NematologyLaboratoryInvestigations PDF
NematologyLaboratoryInvestigations PDF
NematologyLaboratoryInvestigations PDF
LABORATORY INVESTIGATIONS
Morphology and Taxonomy
JONATHAN D. EISENBACK
ISBN: 1-893961-13-3
NEMATOLOGY
LABORATORY
INVESTIGATIONS
JONATHAN D. EISENBACK
Professor
Department of Plant Pathology, Physiology & Weed Science
Virginia Polytechnic Institute & State University
MACTODE PUBLICATIONS
BLACKSBURG, VA
ISBN: 1-893961-13-3
Copyright © 2003
All rights reserved.
Mactode Publications
3510 Indian Meadow Drive
Blacksburg, VA 24060 USA
This book is dedicated to my loving wife,
Marilyn
and my academic mother and the source of most of these investigations,
Hedwig Hirschmann.
John Halbrendt
is acknowledged for his encouragement
to complete this manual.
Preface
Laboratories are very necessary in introductory plant nematology courses. I believe that
they are extremely valuable to the beginning student and well worth the effort that they
require to teach. Laboratory experiences stimulate interest, arouse curiosity, and confirm
information that is presented in lectures. Nematodes are extremely fascinating, and
working with them in the laboratory is bound to excite all but the most uninitiated.
These investigations on the morphology and taxonomy of nematodes can be taught in one
course or individually, according to the instructor’s preference. Teachers are encouraged
to select the exercises that suit the needs of their students, and students are encouraged
to perform additional laboratories on their own initiative. I hope that these laboratories
will accommodate many courses that are taught in a variety of academic settings. Labs
have been selected to provide a basic introduction to plant nematology, as well as more
advanced techniques.
This first volume of exercises covers the morphology and taxonomy of nematodes. More
investigations that deal with many other aspects of nematology are currently being designed
and tested in the classroom. They will be made available in a second volume.
I thank all who have contributed to this manual. In particular, I would like to thank Dr.
Hedwig Hirschmann. She and her laboratory outlines were the inspiration for many of
these laboratory exercises. In addition, I also thank those who trained me as a nematologist,
including Drs. A. C. Triantaphyllou, Ken Barker, James Baldwin, Joseph Sasser, Donald
Schmitt, and Dick Powell at North Carolina State University. Many of these labs were
taught to me by these outstanding nematologists and teachers.
J.D.E.
Contents
1. Preparation of Temporary and Semi-Permanent Mounts........................................1
2. Gross Morphology of Free-Living Nematodes.......................................................7
3. Gross Morphology of Plant-Parasitic Nematodes.................................................12
4. Cuticular Markings.................................................................................................19
5. Cuticular Layers......................................................................................................24
6. Body Wall...............................................................................................................30
7. Hypodermis and Somatic Muscles........................................................................37
8. Stomas.....................................................................................................................42
9. Secernentean Stomas..............................................................................................47
10. Adenophorean Stomas............................................................................................52
11. Esophagi .................................................................................................................56
12. Secernentean Esophagi ..........................................................................................62
13. Adenophorean Esophagi ........................................................................................67
14. Intestine..................................................................................................................71
15. Posterior Gut ...........................................................................................................74
16. Nervous System and Sensory Structures ................................................................78
17. Secretory/Excretory Systems ..................................................................................82
18. Reproductive Systems .............................................................................................85
19. Female Reproductive Systems.................................................................................88
20. Male Reproductive Systems ....................................................................................93
21. Egg Morphology .....................................................................................................96
22. Preparation of Permanent Mounts............................................................................99
23. The Use of Nematological Keys .............................................................................109
24. Proper Use of the Light Microscope ......................................................................114
25. Preparation of Nematodes for SEM........................................................................119
Appendices
1. Glossary of Nematological Terms.........................................................................124
2. Nematode Chart....................................................................................................135
3. Nematological Journals.........................................................................................139
4. Nematological Societies........................................................................................141
5. Nematological Resources......................................................................................142
6. References to Nematological Keys........................................................................148
7. Keys to Females.....................................................................................................166
8. Keys to Males........................................................................................................177
9. Formulas for Fixatives and Stains ..........................................................................187
Investigation 1
Preparation of Temporary and
Semi-Permanent Mounts
Nematodes are small, colorless roundworms that are most often observed with a microscope.
To be seen clearly, they are mounted on a glass slide and covered with a coverslip. In order
to see all of the minute details, the mounts are examined with an oil immersion objective
lens. Usually, several adult specimens of similar appearance in the dissecting microscope
are selected and mounted for identification to species. The specimens are preserved for a
few hours to indefinitely, and, if their structures are clear and clearly displayed, it may be
possible to make an accurate identification. The condition of the specimen, the selection
of the mounting medium, and the orientation of the nematodes are important because
morphological details are easily obscured by improper preparation techniques. Mounts
may be temporary, semi-permanent, or permanent.
The purpose of this lab is to demonstrate techniques to make temporary and semi-permanent
mounts of nematodes. Temporary mounts are made with the specimens mounted in water.
They are observed immediately because they last only for a few hours or less. Semi-
permanent mounts preserve specimens for a few hours to several days. Permanent mounts
can be observed for many years, but they are more difficult to prepare than temporary and
semi-permanent mounts. (See how to make permanent mounts in Investigation 22.)
Nematode species:
Work Sheet
I. Practice making ringed slides for mounting nematodes.
A. Wipe a pre-cleaned slide with a clean cheesecloth. (Do not clean the slide with alcohol
because the nail polish will not stick to the glass properly.)
D. Gently spin the slide in the ringer and touch the loaded brush to the slide using the
scribed circle as a guide. (Angle the brush so that the nail polish is dragged off the
brush onto the turning slide.)
E. Vary the height of the ring to match the different diameters of various nematodes.
F. Allow the ring to dry for at least 30 minutes in a level slide tray before mounting a
specimen. (Keep it covered to prevent dust from ruining the ring.)
Investigation 1 Preparation of Temporary and Semi-Permanent Mounts 3
_______________________________________________________________________
_______________________________________________________________________
A. Pick 10 specimens of each species with a nematode pick and transfer them into a drop
of water on a heavily ringed slide.
B. Gently kill the nematodes over an alcohol flame. Use just enough heat to straighten the
specimens.
C. Gently place a coverslip on top of the drop, and carefully absorb the excess water with
triangular pieces of filter paper. (Placing the coverslip is the hardest part in making
mounts of nematodes.)
D. Seal the slide with a thick layer of nail polish. (Be sure to use enough so that the water
will not evaporate.)
A. Pick 10-15 specimens with a nematode pick and transfer them into a drop of water on
a heavily ringed slide.
B. Gently kill the nematodes over an alcohol flame. Use just enough heat to straighten the
specimens.
C. Transfer 5-7 of the best specimens to the bottom of a drop of 2% formalin on a ringed
slide.
Investigation 1 Preparation of Temporary and Semi-Permanent Mounts 4
D. Gently place a coverslip on top of the drop and carefully absorb the excess formalin
with triangular pieces of filter paper.
E. Seal the slide with a thick layer of nail polish. (Be sure to use enough so that the
formalin will not evaporate.)
F. Label the slide with the name of the genus and species (if known), the number and sex
of specimens, and the date that the slide was made.
G. Observe the specimens under low and high power on the compound light microscope.
H. Make drawings of all of the details that you can see. The instructor will give general
directions for preparing drawings. In some cases it may be necessary to make composite
drawings to show all of the structures that were seen in various specimens.
2. How can you recognize the position in which your specimens are lying? __________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
A. Collect several soil samples from different habitats. (Keep the samples in closed plastic
bags to prevent drying.)
C. Draw off the funnel 24 hours after processing and store the nematodes in a Syracuse
watch dish in the refrigerator. (Stack the dishes, provide them with a cover to prevent
them from drying out, and label them with your name.)
E. Identify the nematodes to genus. (Later labs will provide the necessary information and
keys for identifying the plant-parasitic nematodes to genus.)
F. Each mount should contain at least 5 adult nematodes of the same genus. Make the
mounts carefully; preparations containing air bubbles are useless because specimens
float, dry out quickly, and cannot be observed with oil immersion. The quality of slides
will be evaluated by the instructor.
Investigation 1 Preparation of Temporary and Semi-Permanent Mounts 6
H. Turn in slides of at least 20 different genera as soon as they are made because the
specimens deteriorate rapidly in semi-permanent mounts.
4. What considerations should be taken into account in the collection and storage of soil
and plant tissues that will be used for nematode extraction? _______________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
6. Will a Baermann funnel work equally well for all nematodes? Why or why not?______
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 2
Gross Morphology of
Free-Living Nematodes
Free-living nematodes are common inhabitants of the soil environment. Millions of
individuals can live in the top 20 centimeters of one square meter of soil; they feed on
bacteria, fungi, and dead organic matter. The morphology of the body wall, as well as
the internal anatomy, is easily observed in whole specimens, and even live specimens can
be examined. In order to see the minute details of nematode morphology, it is necessary
to obtain specimens in good physiological condition, to prepare them with a suitable
technique for observation, to adjust the microscope for maximum resolution, to compare
the morphology of several specimens, and to look and look and look.
Nematode species:
Work Sheet
I. Make semi-permanent slides of females and males of Rhabditis sp.
A. Pick 10-15 females and 10-15 males of Rhabditis sp. and transfer them into a drop of
water on a ringed slide.
B. Kill the nematodes gently over the alcohol flame. Use just enough heat to straighten
the specimens.
C. Transfer 5-7 of the best specimens to the bottom of a drop of 2% formalin on a ringed
slide.
D. Gently place a coverslip on top of the drop and carefully absorb the excess formalin
with triangular pieces of filter paper. (Placing the coverslip is the hardest part in making
mounts of nematodes.)
E. Seal the slide with a thick layer of nail polish. (Be sure to use enough so that the forma-
lin will not evaporate.)
1. What are the gross differences between female and male nematodes?______________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 2 Morphology of Free-Living Nematodes 9
II. Study the gross morphology of the Rhabditis females and males.
A. Study the gross morphology of female and male nematodes under low power. Compare
and contrast the differences between the sexes. (Use low power magnification.)
B. Study the gross morphology of the anterior end of the female and the tails of the female
and male. (Use high power magnification.)
C. Make drawings of all of the details that you can see. The instructor may give additional
directions for preparing drawings. In some cases, it may be necessary to make composite
drawings to show all of the structures that are visible. Draw just what you see! Do not
copy the drawing in this investigation or from other sources. The instructor can ensure
that you are correctly seeing and interpreting the specimens by looking at your drawings.
Also, please make your drawings large enough to include all of the minuscule details.
2. Based on the morphology that you observed, do you think that Rhabditis is a plant
Why? __________________________________________________________________
________________________________________________________________________
3. Compare and contrast the differences between the anterior ends of the female
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 2 Morphology of Free-Living Nematodes 10
Fig. 2.1 Gross morphology of a typical soil-borne free-living nematode, female and male
(after Hirschmann).
Investigation 2 Morphology of Free-Living Nematodes 11
4. Compare and contrast the differences between the tails of the female and male. _____
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 3
Gross Morphology
of Plant-Parasitic Nematodes
All plant-parasitic nematodes possess a protusible hypodermic needle-like stylet that
they use to feed on plants. It is usually hollow with few exceptions (Trichodorus and
Paratrichodorus). Most plant-parasitic nematodes are vermiform for their entire life, but
juveniles and females of some species become swollen after they begin feeding on plant
tissues. These saccate females are sedentary parasites, and they sacrifice their mobility to
increase their reproductive capacity. Males of these species remain vermiform and, in most
instances, do not feed as adults.
Usually, the esophagus of plant-parasitic nematodes is less muscular than that of the free-
living nematodes. Often the pumping portion is limited to the median bulb; however, the
esophageal glands are usually very prominent in plant-parasitic nematodes because they
produce the secretions that are necessary for establishing and maintaining the parasitic
relationship. (You may want to compare the morphology of plant-parasitic nematodes with
that of free-living nematodes in Investigation 2.)
Nematode species:
Work Sheet
I. Make semi-permanent slides of females and males of Tylenchorhynchus sp.
(Tylenchorhynchus sp. females and males are representatives of the vermiform
type.)
A. Pick up 20 females and 20 males with the pulp canal file and transfer them to a drop of
water on ringed slides.
D. Transfer each of 6-8 females or males to a drop of 2% formalin on ringed slides and
make them into semi-permanent mounts. Prepare 2-3 slides. Seal the mounts with a
thick ring of nail polish.
E. Study the gross morphology of the Tylenchorhynchus sp. females and males.
F. Make drawings of all of the details that you can see. The instructor will give general
directions for preparing drawings. In some cases, it may be necessary to make composite
drawings to show all of the structures that are visible. Draw just what you see! Do
not copy the drawing in this investigation or from other sources. The instructor can
insure that you are correctly seeing and interpreting the specimens by looking at your
drawings. Also, make your drawings large enough so that you can include all of the
minuscule details.
Investigation 3 Gross Morphology of Plant-Parasitic Nematodes 14
1. Describe the morphology of the stylet of Tylenchorhynchus sp. females and males.
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
2. How many parts of the esophagus can you see in Tylenchorhynchus sp. ? _________
________________________________________________________________________
________________________________________________________________________
3. Where are the openings of the esophageal glands located in the esophagus of
________________________________________________________________________
4. Compare and contrast the tails of females and males of Tylenchorhynchus sp. ______
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 3 Gross Morphology of Plant-Parasitic Nematodes 15
Fig. 3.1 Gross morphology of a typical vermiform plant-parasitic nematode, female and
male (after Hirschmann).
Investigation 3 Gross Morphology of Plant-Parasitic Nematodes 16
III. Examine the morphology of females and males of Meloidogyne sp. (Meloidogyne
sp. females are representatives of the saccate type of plant-parasitic nematodes.)
A. Examine the provided root material infected with Meloidogyne sp. Observe the galled
appearance of the roots and the brownish egg masses on the gall surface.
B. Tease out some Meloidogyne sp. females present in the galls using dissecting needles.
Transfer 10 females with a pipette to a BPI dish containing 2% formalin.
C. Pick out 3-4 Meloidogyne sp. males and transfer them to a drop of water on a slide. Kill
them by gentle heat and make a semi-permanent formalin mount.
D. Examine the females and males at low power on the microscope and make outline
drawings of them.
