905R73108
Biota of Freshwater Ecosystems
Identification Manual No. 10
GENERA OF FRESHWATER NEMATODES (NEMATODA) OF EASTERN NORTH AMERICA
by
V.R. Ferris
J.M. Ferris
J.P. Tj epkema
Department of Entomology
Purdue University
West Lafayette, Indiana 47907
for the
ENVIRONMENTAL PROTECTION AGENCY
EHviRQ'i:'.Ł:r''<,L PROTECTION AGENCY
Library, Resion V
1 North Wacker Drive
Chicago, Illinois 60606
Project # 18050 ELD
Contract # 14-12-894
January 1973
For sale by the Superintendent of Documents, U.S. Government Printing Office, Washington, D.C. 20402
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EPA Review Notice
This report has been reviewed by the Environ-
mental Protection Agency, and approved for
publication. Approval does not signify that
the contents necessarily reflect the views
and policies of the EPA, nor does mention of
trade names or commercial products constitute
endorsement or recommendation for use.
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Protection Agency, Washington, DC 20460.
PROTECTION
11
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FOREWORD
"Genera of Freshwater Nematodes (Nematoda) of Eastern North
America" is the tenth of a series of identification manuals
for selected taxa of invertebrates occurring in freshwater
systems. These documents, prepared by the Oceanography and
Limnology Program, Smithsonian Institution for the Environ-
mental Protection Agency, will contribute toward improving
the quality of the data upon which environmental decisions
are based.
Additional manuals will include but not necessarily be lim-
ited to, freshwater representatives of the following groups:
branchiuran crustaceans (Argulus~), amphipod crustaceans
(Gammaridae), isopod crustaceans (Asellidae), decapod
crustaceans (Astacidae), leeches (Hirudinea), polychaete
worms (Polychaeta), freshwater pianarians (Turbellaria),
dryopoid beetles, and freshwater clams (Sphaeriacea).
111
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ABSTRACT
An illustrated key to 56 genera of freshwater nematodes of eastern
North America is given. Notes are included on the significance of
nematodes in freshwater ecosystems, collecting and isolating nem-
atodes, slide preparations and counting, and identification and use
of the key.
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CONTENTS
Section Page
I Introduction 1
Collecting and Isolating Aquatic Nematodes 3
Techniques for Preparing Nematode Slides
and for Counting 6
Identification and Use of the Key 9
II Key to Genera of Freshwater Nematodes of
Eastern North America 13
III Classification of Genera Included in Key 29
IV Acknowledgements 31
V References 33
VI Glossary 35
VII Index of Scientific Names 37
VII
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FIGURES
Page
1 Amphid shapes: Achromadora sp., Prodesmodora sp.,
Monhystera sp., Plectus sp., Anaplectus similis, and
Tobrilus sp. 9
2 Esophagus shapes: Hirschmanniella sp., Aphelenchoides
sp. ,Rhabditis sp., BwtZeriwe sp., Plectus sp.,
Leptolaimus sp., Achromadora sp., Cylindrolaimus sp.,
Ironus sp., Aphanolaimus sp., Alaimus sp., and Tripyla sp. 10
3 Tail shapes: Mesodorylaimus sp., Aphelenchoides clarus,
Eudorylaimus meridionalis3 Achromadora sp., Tobrilus
sp., Monhystera sp., and Labronema thornei. 11
4 Heads: 4nonc7ms sp., Prismatolaimus sp., Butlerius sp.,
and Mononchoides sp. 13
5 Heads: Anatonchus sp., AfononcTmZ-us sp., and Miconchus
trionchus; female tail: Mononahulus sp. 14
6 Heads: Prionchulus punotatus, Mononehus popHlatus, Nygo-
laimus sp., Thornia sp., Tylenchus cylindrious3 and
Mylonchulus braehyuris; anterior parts: Nygolaimus
sp. and Thornia sp.; female tail: Thornia sp. 15
7 Heads: Ldbronema thorneis Paractinolaimus sp., Oxydivus
oxycephalus, Eudorylaimus meridional-is3 Aulolaimoides
elegans, Mesodoryla-imus sp., Laimydorus sp., Atylenchus
sp., and Dorylaimus sp.; anterior parts: Labronema
thorneij Oscyd-irus sp., Eudorylaimue mevidLonalis3 and
Aulolaimoides elegans; male tail: Mesodovylaimus sp. 16
8 Heads: Hemicycliophora vidua, Hirschmanniella sp., Aphelen-
ohoides sacoharij Ethmolaimus sp., Achromadora sp., and
Chromadorita leuckarti; anterior part: Tylenahus sp.;
female tails: Tylenchus exiguus and Hirschmanniella sp.;
male tails: Hirschmanniella sp. and Aphelenohoides
saochari; female gonads: Hirsehmanniella sp. and
Aphelenohoides sp. 19
9 Heads: Anaplectus similis, Monochromadora sp., Prodesmodora
sp., and Khabdolaimus sp.; anterior parts: Monoohromadora
sp., Prodesmodora sp., and Rhabdolaimus sp.; tails:
Monochromadora sp. and Rhabdolaimus sp. 21
10 Heads: Monhystrella sp., Leptolaimus sp., Cylindrolaimus
sp., and Plectus sp.; female gonads: Monhystrella sp.,
Leptolaimus sp., and Cylindrolaimus sp.; tails: Leptol-
aimus sp. and PZectus sp. 22
11 Heads: Aphanolaimus sp., Paraphanolaimus sp., Paracyath-
olaimus truncatus, Monhystera sp., Tobrilus sp.,
Tripyla sp., Teratocephalus sp., Euteratocephalus sp.,
Bastiania exilis, Amphidelus hortensis, and Alaimus
primitivus; posterior part: Morihystera sp.; female
gonads: Tripyla sp. 23
Vlll
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FIGURES - continued
Page
12 Heads: Ivonus sp., Cryptonehus sp., and Goffartiasp.;
female gonads: Ironus sp.; female tails: Cylindrola-irms
sp. and Cryptonchus sp. 25
13 Heads: Odontolaimus sp., Chronogaster sp., Fthabditis sp.,
and AaTdbeloides nannus; anterior part: Chronogastev sp. 26
14 Anterior part: Aarobeloides sp.; female tails: A.cvdbe1oides
nannus, Euoephalobus oxyuroides, and CephaLobus sp.;
heads: TSucephalobus oxyuroides and Cephalobus sp. 27
IX
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SECTION I
INTRODUCTION
Fifty-six genera of freshwater nematodes are included in this key. It
is reasonable to assume that knowledge about such a large group of
animals would be important and useful to the understanding of fresh-
water ecosystems. Nevertheless, nematodes have been overlooked or
avoided by most aquatic biologists, probably because they are small
and somewhat difficult to handle. They are frequently very numerous,
although their total biomass may be relatively low because of their
small size. However, their role in aquatic ecosystems may be much
greater than their biomass would indicate since they have a very high
metabolic rate.
Nematodes are found in soil and marine habitats as well as all fresh-
water habitats. Some nematodes are capable of living in both soil
and freshwater whereas others are found only in freshwater or only
in soil. The orders Tylenchida and Dorylaimida are primarily found
in soil and have only a few freshwater representatives. Other orders
of nematodes are better represented in freshwater than in soil, and
are even more richly represented in marine habitats. Parasitic
nematodes occur on or in many aquatic animals, but are not included
in this key, as they are usually not collected in the same manner as
free-living nematodes or nematodes parasitic on plants.
