Practical Lesson 3
Extracting Nematodes from Plant Tissues
Direct Examination of Infected Plant Tissues
Symptomatic plant tissues are usually examined with the aid of a dissecting
(stereoscopic) microscope at magnifications of 10 to 50x to detect the presence of plantparasitic nematodes. Inverted compound microscopes are becoming more widely used.
1- Gently wash small roots, stems, leaves, buds, or other plant samples to remove loose
soil and other debris.
2- Transfer a washed sample to a plastic Petri dish or Syracuse dish containing 2 - 4 ml
water, for examination under a dissecting microscope.
3- While observing the operation through a dissecting microscope, cut the sample into
smaller pieces (1 - 2 cm long), and dissect galls, if present. Carefully slice, shred, or
dissect the tissues with strong dissecting needles or a scalpel (Fig. 6 A), and note the
presence and position of nematodes in the plant tissues. If no nematodes can be teasedout and none are seen on the bottom of the dish, reexamine it an hour or several hours
later.
Most plant-parasitic nematodes found in or on aboveground parts of plants
emerge in a few minutes and can be found moving about on the bottom of the
dish.
In root samples, large, sedentary endoparasitic nematodes may be seen attached to
the surface of roots or in dissected tissue. Semi-endoparasites and firmly attached
ectoparasites can be seen attached to the surface of roots (Fig. 6 A). Ectoparasitic
nematodes may be found on the bottom of the dish if the sample was gently and
not excessively washed (Fig.6 B).
4- In the shallow water near the center of the dish., loop an individual nematode over a
thin needle, such as a 00 insect mounting pin, dentist's pulp probe glued to the end of a
needle holder, or sharpened bamboo splinter (Figs. 7 and 8), and transfer it to a drop of
water on a glass slide (Fig. 8). Large, sedentary nematodes and cysts can be very
gently grasped with a fine forceps. Dissection and storage in a saline solution (0.9%
sodium chloride) helps to reduce the osmotic effect of water, which can cause female
nematodes to burst.
5- Prepare a temporary slide mount for viewing under a compound microscope at a
magnification of 100x or more to observe morphological and anatomical details
necessary to confirm the identity of the nematodes. A detailed description of the
preparation of temporary slide mounts is given in the practical lesson 4.
BAERMANN FUNNEL TECHNQUE
(See practical lesson No. 2.)
Fig. 6. Washed roots in water in a plastic Petri dish, viewed through a
dissecting microscope at a magnification of about 15x. Dissecting
needles are used to move the roots to better view free and embedded
nematodes. Needles, sharp pointed scalpels, and forceps are used to
remove endoparasitic nematodes: A) cyst nematodes attached to
roots. B) Ectoparasitic nematodes on the floor of the Petri dish.
(Shurtleff et al., 2000)
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Fig 7. Nematode transfer pick
prepared by splitting and sharpening a small section of bamboo from
just beneath the hard outer surface
of the cane. The section is placed in
a needle holder and brought-to a
fine point with a sharp knife,
scalpel, or razor blade. The final
sharpening is done under a stereoscopic dissecting microscope, until
the point consists of only two or
three fibers.
Fig 8. Bamboo pick used to transfer
vermiform nematodes from shallow water
in a Petri dish to a drop of water on a glass
slide: A) The tip of the pick is placed under
a nematode, slowly raised to the surface,
and then quickly removed to the water. A
magnification of 20x is optimal. B) The tip
of the pick, with the nematode, is placed in
a drop of water on a glass slide, and the
pick is moved in the water to dislodge the
nematode.
(Shurtleff et al., 2000)
Emergence in a Viewing Dish
Cut infected leaf or stem tissue into 8-mm or smaller squares, and cut roots into
pieces 8 mm long or less. Cover them with water in a shallow viewing dish.
Endoparasitic migratory nematodes will wiggle out of infected tissue and can be viewed
directly in the dish, often in less than an hour, or after soaking overnight. Species that can
be extracted by this technique include foliar nematodes (Aphelenchoides spp.), the
pinewood nematode (Bursaphelenchus xylophilus), burrowing nematodes (Radopholus
spp.), and the coconut red ring nematode (Rhadinaphelenchus cocophilus).
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Shaker agitation
Migratory endoparasitic nematodes (e.g., Pratylenchus) also emerge from
chopped roots in a flask or jar of water under continuous agitation in a shaker (motordriven, wrist-action, or other type). Sufficient agitation is required for adequate aeration.
The shaking may continue for several days. Extraction is improved if chopped roots are
shaken in water with ethoxyethyl mercuric chloride (2 ppm) and dihydrostreptomycin
sulfate (50 ppm).