E. Make drawings of all of the details that you can see. The instructor will give general
directions for preparing drawings. In some cases, it may be necessary to make composite
drawings to show all of the structures that are visible. Draw just what you see! Do
not copy the drawing in this investigation or from other sources. The instructor can
insure that you are correctly seeing and interpreting the specimens by looking at your
drawings. Also, make your drawings large enough so that you can include all of the
minuscule details.
B. Observe the sexual dimorphism in Meloidogyne: Females are pyriform, males are
fusiform.
5. What is the significance of the swelling of the females of Meloidogyne sp. ? ________
________________________________________________________________________
________________________________________________________________________
6. Compare and contrast the body shape of females and males of Meloidogyne sp. ____
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 3 Gross Morphology of Plant-Parasitic Nematodes 17
7. Explain the trade-off between the loss of mobility and the increase in body size in
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 4
Cuticular Markings
Nematodes are decorated with various types of cuticular markings that are useful for
identification. These markings are either ornamental or sensory; however, ornamental
does not imply lack of function. Cuticular markings are generally external, although a
few markings occur below the surface, and may be considered as important morphological
characters that are useful for the description and subsequent identification of nematode
species. These markings are sometimes difficult to see and can be enhanced with proper
lighting techniques on the microscope.
For some characters, there is a progression from one state to the next. For example,
transverse markings vary from none, to slightly indented, to moderately deep, to very deep.
These different character states are not clearly defined, nor are they distinctly separated
from each other. Therefore, much experience may be necessary to acquire the ability to
accurately identify a particular character state.
Nematode species:
Work Sheet
I. Examine the progression of transverse markings from striae to apparent
segmentations and make drawings of the following:
D. Modification of annulation: Heterodera sp. Make a sketch of the zig-zag pattern of the
cyst wall. Also notice the punctations in the inner layer of the cyst wall.
Fig. 4.1. Progression of transverse body markings from striae to apparent segmentation
and other variations. A. Striae in Pratylenchus sp. (after Corbet) B. Annulation
in Hoplolaimus sp. (after Sher) C. Apparent segmentation in Mesocriconema sp.
(after Orton Williams) D. Zig-zag like pattern in the body wall of cyst nematodes,
Globodera and Heterodera spp. (after Hirschmann)
Investigation 4 Cuticular Markings 21
1. Explain the differences between striation, annulation, and apparent segmentation. ___
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
II. Study the various types of longitudinal markings and make drawings of the
following:
A. Study the longitudinal ridges and annules that combine to form tesselation in
Tylenchorhynchus sp.
C. Draw the tail of a Hoplolaimus sp. male and note the difference between the morphology
of the bursa (caudal alae) of Hoplolaimus sp. and Rhabditis sp.
2. Compare and contrast the morphology of the mail tail of Hoplolaimus and Rhabditis
spp. ___________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 4 Cuticular Markings 22
Fig. 4.2 Various types of longitudinal markings. A. Tesselation and four lateral lines in
Tylenchorhynchus sp. B. Ten lateral incisures in Aphelenchus sp. (after Goodey &
Hooper) C. Cross-section of the body wall of Hoplolaimus sp. D. Cervical alae of
Oesophagostomum sp. E. Tail of Rhabditis sp. (lateral view) F. Tail of Hoplolaimus
sp. (lateral view) (C-F After Hirschmann)
Investigation 4 Cuticular Markings 23
III. Observe and draw the punctations in the cuticle of a marine nematode.
IV. Examine the lips and their modifications and make drawings of the following:
A. Study the lips and their papillae of Rhabditis sp. in side view.
B. Note the labial probolae which are modifications of the lips in Acrobeles sp.
Fig. 4.3 Lips and their modifications. A. Lips of Rhabditis sp. (after Hirschmann) B.
Probalae of Acrobeles sp. (after Thorne) C. Heavy cephalic framework of Hoplolaimus
sp. (after Sher) D. Moderate framework of Pratylenchus sp. (after Corbett)
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 5
Cuticular Layering
The cuticle of most nematodes is often arranged into four fundamental layers: epicuticle,
exocuticle, mesocuticle, and endocuticle. Both the external and internal cuticular
structures are important taxonomic characters. In the root-knot nematodes, Meloidogyne
spp., the cuticle surrounding the vulva and anus forms a unique fingerprint-like pattern that
is useful for the identification of the four most common species. Preparing these patterns
for observation is difficult and tedious but not impossible.
Nematode species:
Work Sheet
I. Examine the cuticular layering of an animal-parasitic nematode (Ascaris
lumbricoides).
B. Make drawings of a close-up view of a transverse section of the cuticle (use oil
immersion).
Fig. 5.1 Transverse section of cuticle of Ascaris lumbricoides (After Hirschmann, 1960).
A. Transfer several brown cysts to a Syracuse watch glass filled with 2% formalin.
B. Select individual brown cysts and make thin transverse sections of the cyst wall with
the eye knife in a drop of lactophenol on a plastic dish.
D. Make a drawing of a close-up view of a transverse section of the cuticle (use oil
immersion).
Investigation 5 Cuticular Layering 26
A. Select galls with mature females and place in Syracuse dish with tap water. (Single
galls are preferable to compound galls.) Pick out 6-8 adult females by teasing the root
tissue apart with forceps and half spear.
B. Cut off the neck of the female with the eye knife and gently push out the body contents
with the forceps.
C. Place the cuticle in a drop of 45% lactic acid in a plastic Petri dish. Collect 6 to 8
cuticles in the drop and let them stand for 15 - 30 minutes before cutting.
Investigation 5 Cuticular Layering 27
D. Cut the cuticle in half (equatorially) with an eye knife in a drop of 45% lactic acid.
E. Trim the cuticle with the perineal pattern so that it is square. Cut 6 to 8 patterns. (Cut
only on the surface of a plastic Petri dish to prevent dulling the cutting edge.)
F. Thoroughly clean debris from the perineal pattern with a nematode pick.
G. Transfer the perineal pattern with a nematode pick to a drop of glycerin on a clean
glass microscope slide. Align the perineal patterns so that they are in a straight line
and the anus is oriented down. Place the interior surface of the cuticle against the
glass and press the pattern gently against the glass with a nematode pick.
H. Gently place the coverslip in a drop of glycerin. Absorb excess glycerin with a small
triangular piece of filter paper, or additional glycerin can be added by placing a small
drop on the edge of the coverslip.
I. Seal the coverslip with nail polish and label the slide.
I
Investigation 5 Cuticular Layering 28
IV. Observe and draw the perineal patterns of the four most common Meloidogyne
spp. (Fig. 5.2)
Fig. 5.2 Perineal patterns of the four most common Meloidogyne species. A. M. incognita
B. M. javanica C. M. arenaria D. M. hapla (after Eisenback)
Investigation 5 Cuticular Layering 29
1. Describe the major identifying characters for each of the four common species of
Meloidogyne.
M. incognita _________________________________________________________
_____________________________________________________________________
_____________________________________________________________________
_____________________________________________________________________
M. javanica __________________________________________________________
_____________________________________________________________________
_____________________________________________________________________
_____________________________________________________________________
M. arenaria __________________________________________________________
_____________________________________________________________________
_____________________________________________________________________
_____________________________________________________________________
M. hapla ____________________________________________________________
_____________________________________________________________________
_____________________________________________________________________
_____________________________________________________________________
Investigation 6
Body Wall
Nematodes are multicellular animals (metazoans) with true tissues (umetazoans) that have
a true mesoderm with a persistent blastocoel (= pseudocoel) between the gut and body
wall. They are triploblastic, having three germcell layers including ectoderm (cuticle and
hypodermis), mesoderm (somatic muscles), and endoderm (intestine). The body plan of
nematodes consists of a tube within a tube. The body wall forms the outer tube, and the
digestive system forms the inner tube. Animals with a similar body plan include other
psuedocoelmates such as the Nematomorpha (horsehair worms), Entoprocta (entoprocts),
Rotifera (rotifers), Gastrotricha (gastrotrichs), and Kinorhyncha (kinorhynchs).
Nematode species:
Immersion oil
Work Sheet
I. Examine cross-sections of Ascaris lumbricoides with a microscope, studying the
body wall and internal organs.
A. Make a sketch of the esophagus cross-section of Ascaris. Pay particular attention to the
well-developed hypodermis.
B. Study the gross morphology of the cross-section of the Ascaris lumbricoides female and
draw a general view under the low power of the microscope.
C. Draw close-up views of the following using the high power of the microscope:
1. Two or three of the coelomyarian muscle cells.
2. Section of the intestinal wall.
3. Growth zone of the ovary.
4. Uterus with eggs.
D. Study the gross morphology of the cross-section of the Ascaris lumbricoides male and
draw a general view under the low power of the microscope.
1. How many somatic muscle cells are contained in one cross-section of Ascaris
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 6 Body Wall 32
Fig. 6.1 Cross-section of Ascaris lumbricoides body wall at the level of the esophagus
(after Hirschmann).
3. Why are the subdorsal somatic muscle sectors innervated by the dorsal nerve cord
and the subventral somatic muscle sectors innervated by the ventral nerve chord? ______
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 6 Body Wall 33
Fig. 6.2 Cross-section of Ascaris lumbricoides female body wall (after Hirschmann).
6. Why are there so many sections cut through the ovary in the cross section of the body
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 6 Body Wall 34
Fig. 6.3 Cross-section of Ascaris lumbricoides male body wall (after Hirschmann).
7. Compare and contrast the cross-section of the body wall of the female and male
________________________________________________________________________
_______________________________________________________________________
________________________________________________________________________
_______________________________________________________________________
Investigation 6 Body Wall 35
Fig. 6.4 Cross-sections of Ascaris lumbricoides body wall, ovary, uterus, and testis (after
Hirschmann).
8. How many sublayers can you discern in the cross-section of Ascaris lumbricoides?___
________________________________________________________________________
9. What is the function of the bacillary layer of the intestine in Ascaris lumbricoides ? _
________________________________________________________________________
________________________________________________________________________
Investigation 6 Body Wall 36
10. What is the purpose of the muscle layer in the uterus of Ascaris lumbricoides ?_____
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 7
Hypodermis and
Somatic Muscles
The hypodermis is a group of cells or a syncytium that secretes the cuticle. In addition, it
contains the nerve chords, canals of the excretory/secretory system, and sensory structures.
The hypodermis lies between the cuticle and the somatic muscles. It usually forms four
longitudinal chords that divide the somatic muscles into four sections. Members of
the nematode class Adenophorea are characterized by the occurrence of glands in the
hypodermis, whereas they are absent in Secernentea.
Nematode species:
Work Sheet
I. Observe the inflated cuticle of Paratrichodorus sp. and make an outline sketch of
the whole nematode.
II. Note the cuticular sheath of Hemicycliophora sp. and draw the tail region where
the sheath is distinct.
1. Compare and contrast the inflated cuticle of Paratrichodorus with the cuticular
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 7 Hypodermis and Somatic Muscles 39
III. Draw the caudal glands and the spinneret of Mononchus sp.
2. What is the function of the caudal glands and spinneret in Mononchus sp.? _______
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
belong? _________________________________________________________________
IV. Observe the thick hypodermis with numerous nuclei (stained with acid fuchsin-
lactophenol) in the fourth-molt female of Heterodera lespedezae.
4. Why does the hypodermis become thicker during the molting process? ____________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
V. Stain males of Rhabditis sp. with 1% propionic orcein and observe the nuclei in
the ventral hypodermal chord. Note also the muscle nuclei. Specimens should be
in lateral position.
A. Procedure: Kill specimens in water by gentle heat. Transfer them to a drop of orcein
on a ringed slide. Put coverslip and heat 2-3 seconds over alcohol flame.
B. Remove excess stain.
C. Seal mount with nail polish.
Investigation 7 Hypodermis and Somatic Muscles 40
6. Why is it necessary for the specimen to lie in a lateral position in order to see the
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
VI. Note the accumulation of glycogen (stained purple) in the noncontractile part of
the muscles of Ascaris sp.
8. Why does glycogen accumulate in the noncontractile portion of the somatic muscles?
________________________________________________________________________
________________________________________________________________________
Investigation 7 Hypodermis and Somatic Muscles 41
Fig. 8.2 Transverse section of muscle cell of Ascaris lumbricoides. A. Section through
somatic muscles and hypodermal chord B. Fine structure of contractile region
of muscle cell C. Enlargement of one ridge showing the arrangement of the
myofilaments (after Rosenbluth)
Investigation 8
Stomas
The stoma or buccal cavity of nematodes is extremely variable in morphology and size, and
thus very valuable for systematics and taxonomy. The structure of the stoma provides much
information about the feeding habit and mode of parasitism in the case of plant-parasitic
nematodes. All plant-parasitic nematodes are armed with a protusible hypodermic needle-
like stylet. The stylet enables the nematode to become a plant parasite. Not all nematodes
that possess a stylet are plant parasites; however, all plant-parasitic nematodes have a stylet.
Nematodes with very long stylets are usually deep feeding ectoparasites; those with short
stylets are either endoparasites that migrate into plant tissue in order to feed, or they may
be shallow feeding ectoparasites.
Nematode species:
Work Sheet
I. Study the different stoma types and make drawings of each:
A. Adenophorea:
3. Dorylaimus sp. Provided with a protrusible odontostyle which may have originated
from a subventral tooth (not hollow-Nygolaimus) but through further development
became axial and hollow. The odontostyle is formed from a single cell in the wall of
the esophagus during postembryogenesis.
B. Secernentea:
1. Rhabditis sp. Stoma shows division into cheilostom, protostom, and telostom.
3. Pratylenchus sp. The meta- and telo-rhabdions fuse to form a protrusible stylet
(stomatostyle). The metarhadbions give rise to the conical part of the stylet, the
telorhabdions to stylet shaft and knobs. The cheilo-rhabdions, together with the pro-
meso-rhabdions, provide the guiding apparatus. The stomatostyle is formed from
myoepithelial cells of the anterior esophagus in situ during postembryogenesis.
1. What is the purpose of the dorsal tooth in the stoma of Mononchus spp.? __________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
2. Compare and contrast the function of the protusible spear in Dorylaimus spp. ______
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 8 Stomas 46
Fig. 8.3 Scanning electron micrographs of excised stylets of various nematodes including
mainly plant parasites (after Eisenback and Rammah).