The only major systematic publications on North American aquatic
nematodes are those of N.A. and M.V. Cobb (1913, 1914, 1915). How-
ever, samples of aquatic nematodes seldom yield species which cannot
be placed in a known genus, although undescribed species may be very
common. Many of the genera of aquatic nematodes are very widely dis-
tributed, and at least some may be cosmopolitan in distribution.
Nematodes feed on a wide variety of organisms, but apparently do not
feed on dead organic matter. Stylet-bearing nematodes feed on many
types of higher and lower plants and on small animals by puncturing
them and drawing out the liquid contents. Nematodes with simple, un-
armed stomata probably feed mainly on small unicellular organisms such
as bacteria. Some nematodes have large teeth which they use to attack
other small animals including other nematodes.
A recent study in our laboratory (Ferris et al., 1972) has indicated
that an increased understanding of disturbances to aquatic habitats
can be obtained by a study of nematode communities of the habitats.
In our study of small freshwater streams, nematode species of the
genera Morihystera, Mesodorylaimus and Tylenchus proved to be especially
numerous in certain of the stream sites, with species of the genera
Aerobeloides, Tobrilis^ Monondhoides and Goffartia also numerous at
some sites.
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It is hoped that this key will make possible more extensive use of data
on nematode community structure by persons concerned with evaluating
water resource environments. This kind of analysis can be a. useful and
practical tool, particularly when used in combination with other avail-
able techniques, for interpreting ecological conditions and providing
indices of change.
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COLLECTING AND ISOLATING AQUATIC NEMATODES
Nematodes can be found in or on all kinds of benthic substrates. Since
they are too small to be collected individually in the field, samples of
substrate are usually taken to the laboratory for processing. Nematodes
may also be found in small numbers floating in water even though they
are primarily associated with benthic material. Floating nematodes have
been collected by passing water through fine screens (U.S. Standard
Series #400 sieve), the nematodes and associated debris accumulating on
the screen (Faulkner and Bolander, 1966).
Nematodes must be separated from all particulate matter from their habi-
tat before they can be examined. Even tiny particles of soil or debris
will obscure the morphologic details of these microscopic organisms.
They may be isolated by picking them out of small samples of substrate
using a finely pointed bamboo or nylon needle ("pick") while observing
the manipulations under a stereoscopic microscope. However, hand sort-
ing is used when only a few nematodes are needed since it is very time
consuming.
During the processing of large samples (500-1000 cc) of particulate mat-
ter consisting of silt, clay, sand and organic matter, nematodes may be
partially separated from this debris by a combination of decanting and
sieving. To process a sample in this manner, soil or sediment is mixed
with water, allowed to stand for 30 seconds to allow heavier particles
to settle, and the supernatant (containing the nematodes) is poured
through a sieve. Various sizes of sieves are used to remove nematodes
from the water. All aquatic nematodes will pass through a U.S. Standard
Series #10 sieve, so this size is used to remove floating organic matter.
Very large nematodes (2-4 mm in length) will collect on a #25 sieve. A
#325 sieve will catch the smallest (0.5-0.1 mm) nematodes unless they
pass through head or tail first. By passing water containing suspended
nematodes through a #325 sieve several times, almost all the nematodes
will be caught. Processing soil in this manner will not separate the
nematodes from all particles, but it does concentrate them so that they
can be more effectively isolated with a Baermann funnel. A Baermann
funnel is made by slipping a short piece of rubber tubing on the stem of
a 100-150 mm funnel. The tubing is closed with a Day-type pinch clamp
so that the funnel will hold water. Nematodes and substrate are placed
in the water on top of a paper handkerchief (any commercial brand paper
handkerchief or tissue which possesses sufficient wet-strength not to
disintegrate may be used) or muslin square supported by a wire basket in
a funnel filled with water to a level that just covers the sample.
Twenty-four hours later the nematodes which have fallen into the stem of
the funnel after migrating through the tissue or muslin can be removed
by opening the pinch clamp and collecting 5-10 ml of water. Very small
samples of soil or samples of substrate such as twigs or small stones
with nematodes on their surfaces can be placed directly in Baermann fun-
nels to isolate the nematodes.
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Steps used in isolating nematodes by decanting, sieving and the Baermann
funnel may vary, but the following procedure, used in our laboratory,
works well for routine mass collections:
Materials:
2 ten quart buckets labelled "A" and "B"
2 flat pans 18-25 cm diam. X 6-8 cm high
3 250 ml beakers
2 Baermann funnels for each sample processed
1 series of U.S. Standard Series sieves: #10, #25,
#100, and #270
Steps:
1. Place 500-1000 cc of sediment or soil in bucket A. Fill bucket one-
fourth or less with water and break up all lumps. Allow material
to stand 30 seconds and then decant supernatant through the #10
sieve into bucket B. Repeat this operation, washing all fine par-
ticles from the coarse material in bucket A, until bucket B is
filled to within 5-10 cm of the top. Discard material remaining
in the bottom of bucket A and on the #10 sieve. Rinse bucket A
to clean.
2. Pour the contents of bucket B through a #25 sieve, catching the
water passing through the sieve in bucket A. Decanting must be
stopped before large soil particles collect on the sieve. Invert
the sieve and flush material caught on the sieve into one of the
pans. After a short settling period, pass the water in the pan
through the sieve again and catch this water in bucket A. Rinse
the residue on the #25 sieve into a clean pan using 250 ml of
water or less. Then pour the contents of the pan into a 250 ml
beaker and set it aside to permit nematodes to settle to the
bottom. Rinse bucket B to clean.
3. Repeat step 2 using a #100 sieve, pouring the contents of bucket
A through the sieve and catching the water again in bucket B.
Wash the residue on the #100 sieve into a pan, then pass the
contents of the pan through the #100 sieve again (catching this
water in bucket B). Re-suspend material caught on this sieve in
250 ml of water or less and place in a second beaker, to allow
all nematodes to settle to the bottom.
4. To obtain the remainder of the nematodes, pour the contents of
bucket B through the #270 sieve, catching the water in bucket A.
After washing the material caught on the #270 sieve into a pan,
pass the water in bucket A through the #270 sieve again (this
water may now be discarded). Wash the material on the sieve into
the same pan used for the first material caught on the #270. In
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all instances, pouring from a bucket through a sieve should be
stopped before the silty material in the bottom of the bucket
is decanted onto the sieve. This remaining silt is discarded
when the bucket is rinsed.
5. The nematodes, now in the water in the pan, are further con-
centrated by pouring the water through the #270 sieve, catching
it in the second pan. Repeat this procedure two more times,
each time pouring the water through a different area of the
same #270 sieve. The residue now caught on the sieve is washed
off the sieve and back into the pan using 250 ml or less of water.
Allow this to settle 30 seconds and decant into a third 250 ml
beaker where nematodes will settle to the bottom.
6. After the contents of the three 250 ml beakers have settled (for
about an hr.) carefully decant the supernatant.
7. Pour some of the residue from the beaker in which material from
the #25 sieve was saved into a Syracuse watch glass or other
flat-bottomed dish and examine with a stereoscopic microscope
at 30-60 X magnification. Large nematodes may be picked out of
the dish with a finely pointed pick and transferred to a vial
of water. Examine all the material from this beaker, picking
out all the large nematodes observed.
8. To complete the nematode separation procedures, the residues
from the #100 and #270 sieves (now in beakers) are each placed
separately on tissue or muslin in Baermann funnels, in the
manner described previously. Nematodes are removed from the
funnels at 24 and 48 hours by opening the pinch clamp and drawing
off 5-10 ml of fluid into a vial. It may be necessary to add
additional water to the funnel to keep the material submerged.
The nematodes should now be free of silt and organic matter,
ready for preservation and observation.
