and replaced by another secreted by the hypodermis.
Sometimes two molts may occur almost simultaneously,
as in postparasitic juvenile Mermithidae and occasionally
the shed skin is retained around the body of the next stage
(Fig. 9.25b). Free-living nematodes generally complete
their development rapidly in comparison to other metazo-
ans. This can be 3–5 days for rhabditids (under optimum
conditions) and generally from 1–6 weeks for most other
freshwater forms. Therefore, nematode populations can
turn over very quickly, as described in an energetic study
by Schiemer[87–88].
Aside from premolt and quiescent stages, most nema-
todes feed continuously throughout the growth period.
Most freshwater nematodes take food in through the mouth
and absorb nutrients into the intestinal cells. During their
parasitic development in the hemolymph of invertebrates,
mermithids absorb nutrients directly through their body
walls. Nematode growth entails both enlargement and mul-
tiplication of cells. Growth is frequently nonproportional
from stage to stage. For example, the intestine often will
grow faster than the pharynx, so the proportion of the two
organs differs in each stage. This information can be used to
determine nematode stages, along with gonad development.
The types of food vary among the freshwater nematode
groups. Microbotrophic forms ingest bacteria, algae, single-
celled fungi, and protozoa. Predaceous species attack small
metazoans such as nematodes, annelids, early insect stages,
and molluscs. Many are omnivorous and will consume
microbial life as well as metazoans. Plant parasites feed
on the cytoplasm of higher plants, algae, and filamentous
fungi. Parasites of invertebrates absorb nutrients from the
hemolymph of various taxa, especially insects in the case
of the Mermithidae and snails in the case of Daubaylia.
Juveniles or adults of vertebrate parasites may occur in the
alimentary tracts or body cavities of insects (Fig. 9.12),
fish, aquatic birds, or amphibians. Only the egg stages are
free-living, and these are not likely to be encountered or
recognized during routine collecting activities.
The dependency of nematodes on moisture has brought
a tremendous selection pressure to withstand periods of
desiccation. Some nematodes form resistant stages; while
in others, all life stages can tolerate drought conditions for
various periods. Resistant stages include the egg and the
second- through fourth-stage juveniles. Some nematodes
FIGURE 9.11 (A) Various eggs and juvenile stages of freshwater
nematodes developing in a decaying, submerged leaf. (B) Adults of
Pheromermis pachysoma (Mermithidae) in a California springbed; these
nematodes have emerged from parasitized queen wasps visiting the spring
and will undergo molting, mating, and oviposition in the spring. (C)
Preparasitic mermithid juvenile using its stylet (arrow) to break through
the egg shell.
FIGURE 9.10 Egg clusters of a mermithid nematode that are attached
to rocks and debris in fast-flowing streams.
Chapter | 9 Nematoda and Nematomorpha 247
have been maintained for years (up to 25 years) in anabio-
sis and then revived in water.
Very little is known about the resistant stages, dispersal,
and survival of freshwater nematodes. Jacobs[41] mentions
the “ability for passive dispersal” of resistant forms, but does
not discuss how this dispersal occurs. It is very likely that
the eggs and possibly some juvenile stages of forms adapted
to semitemporary water sources can enter a resistant, qui-
escent stage capable of surviving desiccation and possi-
bly temperature extremes; these have been demonstrated
in some soil and plant nematodes. Passive dispersal may
occur when these resistant stages are blown by heavy winds
or washed to new areas by flash floods. The possibility
of freshwater nematodes being carried in mud attached to
the body parts of various water-frequenting animals such
as wading and swimming birds is also feasible.
E. Ecology
1.  Habitats of Freshwater Nematodes
Nematodes constitute an important and significant portion
of the zoobenthic community of freshwater habitats. This
community has been separated by benthic ecologists into
three size categories[97]. Animals retained on sieves with
a mesh ranging from 200–2000 m compose the macro-
fauna, those that are retained on sieves with a mesh rang-
ing from 40–200 m comprise the meiofauna, and those
that pass through a 40 m mesh sieve make up the micro-
fauna. Freshwater nematodes covered in the present chap-
ter could be considered as belonging to all groups; the large
free-living stages of the Mermithidae would be considered
macrofauna, the majority of the adults and juveniles of the
other groups would fall under meiofauna, and the young
juveniles and eggs of many forms would fall into the
microfaunal category (Fig. 9.13).
Nematodes constituted 60% of all the benthic metazo-
ans in Mirror Lake, New Hampshire, and are probably the
most abundant benthic animals in freshwater, according
to a study by Strayer[97]. In Mirror Lake, nematode abun-
dance showed a distinct depth distribution, with a strong
minimum at 7.5 m. More than 90% of the nematode bio-
mass at 7.5 m was composed of species of the large preda-
tory nematodes belonging to the genus Mononchus. About
63% of these nematodes lived in the top 2 cm of sediment,
while only 18% penetrated to a depth greater than 4 cm.
Species of the genera Monhystera and Ethmolaimus, which
constituted 55% of the nematodes in Mirror Lake, were
found in the benthic zone. Although the nematodes out-
ranked all other animals in abundance (constituting 59%
of all metazoan individuals in the benthos), they still rep-
resented only 1% of the zoobenthic biomass. Nematodes
normally contribute between 1 and 15% of the zoobenthic
biomass in lakes, and other studies show that they consti-
tute from 40–80% of all meiobenthic animals[36,38,59].
Nematodes that occur in freshwater have traditionally
been called free-living nematodes. The term free-living is
an ecological ranking, which means that these nematodes
have no symbiotic association with multicellular plants or
animals. Such nematodes obtain nourishment from bac-
teria, algae, protozoa, and other unicellular organisms.
Nematodes with this type of nutrition are best referred to
as microbotrophic, although they have been called sapro-
phytic, saprophagous, bacteriophagous, microbivorous,
and microphagous. Such nematodes act as secondary con-
sumers, feeding on bacteria and fungi. They may serve to
maintain these microbial populations although many of
the free-living forms will die if bacterial populations reach
high numbers, which is why most of the normal aquatic
nematodes are absent from areas high in sewage.
FIGURE 9.12 Spirurid nematodes in the abdomen of an adult caddisfly.
FIGURE 9.13 Scene of the bottom of a typical fast-flowing stream
showing several nematode microhabitats. D, streambed containing endo-
benthic forms; E, bed surface containing epibenthic groups; H, rock
surface containing haptobenthic groups; N, Nostac algal pads contain-
ing mermithids that parasitize Cricotopus midges (Chironomidae) living
inside the cyanobacteria.
Ecology and Classification of North American Freshwater Invertebrates
248
The term free-living, however, is also used to refer to
a particular stage in the life cycle of a nematode—often
a stage in the life cycle of a plant-parasitic or animal-
parasitic nematode—when it has either ended or not yet
begun its parasitic relationship. Examples in the freshwater
habitat are the mermithids. The egg, infective second-stage
juvenile, postparasitic juvenile, and adult mermithids are
free-living and nonfeeding. Nourishment is obtained from
the hemocoel of insects only during a portion of their juve-
nile development. Thus, free-living in the present work refers
to those nematodes that have some developmental stages
occurring in freshwater, irrespective of their nutritional
requirements or their status at other developmental stages.
The habitats of freshwater nematodes vary as do the cat-
egories of freshwater resources. Two important requirements
for freshwater nematodes to complete their development are
continuation of the water source and availability of food
and oxygen. Many freshwater nematodes feed on microor-
ganisms that flourish in the gyttja, an organic deposit com-
posed of excretory material from benthic animals. It is most
obvious in lakes and ponds, where it is often combined with
decomposing plant debris. This sediment increases in den-
sity with increasing depth. Core samples are used for quan-
titative measurements of zoobenthos in the gyttja[97].
Annual productivity of freshwater nematodes in an
Austrian alpine lake was studied by Bretschko[5]. In the
deeper benthos below 20 m, there were 2–4 annual genera-
tions with the total yield of nematode biomass estimated at
66 kg per year. It is interesting that the production was higher
in shallow water and during the winter when the lake was
covered with ice. For example, Tobrilus grandipapellatus
was collected at populations of 235,000/m2 in the winter, but
only 60,000/m2 in the summer. This could be related to the
availability of oxygen and density of microbial populations.
Nematodes of all environments, including freshwater, are
among the most numerous animals feeding on both primary
decomposers such as bacteria and fungi, as well as primary
producers such as algae and higher plants[56]. The signifi-
cance of nematodes in the general economy of the ecosystem
is not known, and attempts to estimate their contributions to
energy flow have been made only with terrestrial and marine
forms. Such estimates have used the equation:
C P R E U= + + +
where C is consumption, P is production, R is respiration,
E is ejecta (feces), and U is excretion.
Although freshwater nematodes have been little stud-
ied, estuarine forms have been examined in detail[102].
Some 40 species of nematodes occur on a mud flat in the
Lynher Estuary in southern Britain. The population varied
from 8–9 106 individuals/m2 in winter to nearly 23 106
in late spring (15% of the macrofauna). Warwick and
Price[102] calculated a total annual oxygen (O2) consumption
of 28 L O2/m2/yr, which is equivalent to some 11 g of metab-
olized carbon (C). The annual production was calculated at
about 6.6 g C/m2/yr. Previous calculations with the soil nem-
atode Caenorhabditis briggsae showed that the conversion
of bacteria into nematode tissue varied from 13–20%[56].
The question of nematodes living under anoxic condi-
tions has been addressed by several workers. In deep lakes,
the hypolimnion may become anaerobic for certain periods,
resulting in a reduction of nematode fauna, but not eradica-
tion. In Israel’s Lake Tiberias, large numbers of Eudorylaimus
andrassyi occurred at 43 m during the winter oxygen-free
period[82]. In the Neusiedlersee in Austria, Tobrilus gracilus
occurs in muddy anaerobic zones beyond the limits of emer-
gent vegetation[86]. Most species of nematodes live in the
littoral zone in Mirror Lake, but species of Ethmolaimus,
Monhystera, and unidentified Tylenchidae were very abun-
dant in the anaerobic sediments at a depth of 10.5 m[97].
Extreme habitats for freshwater nematodes include high-
temperature hot springs. Table 9.1 1ists some freshwater
Table 9.1 Survival of freshwater nematodes in
hot-water springs at high temperatures.
Nematode species Temperature
(°C)
Location Reference
Aphelenchoides sp. 61.3 New Zealand Rahm[83]
Aphelenchoides sp. 35.0 New Zealand Winterbourn
and
Brown[105]
Aphelenchus sp. 57.6 Chile Rahm[83]
Doryalimus atratus
Linstow
47.0 Italy Issel[40]
Dorylaimus atratus
Linstow [D. thermae
Cobb]
40.0 Wyoming,
USA
Hoeppli[34]
Dorylaimus atratus
Linstow [D. thermae
Cobb]
53.0 Wyoming,
USA
Cobb, in
Hoeppli[34]
Euchromadora
striata (Eberth)
52.0 Italy Meyl[48]
Monhystera ocellata
Bütschli
52.0 Italy Meyl[48]
Monhystera gerlachii
Meyl
52.0 Italy Meyl[48]
Plectus sp. 57.6 Chile Rahm[83]
Rhabdolaimus
brachyuris Meyl
52.0 Italy Meyl[48]
Theristus pertenuis
Bresslau and
Schuurmans
Stekhoven
52.0 Italy Meyl[48]
Tylocephalus sp. 45.3 New Zealand Winterbourn
and
Brown[105]
Chapter | 9 Nematoda and Nematomorpha 249
nematodes collected at unusually high temperatures. The
listing of 61.3°C for Aphelenchoides sp. is not only a record
for nematodes but for all metazoan life forms!