3. Compare and contrast the stylets of Pratylenchus spp. and Belonolaimus spp. ______
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
4. Compare and contrast the odontostyle of Xiphinema spp. with the stomatostyle of
Belonolaimus spp._________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 9
Secernentean Stomas
The stoma or buccal cavity of Secernentean nematodes provides valuable information
about the feeding habits and modes of parasitism. Some of the free-living and predatory
members of the class have cylindrical stomas, globular bucal cavities which may be armed
with various kinds of teeth, or a combination of the two forming a globular anterior portion
with a cylindrical posterior.
Most of the plant-parasitic nematodes belong to this class and are armed with a unique
protusible, hollow, hypodermic needle-like stylet. The stylet enables the nematode to
establish a host-parasite relationship with the plant. Nematodes with very long stylets are
deep-feeding ectoparasites, and those with short stylets are often endoparasites that migrate
into plant tissue in order to feed, or they feed at the surface on root hairs and epidermal
cells.
Nematode species:
Work Sheet
I. Study the different stoma types in Secernentea and make drawings.
A. Rhabditis sp. Stoma shows division into cheilostom, protostom, and telostom.
B. Acrobeloides sp. Stoma narrow. The protostom is further subdivided into pro-,
meso-, and metastom.
D. Diplogaster sp. Stoma wide. The metastom bears one or two large teeth, further
developmental stages of the minute teeth found on the metastom in Rhabditis and
Panagrolaimus.
E. Pratylenchus sp. The meta- and telo-rhabdions fuse to form a protrusible stylet
(stomatostyle). The meta-rhadbions give rise to the conical part of the stylet, the
telo-rhabdions to stylet shaft and knobs. The cheilo-rhabdions together with the
promeso-rhabdions provide the guiding apparatus. The stomatostyle is formed from
myo-epithelial cells of the anterior esophagus in situ during postembryogenesis.
G. Ditylenchus sp. The stylet is short and delicate with three small stylet knobs.
Fig. 9.1 Different stoma types in the nematode class Secernentea (after Hirschmann).
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
_____________________________________________________________
_____________________________________________________________
_____________________________________________________________
Investigation 9 Secernentean Stomas 50
Fig. 9.2 Cephalic region of a male of soybean cyst nematode, Heterodera glycines (after
Hirschmann).
3. Compare and contrast the style of Aphelenchus spp. with that of Ditylenchus spp. ___
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
4. Compare and contrast the stylet of Hoplolaimus spp. with that of Belonolaimus spp. _
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 9 Secernentean Stomas 51
Fig. 9.3 Scanning electron micrographs of excised stylets of various genera, species, and
life-stages of nematodes.
5. Based on morphology of the stoma, what type of feeding behavior does Acrobeloides
________________________________________________________________________
7. What is the function of the radial blades and basal ring of the cephalic framework of a
8. Where does the stylet opening occur in the Secernentean nematodes? _____________
________________________________________________________________________
Investigation 10
Adenophorean Stomas
The stoma or buccal cavity of Adenophorean nematodes is extremely variable in morphology
and size, and thus very valuable for systematics and taxonomy. The structure of the stoma
provides information about the feeding habit and mode of parasitism. Most of the members
of this class are free-living, predators, or animal parasites. A few of them are plant parasites
which are armed with a protusible hypodermic needle-like stylet. Nematodes with very
long stylets are deep-feeding ectoparasites, and those with short stylets are shallow-feeding
ectoparasites.
Nematode species:
Work Sheet
I. Study the following types of stoma types of Adenophorea and make drawings:
C. Dorylaimus sp. Provided with a protrusible odontostyle which may have evolved from
a subventral tooth (not hollow in Nygolaimus sp.) but through further development
became axial and hollow. The odontostyle is formed from a single cell in the wall of the
esophagus during post embryogenesis.
1. Compare and contrast the stoma of Plectus spp. and Mononchus spp. _____________
________________________________________________________________________
________________________________________________________________________
Investigation 10 Adenophorea Stomas 55
2. Compare and contrast the stylet of Dorylaimus spp. and Xiphinema. spp. __________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
3. Compare and contrast the stylet of Xipinema spp. with that of Longidorus spp.______
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 11
Esophagi
The esophagi of nematodes are diverse structures and useful for identification. Basically
a pumping organ, the esophagus is composed of muscles and gland cells that surround a
triradiate lumen lining with 3-5 interspersed esophageal gland cells. A one-
part esophagus consists of a muscular cylinder with several gland cells contained
within. A two-part esophagus consists of a narrow muscular anterior portion and a
swollen, glandular basal portion. A four-part esophagus contains an anterior corpus, a
swollen metacorpus or median bulb, a narrow isthmus, and an enlarged glandular posterior
bulb. In a three-part esophagus the procorpus and metacorpus are fused together. Specialized
structures for grinding food particles may be found in the basal bulb. The basal gland lobe
may join with the intestine in a line perpendicular to the body or it may overlap it dorsally
or ventrally. An esophageal-intestinal valve connects the esophagus with the intestine. It
is triangular and is often surrounded by a circular sphincter muscle which prevents food
flowing down the esophagus into the intestine from backing up.
Nematode species:
Work Sheet
I. Study the following types of esophagi of Adenophorea and make drawings.
A. Mononchus sp. Cylindrical type. Five uninucleate esophageal glands: one dorsal, 4
subventral. Esophago-intestinal valve triradiate and massive.
C. Plectus sp. Plectoid type. Esophagus consisting of cylindrical corpus and pyriform
bulbar region containing a triradiate, denticulate valve apparatus. Three uninucleate
esophageal glands: 1 dorsal, 2 subventral. Esophago-intestinal valve elongate, conoid.
II. Study the following types of Secerenentean esophagi and make drawings.
A. Rhabditis sp. Rhabditoid type. Esophagus consisting of corpus (which is divided into
cylindrical procorpus and enlarged, distinctly set-off metacorpus),
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 11 Esophagi 60
Fig. 11.2 Nematode esophagi types. A. Cylindrical type (Mononchus sp.) B. Dorylaimoid
type (Dorylaimus sp.) C. Bulboid type (Plectus sp.) D. Rhabditoid type (Rhabditis
sp.) E. Diplogasteroid type (Diplogaster sp.) F. Tylenchoid type (Tylenchorhynchus
sp.) G. Tylenchoid type (Helicotylenchus sp.) H. Tylenchoid type (Neotylenchus sp.)
I. Criconematoid type (Mesocriconema sp.) J. Aphelenchoid type (Aphelenchus sp.)
(A-J after Hirschmann)
Investigation 11 Esophagi 61
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
10. Compare and contrast the various types of tylenchoid esophagi. ________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 12
Secernentean Esophagi
Nematodes within the class Secernentea have various types of esophagi. The major types
include those with one-part that consists of a muscular cylinder with several gland cells con-
tained within; a two-part esophagus that consists of a narrow muscular anterior portion and a
swollen, glandular basal portion; and a four-part esophagus that contains an anterior corpus, a
swollen metacorpus or median bulb, a narrow isthmus, and an enlarged glandular pos-
terior bulb. In a three-part esophagus, the procorpus and metacorpus are fused together.
Specialized structures for grinding food particles may be found in the basal bulb. The
basal gland lobe may join with the intestine in a line perpendicular to the body or it may
overlap it dorsally or ventrally. An esophageal-intestinal valve connects the esophagus
with the intestine. It is triangular and is often surrounded by a circular sphincter mus-
cle which prevents food flowing down the esophagus into the intestine from backing up.
Nematode species:
Work Sheet
I. Study the different types of esophagi in the following Secernentea and make drawings:
1. Compare and contrast the rhabditoid and diplogasteroid types of esophagi. ________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
2. Compare and contrast the diplogasteroid and tylenchoid types of esophagi. _________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 12 Secernentean Esophagi 65
3. Compare and contrast the tylenchoid and aphelenchoid types of esophagi. _________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
4. Compare and contrast the the tylenchoid and criconematoid types of esophagi. _____
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 12 Secernentean Esophagi 66
Fig. 12.2 Nematode esophagi types. A. Rhabditoid type (Rhabditis sp.) B. Dip-
logasteroid type (Diplogaster sp.) C. Tylenchoid type (Tylenchorhynchus sp.)
D. Tylenchoid type (Helicotylenchus sp.) E. Criconematoid type (Mesocrico-
nema sp.) F. Aphelenchoid type (Aphelenchus sp.) (A-F after Hirschmann)
5. Describe some of the variations within the tylenchoid type of esophagi. ____________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
6. What is the purpose of the denticulate basal pump in the rhabditoid type of
esophagus? ______________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 13
Adenophorean Esophagi
The esophagi of nematodes are diverse structures and useful for identification. Basically
a pumping organ, the esophagus is composed of muscles and gland cells that surround a
triradiate lumen lining with 3-5 interspersed esophageal gland cells. A one-part
esophagus consists of a muscular cylinder with several gland cells contained
within. A two-part esophagus consists of a narrow muscular anterior portion and a
swollen, glandular basal portion. A four-part esophagus contains an anterior corpus, a
swollen metacorpus or median bulb, a narrow isthmus, and an enlarged glandular posterior
bulb. In a three-part esophagus, the procorpus and metacorpus are fused together.
Specialized structures for grinding food particles may be found in the basal bulb. The
basal gland lobe may join with the intestine in a line perpendicular to the body, or it may
overlap it dorsally or ventrally. An esophageal-intestinal valve connects the esophagus
with the intestine. It is triangular and is often surrounded by a circular sphincter muscle
which prevents food, flowing down the esophagus into the intestine, from backing up.
Nematode species:
Work Sheet
I. Study the following types of esophagi of Adenophorea and make drawings:
E. Prodesmodora sp. Bulboid type. Esophagus divisible into cylindrical corpus and
pyriform bulbar region containing 3 cuticularized crescentic valve elements. Three
uninucleate esophageal glands: one dorsal, 2 subventral. Esophago-intestinal valve
short and flattened.
F. Plectus sp. Plectoid type. Esophagus consisting of cylindrical corpus and pyriform
bulbar region containing a triradiate, denticulate valve apparatus. Three uninucleate
esophageal glands: 1 dorsal, 2 subventral. Esophago-intestinal valve elongate, conoid.
Investigation 13 Adenophorea Esophagi 69
1. Compare and contrast the mononchoid and dorylaimoid types of esophagi. _________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
2. Compare and contrast the dorylaimoid and bulboid types of esophagi. ____________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 13 Adenophorea Esophagi 70
3. What is the purpose of the tooth in the mononchoid type of esophagus? ___________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
4. Where are the esophageal gland cells in each type esophagus examined? __________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 14
Intestine
The gross morphology of the intestine of various groups of nematodes is remarkably similar.
Generally, the intestine is a simple tube that is formed from a single layer of epithelial cells
and fills the body space between the esophagus and the posterior gut. Several plant-parasitic
nematodes may have an anterior caecum that extends beyond the base of the esophagus,
almost to the level of the nerve ring. A posterior intestinal caecum is extremely rare.
The single layer of epithelial cells often have an inner bacillary layer that greatly increases
the surface area of the lumen. This layer varies from having large, well-defined microvilli
to ones that are fine and compact. The total number of cells and their shapes are responsible
for the overall appearance of the intestine and its lumen. Oligocytous intestines contain up
to 128 longitudinally elongate to rectangular cells. Intestines that have 20-50 hexagonal
cells in circumference are polycytous; they are myriocytous if they are more than 100
cuboidal cells in circumference. Many plant-parasitic nematodes have an intestine which
is a synticium where cell walls are lacking, the lumen is reduced in form or absent, and the
microvilli are less compact or absent.
Nematode species:
Work Sheet
I. Study the intestine of Rhabditis sp., Mononchus sp., Hoplolaimus sp., Belonolaimus
sp., and Meloidogyne males.
A. Note the presence of distinct cells in Rhabditis sp. (oligocytous intestine) and Mononchus
sp. (polycytous intestine), whereas cells are absent in Hoplolaimus sp. and Belonolaimus
sp.
B. Draw and label the sphaerocrystals of rhabditin in the intestinal cells of Rhabditis sp.
and the fat globules in the intestine of Hoplolaimus sp.
D. Observe the anterior caecum in the male root-knot nematode, Meloidogyne sp.
1. How may cells make up the oligocytous intestine seen in Rhabitis sp.?______________
2. How may cells form the polycytous intestine in Mononchus sp.? __________________
________________________________________________________________________
Investigation 14 Intestine 73
Fig. 15.1 Nematode intestine. A-D. Diagrams of the cross-section through Ascaris suum
showing a progessinve increase in magnification (after Bird and Bird) E. Drawing of
a cross-section through the four cells forming the circumference of Trichinella spiralis
(after Bruce)
________________________________________________________________________
5. What is the purpose of the anterior caecum in the male root-knot nematodes? _______
________________________________________________________________________
________________________________________________________________________
Investigation 15
Posterior Gut
The posterior gut of female nematodes is a simple tube that connects the intestine to the
anus. It is lined with cuticle and often has several rectal glands opening into it. The posterior
intestinal cells form a simple intestino-rectal valve that is surrounded by a single sphincter
muscle cell which keeps the valve closed. During defecation, this sphincter muscle relaxes
and allows fecal matter to flow out of the intestine through the anal opening.
In the male, however, the intestine and reproductive system open into a common duct, the
cloaca. The spicules and other accessory copulatory structures are contained within the
cloaca.
2. Draw and label the major parts of the posterior gut and ac-
cessory structures of various male nematodes.
Nematode species:
Work Sheet
I. Examine the rectum and the rectal glands of young females of Rhabditis sp.
II. Observe the 6 large rectal glands in young females of Meloidogyne sp. that have
been stained with acid fuchsin lactophenol.
E. Hoplolaimus sp., spicules free (tylenchoid type), gubernaculum with titillae, capitulum
on ventral cloacal wall.
1. What is the purpose of the rectal glands in females of Rhabditis spp.? _____________
________________________________________________________________________
________________________________________________________________________
2. What is the purpose of the rectal glands in females of Meloidogyne spp.? __________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 15 Posterior Gut 77
Fig. 15.2 Posterior gut. A. Female with anus and rectum B. Male with spicule and cloaca
(after Caveness)
3. Discuss the importance of the six rectal glands in root-knot females. ______________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
5. Compare and contrast the various types of male accessory sructures. _____________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 16
Nervous System and
Sensory Structures
Observation of the nervous system of nematodes is usually limited to the large nerve ring
and associated ganglia that surround the esophagus in the anterior end of the body and
various sensory structures.