Other methods for extraction of nematodes are available, and are de-
scribed in various nematology texts (Thorne, 1961; Southey, 1970).
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TECHNIQUES FOR PREPARING NEMATODE SLIDES AND FOR COUNTING
Identification of nematodes usually requires they be mounted on slides.
They may be mounted temporarily in water or fixative, or permanently in
glycerin. Permanent mounts should be used when extensive study is con-
templated. It is not necessary to stain nematodes for identification
procedures.
In order to preserve nematodes, they must first be killed by gentle
heat. Such a killing procedure is necessary because live nematodes
placed directly in fixative become distorted. Nematodes may be killed
by placing the vial containing them in a water bath at 57°C for 10 min-
utes, or in an oven at 52°C for 15 minutes. If an oven or water bath
is not available, nematodes may be heat relaxed by adding a quantity
of boiling water to an equal quantity of water at room temperature in
a beaker containing the nematodes to be killed. After heat relaxing,
the nematodes should immediately be placed in fixative. The fixative
may be warmed so that it is about the same temperature as the water
containing the killed nematodes. Many different fixatives are avail-
able, but a commonly used one is 5% formalin. For fixing, and also for
storing mass collections, a 10% solution of formalin is added to an
equal quantity of water containing killed nematodes, so that the final
concentration of formalin is 5%. Nematodes for permanent mounts should
be fixed in F.A.A. (8 ml commercial 37% formaldehyde solution, 1 ml
glacial acetic acid, 20 ml 95% ethanol, and 50 ml H20) for at least
two weeks. This period of time insures proper fixation of fine, de-
finitive morphological details.
Temporary mounts can be made by transferring freshly killed nematodes
to a drop of 5% formalin on a slide and placing a cover glass on the
drop. The drop of formalin should not be so large that excessive
amounts of fluid run out from under the cover glass, or so small that
air pockets form under the cover glass. The nematodes should be at
the bottom and middle of the drop on the slide, and three glass rods,
each approximately 1 mm long, should be arranged around them before the
cover glass is lowered. The glass rods, about the same diameter as the
nematodes to be mounted, prevent the cover glass from flattening the
nematodes. A selection of various sized fine glass rods can be obtain-
ed by heating rods of soft lead glass over an alcohol burner and pulling
them apart. To produce very fine rods, hot glass must be drawn out
quickly; slower pulling produces rods of greater diameter. Glass wool
or angel hair may contain the right diameter fibers for some nematodes.
When making temporary mounts, the cover glass should be lowered slowly
to prevent the nematodes from moving to the edge. Next the cover glass
should be tacked down with small drops of ringing compound at several
points around the edge. After these tacks have dried, the remainder
of the cover glass is sealed to the slide with more ringing compound.
Temporary slides can be sealed with wax by lighting a small candle,
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putting it out, and applying the hot wax using the wick of the candle
as a brush. Clear finger nail polish may be used as a ringing mate-
rial. A special ringing compound, called Zut (available from Bennett
Paint Products, Salt Lake City, Utah), is commonly used by nematol-
ogists for ringing temporary as well as permanent slides. Zut is
thinned with butyl acetate to a consistency which is easy to apply
with a #3 or finer camel's-hair brush. However, it should be thick
enough to give a good seal. Brushes may be cleaned with butyl acetate.
For permanent mounts, nematodes previously fixed in F.A.A. are infil-
trated with glycerin. To do this, place the nematodes in an alcohol-
glycerin mixture which contains 1 1/2% glycerin (3 ml glycerin, plus
50 ml ethanol, plus 147 ml H20). Allow the water and alcohol to
evaporate off slowly over a period of about four weeks. If the des-
iccation process takes place too rapidly, the nematodes will collapse.
A convenient way to control the dehydration is to place the 1 1/2%
glycerin solution containing the nematodes in a small watch glass
which holds about 2 ml of fluid. At the start of the dehydration-
infiltration, the watch glass should be filled to the top with the
dilute glycerin mixture. The watch glass can be placed in a con-
tainer such as a preparation dish. A desiccant such as calcium
chloride in a small screw-cap vial with a small hole drilled in it is
placed in the preparation dish with the watch glass. Petroleum jelly
should be applied between the rim and the top of the dish to seal it.
We have obtained good results by placing about a dozen of the small
watch glasses in a square plastic refrigerator-storage container
(sandwich-size) with a very tight lid. One vial of desiccant prepared
as described above is placed in the container with the watch glasses.
At the end of the dehydration period, only a thick film of glycerin
with nematodes remains in the bottom of the watch glass. Infiltrated
nematodes should be stored in a desiccator since glycerin readily
absorbs moisture from the atmosphere.
Permanent slide mounts are made with glycerin-infiltrated nematodes in
a fashion similar to that described above for temporary mounts. From
1-6 nematodes of like diameter are selected for mounting on a single
slide. Special care should be taken in selecting the glass rods for
supporting the cover glass and in using the correct amount of glycerin.
If the rods are larger in diameter than the nematodes, or if too much
glycerin is used, the nematodes will float under the cover glass,
making observations with the oil immersion objective of a compound
microscope extremely difficult. On the other hand, if the diameter of
the rods is too small, the nematodes will be flattened and distorted.
Arrange the nematodes carefully in the center and bottom of a drop of
glycerin with their heads all pointed in the same direction. Use
glycerin which has been stored in a desiccator. Arrange three glass
rods around the nematodes. Warm a cleaned cover glass over an alcohol
lamp and lower it slowly onto the glycerin. Tack the cover down with
Zut. After the tacks have dried, ring the cover glass with more Zut.
Nematode slides should not be stored resting on their edges.
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Cobb metal slide mounts are often used by nematologists for mounting nem-
atodes. These mounts do not break as easily as glass slides and have the
added advantage that they can be stacked and the ringing material of one
slide does not touch the one next to it. In a Cobb mount, the nematodes
are mounted between two cover glasses, and thus can be examined with an
oil immersion objective from either the top or bottom of the slide. A
Cobb slide consists of a 25 mm square #1 cover glass held in place over
a round hole (18 mm diameter) in the center of an aluminum 75 X 25 mm
slide (with rolled edges) by two pieces of cardboard (Mason and Bosher,
1963). An 18 mm circular cover glass is placed on top of the square
glass over the hole in the metal slide. After the Zut has dried, the
edges of the Cobb mount are crimped over the cardboard to hold the square
cover glass firmly in place. (See Thorne (1961) for more details regard-
ing techniques for making slides).
For identification of nematode species in an area under study, slide
mounts should be prepared by one of the methods described above. After
initial identification, however, most species can be recognized subse-
quently at magnifications available on low-power stereoscopic microscopes
(e.g. 30 X and 60 X). This makes possible the counting of individuals
of dominant species (obtained from substrate samples of standardized
sizes) in a petri dish marked off in squares or lines. If the sample
contains too many individuals for accurate counting, a measured portion
or aliquot may be drawn off and additional water or fixative added before
counting the nematodes in the aliquot. The number of individuals in the
entire sample is calculated based on the size of the aliquot and the
numbers of individuals actually counted. Most nematologists prefer to
count more than one aliquot to increase the accuracy of the population
estimate.
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IDENTIFICATION AND USE OF THE KEY
The basic body shape of nematodes is an elongate cylinder with the oral
opening at the anterior end. These organisms are internally nonsegment-
ed, although thickenings of the cuticle may give the appearance of rings
or body segmentation. Several of the sediment inhabiting nematode genera
have readily observed amphids (Fig. 1 A-F), sensory organs located behind
the lips, which are of importance in identification procedures. The
stoma ("mouth") of a nematode, intimately in contact with its food
source, shows a variety of forms and modifications which are used to di-
agnose genera. The esophagus is located in the area between the stoma
and the intestine. Its form and shape (Fig. 2 A-L) is also of
diagnostic importance. The anus, ventral and sub-terminal on nematodes,
serves as a demarcation point for the region referred to as the "tail".