Other specialized habitats encompass brackish and estu-
arine waters, inland saline lakes, cave streams, and associa-
tions with specific aquatic plants. Manmade aquatic habitats
for nematodes include canals, waste water, tap water, wells,
and filter beds of sewage treatment plants. Nematodes found
in these habitats are often soil nematodes that can tolerate
certain aquatic conditions. Freshwater nematodes are rarely
found in phytotelmata, probably because they do not form
associations with insects that can transport them to and
from such microhabitats. However, freshwater mermithid
genera do occur in tree holes, being transported inside
their hosts that breed in the same environment (Fig. 9.14).
One such species is Octomyomermis troglodytis, which
parasitizes the Western tree hole mosquito in California[76].
The eggs of the mermithid can remain viable in the moist
organic matter at the bottom of the tree hole during the dry
summer and then hatch when the rains arrive and mosquito
larvae are available. Another mermithid species occurs in
tree holes in Barro Colorado, Panama[107].
Other specialized habitats are dune lakes (Fig. 9.15),
which contain a variety of freshwater nematodes. While the
larger lakes can remain for several thousand years before fill-
ing up and disappearing, that could possibly be long enough
for selection of specific subspecies adapted to these habitats.
FIGURE 9.14 Collecting nematodes from a water-filled tree hole in the
fork of a live oak in California.
FIGURE 9.15 Dune lakes contain a variety of freshwater nematodes. It is
unknown how nematodes colonize these unique and temporary habitats.
Another special habitat consists of an ecozone in a
stream in the Sierra Nevada of California (Fig. 9.16). This
ecozone contains a unique assemblage of four closely
associated organisms that form a quadritrophic system
involving a virus that infects a mermithid nematode that
parasitizes a chironomid midge that lives and develops
inside a cyanobacterium (Fig. 9.17). A 25-year study of this
unique association in a stream of several km length (2672–
1860 m above sea level) showed that it only occurred along
a 300 m stretch of stream at about 1860 m a.s.l.[75]. No other
system with these four organisms has been reported from
anywhere else in the world. One wonders how many other
unique, intricately balanced associations occur in freshwa-
ter habitats throughout the world.
A unique cave habitat has resulted in what appears to
be the first freshwater nematode specialized for survival in
hydrogen sulfide–rich thermomineral waters. These condi-
tions are found in the Movile cave, located in a limestone
plateau in Southern Dobrogea, Romania[77]. The nema-
tode, Chronogaster troglodytes (Fig. 9.18), lives in floating
microbial mats composed primarily of mycelia of fungi
belonging to the Class Oomycetes and various bacterial
populations. Selection for this habitat could have under-
gone millions of years since a Miocene age for the origin
of Movile cave has been suggested. Also, while members of
the genus Chronogaster are mostly aquatic, they, like many
freshwater nematodes, are quite broad in their selection of
habitat. Other species of Chronogaster have been collected
Ecology and Classification of North American Freshwater Invertebrates
250
nematodes recovered from caves and subterranean waters.
Apparently no nematode genera are restricted to caves or
subterranean waters, since all genera found in these habi-
tats have species occurring in epigean terrestrial or fresh-
water habitats. Of species of cave nematodes collected from
Mexico, seven were freshwater forms, four were soil micro-
botrophs, three were stylet-bearing plant or invertebrate
predators, and five were non-stylet-bearing predators[112].
Previous workers have attempted to separate the
freshwater-dependent nematodes from the facultative forms.
Micoletzky[49] distinguished between completely aquatic,
principally aquatic, amphibious, and principally terres-
trial species; while more recently, Jacobs[41] presented an
ecological classification with stenohygrophilic nematodes
representing the strictly aquatic forms and enhygrophilic
nematodes representing the semiaquatic forms. The strictly
aquatic or stenohygrophilic taxa covered here occur in
many microhabitats (Figs. 9.11a,b, 9.13). As examples,
periphytic forms live among plant roots, bryophilic taxa
associate with moss and liverworts, endobenthic species
thrive within the sediment and bottom debris, epibenthic
groups live on the surface of the bottom, haptobenthic
forms exist on the surface of submerged rocks, debris, and
aquatic plants, and planktonic taxa live continuously in the
water column. The latter group is poorly known and are
mainly found in turbid waters where their occurrence may
represent more a dispersal than a habitat mode.
Many genera of freshwater stenobygrophilic nema-
todes are widely dispersed, occurring on more than one
continent. Some species are eurytopic, occurring in diverse
habitats, while others are stenotopic and restricted to a few
specialized habitats.
Freshwater nematodes have been considered as indica-
tors of water pollution[113]. Heavy metals and other pollutants
settle and are taken up by organic matter in the sediments.
Nematodes ingest this material, and those that are sensitive to
the pollutants may die. Representatives of the Chromodorida
are apparently more sensitive to pollution than those of
the Rhabditida[111]. In extremely polluted waters, only the
Rhabditoidea may be quite abundant. A 2-year study sam-
pling nematodes along two Indiana streams demonstrated
their usefulness in evaluating aquatic disturbance[24].
2.  Dispersal
No distinct dispersal mechanisms, such as a resistant stage
that can survive desiccation and be carried to new areas by
biotic and abiotic agents, are known for any obligate fresh-
water or marine nematode. The question of how freshwater
nematodes are distributed to various water sources has
never been adequately answered. Although it is likely that
waterfowl and other migrating aquatic animals transport
nematodes either externally on their pelage or internally in
their alimentary tracts, this has never been demonstrated.
While it is easy to understand how freshwater nematodes
FIGURE 9.16 This portion of Sagehen Creek in California’s Sierra
Nevada contains a unique quadri-trophic association between a cyanobac-
terium, chironomid midge, mermithid nematode, and nonoccluded virus.
FIGURE 9.17 The mermithid parasite, Gastromermis anisotis, inside
its chironomid midge host (Cricotopus) in Sagehen Creek, California.
from sandy or sandy loam soils, clay soils, pasture soils, and
even forest soils. In fact, one species, C. africana, is men-
tioned as occurring in multiple aquatic and terrestrial habi-
tats[33]. Such plasticity was probably an important feature
in the specialization of C. trogoldytes in its unique under-
ground cave habitat. See Poinar and Sarbu[77] for a list of
Chapter | 9 Nematoda and Nematomorpha 251
FIGURE 9.18 Chronogaster troglodytes, a spe-
cialized aquatic cave nematode from Movile Cave
in Romania. (A) Anterior end with funnel-shaped
mouth cavity (arrow). (B) Head showing stirrup-
shaped amphid with a circular opening (arrow). (C)
Sperm-like bodies (arrows) near the proximal portion
of the female gonad of the hermaphroditic C. troglo-
dytes. (D) Elongate rectum (arrow) in the female of C.
troglodytes.
are introduced into the sea by rivers and periodic flood-
ing, it is hard to see how marine nematodes could be intro-
duced into freshwater habitats. For instance, along the
Pacific northwest coast are dune lakes, some no larger than
10 m across, which contain freshwater nematode genera.
It is difficult to determine the age of these lakes, but they
eventually fill in with sand and debris, passing through
a pond, swamp, and marsh stage. Most are probably no
older than 3000 years. It is possible that some clades of
enoplids, especially those that occur in both freshwater
and marine habitats, were introduced from the sea. Biotic
agents could have been waterfowl since ducks and grebes
occasionally land in these water sources. However, abiotic
dispersal agents could have also moved marine nematodes
inland. Both salt spray and sea foam are blown considera-
ble distances inland during severe winter storms. Sea foam
is formed during intense wave action with high tides (Fig.
9.19). An examination of sea foam collected by the author
during one such storm revealed the presence of a number
of living juvenile and adult nematodes (Fig. 9.20).
3.  Enemies
Perhaps the greatest enemies of freshwater nematodes are
other predaceous nematodes, but other taxa also consume
nematodes. Representatives of the Mononchida, Dory-
laimida, and Enoplida are known to attack and devour a
range of small invertebrates in their surroundings. The
Ecology and Classification of North American Freshwater Invertebrates
252
juveniles and adults of mermithids. Young trout fingerlings
(Fig. 9.21), as well as various suckers (Catostomidae) prob-
ably feed on other freshwater species as well. Certainly, fish
reduce the population of mermithids by feeding on parasi-
tized hosts. Birds also consume adults of aquatic insects
infected with freshwater mermithids. Mermithids associated
with parasitized caddisfly adults (Trichoptera) were recov-
ered from the stomach of eight arctic terns (Sterna para-
disea) in the arctic tundra. One tern contained 40 mermithids,
some still inside the partly digested caddisflies. The terns
were collected in early August, during a peak emergence of
a large limnephilid caddisfly that bred in the tundra ponds
(Eric Hoberg, personal communication, May 10, 1988).
The role of disease organisms in regulating populations
of freshwater nematodes is not known, but the author has
frequently collected specimens of Tobrilus infested with
what appeared to be microsporidian spores (Fig. 9.26c).
Other records of probable protozoan diseases in freshwater
nematodes have been reported in Chromadora, Dorylaimus,
Ironus, Monhystera, Plectus, Theristus, Trilobus, Tripyla,
Prodesmodora, Achromadora, Paraphanolaimus, and
Desmolaimus[72]. Microsporidian pathogens can best be
observed after spore formation. At this time, clusters of
spores or pansporoblasts can be found in various tissues of
the body (Fig. 9.22c,d)[71].
Viruses are probably fairly common in various tis-
sues of freshwater nematodes. Only one has been reported
from a member of this group today and that was infecting
FIGURE 9.19 Sea foam blown up along the partially destroyed fore-
dune during a storm along the Oregon coast.
FIGURE 9.20 Nematodes recovered from sea foam blown up along the
high tide level during a winter storm on the Oregon coast.
FIGURE 9.21 A fingerling rainbow trout consumes freshwater nema-
todes not only from the stream bottom, but also when eating insects para-
sitized by nematodes.
crayfish Pacifastacus leniusculus occasionally consumes
nematodes[27]. The freshwater rhabdocoel turbellarian
Microstomum feeds on nematodes[96], as will the freshwater
nemertean worm, Prostoma[16]. At Sagehen Creek, rainbow
trout (Fig. 9.21) have been observed feeding on postparasitic
Chapter | 9 Nematoda and Nematomorpha 253
FIGURE 9.22 (A) Nonoccluded virus particles in the
hypodermis of the freshwater mermithid, Gastromermis
anisotis (photograph by Roberta Poinar). (B) Electron
micrograph (TM) of a group of nonocculded virus par-
ticles (arrows) developing in G. anisotis (photograph
by Roberta Poinar). (C) Clusters of pansporoblasts con-
taining spores of Microsporidium sp. in the body cavity
of a nematode. (D) Electron micrograph (TM) showing
spores of Microsporidium sp. developing in the hypo-
dermis of a nematode (photograph by Roberta Poinar).
Gastromermis anisotis in a California stream[73]. The virus,
an enveloped icosahedral type, was found in the reproduc-
tive system as well as in the hypodermis, alimentary tract,
muscles, and nerves of adult mermithids (Fig. 9.22a,b).
Virus particles in the testes may be transmitted to the female
during mating. There was some indication that the virus dis-
rupted the normal physiological pathways of the cells.