TRITURATION
Trituration (Fig. 9) is excellent for extracting active nematodes from bulbs,
cloves, corms, storage roots, crowns, leaves, and small plants (e.g., strawberry). Also it
could be used to extract eggs of Meloidogyne spp. from infected roots. However, the
blending action may release toxic substances from the plant that can kill or immobilize
nematodes. Some blended tissues also form a thick, sticky mass that is difficult to sieve
and inhibits the movement of nematodes.
1. Place about 10 g of washed, diced plant material (2 - 3 cm long) in the jar of an electric
food blender, then add 200 ml of water, and run the blender at medium speed for 20 to
50 sec to triturate the tissue without undue damage to the nematodes.
Fig. 9. Extracting nematodes from plant parts by trituration.
(Shurtleff et al., 2000)
2. Pour the slurried mixture through a 20-mesh sieve nested over a 500-mesh sieve.
Thoroughly wash the slurry on the 20-mesh sieve with water from a fogger nozzle or
hose to remove the nematodes (Fig. 9 B). Discard the slurry on the 20-mesh sieve, as
the nematodes will be on the 500-mesh sieve.
3. Backwash the material on the 500-mesh sieve into a 50-ml beaker or viewing dish.
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Soaking in water
Migratory endoparasites (e.g., Pratylenchus and Radopholus), sedentary parasites
in immature stages, and male sedentary parasites emerge from roots soaked in water over
a period over several days.
1. Cut plant tissues such as leaves, stems, and roots (which should be free of debris and
soil) into pieces 5 - 10 cm long. Fibrous roots should be no more than 3 mm in
diameter. Roots of larger diameter and fleshy roots can be split longitudinally to help
nematodes emerge.
2. Place moist plant tissues loosely in a container that can be tightly closed (e.g., a small
bottle, screw-cap jar, Petri plate, or reclosable plastic bag). Add enough water to cover
the plant material. Seal and label the container.
3. Let the sealed container stand for three to four days at room temperature, about 2025°C .
4. Remove nematodes and add fresh water daily, until no more nematodes emerge. Shake
the container gently to wash the plant tissue, and decant the liquid over a coarse (48or 60-mesh) sieve nested over a 325- or 500-mesh sieve. Discard roots or other plant
material collected on the coarse sieve. Collect the nematodes on the fine sieve, holding
it at an angle of 35 to 45° over a 50-ml beaker or collecting dish. Tilt the sieve and
rinse it from the back with a small stream of water (10 to 20 ml) to wash the
nematodes and debris down the side of the sieve and into the beaker or dish.
The addition of 1 to 3% hydrogen peroxide (H2O2) and antibiotics to the water
improves extraction. Hydrogen peroxide raises the level of dissolved oxygen in
the water, while antibiotics inhibit the growth of bacteria and fungi.
Sodium Hypochlorite Method for Extraction of Nematodes eggs
Sodium hypochlorite (NaOCl) could be used to dissolve the egg masses of
Meloidogyne or cysts of Globodera and Heterodera to release eggs, but special efforts
are required to minimize the exposure to NaOCl which adversely affecting the viability of
eggs.
1- Obtain the following equipment:
A- 150- and 600- ml beakers.
B- 200- and 500 mesh (75 and 26 µ) sieves.
C- 5-ml dipper.
D- 0.5% Sodium hypochlorite (NaOCl).
E- Parafilm.
2- For extraction of Meloidogyne eggs, collect and cut infected roots into 1 – 2 cm
segments.
3- Put the root segments in a beaker containing the 0.5% NaOCl, Then tightly close the
beaker with parafilm.
4- Vigorously, shake the beaker manually or by an electric shaker for 2 – 3 minuets.
5- Quickly, pass NaOCl solution through a 200-mesh (75 µ) sieve, nested over a 500-
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mesh (26 µ) sieve to collect freed eggs.
6- Quickly, place the 500-mesh sieve with eggs under a stream of cold water to remove
residual NaOCl (for several minutes).
7- Standardize number of eggs per unit volume.
8- For extraction of Globodera and Heterodera eggs, the same procedure could be used,
but the time of exposure to NaOCl should need to be increased up to 8 – 10 minutes.
References
Ayoub, S.M. 1977. Plant Nematology, An Agricultural Training Aid. Academic Press.
California, 157 pp.
Barker, K.R. 1985. Nematode extraction and bioassays. Pp. 19-38 in: K.R. Barker, C.C.
Carter and J.N. Sasser (eds.). An Advanced Treatise on Meloidogyne. Vol. II.
Methodology. North Carolina State Univ. Graphs. Raleigh, N.C.
Hooper, D.J. 1986. Extarction of nematodes from plant materials. Pp. 51-58 in: J.F.
Southey (ed.). Laboratory Mthods for Working with Plant and Soil Nematodes. Sixth
ed. Her Majesty’s Stationary Office. London.
Shurtleff, M.C. and C.W. Averre III. 2000. Diagnosing Plant Diseases Caused by
Nematodes. American Phytopathological Society. St. Paul, Minnesota. 187 pp.
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