Thus the tail is that portion of a nematode posterior to the anus. The
tail shape (Fig. 3 A-G) is often used to separate nematode genera. Nem-
atodes are biparental with the sexes differing primarily in their sec-
ondary sexual characters: one or two ovaries and a vulva in the female;
one or two testes, one or two spicules, a bursa (not always present) and
a cloaca in the male.
B
Fig. 1 - Amphid shapes: A, head, Aehromadora sp. (multispiral);
B, head Prodesmodora sp. (unispiral); C, head Monhystera sp.
circular); D, head Pleatus sp. (open circle); E, head Anaplectus
similis (slit-like); F, head Tobrilus sp. (stirrup-shaped);
(all X 1000).
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K
B C
H
K
Fig. 2 - Esophagus shapes: A, Hirschmanniella sp. (X 500); B,
Aphelenohoides sp. (X 500); C, Khdbditis sp. (X 500); D, Butler-
ius sp. (X 250); E, Pleatus sp. ()( 250); F, Leptolaimus sp.
(X 500); G, Aehromadora sp. (X 500); H, Cylindrolaimus sp. (X 500);
I, Ironus sp. (X 250); J, Aphanolaimus sp. (X 250); K, Alaimus
sp. (X 250); L, Tripyla sp. (X 250).
10
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Whenever possible, difficult or ambiguous characters have been avoided in
this key. In general, the diagnostic characters used in this key are il-
lustrated with line drawings of representative species. Frequently two
or three characters are used in a couplet, instead of a single character,
to give more confidence in making identifications and also to provide
more information about the traits of each genus. Definitions for terms
which may not be familiar to the general biologist are given in a glos-
sary (p. 35).
B
s
' s
Fig. 3 - Tail shapes: A, Mesodorylaimus sp.; B, Aphelenchoid.es alarus; C,
Eudorylaimus meridional-is; D, Achromadora sp.; E, Tobrilus sp.; F, Mon-
hystera sp.; G, Labronema thornei (all X 500). s=spinneret.
Occasionally a specimen may be encountered which cannot be identified
using this key because it is an unusual form of a known genus or it
belongs to an undescribed genus. Such specimens will probably be rare
since the nematode genera of northeastern United States are fairly well
known. To be certain of identification, specimens should be compared
with figures in the key, and if any doubt remains, they should be checked
against complete descriptions, or they may be sent to an expert for veri-
fication. Many genera of terrestrial nematodes (which are often washed
into aquatic habitats) are not treated in this key, so other references
must be used if identification to genus of these specimens is desired.
Two orders (Tylenchida and Dorylaimida) and two families (Rhabditidae
and Diplogasteridae) which contain many of the genera of terrestrial nem-
atodes are end points in the key. For further information on the Tylen-
chida and Dorylaimida, refer to Zuckerman, Mai and Rohde (1971).
11
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The book "Soil and Freshwater Nematodes" (Goodey, 1963) may be used as a
general reference since it contains descriptions of almost all genera of
terrestrial and aquatic nematodes. However, there are two exceptions.
Laimydorus Siddiqi, 1969, was described after publication of the Goodey
book, and Hirsehmann-iella is listed as Hirsohmannia in Goodey. This book
has a fairly extensive list of references for soil and aquatic nematodes.
The book "Principles of Nematology" by Thome (1961) is an especially
good reference for soil forms. For more extensive listings of taxonomic
works, the check lists of Tarjan (1960, 1967) and Baker (1962) should be
consulted. Helminthological Abstracts is a good source for references
dealing with all phases of nematology.
Identification of aquatic nematodes to species is usually difficult even
for experts, and is complicated by the occurrence of undescribed species.
Since an extensive literature file is required for species identification,
it is suggested that material be sent to a nematode taxonomist, if it is
considered essential that a specific name be given to a specimen. Nema-
tologists who have recently published descriptions of aquatic or free-
living nematode species are good candidates for this service. Needless
to say, as long as a species is recognizable, much valuable ecological
information may be obtained from it, even though it is referred to by a
code designation rather than a Latin name. In the interests of accurate
science, preserved specimens should be retained of all species considered
in any data used in publication.
A list is appended to the key which places each genus in an order and sub-
order of nematodes. The classification is based on that of de Coninck
(1965) and differs somewhat from that of Goodey (1963). Families are not
included because of the frequent changes now occurring in this taxon.
12
-------
SECTION II
KEY TO GENERA OF NEMATODES OF EASTERN NORTH AMERICA
1 Stoma large, cup-shaped; width and depth of stoma at least
one-half of lip region; stoma strongly cuticularized
(Fig. 4 A, B, C: 5 D) 2
Stoma not both wide and cup-shaped and either weakly or
strongly cuticularized 11
2(1) Cephalic setae present (Fig. 4 A, B) 3
Cephalic setae absent (setose papillae may be present) .... 4
3(2) Tail elongate-clavate with spinneret; 4 large cephalic
setae present (Fig. 4 A): Anonahus
Tail filiform and lacking spinneret; 6 large and 4 short
cephalic setae present (Fig. 4 B): Prismatolai-mus
4(2) Esophagus with median and posterior bulbs (Fig. 2 D)
Diplogasteridae 5
[Several genera in this family, in addition to those
included in this key, may occasionally be found in aquatic
habitats. See Goodey (1963) for names of genera and
illustrations.]
Esophagus cylindrical; no median bulb 6
5(4) Stoma very broad and deep; anterior edge of stoma not
marked and not bearing grooves or rib-like structure
(Fig. 4 C): Butlerius
Stoma deep and moderately broad; anterior edge of stoma
bearing rib-like structures; slender tubular section
of stoma extending posterior to large tooth in stoma
(Fig. 4 D): Uononohoid.es
Fig. 4 - A, head Anonchus sp.; B, head Pvismatolaimus sp.; C, head
Butlerius sp.; D, head Mononohoides sp.; (all X 1000).
13
-------
6(4) Stoma with large subventral tooth; dorsal tooth
obscure (Fig. 5 B); spinneret on ventral portion
of tail (Fig. 5 C): Monondhulus
Stoma with large dorsal tooth; size of subventral
teeth variable; spinneret at terminus of tail 7
7(6) Dorsal tooth posteriorly directed in stoma (Fig. 5 A) -.Anatonchus
Dorsal tooth anteriorly directed in stoma 8
8(7) Large dorsal tooth posteriorly placed in stoma; two
large subventral teeth opposite to dorsal tooth
(Fig. 5 D): Miconchus
Dorsal tooth anteriorly placed in stoma; subventral
teeth absent or not large 9
-s
Fig. 5 - A, head Anatonchus sp. (X 500); B, head Mononahulus sp. (X 1000);
C, female tail Monondhulus sp. (X 500), s=spinneret; D, head Mioonohus
trionchus (X 500).