F. Collecting and Rearing Techniques
1.  Sampling Methods
The general composition of the nematode assemblage can
be determined qualitatively by selecting methods appropri-
ate for the water flow and the zone to be sampled. In fast-
flowing streams, nematodes can be collected with a net, an
Ecology and Classification of North American Freshwater Invertebrates
254
instrument to turn over rocks and debris on the streambed,
and a set of sieves (Fig. 9.23). The net is first anchored
with the opening facing upstream. A pick or geologist’s
hammer is used to disturb the streambed slightly upstream
from the net (Fig. 9.23a). The stream washes nematodes
and other debris from the disturbed area into the net. The
contents of the net are then placed in enamel collecting
pans with water (Fig. 9.23b). Water with nematodes and
debris are passed through a series of sieves with openings
ranging from 0.4 to 0.04 mm (Fig. 9.23c). Nematodes are
washed off the surface of the final screen into a collect-
ing pan with a spray of water from the plastic squeeze
bottle (Fig. 9.23d). Nets can be pulled over aquatic plants
to obtain nematodes living on their surfaces. The type of
nematode being collected will determine the mesh size of
the net selected. To obtain a fair sample of all nematodes,
a mesh size no larger than 40 m should be used.
A number of devices have been used to collect freshwa-
ter nematodes[26]. These include dredges, wormnets, bottom
catchers, mudsuckers, swab or sledge trawls, and plankton
nets. Their use depends on the type of material desired. For
sluggish or stationary water sources, samples of the bottom
(mud, sand) can be placed in a water-filled container which
is then shaken, the suspension allowed to settle for 3–4 sec,
and the supernatant then poured into a second container
for extraction. Rocks and other submerged material can
be placed in buckets and the nematodes washed off with a
water spray or by vigorous shaking or brushing.
FIGURE 9.23 Quick method of
collecting nematodes from a stre-
ambed. (A) With the geologist’s
hammer, the streambed is disturbed
upstream from the positioned net.
(B) Nematodes and debris collected
from the streambed are transferred
to an enamel pan filled with water.
(C) Water with nematodes and debris
is passed through a series of sieves
with openings ranging from 0.4 to
0.04 mm.(D) Nematodes are washed
off the surface of the final screen into
a collecting pan with a spray of water
from the plastic squeeze bottle.
Chapter | 9 Nematoda and Nematomorpha 255
Quantitative sampling to evaluate taxonomic compo-
sition, relative abundance, and population dynamics over
time involves the use of augers or probes to remove core
samples of measurable sizes from the beds[97]. Other quan-
titative methods can be devised on the basis of need and
the desired biotype to be investigated[19,35,37]. Techniques
applied to marine nematodes may also be suitable for
freshwater forms. However, remember that every type of
sampling technique has an inherent bias affecting data on
species composition and densities. Also, note that nema-
todes are fragile animals, and sampling and extraction
techniques should be as gentle as possible.
2.  Extraction
The process of extraction involves removing nematodes
from samples collected in various freshwater habitats. The
basic principles of nematode extraction are based on the
fact that most nematodes have a specific gravity of 1.10–
1.14, thus they will slowly sink in freshwater. They are
denser than fine clay particles but lighter than sand. The
extraction methods used for soil and plant-parasitic nema-
todes are wholly adequate for freshwater forms.
Although the majority of freshwater nematodes are too
small to view with the naked eye, the free-living stages
of many aquatic mermithids are large enough to be hand-
picked from the samples. They can be carefully lifted with
a fine needle containing an L-shaped bend at the tip or
with a pair of fine forceps. Other mermithids that are small
enough to escape visual spotting can be collected with the
extraction processes.
One of the oldest and simplest methods of extracting
nematodes from a wide range of samples is the Cobb siev-
ing and gravity method, or a modification of the latter.
This consists of making a water suspension of the nema-
todes and pouring it through a series of screens of different
mesh to collect the nematodes. The suspension is made by
placing the samples in a pail or other suitably large con-
tainer, adding 2–3 L of water (preferably from the original
source), allowing the mixture to set for 10–20 sec to allow
the heavier particles to sink, and then pouring the suspen-
sion containing the nematodes through a series of screens.
If large debris (wood particles, rocks) is present, the sam-
ples should first be poured through a #20–40 mesh sieve
(0.840–0.350 mm mesh). The sieve mesh size will, of
course, influence the size range of nematodes extracted.
If a sample of all available nematodes is desired, then a
final fine screen of #400–500 mesh (0.035–0.026 mm)
can be used. The problem with the fine screens is that they
become easily clogged with debris. Such screens should
be tapped rapidly and gently on the undersurface with
the fingers to assist passage of the water. The nematodes,
together with debris, will be trapped on the surface of the
#400 or 500 mesh sieve. Both can be removed by directing
a small jet of water on the back of the sieve and washing
the contents into a separate container (Fig. 9.23d). Certain
biases are inherent in sieving extraction methods[38]. Since
they may be unsuitable for quantitative studies, the more
tedious method of sorting through unsieved samples under
a dissecting microscope may be desirable[97].
The final step consists of separating the nematodes
from the fine debris. For this, the Baermann funnel method,
or a modification thereof, can be used. All that is needed is
a funnel with a piece of rubber tubing attached to the stem.
The tube should be closed with a screw clamp. The nema-
tode-debris mixture is placed on a fine cloth or facial tis-
sue, which is supported by a piece of screen held in the top,
wide portion of the funnel. Water is slowly added to the
funnel until the nematode-debris mixture is just covered.
The clamp should be opened to eliminate the air trapped in
the funnel stem and to allow a continuous column of water
to flow into the funnel. The nematodes crawl through the
debris and facial tissue, drop through the screen, and settle
at the base of the funnel stem, where they can be drawn off
by releasing the clamp. The nematodes should be drawn
off every 6 hr; the apparatus can be operated in this fash-
ion for several days. Then the nematodes can be counted
or handpicked under a dissecting microscope.
A more recently devised method of isolating nema-
todes from samples involves the principle of increasing the
specific gravity of the solution to make the nematodes float
to the surface. Substances such as sugar, salt, or Ludox can
be added to the nematode-water mixture in the centrifugal-
flotation and sugar-flotation techniques[1]. If sugar is used,
then 673 g added to 1 L of water will result in a specific
gravity of 1.18, which is greater than that of nematodes
(causing them to float on the surface). Detailed instruc-
tions for these and other methods of extracting nematodes
from soil and plant samples are presented elsewhere[1,57].
Quantitative methods of extraction for monitoring nem-
atode populations over a period of time often involve the
use of rather elaborate elutriators, which use an upcurrent
of water to separate nematodes from the medium[95].
3.  Fixing and Mounting
After the nematodes have been extracted, they should be
held in water (preferably water from their original col-
lecting source) prior to fixation. The nematodes should
be killed before being placed in fixative; otherwise they
become distorted and difficult to examine. They are most
easily killed with heat, which also tends to relax and
extend them. Too much heat will destroy the internal tis-
sues, but good results can be obtained by pouring hot
water (60–70°C) over the nematodes. They should then
be transferred to the fixative as soon as possible. The best
fixative for nematodes is TAF (7 mL 40% formalin, 2 mL
triethanolamine, and 91 mL distilled water). However, 3–
5% formalin or 70% ethanol can also be used if TAF is not
available. Some distortion may occur with ethanol, however.
Ecology and Classification of North American Freshwater Invertebrates
256
Nematodes that are to be shipped through the post are best
fixed in DESS, a nonflammable, nontoxic solution that is
acceptable for shipment and suitable for both preserving
taxonomic characters and DNA for molecular analysis[108].
The nematodes should be fixed for at least 2–3 days
before transfer to glycerin for mounting on microscope
slides. Nematodes are most easily conveyed to glycerin
by the evaporation method, which requires little handling.
Fixed specimens are transferred to a dish containing a solu-
tion of 70 mL ethanol (95%), 5 mL glycerol, and 25 mL
water. The dish is partly covered for the first 3 days to allow
the alcohol and water to evaporate at a slow rate; then the
cover is removed for the next 14 days. Finally, the contain-
ers (now with mostly glycerin) are placed in a desiccator or
an oven (35°C) for another 2 weeks to drive out the remain-
ing water. The nematodes will then be in a relatively pure
solution of glycerin and can be mounted directly on micro-
scope slides. Such slides are termed permanent since they
can be kept for years for continuous study. Eventually, the
nematode tissues tend to become transparent. Temporary
slides are made by mounting the nematodes directly after
fixation in the fixing solution (formalin or alcohol). They
may last several months if the ringing seal is tight.
To make permanent slides, follow the following steps
for specimens transferred to glycerin (modified from
Poinar[63]). (1) With a small pointed instrument (dental pulp
canal file, small insect pin mounted on a wooden splint,
needle), transfer the nematode(s) to a small drop of glycerin
placed in the center of a microscope slide. (2) Push the nem-
atode to the bottom of the drop. Then add at three equidis-
tant points around the nematode, small supports (coverslip
pieces, wire) with a width equal to or slightly wider than
that of the nematode. Push these also to the bottom of the
drop. (3) Place a cover slip over the drop and lower it slowly
at a 30° angle so that air bubbles will not become entrapped
in the glycerin as the slide touches the mounting medium.
(4) Add more glycerin, if needed, by placing a small drop at
the edge of the coverslip and allowing it to move under the
glass and spread throughout. If the coverslip is floating on
glycerin, then remove the excess by blotting it with a mois-
tened (water) piece of tissue. (5) Carefully seal the edges of
the coverslip with a ringing compound such as nail polish
or Turtox slide ringing cement (General Biological Supply
House, Chicago, Illinois). (6) Label the slide with informa-
tion regarding the date, locality, and collector.
4.  Culturing
Some free-living, microbotrophic freshwater nematodes can
be cultured in the laboratory when fed from a growing col-
ony of their preferred food (i.e., bacterial or algal species)
and maintained under environmental conditions (tempera-
ture, oxygen) comparable to those present at the collecting
site. Nuttycombe[58] successfully cultured several species of
freshwater nematodes using a wheat grain infusion method.
Between 200 and 300 grains of wheat seeds were added to
250 mL of spring water in a flask. This mixture and a sepa-
rate flask of pure spring water were heated to a boil and
allowed to cool. The spring water was poured into 200 mL
petri dishes with 3–4 grains of the boiled wheat. The dishes
with the wheat seeds were allowed to stand for 2–3 days
before the nematodes were added. The microbotrophic
nematodes then consumed microorganisms brought in with
the nematodes that grew on the wheat seeds.
Postparasitic juveniles of freshwater mermithids can be
held in the laboratory until they mature for identification. Care
should be taken so that both the temperature and the amount
of dissolved oxygen parallel those of the collection site.
Periodic transfers to new containers with freshwater may be
necessary to prevent the buildup of fungi, which will destroy
the nematodes. Petersen[62] devised a method of culturing a
mosquito mermithid parasite (Romanomermis culicivorax)
through its entire life cycle. The mosquito chosen as a labora-
tory host was Culex pipiens quinquefasciatus (Say), because
it could be reared continuously in crowded conditions, was
easily maintained in colony, and was highly susceptible to the
nematode. Postparasitic juvenile mermithids were placed in
wet sand within plastic trays (36 25 100 cm). The post-
parasites molted to the adult stage, mated, and oviposited in
the sand. The eggs, which embryonated in the moist sand,
could be maintained in their resting state for several months.
When the trays were flooded with water, the infective stages
emerged from the sand and were capable of infecting newly
hatched mosquito larvae. The infected insect larvae were
kept in a separate container and fed. By the time the mos-
quitoes were ready to pupate, the nematodes had completed
their development and had started to emerge. The emerging
postparasitic juveniles were then transferred to wet sand to
continue the cycle. Adequate numbers of mermithids were
available for both scientific investigations and biologic con-
trol studies with this rearing method.
G. Identification of Selected Nematoda
Once the specimens are mounted, they can be examined
and identified with a compound microscope. Because iden-
tification involves many fine details, a microscope equipped
with oil immersion should be employed (1000). The use
of dioscopic illumination after Nomarski or differential
interference contrast better reveals minute features of the
cuticle (such as the amphid structure). If possible, living or
just killed nematodes should also be examined under the
microscope. Their movements can be reduced by removing
water from under the coverslip and compressing the nema-
tode between the microscope slide and coverslip. Certain
characters are much clearer on living specimens, and there
is a certain thrill in watching these creatures move under a
microscope that is absent with preserved material. However,
measurements can best be made on fixed nematodes.