Sensory structures are common to all nematodes. Some of them, such as the amphids,
occur in all groups of nematodes, whereas the phasmids are present in members of the class
Secernentea but absent in the class Adenophorea. Nematodes may possess eye spots, touch
receptors, and chemoreceptors in the form of papillae, sensilla, setae, bristles, pores, warts,
and other manifestations. The occurrence and form of many of these sensory structures
are important taxonomic characters. Many, however, are difficult to observe. Patience
and careful observations with constant focusing are often necessary in order to see many
of these structures.
Nematode species:
Work Sheet
I. Observe and draw the nerve ring and associated ganglia.
A. Mononchus sp. Also note the papillary nerves in the lip region.
B. Helicotylenchus dihystera. Stain specimens with propionic orcein. Nuclei of the gan-
glia around the nerve ring and in ventral chord stain red; however, the nerve ring should
remain unstained. Also, observe the hemizonid anterior to the excretory pore.
Investigation 16 Nervous System and Sensory Structures 80
II. Examine the labial papillae, cephalic setae, and the somatic setae.
B. Draconema cephalatum. Examine the somatic setae and the loop-shaped amphids.
A. Compare the spiral type of the Chromadorida (marine nematode) with the circular type
of the Monhsterida (Monhystera sp.) and the question mark type of Plectidae (Plectus
sp.).
B. Trichodorus sp. Study the amphidial opening leading into the amphidial pouch filled
with several sensilla.
A. Ditylenchus sp.
B. Anguina sp.
C. Hoplolaimus galeatus (one on each lateral side, 25-65% and 75-90% from anterior
end).
B. Anaplectus sp. has tuboid sclerotized preanal organs that terminate in glands.
_______________________________________________________________________
_______________________________________________________________________
Investigation 16 Nervous System and Sensory Structures 81
Fig. 16.1 A. Anterior part of Rhaditis terricola (after Hirschmann, modified after Chit-
wood and Chitwood) B. Posterior part of female nematode (after Hirschmann, modi-
fied after Crofton) C. Posterior part of male of Ascaris sp. (after Hirschmann, modi-
fied after Hyman)
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 17
Secretory/Excretory Systems
The secretory/excretory system is diverse in both form and function. In Adenophorean
nematodes, the system is simple and glandular, usually consisting of a single glandular cell
connected to a terminal duct. This normally elongated duct loops within the pseudocoelom
and the glandular tissue. Usually, it is not lined with cuticle and, thus, difficult to see. In
Secernentean systems, the morphology is more complex and diverse. The terminal duct
is usually lined with cuticle and easier to discern. This system is composed of numerous
cells and is characterized by long canals in the lateral chords. Variations in the secretory/
excretory system occur from the loss of some of the component of the complete H-shaped
system.
Nematode species:
Work Sheet
I. Observe and make sketches:
A. Meloidogyne sp. males. Excretory pore located behind the nerve ring; terminal excretory
duct long; lateral canal ending in sinus cell with large nucleus. Note the hemizonid
anteriad of the excretory pore in the hypodermis.
B. Meloidogyne sp. females. The excretory pore is located far anteriad at the height of the
stylet knobs. The terminal duct is long and strongly sclerotized.
C. Plectus sp. Cuticularly lined excretory duct makes 2 loops after entering single excretory
gland cell.
D. Ascaris esophagus cross-section. One large excretory canal located in each of the
lateral chords.
E. Live specimens of Rhabditis sp. in a temporary water mount. The excretory canals are
located within the 2 lateral chords, and proceed anteriad and posteriad. Two ventral
excretory gland cells are present, and a large sinus nucleus is in their vicinity. The
terminal excretory duct is coiled and strongly sclerotized.
________________________________________________________________________
________________________________________________________________________
Investigation 17 Secretory/Excretory System 84
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 18
Reproductive Systems
The reproductive systems of nematodes vary in form and in detail. Basically, the gonads
are tubular structures that are contained within the pseudocoelom. The female reproductive
system can have one or two ovaries; Adenophorean nematodes ususally have one ovary,
whereas Secernentean females usually have two ovaries. In males, there are usually
two testes in Adenophorean species and one testis in Secernentea. Both sexes may have
secondary sexual characters. Males may develop copulatory structures such as bursae,
papillae, gubernaculi, and spicules.
Nematode species:
Work Sheet
I. Study the female reproductive system of the following nematodes and make
drawings:
B. Hoplolaimus sp. - Gonads didelphic (amphidelphic); ovaries not reflexed. Observe the
spermatotheca in line with ovary, and anterior uterus developed as tricolumella.
E. Pratylenchus sp. - Monodelphic gonad (prodelphic); ovary not reflexed. Observe the
round spermatotheca and short postvulval uterine sac.
II. Study the male reproductive system of the following nematodes and make
drawings:
B. Rhabditis sp. - Single (monorchic) reflexed testis; vas deferens with two ejaculatory
glands.
C. Hoplolaimus sp. - Single (monorchic) outstretched testis.
D. Meloidogyne hapla - Two testes (diorchic) oriented parallel. In some specimens only
one testis.
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 19
Female Reproductive System
The reproductive systems of nematodes vary in form and in detail. Basically, the gonad is
a tubular structure that is contained within the pseudocoelom where it may be outstretched
or folded one or more times. It is composed of two basic parts: the ovary and the gonoduct.
The ovary proper is usually divided into three zones: the germinative, growth, and ripening
zones. The gonoduct may have specialized regions as well, including the spermatheca,
oviduct, and uterus. The female reproductive system can have one or two ovaries;
Adenophorean nematodes ususally have one ovary, whereas females of Secernentea usually
have two ovaries.
Nematode species:
Work Sheet
I. Study the female reproductive system of the following nematodes and make
drawings.
B. Dorylaimus sp. - Gonads didelphic (amphidelphic); ovaries reflexed with short germinal
and growth zones.
C. Hoplolaimus sp. - Gonads didelphic (amphidelphic); ovaries not reflexed. Observe the
spermatotheca in line with ovary, and anterior uterus developed as tricolumella.
G. Pratylenchus sp. - Monodelphic gonad (prodelphic); ovary not reflexed. Observe the
round spermatotheca and short postvulval uterine sac.
Investigation 19 Female Reproductive System 90
1. Compare and contrast the ovaries of Diplogaster sp. and Meloidogyne sp. _________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 19 Female Reproductive System 91
Fig. 19.2 Female reproductive system of Meloidogyne javanica. A. One of two gonads
showing the various regions B. Distal end of the germinal zone of the ovary C.
Growth zone of the ovary D. Oviduct and spermatheca E. Lower region of the uterus
with an egg (after Triantaphyllou)
Investigation 20
Male Reproductive System
The reproductive systems of nematodes vary in form and in detail. Basically, the gonad is
a tubular structure that is contained within the pseudocoelom where it may be outstretched
or folded back. There are usually two testes in Adenophorean males and one testis in
Secernentean males. The testis is composed of two basic parts: the testis and the gonoduct.
Usually the testis proper can be divided into two regions: the germinal and growth zones.
The gonoduct usually has a spermatheca (seminal vesicle) and vas defferens, but also
ejaculatory and cloacal glands can also be present. Male copulatory structures may include
bursae, spicules, gubernaculi, papillae and other characters.
Nematode species:
Work Sheet
I. Study the male reproductive system of the following nematodes and make
drawings:
A. Rhabditis sp. Single (monorchic) reflexed testis. Vas deferens with two ejaculatory
glands, testis with short germinal and growth zones.
C. Meloidogyne javanica Two testes (diorchic) oriented parallel; in some specimens only
one testis.
E. Observe the sperm of Rhabditis sp. and Hoplolaimus sp. and compare them with those
of Parascaris equorum.
1. Compare and contrast the testis of Rhabditis sp. with that of Hoplolaimus sp. _______
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 20 Male Reproductive System 95
2. Why do some males of Meloidogyne javanica have two testes, whereas others have
________________________________________________________________________
3. Compare and contrast the sperm of Rhabditis sp. and Parascaris equorum. _________
________________________________________________________________________
________________________________________________________________________
Investigation 21
Eggs
Nematode eggs are usually rounded, oval, or elliptical in shape, and they are surrounded by
three layers. The innermost layer is the lipid layer; the true shell is a chitinous layer which
is secreted by the egg itself and often covered with a vitelline membrane. The outermost
layer is a proteinacious uterine layer which is secreted by the uterine wall. This outer layer
is absent in some nematode eggs. In general, egg morphology is related to the ecology of
the nematode. Some aquatic forms have small spines or hooks, and eggs of some animal
parasites may have specialized structures such as branched polar cords, polar or equatorial
filaments, mammilations, or opercula.
Nematode species:
Work Sheet
I. Observe and draw the morphology of the eggs of several plant-parasitic
nematodes:
A. Belonolaimus sp
B. Hoplolaimus sp.
C. Meloidogyne sp.
D. Pratylenchus sp.
II. Observe and draw the mammilations on the surface of the eggs of Parascaris
equorim.
1. Explain the relationship between the egg morphology and nematode ecology. ______
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 21 Eggs 98
3. Why do many aquatic nematode eggs have small spines and hooks? ______________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
4. What is the purpose of the byssi on the eggs of Mermis subnigrescens? ___________
________________________________________________________________________
________________________________________________________________________
Investigation 22
Preparation of
Permanent Mounts
Long-term collection of nematodes for voucher and type specimens requires special
preparation and mounting techniques. Permanent mounts of nematodes that are well
prepared and mounted in glycerine may be preserved almost indefinitely. The addition
of a general stain such as picric acid or cotton blue can enhance the definition of certain
structures. For plant-parasitic nematodes, the most common fault of permanent mounting
techniques is the dissolution of the stylet shaft and knobs. Likewise, saccate females are
more difficult to preserve and mount because of their awkward shape.
Most permanent mounting techniques are either fast or slow methods. Fast methods usually
require more handling of the specimens but can be as fast as one hour. Slow methods
generally require less handling, but they may take several days or weeks to perform.
Nematode species:
Work Sheet
I. Practice making permanent mounts according to Seinhorstʼs Rapid Method starting
with nematodes that have already been processed into glycerin.
A. Use the provided fixed nematodes and start with step D of the method. Complete the
remaining steps before the next laboratory.
B. Use the provided living nematodes and kill them with hot FP 4:1. Transfer the nematodes
to 2% formalin and fix for 24-48 h; proceed with steps D through G.
II. Prepare permanent mounts according to a method of your choice from those
outlined on the following pages.
D. Place 5-10 nematodes at bottom of glycerin drop. Select only specimens of the same
genus.
E. Place 3 short glass fibers (angel hair) in glycerin drop as coverslip supports. These
fibers should be slightly smaller in diameter than the nematodes.
G. Heat round coverslip gently over alcohol flame and place over glycerin drop with curved
forceps. It is important to center the coverslip properly!
I. After nail polish has dried (15-20 min), remove excess glycerin with small filter paper
strips and ring coverslip completely with a thick seal of nail polish.
J. Examine mounts with the oil immersion objective and select good nematode prepara-
tions.
K. Remove good coverslip mounts from glass slides and place into Cobb slides.
3. Transfer specimens to a solution of 1.25% glycerin in 20% alcohol (add trace of copper
sulfate to prevent fungal growth) contained in a BPI dish or round-bottom glass dish.
4. Place container with nematodes in partially closed petri dish and transfer the latter to a
small airtight desiccator containing calcium carbonate as desiccating agent. Leave for
4-6 weeks at 30-35°C.
5. After complete evaporation of the alcohol, place container with nematodes in calcium
chloride desiccator for 2-3 days.
1. Pick 20-30 live nematodes into 15 drops of tap water in a BPI dish.
2. Place BPI dish in petri dish lines with a moist piece of filter paper and chill nematodes
in the refrigerator at 8°C for at least 30 min.
4. Keep the specimens in the fixative for an additional 48-72 h in the cold.
5. Wash nematodes several times in cold sodium cacodylate buffer, then pick them into
buffer contained in a round-bottom dish, and keep for 24 h in the cold.
8. After complete evaporation of alcohol, place round-bottom dish with nematodes in cal-
cium chloride desiccator for 2-3 days.
Caution: Glutaraldehyde and sodium cacodylate buffer are very toxic substances
and should be handled with extreme caution only under a fume hood.
Investigation 22 Preparation of Permanent Mounts 104
1. Fix female nematodes in FA 4:10 (formalin 10 ml, glacial acetic acid 10 ml, distilled
water 80 ml) or 2% formalin in closed vials for 12-24 h.
Solution__________1_________2________3________4________5___
Glycerin 50 70 80 90 100
Lactophenol 50 30 20 10 0
2. Pipette about 5 ml of hot FP 4:1 fixative quickly over the drop in the dish. Preparation
of hot fixative: Fill a test tube half with FP 4:1 and place in a beaker filled with water.
Heat the water to boiling on a hot plate.
3. Transfer specimens to a small vial filled with cold 2% formalin and fix for 1-2 days.
5. Place dish in a closed desiccator containing about 1/10 of its volume of 96% alcohol,
and leave dish in this saturated atmosphere for at least 12 h in an oven at 35-40°C.
6. Decrease volume if necessary, then top up dish with a solution of 5% glycerin in 96%
alcohol and place dish in a glass container which should be partially closed to allow
slow evaporation. Maintain for at least 8 h at 40°C, or preferably longer, until all alco-
hol has evaporated and the nematodes are in pure glycerin.
7. Transfer nematodes to calcium chloride desiccator and leave for 2-3 days.
Method A
2. Pour about 5 ml of hot FP 4:1 fixative quickly over the drop in the dish.
Preparation of hot fixative: Fill a test tube half with FP 4:1 and place in 250 ml beaker
filled one half of its volume with water. Heat the water to boiling on a hot plate.
3. Transfer specimens to small vial filled with cold FAA and fix for 1-3 days.
4. Pick specimens to a solution of 1.5% glycerin in 20% alcohol con in a small round-bot-
tom glass dish. Fill dish almost to rim with alcohol-glycerin solution and cover tightly
with parafilm. Puncture a few small holes in parafilm and transfer dish to oven at 35-
40°C. Regulate slow evaporation of alcohol by size of parafilm holes. This process
should take at least 2 weeks.