9(8) Stoma with transverse row of denticles opposite to
dorsal tooth (Fig. 6 I): Mylonchulus
No transverse row of denticles opposite dorsal tooth 10
10(9) Longitudinal ridge without denticles opposite dorsal
tooth (Fig. 6 B) : Mononohus
Longitudinal row of denticles opposite dorsal tooth
(Fig. 6 A): Prionchulus
11(1) Stoma armed with spear (Fig. 6 H; 7 C, F; 8 A) or
spear-like tooth (Fig. 6 C) 12
Stoma lacking spear or spear-like tooth 26
14
-------
IT\
Fig. 6 - A, head Prionchulus punctatus (X 750); B, head Monanchus
papifiatus (X 750); C, head Nygolaimus sp. (X 1000); D, anterior
part Nygolaimus sp. (X 250); E, anterior part Thornia sp. (X 250);
F, head Thornia sp. (X 1000); G, female tail Thornia sp. (X 500);
H, head Tylenohus aylindriaus (X 1000); I, head MylanahuluB
braahyurie (X 750).
15
-------
B
H
M :
Fig. 7 - A, head Labronema fhornei (X 1000); B, anterior part Labronema
thornei (X 100); C, head Paraotinolaimus sp. (X 1000); D, head Oxydirus
oxyeephalus (X 1000); E, anterior part Oxydims sp. (X 250); F, head
Eudorylaimus mevidionalis (X 1000); G, anterior part Eudorylaimus
meridionalis (X 100); H, anterior part Aulolaimoides elegans (X 250); I,
head Aulolaimoides elegans (X 500); J, head Mesodorylaimus sp. (X 1000);
K, male tail Mesodorylaimus sp. (X 500); L, head Laimydorus sp. (X 500);
M, head Atylenchus sp. (X 1000); N, head Dorylaimus sp. (X 500).
16
-------
12(11)
13(12)
14(13)
15(14)
16(15)
17(13)
18(17)
Esophagus lacking median bulb; amphid stirrup-
shaped and fairly distinct. Dorylaimida 13
Esophagus with median bulb; amphid obscure. Tylenchida 22
[Several genera in these orders which are not
truly aquatic and are not included in this key,
may be found in aquatic habitats and may be
carried into aquatic situations for which they
are not adapted. For further information on
such genera and illustrations see Thorne (1961)
and Goodey (1963).]
Tails of both sexes short and blunt or elongate, but
not filiform (Fig. 3 C, G) 14
Female or both female and male tail filiform (Fig. 3 A) ... 17
Tail blunt and approximately twice as long as the
anal body diameter (Fig. 6 G); anterior end of
body nearly square (Fig. 6 F); esophagus expands
very sharply at middle (Fig. 6 E): Thornia
Tail pointed or if blunt then distinctly shorter
than twice anal body diameter; anterior end of
body somewhat rounded; esophagus expands gradually 15
Tooth-like spear attached to stoma wall; stoma wall
cuticularized and well separated from tooth
(Fig. 6 C); esophagus often expanded anterior to
middle (Fig. 6 D): Nygolaimus
Spear present, located centrally in stoma; walls of
stoma weakly developed and barely visible; esophagus
usually expanded at middle
16
Anterior portion of esophagus considerably narrower
than posterior portion (Fig. 7 G); total length
usually 2 mm or less (Fig. 7 F): Eudorylctimus
Anterior of esophagus nearly as wide as posterior
portion (Fig. 7 B); total body length usually
greater than 2 mm (Fig. 7 A): Labronema
Enlarged basal portion of esophagus short, barely
longer than wide (Fig. 7 H); weakly developed
spear surrounded by faint cuticularized ribs
(Fig. 7 I):
Basal portion of esophagus considerably longer than
wide; spear not surrounded by ribs
Aulolaimo-ides
18
Four large teeth or thick cuticularized framework
surrounding spear; denticles present in cavity
around spear (Fig. 7 C): Paraotinolaimus
No large teeth or thick cuticularized framework surrounding
spear; no denticles in cavity around spear 19
17
-------
19(18) Esophagus surrounded by muscular sheath (Fig. 7 E);
spear length approximately equal to lip width
(Fig. 7 D); both male and female tail filiform: Oxydirus
Esophagus not surrounded by muscular sheath; spear
longer than lip width; male tail short and blunt
(Fig. 7 K) 20
20(19) Cuticle with longitudinal ridges (Fig. 7 N); total
body length greater than 2 mm: Dorylcn-mus
Cuticle without longitudinal ridges; total body length
greater or less than 2 mm 21
21(20) Total length usually less than 2 mm; spear guiding ring
usually single (Fig. 7 J): Mesodovylaimus
Total length usually greater than 2 mm; spear guiding ring
usually double (Fig. 7 L): Laimydorus
(very close or identical to Mesodovy'Laimus')
22(12) Cephalic setae present (Fig. 7 M); cuticle with longi-
tudinal ridges and very coarse annulation (Fig. 7 M):
Atylenchus
Cephalic setae absent; cuticular ridges lacking; cuticle
annulation not coarse 23
23(22) Extra, loose outer cuticle layer; spear long, approximately
same length as esophagus (Fig. 8 A): Henri-aycliophora
No extra loose cuticle layer; spear fairly short and not
nearly as long as esophagus 24
24(23) Tail filiform (Fig. 8 B); esophagus not overlapping
intestine, and median bulb of esophagus small and
ovate (Fig. 8 C): Tylenchus
Tail elongate conical but not filiform (Fig. 3 B; 8 F);
esophagus overlapping intestine, and median bulb of
esophagus prominent and spherical (Fig. 2 A, B) 25
25(24) Spear slender with or without modified basal knobs
(Fig. 8 H); single ovary anterior to posterior
vulva (Fig. 8 I); bursa lacking (Fig. 8 J): Aphelendhoides
Spear stout with large basal knobs (Fig. 8 D); two
ovaries, one anterior and one posterior to median
vulva (Fig. 8 G); bursa present (Fig. 8 E): Hirsofananniella
26(11) Spinneret present (Fig. 3D, E, F) 27
Spinneret lacking or obscure 44
27(26) Esophagus with basal bulb (Fig. 2 E, F, G) 28
Esophagus cylindrical with no basal bulb (Fig. 2 J, L) .... 37
18
-------
B
Fig. 8 - A, head HemLoyoliophora vidua (X 500); B, female tail Tylenohus
exiguus (X 500); C, anterior part Tylenohus sp. (X 500); D, head Hirsah-
manniella sp. (X 1000); E, male tail Hirschmanniella sp. (X 500); F,
female tail Hirsohmanniella sp. (X 500); G, female gonads Hirschmanniella.
sp. (X 100); H, head Aphelenohoides sacchari (X 1000); I, female gonad
Aphelenahoides sp. (X 250); J, male tail Aphelenohoides sacdhari (X 500);
K, head Ethmolaimus sp. (X 1000); L, head Aehromadora sp. (X 1000); M,
head Chromadorita. leuckarti (after Micoletzky, 1925).