The following taxonomic key was developed based on
published contributions of other authors[11,13,21–22,25,28,41,60–61,
98,100]; some of which include separate keys[11,13,21–22,28].
Chapter | 9 Nematoda and Nematomorpha 257
For descriptions of various genera and a listing of species,
Goodey[29] is still very helpful, although now out of date.
The classification system used here follows that found in the
book Freshwater Nematodes[23]. A glossary to nematological
terms was prepared by Caveness[8]. Sixty-six genera of fresh-
water nematodes are included in the present key, and 300–
500 species are probably included within these 66 genera.
1.  Taxonomic Key to Families and Selected 
Genera of Freshwater Nematoda
The following key pertains to families and selected gen-
era of nematodes found in freshwater habitats of North
America. Following this is a separate key to the genera of
aquatic Mermithidae.
1a. Elongated and threadlike, generally over 1 cm in length and usually observable with the naked eye; found on the bottom surface or sev-
eral centimeters within the bed ......................................................................................................................................................................2
lb. Not elongate, generally under 1 cm in length and observable only with a lens; found on all exposed surfaces (rocks, on or in plants) as
well as on the bottom surface and within the bed ..........................................................................................................................................3
2a (1a). Forms usually long (6 cm or more in length), leathery body wall, range in color from light brown to black (Fig. 9.30); hairworms ............
.............................................................................................................................................................phylum Nematomorph (see section II).
2b. Forms usually smaller (under 6 cm in length), body wall fragile, range in color from white to rose, green and yellow (Figs. 9.11b, 9.17,
9.24) ................................................................................................................................................. family Mermithidae (see section VI.B.)
3a (1b). Head of nematode bearing a stylet (Fig. 9.26a,b) ..........................................................................................................................................4
3b. Teeth and other armature may be present but a stylet is absent (Figs. 9.18a, 9.25a) ..................................................................................16.
4a (3a). Stylet knobs usually absent (Fig. 9.26b); if present, then pharynx lacking a valvated metacorpus; pharynx narrow anterior and thickened
posterior at junction with intestine; valvated metacorpus absent ........................................................................order Dorylaimida ..........11
4b. Stylet knobs usually present (Figs. 9.25b, 9.26a); pharynx composed of a valvated metacorpus followed by a slender isthmus and basal
glandular bulb leading into the intestine (Fig. 9.25a) ....................................................................................................................................5
5a (4b). Dorsal gland outlet in precorpus; metacorpus moderate in size (Fig. 9.25b) (75% body width) .....................suborder Tylenchina ........8
5b. Dorsal gland outlet in metacorpus; metacorpus large (at least 75% of body width) (Fig. 9.25c) ....................... order Aphelenchida ........ 6
6a (5b). Stylet usually with faint or inconspicuous knobs (Fig. 9.25c); tail tip usually pointed; males lacking bursa and gubernaculum ..................
................................................................................................................................................................. family Aphelenchoididae...............7
6b. Stylet without knobs; tail tip usually rounded; males with a bursa (Fig. 9.7c) and gubernaculum .................................................................
.....................................................................................................................................family Aphelenchidae.................Aphelenchus Bastian
7a (6a). Tail shape elongate, filiform .....................................................................................................................................................Seinura Fuchs
7b. Tail shape short, conical .............................................................................................................................................Aphelenchoides Fischer
8a (5b). Head bearing distinct setae (Fig. 9.8a,d) ............................................................................family Atylenchidae..................Atylenchus Cobb
8b. Head without setae .........................................................................................................................................................................................9
9a (8b). Procorpus fused with large, oval metacorpus; cuticle strongly annulated; adult female with cuticular sheath and well-developed stylet (Fig.
9.25b) ................................................................................................. family Hemicycliophoridae .................................... Hemicycliophora de Man
9b. Procorpus distinct and narrow before reaching the expanded metacorpus, cuticle not strongly annulated; adult female without cuticular
sheath; stylets may or may not be well developed ...................................................................developed superfamily Tylenchoidea .........10
10a (9b). Large distinct stylet, ovaries paired, amphidelphic (Fig. 9.4); tail tapering but not filiform ...........................................................................
family Pratylenchidae ...............................................................................................................................Hirschmanniella Luc and Goodey
10b. Stylet small and inconspicuous, ovary single, prodelphic; tail filiform ...........................................................................................................
family Tylenchidae ..............................................................................................................................................................Tylenchus Bastian
11a (4a). Pharynx gradually widens toward basal bulb (Fig. 9.5a) .............................................................................................................................12
11b. Pharynx abruptly widens to form the basal bulb that contains a valvular chamber .........................................................................................
family Aulolaimoididae ....................................................................................................................................... Aulolaimoides Micoletzky
12a (11a). Head bearing a narrow protrusible mural spear that is symmetrically pointed at tip ......................................................................................
family Nygolaimidae ......................................................................................................................................................... Nygolaimus Cobb
12b. Head bearing a thick protrusible axial spear that is asymmetrically pointed at tip (sloped on one side) (Fig. 9.26b) ....................................
family Dorylaimidae ................................................................................................................................................................................... 13
13a (12b). Stomal area bearing distinct teeth or denticles around the tip of the stylet ............................. family Actinolaimidae ..............................14
13b. Stomal area lacking distinct teeth or denticles in the stylet tip region ......................................family Dorylaimidae ................................15
14a (13a). Four large teeth together with mural denticles present ................................................................................................ Paractinolaimus Meyl
14b. Four large teeth only present ............................................................................................................................................. Actinolaimus Cobb
15a (13b). Cuticle thick and longitudinally ridged (Fig. 9.7b) ....................................................................................................... Dorylaimus Dujardin
15b. Cuticle smooth ......................................................................................................................................................Mesodorylaimus Andrassy
16a (3b). Amphids minute and borne on the lateral lips, excretory duct cuticularized (canal lined with a fine layer of cuticle); caudal glands absent,
conspicuous cephalic setae absent, bursa present or absent .........................................................................................................................17
FIGURE 9.25 (A) Pharyngeal region of Pellioditis
(Rhabditidae) showing tubular stoma (S), corpus (C),
metacorpus (M), isthmus (I), nerve ring (N), and glan-
dular basal bulb (G). Arrow shows valve in basal bulb.
(B) Outer ensheathing cuticle of Hemicycliophora
(Hemicycliophoridae); note the long stylet (S) with
prominent basal knobs (arrow) and valvated meta-
corpus (M). (C) Anterior portion of Aphelenchoides
(Aphelenchoididae) showing small stylet lacking knobs
(arrow) and large metacorpus (M) with valve. (D)
Reflexed ovary (arrow shows point of reflexion).
FIGURE 9.24 Mermithid nematode emerging from the abdomen of an adult
caddisfly.
Chapter | 9 Nematoda and Nematomorpha 259
16b. Amphids usually enlarged and located on the neck region (Fig. 9.8d), excretory duct rarely cuticularized (except in Plectidae); head
often with conspicuous setae; bursa usually absent .....................................................................................................................................21
17a (16a). Pharynx with an expanded median bulb (metacorpus) containing a longitudinal valve and a glandular nonvalvated basal bulb ...................
................................................................................................................................family Rhabditolaimidae...............Rhabditolaimus Fuchs
17b. Pharynx may or may not contain a metacorpus, but if it does, it does not contain a valve; basal pharyngeal bulb with or without a basal
valve .............................................................................................................................................................................................................18
18a (17b). Pharynx elongate and lacking a valve in the basal bulb or bulb area, slender; slow-moving forms (Fig. 9.26d) ............................................
..................................................................................................................family Daubayliidae ..............Daubaylia Chitwood and Chitwood
18b. Pharynx normal in length, with a basal bulb containing a valve (Fig. 9.25a); stouter, quick-moving forms ..............................................19
19a(18b). Stoma with rhabdions separate, female with a single ovary, bursa absent ............................................................................................ family
Cephalobidae .................................................................................................................................................................................................20
19b. Stoma with rhabdions fused, cylindrical (Fig. 9.25a), female with a single or paired ovaries, bursa usually present (Fig. 9.7c) ..................
............................................................................................................................................................................................family Rhabditidae
[Most of the genera in this large family are soil forms, some of which appear from time to time in aquatic habitats. Mesorhabditis
(Osche), which has a single ovary and a bursa, occurs in sewage beds and run-off water. Pellioditis (Doughtery), which has paired ova-
ries also occurs in aquatic habitats.]
FIGURE 9.26 (A) Anterior portion of
Criconema showing the stomato stylet (arrow
points to enlarged basal knobs). (B) Head of
Dorylaimus (Dorylaimidae) showing the small,
pore-like amphids at the base of the lip region
(arrows) and odontostylet lacking basal knobs.
(C) Tail region of Tobrilus (Tobrilidae) show-
ing spores of a microsporidian parasite filling
the hypodermal tissue (arrows). (D) Adults of
Daubaylia (Daubayliidae) inside an aquatic snail.
Ecology and Classification of North American Freshwater Invertebrates
260
20a (19a). Lateral fields reach to tail tip; female tail rounded ..........................................................................................................Cephalobus Bastian
20b. Lateral fields end at the phasmids; female tail usually pointed ....................................................................................Eucephalobus Steiner
21a (16b). Amphids spiral, loop-like, stirrup-shaped (Fig. 9.8c) or circular (Fig. 9.8d); pharynx usually with a basal bulb ......................................22
21b. Amphids pore-like to pocket-like, pharynx usually without a distinct basal bulb .......................................................................................41
22a (21a). Ovary usually outstretched (reflexed in Prismatolaimus); amphids usually circular (Fig. 9.8d) but may be slit-like ....................................
.........................................................................................................................................................................order Monhysterida..............23
22b. Ovaries reflexed (Fig. 9.4) (except in Odontolaimus, which also has circular amphids), amphids variable (including circular) ...............25
23a (22a). Amphids slit-like, faint; stoma with denticulated cushions (padded areas covered with minute projections) ................................................
............................................................................................................................family Prismatolaimidae.................Prismatolaimus de Man
23b. Amphids circular, distinct, stoma lacking denticulated cushion ......................................................................................................................
........................................................................................................................................................................... family Monhysteridae........24
24a (23b). Stoma shallow; pharynx without distinct basal bulb ........................................................................................................................................
...........................................................................................................................................................................................Monhystera Bastian
24b. Stoma elongate; pharynx with distinct basal bulb (as in Fig. 9.25a) ................................................................................Monhystrella Cobb
25a (22b). Caudal glands and spinneret present (Figs. 9.5b, 9.7d); stoma usually armed with teeth (Fig. 9.8b); knobs, bristles (Fig. 9.8a), or puncta-
tions usually present on cuticle ........................................................................................................................................................................