5. After complete evaporation of alcohol, place container with nematodes in calcium chlo-
ride desiccator for 2-3 days.
Method B
1. Transfer 20-50 live nematodes to centrifuge tube filled with 5 ml water at room tem-
perature. After nematodes have settled, add 5 ml boiling 10% formalin (prepare hot
10% formalin as in A.2), and immediately shake tube to mix contents properly.
2. After nematodes have settled again, remove some of the supernatant fixative and trans-
fer rest of fixative (now 5% formalin) with nematodes to small capped vial. Fix for up
to 1 week in 5% formalin.
3. Pick specimens to a solution of 1.5% glycerin in water (add trace of copper sulfate to
prevent fungal growth) contained in a small round-bottom glass dish. Fill dish almost
to rim with water-glycerin solution and cover tightly with parafilm. Puncture small
holes in parafilm to facilitate slow evaporation of water at room temperature. This pro-
cess should take from 2-4 weeks.
4. After complete evaporation of alcohol, place container with nematodes in calcium chlo-
ride desiccator for 2-3 days.
Method C
1. Pick 20-50 live nematodes and wash them 3 times in tap water. Pick specimens into 15
drops of tap water in a BPI dish.
2. Place BPI dish in closed petri dish lines with a moist piece of filter paper, and chill
nematodes in the refrigerator at 8° C for at least 30 min (nematodes should be com-
pletely straight).
4. Keep the specimens in the fixative for an additional 2-3 days in the cold.
5. Transfer the nematodes to a special processing chamber made from a BEEM capsule.*
Wash the specimens in cold sodium-cacodylate buffer 5 times at 15 min intervals (can
keep in buffer overnight in the cold).
6. Remove specimens from refrigerator and allow to attain room temperature. Then dehy-
drate with an alcohol series of 10, 20, 35, 50, 65, 80, 95 and 100%, 15 min at each step
at room temperature (repeat 3 times in 100% alcohol).
9. After complete evaporation of alcohol, place round-bottom dish with nematodes in cal-
cium chloride desiccator for 2-3 days.
However, no matter how good the key, identification of an unknown nematode eventually
requires a detailed study of original species descriptions, and perhaps even type or voucher
specimens. In order for a correct identification to be made, the specimens have to be plenti-
ful, in good physiological condition, well preserved, properly mounted, and adequate keys
be available.
Nematode species:
Work Sheet
I. Prepare semi-permanent formalin mounts (2% formalin) of the nematodes
provided.
A. Use nail polish rings as supports, square or round coverslips for mounting and seal the
mounts with nail polish.
II. Identify the nematodes to genera and, with the help of published keys, identify one
of the nematodes to species.
Investigation 23 Use of Nematological Keys 111
3. Stylet long, straight; body long, slender; very large nematodes; ........Longidoridae......4
3ʼ. Stylet short, curved, weak; body short, thick;..............Trichodoridae..........Trichodorus
Paratrichodorus
15. Scutellae......................................................................................................................16
15ʼ. Phasmids.....................................................................................................................17
Nematode species:
Work Sheet
I. Adjust your microscope for Köhler illumination.
A. Focus the microscope on a contrasting slide (plant sections stained with safranin and
fast green are excellent) with a low power objective (about 10x).
B. Partially close the substage diaphragm and turn down the substage until the edges of
the diaphragm are in focus; center with adjusting screws on the side of the substage. (Some
microscopes have “pre-centered” con densers that should only be adjusted by a service
person.) Rack up the substage condenser close to the stage. Close the field diaphragm (in
front of the illumination) and move the substage condenser up or down until the edge of
the field diaphragm is in perfect focus. The stained slide should remain in focus. Center
the field diaphragm image by moving the mirror as necessary. (This adjustment can prob-
ably be omitted with a built-in illumination. However, some such microscopes do allow for
minor adjustments of a mirror). Check alignment of the lamp relative to the microscope.
When the field diaphragm is centered, open to just the edge of the field of view.
C. Open the substage diaphragm to the proper degree. This can be done in a very clean
microscope room by removing the eyepiece and opening the diaphragm so that the back
(upper) lens of the objective is just filled with light. A second method does not involve
removing the eyepiece. Simply open the diaphragm to the exact point at which further
opening does not increase the brightening of the field. When locating low contrast material
(i.e., unstained nematodes), it may be necessary to further close the substage diaphragm
to increase contrast. The microscopist should be aware that contrast achieved in this way
reduces resolution. It is poor microscopy to use the position of the substage or condenser
diaphragm or the field diaphragm to regulate brightness. Filters or an illuminator reostat is
needed for this purpose.
Investigation 24 Optimal Use of the Light Microscope 116
D. Minor alignment adjustments will need to be made with objectives of higher magnifica-
tions. Certainly the field and condenser diaphragms will need to be reduced to fit the field,
and slight focusing and centering adjustments may be needed. At very low magnifications,
it may be necessary to swing out a condenser front lens or otherwise disperse light to fill
the field.
The ocular micrometer is used to measure the size of microscopic specimens. Because we
use four different lenses, each with a different power of magnification, and because the
optics of each microscope are slightly different, the ocular micrometer must be calibrated
for each objective and each microscope. This standardization procedure is accomplished
tbrough the use of a precision stage micrometer.
1. Record the serial number of your ocular micrometer. Be sure to always use the same
micrometer throughout the term.
Fig. 24.1. Ocular micrometer calibrated with a stage micrometer. With the 10X objec-
tive, the tenth line on the ocular micrometer is in line with the eleventh line on the stage
micrometer. Since the lines on the stage micrometer are 10 µm apart, (111/10) X 10 = 11
µm per ocular scale division.
Investigation 24 Optimal Use of the Light Microscope 117
4. Remove right eyepiece and insert an ocular micrometer. Handle both the eyepiece and
ocular micrometer with care, and store in a dustfree environment.
6. Determine the length of the black scale on the ocular micrometer by measuring it with
the known units on the stage micrometer. Determine the length of the subunits on the ocu-
lar micrometer scale.
8. Repeat procedure for low power, high power, and oil immersion objectives.
A. Oil immersion lenses are absolutely necessary for the observation of nematodes.
1. Focus on the specimen with the high-dry lens.
2. Rotate the objective turrent half-way between the high-dry and the oil immersion
lenses.
3. Place a small drop of oil on the top of the coverslip over the specimen.
4. Rotate the oil immersion lens into place.
5. Since the specimen will almost be in focus, use the fine focus not to gently bring
the specimen into sharp focus.
6. Immediately after use, clean off the oil with a dry piece of lens tissue paper. At
times it may be necessary to use a small amount of liquid lens cleaner.
A. The microscope, and specifically the lens, must be kept as clean as possible:
1. Remove dust with a camel hair brush.
2. Do not use incompatible immersion oils (some types do not mix).
3. Use lens paper with care to avoid scratches.
4. Do not use chemicals that leave a film on lens in the vicinity of microscopes.
5. Do not smoke near a microscope.
6. Do not use drinks or food where they might accidentally get on a microscope.
7. Mascara is a definite problem on eyepieces.
8. A very dirty microscope, particularly one with dirt inside, should only be
cleaned by a service person.
9. Some solvents may actually dissolve the material seating the lens.
B. Eyeglasses normally should not be used with the microscope. However, nearsighted
persons will need corrective glasses for camera lucida work. Persons with astigmatism
should also wear glasses. High eyepoint eyepieces are available for microscopes of per-
sons who must wear eyeglasses.
C. Microscope safety
1. Always carry a microscope with both hands.
2. Always return the turrrent to low power before removing the specimen.
3. Use only lenspaper with lens cleaning solution on the oil immersion lens.
4. Avoid getting oil on lenses other than the oil immersion lens.
5. Clean oil immersion lenses with lenspaper after each use.
Investigation 25
Preparation of Nematodes
for SEM
Scanning electron microscopy (SEM) is very useful for examining nematodes to see all
the minute details of their surface morphology. Proper preparation is essential because
the morphology of poorly prepared specimens may be obscured by surface precipitation
and distorted by shrinkage or swelling.
Preparing nematodes for SEM involves killing, fixation, dehydration, drying, mounting,
coating, and viewing. All of these steps can be deleterious to the surface morphology by
causing the nematodes to shrink and to produce other artifacts. The best way to insure
that the artifacts are minimal is always to compare the SEM observations with images
seen with a light microscope.
Nematode species:
8-step graded series of ethanol of 5, 10, 20, 25, 50, 75, 90, 95, and 100%
Work Sheet
I. Make a processing chamber for preparing specimens for SEM.
C. Cut out a small square of nylon mesh screen and place it over one end of the capsule.
D. Place the cap into position over the screen to hold it in place.
1. What is the purpose of the processing chamber for preparing nematodes for SEM? __
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Investigation 25 Preparation of Nematodes for SEM 121
Fig. 25.1 SEM processing chamber for handling nematodes. A. BEEM® capsule con-
tainer B. Capsule with conical portion removed and perforated cap and two squares
of nylon mesh screen C. Chamber without cap in a Stendor dish D. Container
with cap and lid of Stendor dish (After Eisenback)
G. Post-fix the specimens in 2% osmium tetroxide for at least 4 hr, overnite if desired.
________________________________________________________________________
Investigation 25 Preparation of Nematodes for SEM 122
________________________________________________________________________
________________________________________________________________________
H. Dehydrate nematodes in a eight-step graded series of ethanol of 5, 10, 20, 25, 50, 75,
90, 95, and 100% ethanol for 15-20 min each.
I. Exchange the 100% ethanol three times to ensure that all of the water is removed.
J. Dry the specimens with a critical point dryer using proper techniques as descibed by
the manufacterer of the drying apparatus. (Arrange a time for using the critical point
dryer with the instructor.)
4. Why is it necessary to remove all of the water from the nematodes for SEM?
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
5. Why does air drying from water cause so much damage to the specimens? _________
________________________________________________________________________
________________________________________________________________________
III. Mount the nematodes on stubs for viewing with the SEM.
A. Stick a small square of double sticky tape onto a SEM veiwing stub.
C. Place nematodes on the tape propped up against the hair. The specimens are best
viewed at a 90°angle with the surface of the stub.
D. Coat the specimens with a 20nm layer of gold/palladium. (Arrange a time for using
the gold coater with the instructor.)
Investigation 25 Preparation of Nematodes for SEM 123
Fig. 25.2. Mounting whole dry specimens on the SEM viewing stub. A. Stub with
square of double sticky tape and short length of human hair B. Closeup of nema-
todes propped up against the hair
6. Why are the specimens coated with a thin layer of gold/palladium? _______________
________________________________________________________________________
________________________________________________________________________
7. Compare and constrast the images of nematodes formed with a light microscope and a
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
________________________________________________________________________
Appendix 1
Glossary of
Nematological Terms
“a” - body length/greatest body width (range).
acrosome - refringent body in sperm of some animal parasites.
alae - expansions or projections formed by a longitudinal thickening of the cuticle; there
are three types: 1) cervical alae - confined to the anterior region of the nematodes
parasitic on animals; 2) caudal alae - occur in the posterior region of males in a
number of genera; 3) longitudinal alae - usually 4 in number and extend the length of
the body sublaterally.
allotype - a paratype of the opposite sex to the holotype.
amalgamate - uniting to form a unit.
ambulatory setae - bristles or setae, sometimes hollow and tube-like, by which some
nematodes “walk” along a surface.
amphid - a chemo-sensory organ, occurring laterally in pairs, located in the anterior
region of the nematodes; among the Secernentea, the amphids appear as small, pore-
like openings on the lips; the openings are post-labial in the Adenophorea, and may
be of varied form and made up of the following structures:
1. amphidial duct - a channel which connects the amphidial pore to the sensilla
pouch.
2. amphidial gland - a microvillous organ enveloping sensilla pouch.
3. amphidial nerve - a nerve innervating the amphid.
4. amphidial pore - the aperture in the cuticle through which the amphid opens
exteriorly.
5. amphidial pouch - (amphidial chamber) - a chamber, usually just behind
amphidial pore, outline often cup-shaped.
6. sensilla pouch - chamber containing nerve processes.
amphidelphic (ovary) - having two ovaries, directed anteriad and posteriad, respectively.
ampulla - widening in canal, forming a reservoir.
aphelenchoid - dorsal gland empties the lumen of esophagus just anterior of the valve of
the median bulb instead of opening near stylet base.
annule - the thickened interval between transverse depression (annulations) in the cuticle.
arcuate - curved like a bow.
areolation - a condition where the transverse body annulation traverses the lateral field.
articulate - jointed, segmented.
attenuate - to thin in consistency.
Appendix 1 Glossary of Nematological Terms 125
lateral field - longitudinal cuticular thickening situated on top of the lateral chords; the
field may be divided by longitudinal striae (= incisures) and at times by transverse
markings (areolation).
lateral guiding pieces - structure that consists of two, very small, lineate, cuticularized
pieces lying lateral to the distal portion of the spicules and joining with the
muscular sheath surrounding the anus (present in Dorylaimidae, Belondiridae and
Leptonchidae).
lectotype - one of a species of syntypes which, subsequent to the publication of the
originial description, is selected and designated through publication to serve as the
type.
leptoderan - caudal alae which do not meet posterior to the tail tip.
lip - cuticular structures (usually six), 2 subdorsal, 2 lateral, 2 subventral, surrounding
the mount opening. (May be fused in pairs.)
longitudinal ridges - raised cuticular areas which extend through the length of the body
and are present on all sides of the nematode.
lumen - the canal or duct of the esophagus.
mammillate - digitate; nipple-shaped protuberances.
marginal muscle fibers - muscle fibers of the esophagus at the apices of the triradiate
lumen which have a suspensory and skeletal function.
matrix - the middle layer of the cuticle.
median bulb - see metacorpus.
meromyarian - an arrangement of the somatic musculature, where only few (2-4)
longitudinal rows of muscle cells are present between each two chords.
metacorpus - the swollen posterior portion of the corpus of the esophagus (sometimes
referred to as the median bulb).
metarhabdions - see rhabdions.
metatype - a specimen compared by the author of the species with the holotype and
determined by him as conspecific with it.
micropyle - the minute opening in the membrane of an egg through which the
spermatozoon enters.
molt - to cast off the cuticle.
monodelphic - possessing one genital tube or ovary.
monorchic - possessing one testis.
monoxenic culture - growing an organism in the presence of another organism.
mucro - a stiff or sharp point abruptly terminating an organ.
mucronate - ending in a sharp point.
mural - resembling a wall.
myofilaments - the thick and thin muscle elements of the contractile region of the
muscle cell.
myriocytous - a very large number of cells (over 8,224) in the intestinal epithelium (the
cells are cuboidal).
neck - that portion of the nema body occupied by the esophagus.