19
-------
28(27)
29(28)
30(29)
31(29)
32(31)
33(32)
34(28)
35(34)
Araphid multispiral (Fig. 1 A), slit-like (Fig. 1 E)
or obscure
Amphid large, unispiral, circular or open circle
(Fig. 1 B, C, D)
Amphid spiral
Amphid slit-like or obscure
29
34
30
31
Dorsal tooth and opposing subventral teeth large;
tubular section of stoma extending posterior
to teeth (Fig. 8 K):
Dorsal tooth large, and opposing subventral teeth
small or obscure; stoma ending just posterior
to teeth; tubular section posterior to teeth
lacking (Fig. 8 L):
Stoma with large sharp dorsal tooth and obscure
subventral teeth (Fig. 8 M); cuticle marked
with punctations:
Stoma with 2 or 3 small teeth or no teeth;
cuticle lacking punctations
Ethnolaimus
Adhromadova
Chromadorita
32
Cephalic setae present; cuticle distinctly
annulated (Fig. 9 A): Anapleetus
Cephalic setae absent; cuticle weakly annulated
or not annulated 33
Inconspicuous peg-like spinneret (Fig. 9 D);
stoma funnel-shaped (Fig. 9 B); basal bulb of
esophagus lacking valves (Fig. 9 C):
Prominent long conical spinneret (Fig. 9 I);
stoma tubular (Fig. 9 G); basal bulb of
esophagus with valves (Fig. 9 H):
Basal bulb of esophagus divided transversely
in two places (Fig. 9 E); anterior end of
of body set off by shallow construction at
level of base of stoma (Fig. 9 F):
Basal bulb of esophagus not divided transversely;
anterior end of body not set off, or set off
at base of lips
Monochromadora
Khabdolaimus
Prodesmodora
35
Cuticle not annulated (Fig. 10 A); single ovary
anterior to posterior vulva (Fig. 10 B):
Cuticle annulated; two ovaries, one anterior and
one posterior to median vulva (Fig. 10 E)
Monhystrella
36
20
-------
H
-s
-s
Fig. 9 - A, head Anaplectus similis (X 1000); B, head Monochromadora sp.
(X 1000); C, anterior part Monochromadora sp. (X 500); D, tail terminus
Monochromadora sp. (X 500); E, anterior part Prodesmodora sp. (X 500);
F, head Prodesmodora. sp. (X 1000); G, head Rhdbdolaimus sp. (X 1000); H,
anterior part fthabdolaimus sp. (X 500); I, female tail Rhabdolaimus sp.
(X 500) 8=spinneret.
36(35)
37(27)
38(37)
Cephalic setae absent; stoma cylindrical (Fig. 10 C);
tail moderately clavate (Fig. 10 D): Leptolairms
Cephalic setae present; walls of stoma converging
slightly posteriorly (Fig. 10 J); tail not clavate
or slightly clavate (Fig. 10 H, I): Pleotus
Spinneret located ventrally on tail (Fig. 5 C);
stoma with large subventral tooth (Fig. 5 B):
Spinneret on tail terminus; no large tooth or
large tooth dorsal in stoma
Mononehulus
38
Anterior end of body fairly narrow and rounded
(Fig. 10 G; 11 A) 39
Anterior end of body broad and square (Fig. 11 D, G) 41
21
-------
39(38) Stoma distinct, elongate, and cylindrical; weak
cuticle annulations (Fig. 10 G); single ovary
(Fig. 10 F):
Stoma a short broad cylinder or obscure; fairly
strong cuticle annulation; two ovaries
Cy lindfo laimus
40
Fig. 10 - A, head Morihystrella sp. (X 1000); B, female gonad Morihystrella
sp. (X 250); C, head Leptolaimus sp. (X 1000); D, female tail Leptolaimus
sp. (X 250); E, female gonads Leptolaimus sp. (X 500); F, female gonad
Cylindrolaimus sp. (X 250); G, head Cylindrolaimus sp. (X 1000); H, tail
terminus Plectus sp. (X 600); I, female tail Pleotus sp. (X 600); J, head
Pleotus sp. (X 1000).
40(39)
41(38)
Stoma obscure; amphid circular and prominent
(Fig. 11 A):
Stoma a short broad cylinder; amphid unispiral
and prominent (Fig. 11 B) :
Aphanolaimus
Paraphano laimus
42(41)
Amphid multispiral; stoma with large dorsal tooth
(Fig. 11 C): Paraayatholaimus
Amphid stirrup-shaped (Fig. 1 F), circular
(Fig. 1 C) or obscure; dorsal tooth lacking
or inconspicuous 42
Amphid circular (Fig. 11 D); single outstretched
ovary anterior to posterior vulva (Fig. 11 E): Monhystera
Amphid stirrup-shaped or obscure; two ovaries, one
anterior and one posterior to median vulva (Fig.
11 H) 43
22
-------
G
K
Fig. 11 - A, head Aphanolaimus sp. (X 1000); B, head Paraphanolaimus sp.
(X 1000); C, head Paracyatholaimus truncatus (after Cobb, 1914); D, head
Monhystera sp. (X 1000); E, posterior part Morihystera sp. (X 100); F,
head Tobrilus sp. (X 1000); G, head Tripyla sp. (X 1000); H, female
gonads Tripyla sp. (X 100); I, head Teratocephalus sp. (X 1000); J, head
Euteratoeephalus sp. (X 1000); K, head Bastiania exi-lis (X 1000); L,
head Amph-idelus hortensis (X 1000); M, head Alaimus primitivus (X 1000).
23
-------
43(42) Stoma funnel-shaped; amphid stirrup-shaped
(Fig. 11 F): Tobrilus
Stoma obscure, narrow; amphid obscure
(Fig. 11 G): Tripyla
44(26) Six sharply pointed, strongly cuticularized,
incurved lips (Fig. 11 I, J) 45
Lips not pointed, strongly cuticularized, nor
incurved 46
45(44) Cuticle strongly annulated; amphid obscure;
single ovary (Fig. 11 I): Teratocephalus
Cuticle marked only by punctations; amphid
unispiral (Fig. 11 J); two ovaries: Euteratocephalus
46(44) Stoma obscure, slender, weakly cuticularized 47
Stoma not slender or if slender then distinctly
cuticularized 51
47(46) Anterior end of body square; small tooth located
dorsally in stoma (Fig. 11 G): Tripyla
Anterior end of body rounded; no tooth in stoma 48
48(47) Long cephalic setae; distinct cuticular annulation
(Fig. 11 K): Bastiania
No cephalic setae; no cuticular annulation or very
faint annulation % 49
49(48) Enlarged basal portion of esophagus sharply set
off (Fig. 7 H); faint cuticularized ribs
surrounding stoma (Fig. 7 I) : Aulola-imoides
Basal portion of esophagus enlarges gradually
(Fig. 2 K); no faint ribs around stoma 50
50(49) Amphid large and slit-like or crescent-shaped
(Fig. 11 L): Amphidelus
Amphid obscure (Fig. 11 M): Alaimus
51(46) Esophagus cylindrical, lacking basal bulb
(Fig. 2 H, I, J, L) 52
Esophagus with basal bulb (Fig. 2 D; 13 C; 14 A) 54
52(51) Three large, hook-like teeth at anterior end of
stoma (Fig. 12 A); two ovaries, one anterior
and one posterior to median vulva (Fig. 12 B): Ironus
No teeth at anterior end of stoma; single ovary 53
53(52) Terminus of tail bluntly rounded (Fig. 12 C);
circular amphid (Fig. 10 G): Cylindrolaimus
Terminus of tail pointed (Fig. 12 E); amphid
stirrup-shaped (Fig. 12 D): Cryptonohus
24
-------
54(51)
55(54)
Basal bulb of esophagus non-muscular and lacking
valves; median bulb present (Fig. 2 D).Diplogasteridae
[Several genera in this family, in addition to
those included in this key, may occasionally be
found in aquatic habitats. See Goodey (1963) for
illustrations.]
Basal bulb of esophagus muscular and valvate;
median bulb present or absent
55
56
Stoma moderately broad and deep; anterior edge of stoma
bearing rib-like structures; slender tubular section
of stoma extending posterior to large tooth in stoma
(Fig. 4 D); amphipds obscure: Mononahoides
Stoma slender, barrel-shaped or tubular; anterior edge
of stoma without rib-like structures; without tooth
in stoma; large oval amphipds (Fig. 12 F) : Goffart-La
56(54) Amphid large and circular 57
Amphid small and slit-like or pore-like 58
B
Fig. 12 - A, head Ivonus sp. (X 1000); B, female gonads Ivonus sp.