...........................................................................................................................................................................order Chromadorida...........26
25b. Caudal glands and spinneret present or absent; stoma may or may not be armed with teeth; bristles sometimes present on cuticle, espe-
cially around head ........................................................................................................................................ order Araeolaimida.................33
26a (25a). Amphids circular (Fig. 9.8d); cuticular punctations minute ..................................family Microlaimidae ................... Microlaimus de Man
26b. Amphids spiral; kidney shaped or circular; cuticular punctations coarse ....................................................................................................27
27a (26b). Amphids circular or spiral; pharyngeal-intestinal junction large, triradiate ................................................................................................29
27b. Amphids spiral or kidney-shaped, pharyngeal-intestinal junction small, not triradiate ..................................................................................
family Chromadoridae ................................................................................................................................................................................ 28
28a (27b). Amphids represented as a broad transverse slit; five or more preanal genital papillae in male ................................. Chromadorita Filipjev
28b. Amphids represented as a flattened, broken ring; no more than three preanal genital papillae in male ........................ Punctodora Filipjev
29a (27a). Amphids circular ..........................................................................................................................................................................................30
29b. Amphids spiral .............................................................................................................................................................................................31
30a (29a). Stoma cuplike; length shorter than neck width ..................................... family Desmodoridae .......................... Prodesmodora Micoletzky
30b. Stoma tubular; length two or more times neck width ..................................................................................................Odontolaimus de Man
31a (29b). Stoma tubular; there may be three anterior equal teeth in stoma, but single large dorsal tooth is lacking ......................................................
........................................................................................................................................ family Ethmolaimidae............Ethmolaimus de Man
31b. Stoma cup- or funnel-shaped, armed with a prominent dorsal tooth (as in Fig. 9.8b) much larger than any other teeth ............................32
32a (31b). Amphids located at level of stoma, small subventral teeth absent ..............................................................................family Cyatholaimidae
......................................................................................................................................................................... Paracyatholaimus Micoletzky
32b. Amphids located posterior to stoma, one or two small subventral teeth present ..................family Achromadoridae.....Achromadora Cobb
33a (25b). Caudal glands absent; stoma funnel-shaped with separate rhabdions ..........................................................family Teratocephalidae........34
33b. Caudal glands present (Fig. 9.4) or absent; stoma tubular with rhabdions fused ........................................................................................35
34a (33a). Amphids pore-like; cuticular annulation strong ........................................................................................................ Teratocephalus de Man
34b. Amphids spiral; cuticular annulation weak ..........................................................................................................Euteratocephalus Andrassy
35a (33b). Ovaries outstretched; amphids circula ............................ order Araeolaimida. ............... family Diplopeltidae.........Cylindrolaimus de Man
35b. Ovaries reflexed (Figs. 9.4, 9.25d) ...............................................................................................................................................................36
36a (35b). Pharynx usually with a basal valvated bulb (Fig. 9.4) .................................................................................................................................37
36b. Pharynx with or without a basal bulb, if bulb present, then valve absent ....................................................................................................39
37a (36a). Basal bulb of pharynx lacking valves ..........................................................order Plectida......family Aphanolaimidae........Anonchus Cobb
37b. Basal bulb of pharynx with valves (Fig. 9.8a) .............................................................................................................................................38
38a (37b). Pharyngeal basal bulb with postbulbar extension connecting it with the intestine; caudal glands absent .......................................................
...................................................................................................................................... family Chronogasteridae............ Chronogaster Cobb
38b. Phayrngeal basal bulb without postbulbar extension, abutting the intestine; caudal glands present (Fig. 9.4) ...............................................
.............................................................................................................................................................family Plectidae...........Plectus Bastian
39a (36b). Basal bulb lacking ......................................................................................................................family Bastianiidae ........ Bastiania de Man
39b. Basal bulb present ..........................................................................................................................................family Aphanolaimidae........40
Chapter | 9 Nematoda and Nematomorpha 261
40a (39b). Amphids circular; basal pharyngeal bulb absent ........................................................................................................ Aphanolaimus de Man
40b. Amphids spiral, a slight basal pharyngeal bulb present ...................................................................................Paraphanolaimus Micoletzky
41a (21b). Stoma usually oval in outline, heavily sclerotized and armed with one or more teeth, setae and bristles absent .......................................41
41b. Stoma not in the form of a heavily sclerotized oval cavity, teeth or denticles may be present, setae and bristles may be present .............................
(Fig. 9.7a) ........................................................................................................................................................................................................... 42
42a (41b). Cuticle of head double .................................................................................................................................................................................43
42b. Cuticle of head normal .................................................................................................................................................................................44
43a (42a). Setae or bristles present on tail ........................................................................................................................................................................
..................................................................................................................................... family Oncholaimidae.............Oncholaimus Dujardin
43b. Setae and bristles usually lacking on tail .............................................................................................................. family Mononchulidae......
.......................................................................................................................................................................................... Mononchulus Cobb
44a (42b). Stoma heavily sclerotized, cylindrical .........................................................................................................................................................49
44b. Stoma not heavily sclerotized, funnel-shaped or tubular .............................................................................................................................45
45a (44b). Stoma vestigial and unarmed; pharynx base expanded and set off to form a slight, elongate bulb .......................... family Alaimidae......46
45b. Stoma distinct, or if vestigial then armed with an inconspicuous median tooth; pharynx may be expanded at base, but rarely set off from
the remainder in the form of a bulb ..............................................................................................................................................................47
46a (45a). Amphids pore-like, minute ................................................................................................................................................... Alaimus de Man
46b. Amphids cup-shaped, distinct .......................................................................................................................................... Amphidelus Thorne
47a (45b). Base of pharynx expanded to form a valvated bulb; three rodlike thickenings compose posterior part of stoma ..........................................
................................................................................................................................... family Rhabdolaimidae.............Rhabdolaimus de Man
47b. Pharynx cylindrical, not with basal valvated bulb; stoma a simple tube without rodlike thickenings ........................................................48
48a (47b). Three lips; amphids pore-like, minute; stoma cylindrical ...............................................................................................family Tripyloididae
......................................................................................................................................................................................... Tripyloides de Man
48b. Six lips; amphids cup-shaped, distinct; stoma funnel-shaped ....................................................family Tobrilidae .......... Tobrilus Andrassy
49a (44a). Stoma tubular, with two minute teeth at the base; excretory pore opening posterior to head .........................................................................
...............................................................................................................................................family Cryptonchidae...........Cryptonchus Cobb
49b. Stoma tubular, with three anterior hooklike teeth; excretory pore opening in head area family Ironidae ................................ Ironus Bastian
50a (41a). Pharyngeal-intestinal junction tuberculate; dorsal tooth on stoma pointing posteriorally ..............................................................................
............................................................................................................................................... family Anatonchidae.............Anatonchus Cobb
50b. Pharyngeal-intestinal junction not tuberculate; dorsal tooth on stoma pointing anteriorally .............................. family Mononchidae......51
51a (50b). Ventral stomatal ridge with longitudinal row of denticles ................................................................................................. Prionchulus Cobb
51b. Ventral stomatal ridge absent, or if present then unarmed ............................................................................................... Mononchus Bastian
2.  Taxonomic Key to Extant Genera of 
Freshwater Mermithidae
The mermithids represent a family of nematodes that are par-
asitic on other invertebrates. Over half of the described gen-
era parasitize only aquatic insects. Two genera, Aranimermis
and Pheromermis (Fig. 9.11b), enter the immature stages of
aquatic insects that are later consumed by terrestrial arthro-
pods. However, the free-living stages of these and all other
aquatic mermithids (postparasitic juvenile, adults, eggs, and
infective juveniles) are found in the aquatic habitat. Only the
second- and third-stage juveniles occur in insects. Eggs and
infective-stage juveniles are microscopic and normally not
taken in samples, but the postparasitic juveniles and adults
are frequently collected. Since the keys are based on adult
characters, postparasitic juveniles should be held in water
until they molt to the adult stage. One important character in
identifying mermithids is the number of hypodermal cords
present. These can be determined by cutting thin cross sec-
tions with a razor blade by hand and examining them under
the microscope. The hypodermal cords protrude through the
muscle fields in dorsal, ventral, lateral, and often the sub-
dorsal and subventral regions. Only genera known to occur
in North America are included.
1a. Adult cuticle with cross-fibers; thick, robust white nematodes usually found along the edges of bogs, springs, and streams .......................
.......................................................................................................................................................... Pheromermis Poinar, Lane, and Thomas
[Parasites of wasps, ants (Hymenoptera), and horseflies (Tabanidae).]
1b. Adult cuticle without cross-fibers, robust or slender, white, green, pink, brown, or yellow nematodes found in lake, stream, and pond
beds ................................................................................................................................................................................................................2
2a. With four cephalic papillae .........................................................................................................................................Pseudomermis de Man
[synonym: Tetramermis Steiner]
Ecology and Classification of North American Freshwater Invertebrates
262
2b. With six cephalic papillae ..............................................................................................................................................................................3
3a (2b). With single or fused (rare) spicules ...............................................................................................................................................................4
3b. With paired, separate spicules ........................................................................................................................................................................8
4a (3a). Spicule medium to long, more than twice anal body width ...........................................................................................................................5
4b. Spicule short, less than twice anal body width ..............................................................................................................................................6
5a (4a). Mouth terminal; spicules J-shaped; vulval flap present ....................................................................................Lanceimermis Artyukhovsky
[Parasites of true midges (Chironomidae).]
5b. Mouth normally shifted ventrally; spicule curved but not J-shaped; vulva flap absent .........................................Gastromermis Micoletzky
[Parasites of blackflies, true midges (Fig. 9.17), and mayflies.]
6a (4b). Spicule shorter than anal body width; amphids small .................................................................................................Perutilimermis Nickle
[Parasites of mosquitoes (Culicidae).]
6b. Spicule longer than anal body width; amphids medium to large ...................................................................................................................7
7a (6b). Tail pointed; eight hypodermal cords ................................................................................................................................Hydromermis Corti
[Parasites of true midges and mosquitoes.]
7b. Tail rounded; six hypodermal cords ............................................................................................................................... Limnomermis Daday
[Parasites of true midges and blackflies (Simuliidae).]
8a (3b). Vagina straight or nearly so, barrel- or pear-shaped ......................................................................................................................................9
8b. Vagina S-shaped, U-shaped, or elongate with both ends curved .................................................................................................................13
9a (8a). Spicules shorter than cloacal body width .....................................................................................................................................................10
9b. Spicules equal to or longer than cloacal body width ...................................................................................................................................11
10a (9a). Head expanded into a bulb-like shape with very thick cuticle; male lacking lateral-dorsal genital papillae ..................................................
......................................................................................................................................................................................Capitomermis Rubtsov
[Parasites of true midges.]
10b. Head not expanded into a bulb-like shape with thick cuticle; male with genital papillae on lateral-dorsal surface .......................................
....................................................................................................................................................................................Heleidomermis Rubtsov
[Parasites of biting midges (Ceratopogonidae).]
lla (9b). Six hypodermal cords ......................................................................................................................................................Mesomermis Daday
[Parasites of blackflies.]
11b. Eight hypodermal cords ...............................................................................................................................................................................12
12a (1lb). Spicules 2–4 times anal body width ........................................................................................................................... Romanomermis Coman
[Parasites of mosquitoes.]
12b. Spicules 1–2 times anal body width ........................................................................................................................ Octomyomermis Johnson
[Parasites of mosquitoes and true midges.]
13a (8b). Eight hypodermal cords ...............................................................................................................................................................................16
13b. Six hypodermal cords ..................................................................................................................................................................................14
14a (13b). Spicule length less than two times anal diameter ....................................................................................................Strelkovimermis Rubtsov
[Parasites of true midges and mosquitoes.]
14b. Spicule length three or more times cloacal diameter ...................................................................................................................................15
15a (14b). Spicules 10 or more times body width at cloaca; vagina elongate with bends at both ends; postparasitic juveniles with distinct tail
appendage .................................................................................................................................................. Drilomermis Poinar and Petersen
[Parasites of diving beetles (Dytiscidae).]
15b. Spicules 3–10 times body width at cloaca; vagina elongate with 3–6 irregular bends; postparasitic juveniles with indistinct or no tail
appendage ....................................................................................................................................................Aranimermis Poinar and Benton
[Parasites of spiders (Arachnida).]