Appendix 1 Glossary of Nematological Terms 130
PUNCTODERA
P. punctata...................................Grass cyst nematode...............Wheat, other small grains
NACOBBUS
N. batatiformis.................................................................................................... Sugarbeet
SPHAERONEMA
S. sasseri........................................................................................... Red spruce, Fraser fir
TYLENCHULUS
T. semipenetrans.............................Citrus nematode.......................................Citrus, olive
Appendix 2 Nematode Chart 137
ECTOPARASITIC NEMATODES
Nematodes that feed on the root surface and normally do not enter the root tissue
BURSAPHELENCHUS
B. xylophilus.............................Pinewood nematode...........................................Pine trees
DITYLENCHUS
D. dipsaci....................................Stem and Bulb, or.........................Alfalfa, clover, onion,
Teasel nematode garlic, sweet potato,
strawberry, nursery
Appendix 3
Nematological Journals
*Afro-Asian Journal of Nematology (1990-1996)
Annals of Applied Biology
Annals of the Phytopathological Society of Japan
Biological and Cultural Tests for Control of Plant Diseases
Bulletin OEPP = EPPO Bulletin
Canadian Journal of Zoology
Comparative Parasitology (2000)
Fitopatologia
*Fundamental and Applied Nematology (until 1991 Revue de Nématologie)
Fungicide & Nematicide Tests
*Indian Journal of Nematology (1971)
International Journal of Nematology (until 1996 Afro-Asian Journal of Nematology)
*Japanese Journal of Nematology (1972)
*Journal of Nematology (1969)
*Journal of Nematode Morphology and Systematics (1999)
*Nemapix (1998)
*Nematologica (1956-1998)
*Nematologica Mediterranea (1973)
* Nematology: International Journal of Fundamental and Applied Nematology (1999)
*Nematropica (1971)
Netherlands Journal of Plant Pathology
*Pakistan Journal of Nematology (1983)
Parasitology
Phytoparasitica
Phytopathologica Mediterranea
Phytopathologische Zeitschrift (Journal of Phytopathology)
Phytopathology
Phytophylactica
Phytoprotection
Plant Disease (formerly Plant Disease Reporter)
Plant Pathology
Proceedings of the Helminthological Society of Washington (until 2000 = Comparative
Parasitology
*Revue de Nematologie (1978-1991)
*Russian Journal of Nematology (1993)
Rivista di Patologia Vegetale
Systematic Parasitology
*specializes exclusively in plant nematology
Appendix 3 Nematological Journals 140
Bibliography of Agriculture
Biological Abstracts
Bio Research Index
Bio Systematic Index
Commonwealth Agricultural Bureaux (CAB International) Abstract Journals
Current Contents (Life Sciences)
Dissertation Abstracts
Helminthological Abstracts, Series B
Nematological Abstracts (Helminthological Abstracts, Series B: Plant Nematology.)
Science Citation Index
Zoological Record
Appendix 4
Nematological Societies
PROFESSIONAL SOCIETIES FOR NEMATOLOGISTS
Dropkin, V. H. 1989. Introduction to Plant Nematology, 2nd ed. John Wiley & Sons: New
York.
Eisenback, J. D. 2002. Identification Guides for the Most Common Genera of Plant-Par-
asitic Nematodes. Mactode Publications: Blacksburg, Viginia.
Evans, K., D. Trudgill, and J. M. Webster, (eds.). 1993. Plant Parasitic Nematodes in
Temperate Agriculture. CAB International: Wallingford, United Kingdom.
Fenoll, C., F. Grundler, F. M. W. and S. A. Ohl, (eds.). 1997. Cellular and Molecular
Aspects of Plant-Nematode Interactions. Kluwer Academic: Dordrecht.
Fortuner, R. 1988. Nematode Identification and Expert System Technology. NATO
Workshop, Raleigh. Plenum Press: New York and London.
Fortuner, R., E. Geraert, M. Luc, A. R. Maggenti, and D. J. Raski. 1988. A Reappraisal
of Tylenchina (Nemata). ORSTROM, France.
Gaugler, R., and H. K. Kaya. 1990. Entomopathogenic Nematodes in Biological Control.
CRC Press: Boca Raton, Florida.
Gommers, F., and P. W. Th. Maas, (eds.). 1992. Nematology: from Molecule to
Ecosystem. Proceedings of the Second International Nematology Congress 1990
Veldhoven. European Society of Nematologists.
Gubina, V. G. 1988. Nematodes of Plants and Soils: the Genus Ditylenchus. Saad Publi-
cations: Karachi, Pakistan.
Hickey, K. D. 1986. Methods for Evaluating Pesticides for Control of Plant Pathogens,
Fungicides, Nematicides and Bactericides. American Phytopathological Society: St.
Paul, Minnesota.
Hunt, D. J. 1994. Aphelenchida, Longidoridae and Trichodoridae: Their Systematics and
Bionomics. CAB International: Wallingford, United Kingdom.
Jairajpuri, M. S., and W. Ahmad. 1992. Dorylaimida - Free-living, Predaceous and Plant-
parasitic. E. J. Brill: Leiden.
Jepson, S. B. 1987. Identification of Root-knot Nematodes. CAB International:
Wallingford, Oxon, United Kingdom.
Khan, M. W., (ed.). 1993. Nematode Interactions. Chapman & Hall: London, United
Kingdom.
Khan, M. W., and Sharma, P. D., 1996. Root-knot Nematodes in India. Venus Publishing
House: New Delhi, India.
Lamberti, F., and Taylor, C.E. (eds.). 1979. Root-knot Nematodes. Academic Press. New
York.
Lamberti, F., C. de Giorgi, and D. McK. Bird. 1996. Advances in Molecular Plant
Nematology. NATO ASI Series A., Vol. 268. Plenum Publishers: New York.
Lamberti, F. D. Bird, and C. de Giorgi. 1994. Advances in Molecular Plant Nematology.
Kluwer Academic Publishers:
Lamberti, F., C. E. Taylor, and J. W. Seinhorst. 1975. Nematode Vectors of Plant Viruses.
Plenum Press: London and New York.
Lorenzen. S. 1994. The Phylogenetic Systematics of Freeliving Nematodes. The Ray
Society: Andover.
Appendix 5 Nematological Resources 144
Poinar, G. O., and H. B. Jansson. 1988. Disease of Nematodes, Volume I. CRC Press:
Bocan Raton, Florida.
Poinar, G. O., and H. B. Jansson. 1988. Disease of Nematodes, Volume II. CRC Press:
Bocan Raton, Florida.
Riggs, R. D. (ed.) 1982. Nematology in the Southern United States. Southern Cooperative
Series Bulletin 276: Fayetteville. Arkansas.
Riggs, R. D., and J. A. Wrather, (eds.). 1992. Biology and Management of Soybean Cyst
Nematodes. American Phytopathological Society: St. Paul, Minnesota.
Santos, M. S. N. de A., I. M. de O. Abrantes, D. J. F. Brown, and R. M. Lemos, (eds.).
1997. An Introduction to Virus Vector Nematodes and their Associated Viruses.
Universidade de Coimbra: Portugal.
Sasser, J. N. 1989. Plant-Parasitic Nematodes: The Farmer’s Hidden Enemy. North
Carolina State University, Raleigh, North Carolina.
Sasser, J. N., C. C. Carter, and K. R. Barker. 1985. An Advanced Treatise on
Meloidogyne. Vol. I Biology and Control. Vol. II Methodology. International
Meloidogyne Project. North Carolina State University, Raleigh, North Carolina
Sauer, M. R. 1985. A Scanning Electron Microscope Study of Plant and Soil Nematodes.
Commonwealth Science Industry Research Organization, Australia.
Sharma, S. B. (ed,) 1998. The Cyst Nematodes. Kluwer Academic Publishers: Dordrecht.
Sharma, R., and G. Swarup. 1988. Pathology of Cyst Nematodes. Malhotra Publishing
House: New Delhi.
Shurtleff, M.C., and Averre. C. W. 2000. Diagnosing Plant Diseases Caused by
Nematodes. American Phytopathological Society: St. Paul Minnesota.
Siddiqi, M. R. 2000. Tylenchida: Parasites of Plants and Insects. CAB International:
Wallingford, United Kingdom.
Simoes, N., N. Boemare, and R. Ehlers, (eds.). 1988. Entompathogenic Nematodes-
Pathogenicity of Entomopathogenic Nematodes Versus Insect Defence
Mechanisms: Impact on Selection of Virulent Strains. Office for Official
Publications of the European Communities.
S’Jacob, J. J., and J. van Bezooijin. 1984. Manual for Practical Work in Nematology.
Landbouwhogeschool, Wageningen.
Southey, J. F., (ed.). 1978. Plant Nematology. GD 1, MAFF, HMSO, London.
Southey, J. F. 1986. Laboratory Methods for Work with Plant and Soil Nematodes, 6th ed.
Reference Book 402. Ministry of Agriculture, Fisheries, and Food, HMSO:
London, United Kingdom.
Starr, J. L., (ed.). 1990. Methods for Evaluating Plant Species for Resistance to Plant
Parasitic Nematodes. Society of Nematologists, Hyattsville, Maryland.
Starr, J. R., R. Cook, and J. Bridge. 2002. Plant Resistance to Parasitic Nematodes. CABI
Publishing: Wallingford, United Kingdom.
Stirling, G. R., 1991. Biological Control of Plant Parasitic Nematodes. CAB
International: Wallingford, United Kingdom.
Stone, A. R., H. M. Platt, and L. F. Khalil. 1983. Concepts in Nematode Systematics.
Appendix 5 Nematological Resources 146
CD ROM RESOURCES
PICTURE COLLECTIONS
Eisenback, J. D., and Ulrich Zunke. 1997. Nemapix Volume 1. Mactode Publications:
Blacksburg, Virginia.
Eisenback, J. D., and Ulrich Zunke. 1999. Nemapix Volume 2. Mactode Publications:
Blacksburg, Virginia.
Eisenback, J. D., and Ulrich Zunke. 2002. Nemapix Volume 3. Mactode Publications:
Blacksburg, Virginia.
SLIDE SERIES
Society of Nematologists, Slide Set Nematology, 100 Slides. T.A. Niblack, Department of
Plant Pathology, University of Missouri, Columbia, Missouri.
Högger, C.H. FAP Dia-Serien 1986: Nr. 3 Nematoden: Formenvielfalt und Antagonisten;
Nr. 4 Nematoden an Getreide; Nr. 5 Nematoden an Kartoffeln; Nr. 6 Nematoden an
Zucker- und Futterrüben, je 20 Dias, Legende, FAL Zürich-Reckenholz.
De Grisse, A. 1976. Rasterelektronenmikroskop-Aufnahmen (SEM) von Nematoden. 113
Dias. C. De Grisse-Helleybuck, Schoonmeerstr. 37, B-9000 Ghent, Belgien.
Society of Nematologists. 1990. Slide Set: Parasites and Predators of Plant-Parasitic
Nematodes. 60 slides. Society of Nematologists Biological Control Committee.
VIDEOS
Eisenback, J. D., and E. C. McGawley. 1996. A Videocassette for Teaching the Iden-
tification of the Most Common Genera of Plant-Parasitic Nematodes. Mactode
Publications: Blacksburg, Virginia.
Appendix 6
References to
Nematological Keys
Aglenchus Andrássy, 1954
Andrássy, I. 1980. The genera and species of the family Tylenchidae Oerley, 1880
(Nematoda). Acta Zoologica Academiae Scientiarum Hungaricae 26:1-20.
Nama, H. S., and Soni, G. R. 1981. Taxonomy of some species of the genus
Aphelenchus Bastian, 1865 (Aphelenchoidea) with a key. Proceeding of the
Indian Academy of Parasitology 2:107-110.
Anderson, R. V., and D. J. Hooper. 1980. Diagnostic value of vagina structure in
the taxonomy of Aphelenchus Bastian, 1865 (Nematoda: Aphelenchidae) with
a description of A. (Anaphelenchus) isomerus n. subgen., n. sp. Canadian
Journal of Zoology 58:924-928.
Karegar, A., and E. Geraert. 1997. The genus Basiria Siddiqi, 1959 (Nematoda:
Tylenchida). II. Species with four lateral lines and anterior median bulb.
Nematologica 43:383-406.
Karegar, A., and E. Geraert. 1998. The genus Basiria Siddiqi, 1959 (Nemata:
Tylenchida) IV. General discussion, genus diagnosis and key to the species.
Nematologica 44:1-13.
Sultan, M. S., and M. S. Jairajpuri. 1981. Two new species of the genus
Cephalenchus (Goodey, 1962) Golden, 1971 with a key to species. Indian
Journal of Nematology 11:165-171.
Mizukubo, T., and N. Minagawa. 1985. Taxonomic study of the genus Cephalenchus
(Nematoda:Tylenchida) from Japan. Descriptions of three new species and
records of C. planus Siddiqui & Khan with a key to species. Japanese Journal
of Nematology 15:26-40.
Raski, D. J., and E. Geraert. 1986. Descriptions of two new species and other
observations on the genus Cephalenchus Goodey, 1962 (Nemata: Tylenchida).
Nematologica 32:56-78.
Geraert, E., and D. J. Raski. 1988. Study of some Aglenchus and Coslenchus
species (Nematoda:Tylenchida). Nematologica 34:6-46.
Golden, A., and W. Friedman. 1964. Some taxonomic studies on the genus
Criconema (Nematoda: Criconematidae). Proceedings of the Helminthological
Society of Washington 31:47-59.
Raski, D. J., and E. Geraert. 1986. Review of the genus Filenchus Andrássy, 1954
and descriptions of six new species (Nemata: Tylenchidae). Nematologica
32:265-311. (Synonymy of Ottolenchus, Dactylotylenchus, Lambertia,
Duosulcius, Zanenchus, and Discotylenchus with Filenchus)
Appendix 6 References to Nematological Keys 153
Ebsary, B. A., and R. V. Anderson. 1982. Two new species of Hirschmaniella Luc
and Goody, 1963 (Nematoda: Pratylenchidae) with a key to the nominal
species. Canadian Journal of Zoology 60:530-534.