(X 100); C, female tail Cylindrolaimus sp. (X 600); D, head Cryptonchus
sp. (X 1000); E, female tail Cryptonchus sp. (X 500); F, head Goffartia
sp. (X 1000).
57(56) Stoma elongate, nearly cylindrical, and strongly
cuticularized (Fig. 13 A); two ovaries, one anterior
and one posterior to median vulva: OdontoZaimus
Stoma fairly short, funnel-shaped, and moderately
cuticularized (Fig. 10 A); single ovary anterior
to posterior vulva (Fig. 10 B): Morihystrella
25
-------
58(56)
59(58)
Cephalic setae present (Fig. 13 B); esophagus
uniform in thickness anterior to basal
bulb (Fig. 13 C); body slender and tapering
very gradually anteriorly and posteriorly:
Cephalic setae absent; esophagus narrows between
anterior portion and basal bulb (Fig. 2 C;
14 A); body spindle-shaped, considerably
thicker in middle than at anterior and
posterior ends
Chronogaster
59
Stoma elongate, open, cylindrical (Fig. 13 D);
two ovaries, one anterior and one posterior
to median vulva; or single ovary anterior to
posterior vulva; cuticle weakly annulated:
[This family is mainly terrestrial, but
includes several genera which may occasionally
be found in aquatic habitats. See Fig. 13 D
for Rhabditis sp. and Goodey (1963) and Thome
(1961) for illustrations of additional genera.]
Stoma with small widely separated plates at anterior
end followed by a slender moderately cuticular-
ized tube (Fig. 13 E; 14 E); median vulva;
single ovary directed first anterior to vulva
then reflexed and extending posterior to vulva;
cuticle distinctly annulated
Rhabditidae
60
P-
Fig. 13 - A, head Odontolaimus sp. (X 1000); B, head Chronogaster sp.
(X 1000); C, anterior part Chronogaster sp. (X 250); D, head Rhabditis
sp. (X 1000); E, head Acrdbeloides nannus (X 1000) p=probolae.
26
-------
60(59) Three cuticularized plates or probolae extending
beyond lip region (Fig. 13 E); esophagus
broadly expanded in middle (Fig. 14 A); tail
blunt (Fig. 14 B):
No cuticularized plates or probolae extending
beyond lip region; esophagus not expanded in
middle; tail blunt or pointed
Acfobeloides
61
61(60) Tail pointed (Fig. 14 C, D):
Tail blunt (Fig. 14 E, F):
Eucephalobus
Cephalobus
Fig. 14 - A, anterior part Aorobeloides sp. (X 500); B, female tail
Aerobeloid.es nannus (X 600); C, head Euoephalobus oxyuroides (X 1000);
D, female tail Euaephalobus oxyuroides (X 600); E, head Cephalobus sp.
(X 1000); F, female tail Cephalobus sp. (X 500).
27
-------
SECTION III
CLASSIFICATION OF GENERA INCLUDED IN KEY
Phylum NEMATODA
Class ADENOPHOREA
Chromadorida
Araeolaimida
Monhysterida
Desmodorida
Enoplida
Dorylaimida
Chromadorina
Cyatholaimina
Araeolaimina
Monhysterina
Desmodorina
Enoplina
Dorylaimina
Chromadorita
Ethnolaimus
Aohvomadora
Monoohromadora
Paraeyatholaimus
Anaplectus
Anonohus
Aphanolaimus
Bastiania
Ch.Tonoga.ster>
Cylindrolaimus
Euteratoeephalus
Leptolaimus
Paraphanolaimus
Plectus
Teratocephalus
Morihystera
Monhystrella
Odontolaimus
Prodesmodora
Cryptonchus
Ironus
Prismatolaimus
Rhdbdolaimus
Tobrilus
Tripyla
Aulolaimoides
Dorylaimus
Eudorylaimus
Labronema
Laimydorus
Mesodory laimus
Oxydirus
Nygolaimus
Paractinolaimus
Thornia
29
-------
Mononchida
Alaimina
Mononchina
Class SECERNENTEA
Tylenchida
Bathyodontina
Tylenchina
Rhabditida
Aphlenchina
Rhabditina
Alaimus
Amphidelus
Anatonchus
Miconchus
Mononohus
Mylonchulus
Priondhulus
Mononahulus
Atylenahus
Hemicyo Hophora
Hirschmanniella
Tylenchua
Aphelenehoides
Acrobeloides
Butlerius
Cephalobus
Euaephalobus
Goffartia
Mononahoi-des
Rhabditis
30
-------
SECTION IV
ACKNOWLEDGEMENTS
The authors acknowledge the assistance of Dr. S. R. Johnson and Mr. C. A.
Callahan, who prepared many of the sketches adapted from our catalogues
for use in this key; and to Mr. Lu-Hong Wang who inked the sketches.
Previous studies which made possible the preparation of the key were
supported (in part) by National Science Foundation Grant GZ-416, and by
Office of Water Resources Research Project No. A-015-IND (Agreement No.
14-31-0001-3514).
31
-------
SECTION V
REFERENCES
Baker, A. D. 1962. "Check lists of the nematode superfamilies Dorylai-
moidea, Rhabditoidea, Tylenchoidea, and Aphelenchoidea." E. J. Brill
Brill, Leiden, 261 pp.
Cobb, M. V. 1915. Some freshwater nematodes of the Douglas Lake region
of Michigan, U.S.A. (Descriptions by N. A. Cobb). Transactions of
the American Microscopical Society, 34: 21-47.
Cobb, N. A. 1913. New nematode genera found inhabiting freshwater and
non-brackish soils. Journal of the Washington Academy of Sciences,
3: 432-444.
1914. The North American free-living freshwater nematodes. Contri-
butions to a science of nematology, 2. Transactions of the American
Microscopical Society, 33: 69-134.
de Coninck, L. 1965. Systematique des Nematodes. p. 586-681. In P.P.
Grasse" (ed.) Traite de Zoologie. Anatomie, Systematique, Biologie,
4 (2), 731 pp.
Faulkner, L. R. and W. J. Bolander. 1966. Occurrence of large nematode
populations in irrigation canals of south central Washington.
Nematologica, 12: 591-600.
Ferris, V. R., J. M. Ferris and C. A. Callahan. 1972. Nematode community
structure a tool for evaluating water resource environments. Purdue
University Water Resources Research Center, Technical Report, 30:
1-40.
Goodey, T. 1963. "Soil and freshwater nematodes." 2nd ed. revised by
J. B. Goodey, Methuen, London, 544 pp.
Helminthological Abstracts. 1935-1969. Vols 4-38.
Helminthological Abstracts. 1970- . Series A. Animal and Human Helmin-
thology, Series B. Plant Nematology. Vol. 39- .
Mason, E. B. B. and J. E. Bosher. 1963. Combination die for making alumi-
num micro slides. Proceedings of the Helminthological Society of
Washington, 30: 19-20.
Siddiqi, M. R. 1969. Mumtazium rmmtazae n. gen., n. sp. (Nematoda:
Tylencholaimidae) with the proposal of Laimydorus n. gen. (Thomene-
matidae). Nematologica, 15: 234-240.
Southey, J. R. (ed.) 1970. Laboratory methods for work with plant and
soil nematodes. Great Britain Ministry of Agriculture, Fisheries
and Food, Technical Bulletin No. 2, 282 pp.
Tarjan, A. C. 1960. "Check list of plant and soil nematodes." University
of Florida Press, Gainesville, 200 pp.