16a (13a). Vagina elongate, slightly curved; spicules shorter than anal body diameter ............................................. Culicimermis Rubtsov and Isaeva
[Parasites of mosquitoes.]
16b. Vagina S-shaped, ends distinctly curved; spicules range from shorter than anal body diameter to about 10 times tail diameter ...............17
17a (16b). Spicule length equal to or shorter than tail diameter; amphids small; cephalic crown well developed ......................Empidomermis Poinar
[Parasites of mosquitoes.]
17b. Spicule length between 1 and 10 times anal diameter; amphids medium-large, cephalic crown absent or only slightly developed ..............
............................................................................................................................................................................................. Isomermis Coman
[Parasites of blackflies.]
Chapter | 9 Nematoda and Nematomorpha 263
II. NEMATOMORPHA
A. Introduction
The phylum Nematomorpha comprises the freshwater
(Gordiaceae) and marine (Nectonematoidea) hairworms.
All hairworms have parasitic lifestyles. Although freshwater
hairworms usually develop in large terrestrial invertebrates
(mostly insects), they all enter freshwater to mate, ovi-
posit, and produce infective larvae, which in turn encyst in
various aquatic invertebrates and amphibians. Although the
group is worldwide (except Antarctica), fewer than 500 spe-
cies have been described and less than 20 species are know
from North America[66]. Recorded developmental hosts for
the North American species are restricted to members of the
insect orders Orthoptera and Coleoptera (Table 9.2). Beetles
in the family Carabidae appear to be one of the most com-
mon hosts. In fact, a world survey showed that at least 70
species of carabids served as developmental hosts to repre-
sentatives of five hairworm genera[81].
The life cycle of hairworms requires both a paratenic
and a developmental host, as well as a water source. Also,
in contrast to the mermithids that can develop in small
Diptera like midges and blackflies, hairworms require
larger hosts and can only utilize small Diptera as paratenic
hosts. Thus, their existence is much more tenuous than that
of the nematodes, and many species have probably become
extinct, especially locally, over the past 100 years.
FIGURE 9.27 The oldest known hairworm fossil, Cretachordodes bur-
mitis Poinar and Buckley (2004), in Early Cretaceous amber from Burma
(100 mya).
Table 9.2 List of insect families with representatives
reported to serve as developmental hosts of North
American hairworms.
Phylum Arthropoda
Class Insecta
Order Orthoptera
Family Acrididae
Family Blattidae
Family Gryllacrididae
Family Gryllidae
Family Stenopelmatidae
Family Tettigonidae
Order Coleoptera
Family Carabidae
1A problem solvable only by drastic action as demonstrated by Alexander
the Great cutting the knot that could not be untied, which bound a chariot
to a pole at Gordium, the capital of Phrygia, in 333 BC.
Some myths surround this group, perhaps associated
with the proverbial gordian knot1. One myth, still believed
by some today, is suggested by the common name “hair-
worms” or “horsehair worms” indicating that they were
supposed to have arisen from horsehairs that fell into
water. This belief was scientifically disproved in the late
1800s by Leidy[42] when he observed horsehairs placed in
water over a period of many months without “…having
had the opportunity of seeing their vivification.
Adult hairworms have been associated with the diges-
tive and urogenital tract of humans[104], and larval hair-
worms will burrow into a wide range of invertebrate and
vertebrate tissues, including human facial tissue, some-
times resulting in orbital tumors[104].
B. Phylogeny and Fossil Record
The relationship of hairworms to other invertebrate groups
is still under scrutiny. Since there are no nonparasitic
members in the group, it is difficult to determine the mor-
phological characters of the primitive hairworm or com-
pare them with free-living groups. Host selection provides
no clues since many hairworms can develop in hosts of
different insect orders.
Fossil hairworms are rare, with the earliest reported,
Cretachordodes burmitis, described from 100-million-year-
old Lower Cretaceous Burmese amber[67] (Fig. 9.27). The
Eocene fossil Gordius tenuifibrosus[101] (identified from
a 15 mm fragment of subcuticular tissue in the Eocene
brown coals of the Geisel Valley near Halle, Germany)
is now questionable in light of similar characters found
on members of the Mermithidae[65] . The only unequivo-
cal Tertiary record of fossil Nematomorpha are two hair-
worms, Paleochordodes protus Poinar[65], emerging from
a cockroach in Dominican amber dated at 15–45 million
years (Fig. 9.28). While it has been proposed that fossil
palaeoscolecid worms are larval hairworms[106], similarities
Ecology and Classification of North American Freshwater Invertebrates
264
Table 9.3 Systematic arrangement of the nematode genera (include all families and/or genera cited in this chapter).
Phylum Nematoda Family Ethmolaimidae (Ethmolaimus)
Class Enoplea Family Achromadorindae (Achromadora)
Order Enoplida Family Cyatholaimidae (Paracyatholaimus)
Family Oncholaimidae (Oncholaimus) Order Desmodorida
Family Ironidae (Ironus) Family Desmodoridae (Prodesmodora)
Family Tripyloididae (Tripyloides) Family Microlaimidae (Microlaimus)
Family Alaimidae (Alaimus, Amphidelus) Order Monhysterida
Order Triplonchida Family Monhysteridae (Monhystera, Monhystrella)
Family Prismatolaimidae (Prismatolaimus) Order Araeolaimida
Family Tobrilidae (Tobrilus) Family Diplopeltidae (Cylindrolaimus)
Family Tripylina (Tripyla) Order Plectida
Order Dorylaimida Family Aphanolaimidae (Anonchus, Aphanolaimus,
Paraphanolaimus)
Family Aulolaimoididae (Aulolaimoides) Family Rhabdilaimidae (Rhabdolaimus)
Family Nygolaimidae (Nygolaimus) Family Odontolaimidae (Odontolaimus)
Family Dorylaimidae (Dorylaimus, Mesodorylaimus) Family Chronogasteridae (Chronogaster)
Family Actinolaimidae (Actinolaimus, Paractinolaimus) Family Plectidae (Plectus)
Family Qudsianematidae (Eudorylaimus) Family Metateratocephalidae (Euteratocephalus)
Order Mononchida Order Rhabditida
Family Cryptonchidae (Cryptonchus) Family Teratocephalidae (Teratocephalus)
Family Anatonchidae (Anatonchus) Family Spiruridae
Family Mononchidae (Mononchus, Prionchulus) Family Cephalobidae (Cephalobus, Eucephalobus)
Order Mermithida Family Daubayliidae (Daubaylia)
Family Mermithidae (Aranimermis, Capitomermis,
Cretacimermis, Culicimermis, Drilomermis, Empidomermis,
Gastromermis, Heleidomermis, Heydenius, Hydromermis,
Isomermis, Lanceimermis, Limnomermis, Mesomermis,
Octomyomermis, Perutilimermis, Pheromermis,
Pseudomermis, Romanomermis, Strelkovimermis)
Family Aphelenchidae (Aphelenchus)
Family Aphelenchoididae (Aphelenchoides, Seinura)
Family Hemicycliophoridae (Hemicycliophora)
Family Atylenchidae (Atylenchus)
Family Tylenchidae (Tylenchus)
Family Pratylenchidae (Hirschmaniella)
Family Diplogasteridae (Rhabditolaimus)
Class Chromodorea Family Mesorhabditidae (Mesorhabditis)
Order Chromadorida Family Rhabditidae (Caenorhabditis, Pellioditis)
Family Chromadoridae (Chromadorita, Punctodora)
Chapter | 9 Nematoda and Nematomorpha 265
between these two taxa are only superifical and pro-
vide no evidence for phylogenetic relationship[65]. The
Nematomorpha probably originated in the Early Paleozoic
as an offshoot from now extinct lines. Chitwood[12] con-
sidered that Nematomorpha had the closest ties (albeit dis-
tant) with nematodes and rotifers (the Acanthocephala and
Echinodermata were tied for the next closest kin). Using
cladistic methods, Schmidt-Rhaesa[91] also concluded that
nematomorphs are closest to nematodes, as confirmed by
a molecular study[2]. However, no nematode stages exist
that resemble hairworm larvae. While both nematodes and
hairworm larvae possess a triradiate pharynx (Fig. 9.29),
rotifers and gastrotrichs also have that character. The sig-
nificance of molting has yet to be determined since no
molts have been reported during the growth phase of any
nematomorph, while they are consistent throughout the
nematodes. The body plan of the Nematomorpha is unique
in the animal kingdom and even the sperm (Fig. 9.33) is
considered aberrant and is quite different from those of
the nematodes (Fig. 9.9); this supports arguments that this
group separated quite early from the other members of the
Aschelminthes and evolved independently thereafter[44].
FIGURE 9.28 Two specimens of Paleochordodes protus Poinar (1999)
that were in the process of emerging from their cockroach host in 15–45-
million-year-old Dominican amber.
FIGURE 9.29 An electron micrograph (TM) showing the triradi-
ate pharynx of the larva of Neochordodes occidentalis. Photograph by
Roberta Poinar.
FIGURE 9.30 Adult hairworms may attach themselves to various
objects, especially in flowing streams.
C. Morphology and Physiology
It is the free-living adults of hairworms that are normally
described because these stages are most frequently encoun-
tered in sampling. They are dark (rarely white), slender
worms ranging in length from several centimeters to 1 m
and in width from 0.25 to 3 mm (Figs. 9.30, 9.31, 9.45).
The color of most adults varies from yellowish to black.
Although the anterior end is generally attenuated, both tips
tend to be obtusely rounded or blunt (divided in some males
and females). Because the cuticle is normally opaque, it is
impossible to examine the internal organs through the body
wall. Ultrastructural studies of a North American Gordius
revealed some interesting morphological features[20]. The
cuticular structures that are such important taxonomic
Ecology and Classification of North American Freshwater Invertebrates
266
characters are actually sculpturing on the surface of the
thin, superficial epicuticle. This epicuticle is normally cris-
crossed by grooves or furrows, leaving small elevations of
irregular areas (areoles) between them. The surfaces of the
areoles may be smooth (Fig. 9.47b) or may bear setae (bris-
tles) or cuticular projections (tubercles) (Fig. 9.46a,c,d),
arranged singly or in clusters. These bristles and tubercles
may also occur in the interareolar furrows. Additional
observations on the cuticular structure of hairworms include
investigations of Paragordius varius by Zapotosky[109],
Parachordodes spp. by Smith[93], G. aquaticus difficilis
by Smith[94] , and Chordodes morgani by Chandler and
Wells[92]. Many of these studies[92] involve the use of the
scanning electron microscope (SEM), which is a useful tool
for examining details of the surface sculpturing on the epi-
cuticle (Fig. 9.32). While SEM reveals many of the details
FIGURE 9.31 The hairworm, Parachordodes tegonotus Poinar et al.
(2004), emerging from the anus of its carabid beetle developmental host.
FIGURE 9.32 A scanning electron micrograph (SEM) of the poste-
rior end of a male hairworm showing the subterminal cloacal opening.
Photograph courtesy of Clay Chandler.
of the body surface and is excellent for morphological char-
acterization, it is not available to many researchers and,
therefore, is not a practical tool for routine identification.
The cuticle, beneath the epicuticle, is composed of
many crossing layers of cylindrical, nonperiodic collagen-
ous fibers (Fig. 9.42b). Beneath the cuticle is the epidermal
layer, constructed of a single layer of interdigitated cells.
The musculature consists only of overlapping, longitudinal,
flat cells; circular muscles are absent. Ultrastructural details
of the musculature of G. aquaticus and Nectomena muni-
dae are provided by Schmidt-Rhaesa[90]. The pseudocoel is
nearly filled with mesenchymal cells containing large clear
vacuoles, which give the tissue a foamy appearance. These
cells are embedded in a supporting collagenous matrix.