Sivakumar, C. V., and E. Khan. 1982. Description of Hirschmaniella kaverni sp. n.
(Radopholidae: Nematoda) with a key for identification of Hirschmaniella spp.
Indian Journal of Nematology 12:86-90.
Andrássy, I. 1981. The genera and species of the family Tylenchidae Oerley,
1880 (Nematoda). The genus Malenchus Andrássy, 1968. ). Acta Zoologica
Academiae Scientiarum Hungaricae 27:1-47.
Geraert, E., and D. J. Raski. 1986. Unusual Malenchus species (Nemata:
Tylenchidae). Nematologica 32:27-55.
Taylor, A. L., and J. N. Sasser. 1978. Biology, identification and control of root-knot
nematodes (Meloidogyne species). Raleigh: North Carolina State University
Graphics.
Eisenback, J. D., H. Hirschmann, J. N. Sasser, and A. C. Triantaphyllou. 1981. A
guide to the four most common species of root-knot nematodes (Meloidogyne
spp.), with a pictorial key. Raleigh: North Carolina State University.
Ebsary, B. A., and E. S. Eveleigh. 1983. Meloidogyne aquatilis n. sp. (Nematoda:
Meloidogynidae) from Spartina pectinata with a key to the Canadian species of
Meloidogyne. Journal of Nematology 15:349-353.
Hewlett, T. E., and A. C. Tarjan. 1983. Synopsis of the genus Meloidogyne Goeldi,
1887. Nematropica 13:79-102.
Jepson, S. B. 1987. Identification of root-knot nematodes (Meloidogyne species).
Wallingford, UK: CAB International. 265 p.
Eisenback, J. D., and H. H. Triantaphyllou. 1991. Root-knot nematodes:
Meloidogyne species and races. Pp. 191-274 in W. R. Nickel, ed. Manual of
agricultural nematology. New York: Marcel Dekker.
Sher, S. A. 1970. Revision of the genus Nacobbus Thorne and Allen, 1944
(Nematoda: Tylenchoidea). Journal of Nematology 2:228-235.
Powers, T. O., J. G. Baldwin, and A. H. Bell. 1983. Taxonomic limits of the genus
Nagelus (Throne and Malek, 1968) Siddiqi, 1979 with a description of
Nagelus borealis n. sp. from Alaska. Journal of Nematology 15:582-593.
(Tabular information)
Khan, F. A., and A. M. Khan. 1975. Three new species of Neopsilenchus Thorne &
Malek, 1968 (Nematoda: Psilenchinae). Indian Journal of Nematology 5:15-
21.
Sultan, M. S., I. Singh, P K. Sakhuja. 1987. Plant parasitic nematodes of the Punjab,
II. Neopsilenchus longicaudatus n. sp. (Nematoda: Tylenchidae). Indian
Journal of Nematology 17:330-332.
Shahina, F., and Maqbool, M. A. 1990. Studies on the genus Neopsilenchus
(Nematoda: Tylenchidae) with description of Acusilenchus n. subgen and three
new species. Pakistan Journal of Nematology. 8:49-63.
Ottolenchus Husain & Khan, 1967 (Wu, 1970) Golden, 1971 synonymized this
subgenus with Aglenchus, Andrássy, 1954
Kheiri, A. 1970. Two new species in the family Tylenchidae (Nematoda) from Iran,
with a key to Psilenchus De Man, 1921. Nematologica 16:359-368.
Brzeski, M. W. 1989. Notes on the genus Psilenchus de Man, 1921, with description
of P. klingleri sp. n. (Nematoda: Tylenchidae). Annales Zoologici 43:51-69.
Kleynhans, K. P., and P. Cadet. 1994. Trophurus deboeri n. sp. from sugarcane
soil in Barbados and key to the species of the genus Trophurus Loof, 1956
(Nematoda: Belonolaimidae). Fundamental and Applied Nematology 17:225-
230.
Appendix 6 References to Nematological Keys 161
Loof, P. A. A., and M. Luc. 1993. A revised polytomous key for the identification of
species of the genus Xiphinema Cobb, 1913 (Nematoda: Longidoridae) with
exclusion of the X. americanum-group: Supplement I. Systematic Parasitology
24:185-189.
Loof, P. A. A., M. Luc, and P. Baujard. 1996. A revised polytomous key for the
identification of species of the genus Xiphinema Cobb, 1913 (Nematoda:
Longidoridae) with exclusion of the X. americanum-group: Supplement 2.
Systematic Parasitology 33:23-29.
Robbins, R. T., D. J. F. Brown, J. M. Halbrendt, and T. C. Vrain. 1996. Compendium
of juvenile stages of Xiphinema species (Nematoda: Longidoridae). Russian
Journal of Nematology 4:163-171.
This list of keys and references for identifying species of selected genera of plant-
parasitic nematodes was compiled to assist students and professionals in the discipline of
nematology. It is not comprehensive but was compiled as a current source of information
and initial perspective for those interested in species identification of selected plant-
parasitic nematode genera. Nomenclature and systematics follow that of Ebsary, 1991,
and genera selected are primarily those in Mai, W. F., and P. G. Mullin, with H. H. Lyon
and K. Loeffler. 1996. Plant-parasitic nematodes: A pictorial key to genera. Ithaca,
NY: Cornell University Press. We added older references where most appropriate and
included some references that are not keys, where keys are unavailable or if particularly
useful as supplemental references. For information on older references and on genera
not included herein, see E. C. Bernard and S. M. Baird. 1983. Bibliography of keys for
the identification of plant-parasitic and free-living terrestrial nematodes. Nematology
Newsletter 29(4):9-18. Detailed study should encompass original species descriptions.
We recommend the following references for studies of the principles of taxonomy behind
many of the keys and for general review:
Ebsary, B. A., 1991. Catalog of the Order Tylenchida (Nematoda). Publication 1869/
B. Ottawa, Canada: Canada Communication Group-Publishing. 196 p.
Nickle, W. R., ed. 1991. Manual of agricultural nematology. New York, NY: Marcel
Dekker. 1035 p. (A valuable reference that has many species keys)
Appendix 6 References to Nematological Keys 165
This list of references to the keys was prepared by:
Z. A. Handoo and A. M. Golden, USDA, ARS, Nematology Lab, Bldg. 011A, Rm.
159, BARC-West, 10300 Baltimore Avenue, Beltsville, MD 20705-2350
The authors thank Dr. E. C. Bernard for his review of this submission.
Appendix 7
Key to Females
1. Body greatly swollen; adult female immobile, completely or partially embedded in
plant root..............................................................................................................................2
1. Body relatively slender; mobile animals.........................................................................9
9. On the front side of the head, there is a mouth stylet or spear that can be projected....10
9. Head without mouth stylet or spear...............................................................................63
10. Mouth with tylenchid spear; esophagus usually with a median bulb...........................11
10. Mouth with dorylaimid spear; esophagus cylindrical or bottle-shaped, without
median bulb.......................................................................................................................35
17. Gradual transition between procorpus and median bulb, basal bulb strongly reduced;
cuticle often conspicuously heavily ringed........................................................................18
17. Distinct constriction present between procorpus and median bulb..............................21
18. Cuticle not conspicuously coarsely ringed; constriction between median and basal
bulb is relatively long.................................................................................Paratylenchidae
18. Cuticle not conspicuously coarsely ringed, already visible at 40 x magnification;
median bulb not much constricted, fused with the basal bulb...........................................19
19. Cuticle consists of 2 layers, outer layer as-it-were forming an extra cuticle which
surrounds the body like a sheath........................................................................................20
19. Cuticle consisting of one layer; body plump and heavily ringed, rings often
ornamented .................................................................................................Criconematidae
20. Stylet knobs pointed forward; body plump, ratio a smaller than 20; vulva at
over 90% (presumably of body length).................................................Hemicriconemoides
20. Stylet knobs rounded; body relatively slender, ratio a greater than 20;
vulva at most at 90%............................................................................Hemicycliophoridae
23. Median bulb strongly muscular and conspicuously well developed; already
conspicuous at low magnification; duct of dorsal esophageal gland opens into median
bulb.....................................................................................................................................24
23. Median bulb developed normally; duct of the dorsal esophageal gland opens behind
the mouth stylet in the lumen of the esophagus.................................................................26
Appendix 7 Key to Females 169
26. Head skeleton strongly developed; mouth spear conspicuously robust; tail usually
shorter than 2.5 anal body diameters............................................................Pratylenchidae
26. Head skeleton poorly developed; mouth spear slender; tail usually longer than 2.5
anal body diameters, rarely shorter....................................................................................27
28. Mouth spear long, longer than 40% of the distance from the front of the body to the
median bulb.......................................................................................................Tylodoridae
28. Mouth spear at most 30% of the distance from the front to the median bulb..................
...........................................................................................................................Tylenchidae
35. Long, slender nematodes, greater than 2 mm; a ratio greater than 60 with a strongly
elongated straight stylet, longer than 55 µm...................................................Longidoridae
35. Stylet shorter than 55 µm, or when longer than the nematode is shorter than 2 mm;
a ratio less than 50..............................................................................................................36
Appendix 7 Key to Females 170
36. Stylet relatively long and strongly bent; tail virtually absent..................Trichodoridae
36. Stylet straight; tail present...........................................................................................37
39. Oral cavity with 4 large teeth; in the wall of the oral cavity there is also a number of
smaller teeth; tail long, head weakly set off, defined.................................Paractinolaimus
39. Oral cavity without large teeth; wall of oral cavity with 6 sclerotized ribs; tail short,
head strongly set off....................................................................................Carcharolaimus
40. Lip region set off by a deep constriction; lip region disc- or sucker-shaped; lateral
field usually with a conspicuous series of glands..............................................................41
40. Lip region not disc- or sucker-shaped.........................................................................42
44. Small, plump nematodes, with stylet made up of several components but often
invisible because of grains on the body, with cuticle often loosely draped around the body
...............................................................................................................Diphtherophoridae
44. Body without grains and with a normal cuticle...........................................................45
45. The posterior widened part of the esophagus is surrounded by a sheath of muscle.....
..........................................................................................................................Belondiridae
45. Esophagus proximally not surrounded by a muscle sheath........................................46
46. Stylet surrounded by a membranous guide ring; several layers of cuticle often easily
discernible o the tail....................................................................................Aporcelaimidae
46. Guide ring not membranous; cuticular layers on the tailʼs tip not easily discernible...
............................................................................................................................................47
Appendix 7 Key to Females 171
47. Stylet asymmetrical, the dorsal side of the stylet is longer than the ventral side; stylet
continuation is curved and surrounded by dense esophageal tissue.............Dorylaimoides
47. Stylet, if asymmetrical, not with a stylet extension that is curved and surrounded by
dense esophageal tissue......................................................................................................48
48. Proximal part of esophagus short and swollen, pear-shaped; transition of intestine to
prerectum is situated close to the vulva..............................................................Leptonchus
48. Prerectum shorter, proximal part not swollen pear-shaped..........................................49
49. Stylet extension with knob-shaped outgrowths; body length shorter than 1 mm; lip
region set off like a cap................................................................................Tylencholaimus
49. Stylet extension without knob-shaped outgrowths, if with knob-stylet extension, then
the lip region is not set off like a cap or the body length is greater....................................50
55. Tail bluntly rounded, cylindrical; lip region not set off; stylet extension weakly
flanged......................................................................................................................Thornia
55. Tail not cylindrical, stylet extension not flanged...................................Tylencholaimus
57. The wide proximal part of the esophagus is surrounded by a muscle sheath..............57
57. Esophagus not surrounded by a muscle sheath............................................................59
62. Tail bluntly rounded; a few light-refracting particles around the mouth
opening...........................................................................................................Pungentus
62. Tail long or cone-shaped; no refractory particles around oral
cavity..................................................................................................Thornenematidae
63. Body caterpillar-shaped with alternating wide and narrow rings; amphids bladder-
shaped........................................................................................................Desmoscolecidae
63. Body not caterpillar-shaped.........................................................................................64
64. Body asymmetrical, tight body half displays a fine net structure and/or a series of fins
or small shields............................................................................................Bunonematidae
64. Body asymmetrical without conspicuous fins or small shields....................................65
68. Oral cavity tubular and with stylet knobs; esophagus tylenchid.........Tylopharyngidae
68. Oral cavity without stylet knobs..................................................................................69
69. Oral cavity with a movable tooth dorsally; right subventrally a large tooth, and left
subventrally a smooth or toothed small plate........................................Neodiplogasteridae
69. Oral cavity with an immovable tooth dorsally; left and right metastoma swellings are
identical to each other..................................................................................Diplogasteridae