1967. "Supplement (1961-1965) to the check list of plant and soil
nematodes." University of Florida Press, Gainesville, 115 pp.
Thome, G. 1961. "Principles of nematology." McGraw-Hill Book Co., Inc.,
New York, 553 pp.
Zuckerman, B. M., W. F. Mai and R. A. Rohde. 1971. "Plant Parasitic Nem-
atodes." Vol. 1. Academic Press, New York. 345 pp.
33
-------
SECTION VI
GLOSSARY
amphid one of a pair of organs that open laterally on either side of
the anterior end of the body (Fig. 1).
annulations transverse grooves circling the body externally in the
cuticle at regular intervals (Fig. 14 B-F).
anterior toward the front of the body.
basal bulb enlargement of the esophagus at the posterior end of the
esophagus (Fig. 2 E).
cephalic setae bristle-like, elongate cuticular structures at the
anterior end of the body (Fig. 4 A, B).
clavate club-shaped.
cuticle non-cellular external covering of the body; also lining
certain structures such as the stoma.
denticles small teeth located in stoma (Fig. 6 I).
dorsal -- top side of the nematode body; side of body opposite the side
bearing the anus and vulva.
esophagus muscular tube leading from the stoma to the intestine
(Fig. 2).
filiform very slender and thread-like (Fig 3 A shows a filiform tail).
guiding ring cuticularized ring surrounding the spear (Fig. 7 A, J, L),
lips six (or three) lobes arranged radially around the anterior stomal
opening (Fig. 14 C).
median bulb enlargement of esophagus approximately midway between the
anterior and posterior ends of the esophagus (Fig. 2 A, B).
ovary the reproductive gland of the female, often paired, which pro-
duces the ova (Fig. 10 B, E, F).
papillae minute nipple-like projections of cuticle on surface of body
(Fig. 5 A for papillae on lips).
posterior toward the back of the body or the tail.
35
-------
pimotations small pits or depressions in the cuticle, usually round
(Fig. 8 L, M).
spear a hollow, elongate structure in the stoma used to puncture and
feed on various food sources (Fig. 2 A; 7 A).
spiaules male intromittent organs, usually paired, and extrusible
through the cloacal opening (Fig. 7 K).
spinneret a single duct opening externally on the tail; usually well
cuticularized (Fig. 3 D - F; 5 C).
stoma the mouth cavity anterior to the esophagus.
subventral on either side of the ventral portion of the body.
tail portion of body posterior to the anus (Fig. 3).
tooth pointed cuticular projection of stoma wall (Fig. 4 C, D; 5 A,
B, D).
ventral bottom side of nematode body on which the vulva and anus are
located.
vulva female genital opening (Fig. 10 B, E, F).
36
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SECTION VII
INDEX OF SCIENTIFIC NAMES
Achromadora, 20,29; Fig.1,2,3,8
AoTtibeloides3 27,1,30; Fig.13,14
Adenophorea, 29
Alaimina, 30
Alaimus, 24.,30; Fig.2,11
Amphidelus, 24,30; Fig.11
Anapleatus, 20,29; Fig.1,9
Anatonohus, 24,30; Fig.5
Anonchus, 13*29; Fig.4
Aphanolaimus, 22,29; Fig.2,11
Aphelenchina, 30
Aphelenchoides, 25,30; Fig.2,3,8
Araeolaimida, 29
Araeolaimina, 29
Atylenehus, 25,30; Fig.7
Aulolaimoides, 27,24,29; Fig.7
Bastiania, 24,29; Fig.11
Bathyodontina, 30
brachyuris, Mylonahulus, Fig.6
Butleriue, 22,30; Fig.2,4
Cephalobus, 27,30; Fig.14
Chromadorida, 29
Chromadorina, 29
Chromadorita, 20,29; Fig.8
Chronogaster, 26,29; Fig.13
clarus, Aphelenchoides, Fig.3
Cryptonohus, 24,29; Fig.12
Cyatholaimina, 29
cylindriaus, Tylenchue, Fig.6
Cylindrolaimus, 22,24,29; Fig.
2,10,12
Desmodorida, 29
Desmodorina, 29
Diplogasteridae, 23,25,11
Dorylaimida, 27,1,11,29
Dorylaimina, 29
Dorylaimus, 18,29; Fig.7
elegans, Aulolaimoides, Fig.7
Enoplida, 29
Enoplina, 29
Ethnolaimus, 20,29; Fig.8
Eucephalobus, 27,30; Fig.14
Eudorylaimus, 27,29; Fig.3,7
Eutevatooephalus, 24,29; Fig.11
exiguus, Tylenohus, Fig.8
exilis, Bastiania, Fig.11
Goffartia, 25,1,30; Fig.12
HemiayaHophora, 25,30; Fig.8
Hirschnannia, 12
Hirschmanniella, 25,12,30; Fig.2,8
hortensis, Amphidelus, Fig.11
Ironus, 24,29; Fig.2,12
Labronema, 27,29; Fig.3,7
Laimydorus, 25,12,29; Fig.7
Leptolaimus, 21,29; Fig.2,10
leuakart-i, Chromadorita, Fig.8
meridional-is, Eudorylaimus, Fig. 3,7
Mesodorylaimus, 25,1,29; Fig.3,7
Miconchus, 24,30; Fig.5
Monhystera, 22,1,29; Fig.1,3,11
Monhysterida, 29
Monhysterina, 29
Monhystrella, 20,25,29; Fig.10
Monochromadora, 20,29; Fig.9
Mononchida, 30
Mononchina, 30
Mononchoides, 1Z,25,1,30; Fig.4
MononahuluSf 24,22,30; Fig.5
Mononahus, 24,30; Fig.6
Mylonchulus, 24,30; Fig.6
nannus, Acrobeloides, Fig.13,14
Nematoda, 29
Nygolaimus, 27,29; Fig.6
Odontolaimus, 25,29; Fig.13
oxycephalus, Oxydirus, Fig.7
Oxydirus, 18,29; Fig.7
oxywfoides, Eueephalobus, Fig. 14
papillatus, Mononchus, Fig.6
Paraotinolaimus, 17,29; Fig.7
Paracyatholaimus, 22,29; Fig.11
ParaphanolaimuSj 22,29; Fig.11
Plectus, 22,29; Fig.1,2,10
primitivus, Alainrus, Fig. 11
Prionchulus, 24,30; Fig.6
Prismatolaimus, 13,29; Fig.4
Prodesmodora, 20,29; Fig.1,9
punctatus, Prionchulus, Fig.6
Rhabditida, 30
Rhabditidae, 25,11
Rhabditina, 30
37
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Rhabditis, 30; Fig.2,13
Rhabdolaimus, 20,29; Fig.9
saoohari, Aphelenohoides, Fig.8
Secernentea, 30
similis, Anapleotue, Fig.9
Tevatocephalus, 24,29; Fig.11
thornei, Ldbvonema, Fig.3,7
Thormia, 17,29; Fig.6
Tobrilus, 24,1,29; Fig.1,3,11
trionohuSf Mioonohus, Fig.5
Tripyla, 24,29; Fig.2,11
truncatus, ParaGyccfholaimus, Fig. 11
Tylenchida, 37,1,11,30
Tylenchina, 30
Tylenohue, 18, 1,30; Fig.6,8
vidua, Eemioyoliophova, Fig.8
38
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, BIOTA OF FRESHWATER ECOSYSTEMS IDENTIFICATION MANUAL $ &*;>ortDst
NO. 10 Genera of freshwater nematodes (Nematoda) of eastern c
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Ferris, V. R., Ferris, J. M., Tjepkema, J. P.
Department of Entomology
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