The mouth is located at the calotte or anterior tip of the
body. This region is often lighter in color than the rest of
the body and contains the nonfunctional pharynx, which, in
turn, leads into a degenerate intestine lined with epithelium
one cell in thickness. The gut epithelial cells of Gordius
contain numerous microvilli projecting into the lumen[20].
In both sexes, the genital ducts empty into the intestine,
forming a cloaca lined with cuticle (Fig. 9.49a). Because
adults do not feed, the cloaca is probably used solely for
reproductive purposes. All nematomorphs are amphimictic.
Males have paired cylindrical testes, each of which con-
nects with the cloaca via a separate sperm duct. Females
possess paired ovaries, and the eggs, after passing through
separate oviducts, enter the cloaca independently.
Adults become sexually mature soon after emerg-
ing from their hosts and copulation occurs after the male
coils its posterior end around the terminus of the female.
The spermatozoa are either deposited as a spermatophore
on the female terminus, from where they migrate into the
cloaca, or are deposited directly into the female cloaca.
Ultrastructural studies of the sperm show that they are com-
posed of a main cell body attached to a nuclear portion
containing an elongated nucleus with a helical configura-
tion[44,89]. There are unique features of hairworm sperm,
one of which is the presence of a multivesicular complex
composed of numerous flattened pockets, each filled with
a mucopolysaccharide-appearing substance that surrounds
the nuclear portion of the spermatozoan (Fig. 9.33). The
eggs are deposited singly or in clusters, which often form
elongate strings held together by secretions produced by the
antrum (anterior portion of the female cloaca) (Fig. 9.34).
In certain species occurring in fast-flowing waters, oviposi-
tional modifications occur. Eggs of Euchordodes nigromac-
ulatus were found attached to rocks and other submerged
debris in a New Zealand stream (G. Poinar, personal obser-
vation). This unique manner of oviposition explains how
certain hairworms can maintain populations in fast-flowing
streams. Other investigators have noted eggs attached
to twigs and other stationary objects in the water[4]
(D.J. Watermolen, personal observation).
Chapter | 9 Nematoda and Nematomorpha 267
The preparasitic stage that hatches from the egg is mor-
phologically quite different from the adult worm and can,
therefore, properly be called a larva. [Recall that the term
larva implies that some type of metamorphosis occurs before
the adult stage.] The larva consists of a presoma or presep-
tum (featuring an evaginable proboscis armed with cuticu-
lar spines) and a body or postseptum containing adult tissue
primordia (Figs. 9.35 and 9.36). Ultrastructural studies of
the larvae of Paragordius varius demonstrated quite clearly
the nature of the proboscis and spines[109–111]. It is unfortu-
nate that so few larvae of described nematomorph species
are known, since they possess their own specific characters
which could be of taxonomic value[4,69]. Other aspects on the
biology of the Nematomorpha are summarized elsewhere[31].
D. Development and Life History
The four stages in the life history of nematomorphs are the
egg, the preparasitic larva that hatches from the egg, the par-
asitic larva that develops within an invertebrate, and the free-
living, aquatic adult. Parasitic larvae of all Nematomorpha
mature only within invertebrates, which can be called the
developmental or definitive hosts. A second type of host
involved in the life cycle of probably the great majority of
hairworms is a transport or paratenic host. The preparasitic
larva enters the hemocoel and internal tissues of this host
but does not develop further until the paratenic host is eaten
by a scavenger or predator. The paratenic host is usually
an invertebrate but can also be a vertebrate (e.g., tadpoles,
fish, etc.). Cysts in a host may be composed of overlapping
layers (Fig. 9.42b), as noted when larvae of Neochordodes
occidentalis encysted in tadpoles (Hyla regilla)[69].
FIGURE 9.33 Electron micrographs (TM) of sperm cells of Neochordodes occidentalis (Montgomery) (photographs by Roberta Poinar).
(A) Longitudinal section. M, main cell body; N, nucleus. (B) Cross section. N, nucleus; G, glycogen deposit surrounding nucleus; O, membraneous
organelles forming the multivesicular complex.
FIGURE 9.34 Strings of white egg masses deposited by female hair-
worms in a roadside ditch filled with water.
Ecology and Classification of North American Freshwater Invertebrates
268
3 sp
2 sp
1 sp
pr
p
r.m
g.d
p.g
st
B
g
i.g
i
a
tm
26µ
FIGURE 9.35 Preparasitic larva of Neochordodes occidentalis
(Montgomery). Lateral and ventral views. P, presoma (preseptum); B,
body (postseptum); a, anus; g, globule; g.d, gland duct; i, intestine; i.g,
intestinal gland; m, mesenchyme cells; p.g, preintestinal gland; pr, pro-
trusible proboscis; r.m, retractor muscles; st, stylets; sp, circulet of spines;
t, tail spines (modified from Poinar and Doelman[69]).
FIGURE 9.36 An electron micrograph (TM) of an encysted larva of
Neochordodes occidentalis. Photograph by Roberta Poinar.
Earlier studies on life cycles of hairworms did not always
clearly distinguish between paratenic and developmental
hosts. While three types of life cycles have been described
for nematomorphs, the first two need verification. Dorier[18]
reported that eggs of Gordius hatched in water and the
preparasitic larva were ingested by and developed in caddisfly
larvae (Stenophylax). He also reported an indirect-free-living
stage in a species of Gordius that infected millipedes. After
the preparasitic larva hatched from the egg in this spe-
cies, it encysted on submerged leaves or other detritus. The
definitive host then became infected by ingesting cysts on
the exposed vegetation after the water evaporated. For this
cycle to be effective, the cysts would have to be resistant to
desiccation. Neither of these types of developmental cycles
has been confirmed. The most common life cycle can be
called indirect-paratenic, where the hatched preparasitic larva
is ingested by an invertebrate or vertebrate. Hatching occurs
when the mature preparasitic larva forces itself through the
wall of the egg (Fig. 9.37). The parasite burrows into the tis-
sues of this paratenic host and then ceases development after
encysting. Only when the paratenic host is eaten by a predator
(carabid beetle) or omnivore (various orthopteran insects) does
the parasite initiate development. This type of cycle occurs
in the great majority of investigated hairworms, including a
Chordodes developing in praying mantids that ingest infected
mayfly paratenic hosts[39], a Gordius developing in the aquatic
beetle Dytiscus that feeds on infected tadpole (Rana tempo-
raria) paratenic hosts[3], Euchordodes nigromaculatus devel-
oping in stenopelmatid crickets that fed on infected stoneflies,
mayflies, or caddisflies[64], Gordius robustus developing in
crickets that were fed infected meal worms[30], and probably
Neochordodes occidentalis which readily infects and encysts
in mosquitoes and other aquatic insects[69].
Hairworm larvae hatch by forcing their stylets against
the egg shell (Fig. 9.37). Once inside a paratenic host, lar-
vae penetrate hosts tissues with spines and stylets (Figs.
9.38, 9.39, 9.40). No evidence shows that the prepara-
sitic larvae can burrow directly through the outer body
wall of either the paratenic or definitive host. In all cases,
they enter by way of mouth and encyst in the midgut or
FIGURE 9.37 Preparasitic larva of Neochordodes occidentalis
(Montgomery) forcing its way out of the egg shell.
Chapter | 9 Nematoda and Nematomorpha 269
FIGURE 9.38 Three larvae of Neochordodes occidentalis penetrating
into the tissue of their paratenic host, a mosquito larva.
FIGURE 9.39 Forward-facing view of a larva of Neochordodes occi-
dentalis penetrating into the tissue of its mosquito larva paratenic host.
Note the circlet of six spines.
FIGURE 9.40 Larva of Neochordodes occidentalis attempting to pene-
trate the peritrophic membrane of its mosquito larva paratenic host. Note
the extended stylets.
larvae is humoral melanization which may occur immediately
after entry as in the case of N. occidentalis invading mosquito
larvae (Fig. 9.43)[69] or after the larvae have encysted in the
internal tissues of the paratenic host, as with hairworm larvae
in caddisflies (Fig. 9.44), mayflies, stoneflies, and especially
chironomid larvae[64]. Such melanization reactions often
result in the death of invading larvae. When the host reaction
occurred after encystment, however, the parasites may have
died of other causes and were then melanized.
When parasitic development has been completed and
the hairworm is ready to emerge, the host must come into
contact with water. This poses no problem for parasites in
aquatic hosts but can present obstacles when the host is a
burrow through the peritrophic membrane and midgut wall
and encyst in the tissues of the body cavity (Fig. 9.41).
Structures considered as possible sensory organs in the
anterior region of the larvae may explain orientation of the
parasite in regards host tissue (Fig. 9.42a). In cases of mass
infection as demonstrated in laboratory studies with larvae
of N. occidentalis invading mosquito larvae, the tissues
may be damaged to such a degree that the host dies[69].
In cases where the primary paratenic host is eaten by an
insect predator which is not suitable for further parasitic
development, the hairworm larvae may be able to reencyst
in the predator. Encysted larvae in dobsonflies probably
represented cases of secondary paratenic hosts[64].
Host immunity to hairworms has thus far been documented
only in the case of larvae entering or remaining in paratenic
hosts. The usual type of host reaction to such parasitic
FIGURE 9.41 Larva of Neochordodes occidentalis encysted in the body
cavity of its mosquito larva paratenic host. Note the large cyst enclosing
the larva.
Ecology and Classification of North American Freshwater Invertebrates
270
FIGURE 9.45 Chordodes morgani emerging from a cockroach.
FIGURE 9.44 Larva of a caddisfly that served as a natural paratenic host
to hairworms. Members of the Trichoptera are important paratenic hosts
for hairworms as well as developmental hosts for mermithid nematodes.
FIGURE 9.42 Electron micrographs (TM) of an encysted larva of
Neochordodes occidentalis. (A) Anterior region of presoma showing
an area (between the arrows) that could serve as a sensory organ. (B)
Detail of the cuticle of the presoma adjacent to the multilayered cyst wall
formed in a tadpole. Photographs by Roberta Poinar.
FIGURE 9.43 Melanin deposited on two unencysted larvae of Neochordodes
occidentalis as a defense reaction by a mosquito larva paratenic host.
terrestrial arthropod. Although submergence may occur acci-
dentally, it is more likely that the host suddenly has a desire
to reach water. This may result from partial desiccation due
to the actions of the parasite or be a consequence of some
abnormal stimuli from parasite products that affect physi-
ological centers of the host. When the host enters water, the
hairworms emerge (Figs. 9.31, 9.45) and either slowly sink
or initiate undulating body movements (Fig. 9.30). Males
Chapter | 9 Nematoda and Nematomorpha 271
are considered more active than females. As the parasites
mature in the host, they change from a light, cream color to
a yellowish brown or dark brown color. At the time of emer-
gence, most have already turned their natural color; but in
some cases, further darkening occurs after emergence.
Since there are so few records of known gordiids from
identified hosts, very little is known about host selection
and specificity. Some hairworms are probably restricted
developmentally to certain invertebrate genera, while oth-
ers may parasitize a wide range of hosts. Most definitive
hosts are medium- to large-bodied predaceous or omnivo-
rous arthropods. The great majority of freshwater nemato-
morphs have been collected from representatives of the
insect orders Coleoptera (Fig. 9.31) and Orthoptera (Fig.
9.45). A list of insect families known to contain develop-
ing stages of North American hairworms is presented in
Table 9.2. The diversity of paratenic hosts is quite great,
extending from trematodes[17] to vertebrates. Preparasites
will attempt to burrow and encyst into the soft tissues of
any organism ingesting them (Table 9.3).
The natural enemies of hairworms are poorly known.