72. Front part of esophagus surrounding the oral cavity sharply set off by a constriction...
....................................................................................................................Ethmolaimidae
72. Front part of esophagus not sharply set off................................................................73
74. Oral cavity with a dorsal and several subventral teeth; distal part of esophagus
symmetrical................................................................................................Chromadoridae
74. Dorsal tooth is actually a continuation of the esophageal tissue; subventral teeth
inconspicuous; distal part of esophagus asymmetrical........................Hypodontolaimidae
78. Oral cavity uniformly tubular; tail glands lacking; amphids inconspicuous; head setae
absent................................................................................................................................79
78. Oral cavity narrowing at the base; tail glands and drainage duct present; amphid
halfway oral cavity, round or slit-shaped; neck wings present or absent; head setae often
present....................................................................................................................Plectidae
79. Oral cavity not fused, consisting of separate rings; esophageal body with three
consecutive swellings.........................................................................23...Alloionematidae
79. Walls of oral cavity fused; body of esophagus at most regularly swollen..................80
83. Tail drawn out like a thread; oral cavity small with barely sclerotized walls; head
setae present.............................................................................................Prismatolaimidae
83. Tail never drawn out like a thread; oral cavity large and with strongly sclerotized
walls; head setae absent.....................................................................................................84
84. Base of oral cavity flat; base of esophagus with three small glands; usually three
identical teeth in oral cavity...........................................................................Anatonchidae
84. Base of oral cavity funnel-shaped; base of esophagus without glands; subventral tooth
or teeth, if present, never identical to dorsal tooth.........................................Mononchidae
86. Amphids absent, tail bluntly rounded; oral cavity tubular, cuticle very finely punctate;
ratio b greater than 10.........................................................................................Isolaimidae
86. Amphids spiral or funnel-shaped; ratio b smaller than 10...........................................87
92. Oral cavity formed by a long thin tube which is at least 1/3 as long as the distance
from the front to the esophageal base.............................................................Aulolaimidae
92. Oral cavity shorter........................................................................................................93
94. Head with four setae; body conspicuously ringed; tail never spatula-shaped; behind
the basal bulb the esophagus does not continue in the intestine........................................96
94. Head with 10 setae, 6 long ones and 4 shorter ones, or without setae; body not
conspicuously ringed; tail spatula-shaped; behind the basal bulb the esophagus narrows
and continues partially into the intestine esophagus with a basal bulb.........Leptolaimidae
97. Cuticle ringed; oral cavity closed at rest, but a tooth is recognizable in the closed oral
cavity....................................................................................................................Tripylidae
97. Cuticle smooth.............................................................................................................98
102. Anterior gonad flipped (turned around) and continuing to behind the vulva...........103
102. Anterior gonad, if flipped, not continuing to behind the vulva................................105
104. Anterior gonad continues to behind the vulva, where it bends sharply; cuticle usually
coarsely ringed; oral cavity long and narrow, regularly sclerotized; cheilostoma usually
wider than the other elements; edge of head often indented, lips with or without
appendages......................................................................................................Cephalobidae
104. Anterior gonad continues beyond the vulva, and has no sharp bend; cuticle finely
ringed; oral cavity consisting of cheilo-, pro-, and mesostome, which together form a
more or less rectangular cavity; pro- and mesostome are strongly sclerotized; lips never
with appendages .......................................................................................Panagrolaimidae
Appendix 7 Key to Females 176
109. At the front of the head there are four long thin hairs; the bulb consists of two parts,
in the anterior part a number of small teeth are present; the esophagus continues behind
the posterior bulb into the intestine.................................................................Chronogaster
109. Head without long thin hairs; esophagus not continuing behind the posterior bulb......
..........................................................................................................................................110
112. On the dorsal wall of the oral cavity there is a large tooth; anterior gonad turned
around, head setae absent; amphid cup-shaped and inconspicuous....................Campydora
112. Oral cavity without teeth; gonad stretched out; head setae present; amphid
conspicuous and round.....................................................................................Monhystrella
114. Esophagus consists of two parts, the anterior part is muscular, the posterior part
consists of gland tissue; head setae absent; there are a number of small teeth in the oral
cavity....................................................................................................Odontopharyngidae
114. The esophagus is not differentiated; head setae present; oral cavity at most with an
inconspicuous tooth..........................................................................................................115
116. On the dorsal wall of the oral cavity there is a large tooth; head setae absent, amphid
inconspicuous, cup-shaped.................................................................................Campydora
116. Oral cavity without teeth; head setae present; amphid round and conspicuous.......117
117. Cuticle distinctly ringed; gonad located to the left of the intestine................Xyalidae
117. Cuticle smooth; gonad located to the right of the intestine..................Monhysteridae
Translated from Dutch by Dr. Anton Baudoin from the excellent resource written by Dr.
Tom Bongers. (Used with permission.)
2. Tylenchid stylet present, esophagus usually with a median bulb with valve apparatus;
ventral supplements lacking, a bursa is usually present.......................................................3
2. Dorylaimid spear present; esophagus usually bottle-shaped; a median bulb is lacking;
supplements virtually always present, a bursa virtually always lacking...........................86
4. Bursa absent....................................................................................................................5
4. Bursa present.................................................................................................................11
13. Body conspicuously slender, strongly reminiscent of a glass fiber; the body behind the
anus is conspicuously narrower; bursa lobed........................................Ecphyadophoridae
13. Body plump, ratio a smaller than 100..........................................................................14
16. Median bulb strongly developed; duct of the dorsal esophageal gland opens just
before the valve apparatus into the lumen of the esophagus; bursa rudimentary..................
.....................................................................................................................Paraphelenchus
16. Median bulb not conspicuously muscular, the bursa is well developed, the anterior
part of the esophagus is swollen in the shape of a spool; a genuine median bulb is lacking.
............................................................................................................................................17
17. The anterior part of the esophagus is swollen in the shape of a spool; a genuine
median bulb is lacking.................................................................................Neotylenchidae
17. Median bulb well developed........................................................................................18
19. Spicula widened at the front; tail cone-shaped, bursa reaches almost to the top of the
tail.......................................................................................................................Anguinidae
19. Spicula not widened at the front; tail long drawn out; bursa adanal............................20
20. Stylet knobs absent; phasmid on the tail; amphids slit-shaped, located at the side of
the lip region.....................................................................................................Psilenchidae
20. Stylet knobs present; phasmid at the middle of the body; amphids pore-shaped, inside
view virtually invisible.......................................................................................................21
21. Mouth spear long, reaching to 40% of the distance from the front of the lip region to
the median bulb.................................................................................................Tylodoridae
21. Mouth spear shorter, at most to 1/3 of the distance to the median bulb......Tylenchidae
26. Lip region dorsally move strongly developed than ventrally, spicule shorter than 20
mm.......................................................................................................................Hoplotylus
26. Lip region symmetrical, spicule longer than 20 mm.................................Telotylenchus
32. Dorsal tooth in the oral cavity immobile, the two subventral swellings of the
metastoma are identical...............................................................................Diplogasteridae
32. The dorsal tooth is mobile and protrusible, the right swelling of the metastoma
contains a large tooth as well, the left swelling is smooth or carries a small toothed
plate.......................................................................................................Neodiplogasteridae
33. Body caterpillar-shaped, consisting of alternating wide and narrow rings; amphids
bladder-shaped..........................................................................................Desmoscolecidae
33. Body not consisting of alternating wide and narrow rings..........................................34
34. Body asymmetrical, the right half of the body displays a fine net structure and/or a
series of small shields or fins......................................................................Bunonematidae
34. Body symmetrical........................................................................................................35
Appendix 8 Key to Males 180
44. Oral cavity tubular, with stylet knobs, tail drawn out like a thread
..................................................................................................................Tylopharyngidae
44. Oral cavity without sytlet knobs, tail not drawn out like a thread...............................45
45. Amphids absent, oral cavity long and undifferentiated; large nematodes, longer than 3
mm, with a bluntly rounded tail and a conspicuously short esophagus, b ratio greater than
10........................................................................................................................Isolaimidae
45. Amphids present, b ratio smaller than 10, tail not bluntly rounded and body shorter
than 3 mm...........................................................................................................................46
50. The dorsal and subventral teeth are equally large, the part of the esophagus
surrounding the oral cavity is symmetrical.................................................Chromadoridae
50. The dorsal tooth is small, it is in fact a continuation of the esophageal tissue; the
subventral teeth are barely developed. The dorsal sector of the esophagus is wider than
the ventral sector in front.....................................................................Hypodontolaimidae
57. Bulb with valve apparatus. The esophagus does not continue into the intestines behind
the posterior bulb....................................................................................................Plectidae
57. Bulb if present, without valve apparatus. The esophagus has a tongue-shaped
extension behind the posterior bulb..............................................................Leptolaimidae
60. Oral cavity small with an inconspicuous small tooth at the base. Tail drawn out like a
thread.......................................................................................................Prismatolaimidae
60. Oral cavity large with sclerotized walls and one or a few large teeth, tail never drawn
out like a thread..................................................................................................................61
61. Base of oral cavity funnel-shaped, the dorsal tooth is not identical to the subventral
teeth; base of the esophagus contains no bladders..........................................Mononchidae
61. The base of the oral cavity is flattened, the dorsal tooth is identical to the two
subventral teeth; the base of the esophagus contains three bladders..............Anatonchidae
62. Oral cavity tube-shaped and longer than half of the actual esophagus.....Aulolaimidae
62. Oral cavity, if tube-shaped, shorter..............................................................................63
65. Amphids inconspicuous, chalice-shaped. Oral cavity closed at rest, but a tooth is
discernible in the oral cavity. Body spiral-shaped after heat fixation..................Tripylidae
65. Amphids round and conspicuous, oral cavity without teeth.....................Diplopeltidae
71. Tail with tail glands and a terminal drainage duct ......................................................72
71. Tail glands and drainage duct absent...........................................................................73
Appendix 8 Key to Males 183
72. Setae absent; amphid inconspicuous; oral cavity without cross ridges..........................
..................................................................................................................Rhabdolaimidae
72. Setae on the head and scattered over the body; amphids round; oral cavity with three
cross ridges...................................................................................................Linhomoeidae
84. Oral cavity large, with small teeth; right spicule degenerated......Odontopharyngidae
84. Oral small or absent, both spicules equally large........................................................85
Appendix 8 Key to Males 184
85. Body and head conspicuously slender (a ration greater than 60); amphid slit- or pore-
shaped, at several body diameters behind the front of the body...........................Alaimidae
85. Body not conspicuously slender, amphid not discernible in side view............................
......................................................................................Sphaeronema or Paratylenchidae
86. A conspicuous suction-cup-shaped lip disc at the anterior side of the body....................
........................................................................................................................Discolaimidae
86. No suction-cup-shaped outgrowth of the lip region.....................................................87
87. At the front end of the spear, there is a large oral cavity; on its base there are four
large teeth and on the walls a large number of small teeth and 24 longitudinal ridges.........
......................................................................................................................Actinolaimidae
87. Oral cavity with teeth lacking......................................................................................88
88. Spear and other body content barely discernible since they are obscured.......................
...............................................................................................................Diphtherophoridae
88. Body content easily visible..........................................................................................89
89. Tail long; c ratio smaller than 15 and cʼ ratio greater than 2........................................90
89. Tail short; c ratio greater than 15 and cʼ less than 2.....................................................95
94. Cuticle in the oral cavity swollen in the shape of a barrel, c ratio greater than 9............
....................................................................................................................Chrysonemoides
94. Cuticle in the oral cavity not swollen in the shape of a barrel, c ratio less than 9...........
.....................................................................................................................Prodorylaimium
95. Stylet and stylet extension fused into a curved spear; in front of the anus there are
three supplements, there are a few ventral papillae in the esophageal region. Bursa
sometimes present..........................................................................................Trichodoridae
95. Spear straight, no ventral papillae in esophageal region..............................................96
Appendix 8 Key to Males 185
98. Spear is inserted into the ventral wall of the oral cavity, is dorsally grooved, and with
a basal outgrowth.................................................................................................Sectonema
98. Spear axial....................................................................................................................99
100. Spicule longer than 80 µm; spear opening 30% of the spear length..........Axonchium
100. Spicule shorter than 70 µm; spear opening 50%; base of esophagus not surrounded
by a muscular sheath.......................................................................................Aporcelaimus
102. Spear curved like a sickle body conspicuously narrowed at the anterior side;
body width at the base of the esophagus is 4.5 times the width at the lips; guide ring
membranous, folded; twenty supplements of which one is isolated by itself........................
......................................................................................................................Paraxonchium
102. Body not conspicuously narrowed...........................................................................103
105. Spear embedded/inserted in right subventral wall of the oral cavity; base of
esophagus surrounded by a muscular sheath; supplements poorly developed......................
........................................................................................................................Nygolaimidae
105. Spear axial................................................................................................................106
107. Stylet continuation curved; esophageal tissue swollen behind the spear.......................
.......................................................................................................................Dorylaimoides
107. Stylet continuation not curved; esophagus not swollen distally..............................108
Appendix 8 Key to Males 186
108. Guide ring membranous; spear opening greater than 50%; gubernaculum absent;
three separate cuticular layers easily discernible........................................Aporcelaimidae
108. Tail cone-shaped, rounded or pointed, never broadly rounded................................109
110. Oral cavity shielded by six small shields; 19-21 grouped supplements; body longer
than 2 mm...........................................................................................................Labronema
110. Oral cavity without shields.......................................................................................111
111. Tail with a number of cuticular granules; lip region not set off; guidering consisting
of a muscular collar...................................................................................................Thonus
111. Tail without granules............................................................................Thornematidae
Translated from Dutch by Dr. Anton Baudoin from the excellent resource written by Dr.
Tom Bongers. (Used with permission.)
FA 4:1
Formalin (40% formaldehyde).........................................................................10 ml
Glacial acetic acid...............................................................................................1 ml
Distilled water..................................................................................top up to 100 ml
FA 4:10
Formalin (40% formaldehyde).........................................................................10 ml
Glacial acetic acid.............................................................................................10 ml
Distilled water...................................................................................................80 ml
FAA
95% ethanol......................................................................................................20 ml
Formalin (40% formaldehyde)...........................................................................6 ml
Glacial acetic acid...............................................................................................1 ml
Distilled water...................................................................................................40 ml
Formalin (2%)
Formalin (40% fromaldhyde).............................................................................5 ml
Glycerol..............................................................................................................2 ml
Distilled water...................................................................................................60 ml
FP 4:1
Formalin (40% formaldehyde)..........................................................................10 ml
Propionic acid.....................................................................................................1 ml
Distilled water...................................................................................................89 ml
Appendix 9 Formulas for Fixatives 188
Hechler’s Solution
Formalin (40% formaldehyde)...........................................................................4 ml
Glycerol..............................................................................................................6 ml
Distilled water...................................................................................................90 ml
Lactophenol
Liquid phenol................................................................................................20 parts
Lactic acid (45%)..........................................................................................20 parts
Glycerine.......................................................................................................40 parts
Distilled water...............................................................................................20 parts
Propionic-orcein
Orcein stain (natural or synthetic)......................................................................2.2 g
Glacial propionic acid..................................................................................100.0 ml
Boiled gently for 20 minutes (Warning: boiling can suddenly become violent)
Cool the solution to room temperature
Diluted with distilled water.............................................................................100 ml
Seinhorst I
Ethanol (95%)...............................................................................................20 parts
Glycerine .........................................................................................................1 part
Distilled water...............................................................................................79 parts
Seinhorst II
Ethanol (95%)...............................................................................................95 parts
Glycerine ........................................................................................................5 parts
TAF
Formalin (40% formaldehyde)...........................................................................7 ml
Triethanolaimine.................................................................................................2 ml
Distilled water...................................................................................................91 ml
Note: All solutions are very toxic and should be handled only with pro-
tective gloves, goggles, etc. and under a fume hood.