Predation by rock bass, brown trout, crayfish[14], and brook
trout (Fig. 9.21)[92] have been documented. Birds may
occasionally feed on hairworms, as reported for a tapaculo
(Scleorchious rubecula) in Chile which brought an adult
Gordius to its nestlings (Mary F. Willson, personal corre-
spondence, 1994).
E. Sampling
Except for the marine genus Nectonema, all known repre-
sentatives of the Nematomorpha occur in freshwater. Their
habitats range from watering troughs and puddles to rivers,
lakes, subterranean streams, and even the Great Lakes[15,103].
No special sampling technique has been developed for hair-
worms. They are large enough to be seen with the naked
eye and can be netted or lifted out of the water by hand.
Stream forms often accumulate in slower-moving water lat-
eral to the main current (Fig. 9.30). Holes dug into small
streamlets will often catch and hold adults that are being
carried downstream and nets can be placed across small
streams to collect the unattached adults. Late summer and
spring are the best times to look for the free-living adults.
Although the long, whitish egg strands (Fig. 9.34) of some
species can sometimes be spotted in water, the preinfec-
tives are too smal1 to be found and are usually noted only
by chance when examining the paratenic hosts. Perhaps dis-
secting paratenic hosts and searching for encysted larvae is
the easiest way to determine if hairworms are present in an
aquatic system. The eggs of those species adapted to fast-
flowing streams can be spotted on surfaces of rocks and
other debris. Growing larvae in developmental hosts can be
found throughout the year but are much less frequent than
are encysted larvae in paratenic hosts and are nearly impos-
sible to keep alive if they are removed from their host prior
to completion of their development. No methods are avail-
able for in vitro development or culture of nematomorphs.
May[47] monitored development by injecting preparasitic
larvae of Gordius into the body cavity of long-horned grass-
hopper hosts (Tettigonidae). However, because of the frag-
ile character of preparasitic forms and the relatively long
period of parasitic development (several months) coupled
with the problem of maintaining the hosts, few workers
have even bothered to culture hairworms in vivo.
F. Identification of Nematomorpha Genera
Adult hairworms can be preserved in 5% formalin or 70%
alcohol for identification. All diagnostic characters are exter-
nal features associated with the head, tail, and epicuticular
surfaces. The extremities can be removed and mounted in
lactophenol or glycerin after dehydration in an alcohol series.
For cuticular examination, small slivers of epicuticle can be
removed from the midbody region with a razor blade and
placed in lactophenol or dehydrated in glycerin. The underly-
ing epidermis and muscle tissue can then be scraped away and
the cuticular slice mounted (outer surface up) in lactophenol
or glycerin. This will expose the areoles and their ornamen-
tation (Figs. 9.46, 9.47). Some workers prefer clearing sec-
tions in xylene before mounting them. As mentioned above,
the scanning electron microscope can reveal cuticular details;
however, it is not practical for field biologists. Adequate diag-
nostic characters can normally be observed at least with oil
immersion objectives (1000x). In a study comparing the light
and scanning electron microscope in differentiating cuticu-
lar characters separating Parachordodes lineatus and P. vio-
laceus, Smith[93] noted that while details were clearer with
SEM, the light microscope revealed the essential characters
needed to separate species. Success in large part with the light
microscope depends greatly on skill in preparing the sam-
ple. Both males and females should be available for accurate
identification; however, both sexes may not be present. Some
males can be identified alone using the following key.
The characters used in the following taxonomic key fol-
low that of Poinar and Chandler[68] and incorporates contri-
butions of numerous authors[7,9,10,43,47,50–53,61,69,78,84,92–94,99].
1.  Taxonomic Key to Genera and Species of North American Hairworms
1a. Cuticle smooth, lacking areoles or with flat, smooth areoles; male tail bilobed (as in Figs. 9.48a,b,c, 9.50a–c,f–k), with a postcloacal or
precloacal ridge or crescent; female tail entire (as in Fig. 9.49a) ................................. family Gordiidae . .......................Gordius L.......... 2
1b. Cuticle with distinct areoles usually ornamented with bristles or tubercles; male tail entire (Figs. 9.48d, 9.50e); or if bilobed, then with-
out a postcloacal (Fig. 9.50a) or precloacal ridge or crescent (as in Fig. 9.50c); female tail entire or trilobed (Fig. 9.49b) ........................5
Ecology and Classification of North American Freshwater Invertebrates
272
FIGURE 9.46 (A) Surface view of
areoles with minute spines present,
especially in the furrows between
the flattened areoles. (B) Surface
view of the areoles of Parachordodes
tegonotus showing the presence of
a megareoles. (C) Surface view of
the areoles of Gordionus diblastus
(Orley) showing variation in areole
size and distribution. (D) Lateral
view of the raised areoles of G.
diblastus showing size variability.
2a (1a). Male cuticle with large white spots; broad semicircular postcloacal crescent present (Fig. 9.50a); elongate white triangular streak from
neck to anterior fourth of body in both sexes; head tapers (spindle-shape) ................................................................ Gordius attoni Redlich
2b. Male cuticle with or without white spots, lacking an elongate white triangular streak; head variable .........................................................3
3a (2b). Male with both a precloacal semicircular hairline and a postcloacal ridge (Fig. 9.50b); areoles present or absent on females .....................
....................................................................................................................................................................... Gordius difficilis (Montgomery)
3b. Male with either a postcloacal or a precloacal ridge, but not both ................................................................................................................4
4a (3b). Male with a precloacal ridge (Fig. 9.50c) areoles confluent, interconnected .............................................Gordius alascensis Montgomery
4b. Male with a postcloacal ridge (Fig. 9.50a); areoles absent ........................................................................................ Gordius robustus Leidy
5a (1b). Female tail trilobed (Fig. 9.49b); male tail bilobed with length of lobes greater than twice their width (Fig. 9.48b); head end obliquely
truncated ..............................................................................................................................................................Paragordius varius (Leidy)
5b. Female tail entire; male tail bilobed with length of lobes equal to or less than twice their width (as in Fig. 9.50a–c), or male tail entire
(Figs. 9.50e, 9.48d) ........................................................................................................................................................................................6
6a (5b). Male tail bilobed ..........................................................................................................................................................................................12
6b. Male tail entire (Fig. 9.48d), lobes absent or abbreviated ..............................................................................................................................7
7a (6b). Cuticle with normally one kind of areole ...................................................................................................... Neochordodes Carvalho.........8
7b. Cuticle with two or more types of areoles .....................................................................................................................................................9
Chapter | 9 Nematoda and Nematomorpha 273
FIGURE 9.47 (A) Surface view of
the areoles on Euchordodes nigromac-
ulatus showing larger areoles on the
dorsal side of the body. (B) Edge of
the flattened areoles of Euchordodes
nigromaculatus. (C) Surface view of
the areoles of Neochordodes occi-
dentalis. (D) Bristles surrounding the
cloaca of Parachordodes tegonotus.
8a (7a). Areoles containing short bristles and tubercles (Fig. 9.47c) (some specimens possess a smaller type of areole) ..........................................
......................................................................................................................................................Neochordodes occidentalis (Montgomery)
8b. Areoles lacking bristles or tubercles ................................................................... Neochordodes californensis de Miralles and de Villalobos
9a (7b). Areoles with prominent tubercles, crowns or papillae; female cloacal aperture terminal ...................... Chordodes morgani (Montgomery)
9b. Areoles without prominent tubercles, crowns or papillae; female cloacal aperture subterminal .............. Pseudochordodes Carvalho......10
10a (9b). Smaller areoles with knobs on surface; arranged randomly ..................................................... Pseudochordodes gordioides (Montgomery)
10b. Smaller areoles without knobs on surface, arranged in uneven rows ..........................................................................................................11
11a (10b). Large elevated areoles in clusters of usually two ................................................................................... Pseudochordodes manteri Carvalho
11b. Large elevated areoles in clusters of three and four ........................................................... Pseudochordodes taxanus Schmidt-Rhaesa et al.
12a (6a). Areoles of two types, large megareoles with a central pore or tubercle and small microareoles lacking a pore or tubercle (Fig. 9.46b);
broad hair line of short simple and branched hairs beginning anterior to cloacal aperture but not reaching to point of bifurcation of tail
lobes (Figs. 9.47d, 9.50h) ......................................................................................... Parachordodes tegonotus Poinar, Rykken and LaBonte
12b. Areoles of one (microareolar) type, but of various sizes ....................................................................................... Gordionus Müller.........13
Ecology and Classification of North American Freshwater Invertebrates
274
FIGURE 9.48 Tails of male hair-
worms: (A) Gordius dimorphus; (B)
Paragordius varius; (C) Gordionus
diblastus; (D) Neochordodes
occidentalis.
13a (12b). Oblique rows of single and/or branched hairs flanking cloacal aperture (Fig. 9.50f,g); areoles closely opposed, may produce longitudinal
ridges ............................................................................................................................................................................................................14
13b. No such hairs on lateral body surface flanking cloacal aperture .................................................................................................................15
14a (13a). Body dark brown; areoles irregularly polygonal, not arranged in rows, well separated from one another by distinct interareolar region;
interareolar spaces with small spinules or papillate processes; areoles not showing sexual dimorphism; hairline on male tail not reaching
point of bifurcation of tail lobes (Fig. 9.50f); calotte white ............................................................................... Gordionus violaceus (Baird)
14b. Body light yellow to deep buff; areoles closely opposed without intervening spaces, having a tendency to group themselves in longitudi-
nal rows in males (sexual dimorphism); hairline extending beyond point of bifurcation of tail lobes (Fig. 9.50g); calotte dark or absent ...
...............................................................................................................................................................................Gordionus lineatus (Leidy)
Chapter | 9 Nematoda and Nematomorpha 275
FIGURE 9.49 Tails of female hair-
worms: (A) Parachordodes tegono-
tus; (B) Paragordius varius.
15a(13b). Tubercles located behind male cloacal aperture (Fig. 9.50g,h); tale lobes of male shorter than, equal to or longer than wide; female tail
not swollen; head variable ............................................................................................................................................................................16
15b. No tubercles behind male cloacal aperture (Fig. 9.50i); tail lobes of male longer than wide; female tail swollen; head flattened ................
..........................................................................................................................................................Gordionus platycephalus (Montgomery)
16a (15a). Tail lobes of males short (Fig. 9.50j), length equal to or shorter than width; black ring surrounds cloacal aperture; postcloacal integumen-
tary ridge; tubercles extend anterior to cloacal aperture; areoles more or less confluent, tending to merge with one another and produce
transverse rows, head usually cylindrical ...............................................................................Gordionus densareolatus (Montgomery 1898)
16b. Tail lobes of males longer than wide (Fig. 9.50k); no black ring around cloacal aperture; postcloacal integumentary ridge absent; tuber-
cles not extending anterior to cloacal aperture .............................................................................................................................................17
17a (16b). Areoles elongated, well separated from one another; few interareolar structures (tubercles) .........................................................................
..........................................................................................................................................................Gordionus longareolatus (Montgomery)
17b. Areoles rounded to polygonal, numerous bristles in interareolar spaces ................................. Gordionus sinepilosus Schmidt-Rhaesa et al.
Ecology and Classification of North American Freshwater Invertebrates
276
FIGURE 9.50 Male tails of North American hairworms.
H 5 hairline; CA 5 cloacal aperture; PE 5 precloacal
ridge or crescent; PO 5 postcloacal ridge or crescent;
S 5 semicircular hairline; T 5 tubercles (modified from
Poinar and Chandler100)
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... Gelişimlerini tamamlayabilmek için bir konağa ve sucul yaşam alanına ihtiyaç duyarlar. Ayrıca gelişimlerinde paratenik arakonak da kullanabilirler [11]. ...
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