Front Cover
 Title Page
 Table of Contents
 Basic biology of plant parasitic...
 Symptoms of nematode damage
 Nematode extraction
 Direct examination of plant...
 Handling, fixing and staining...
 Estimation of nematode density
 Damage analyis
 Appendix 1 : Examples of nematode...
 Appendix 2 : Basic identification...
 Appendix 3 : Score sheets for measuring...
 Back Cover

Title: Practical plant nematology : a field and laboratory guide
Full Citation
Permanent Link: http://ufdc.ufl.edu/UF00077505/00001
 Material Information
Title: Practical plant nematology : a field and laboratory guide
Physical Description: Book
Language: English
Creator: Coyne, D. L.
Publisher: International Institute of Tropical Agriculture (IITA)
Publication Date: 2007
Subject: Africa   ( lcsh )
Farming   ( lcsh )
Spatial Coverage: Africa -- Benin
 Record Information
Bibliographic ID: UF00077505
Volume ID: VID00001
Source Institution: University of Florida
Holding Location: African Studies Collections in the Department of Special Collections and Area Studies, George A. Smathers Libraries, University of Florida
Rights Management: All rights reserved by the source institution and holding location.
Resource Identifier: isbn - 978-131-294-7


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Table of Contents
    Front Cover
        Front cover
    Title Page
        Page i
        Page ii
    Table of Contents
        Page iii
        Page iv
        Page v
        Page vi
        Page 1
        Page 2
    Basic biology of plant parasitic nematodes
        Page 3
        Page 4
        Page 5
        Page 6
        Page 7
        Page 8
        Page 9
        Page 10
    Symptoms of nematode damage
        Page 11
        Page 12
        Page 13
        Page 14
        Page 15
        Page 16
        Page 17
        Page 18
        Page 19
        Page 20
        Page 21
        Page 22
        Page 23
        Page 24
        Page 25
        Page 26
        Page 27
        Page 28
        Page 29
        Page 30
    Nematode extraction
        Page 31
        Page 32
        Page 33
        Page 34
        Page 35
        Page 36
        Page 37
        Page 38
        Page 39
        Page 40
        Page 41
        Page 42
        Page 43
        Page 44
        Page 45
        Page 46
        Page 47
        Page 48
        Page 49
        Page 50
    Direct examination of plant tissue
        Page 51
        Page 52
    Handling, fixing and staining nematodes
        Page 53
        Page 54
        Page 55
        Page 56
        Page 57
        Page 58
        Page 59
        Page 60
    Estimation of nematode density
        Page 61
        Page 62
    Damage analyis
        Page 63
        Page 64
        Page 65
        Page 66
    Appendix 1 : Examples of nematode genera and species known to be important crop pests worldwide
        Page 67
        Page 68
        Page 69
        Page 70
        Page 71
    Appendix 2 : Basic identification of nematodes
        Page 72
        Page 73
        Page 74
    Appendix 3 : Score sheets for measuring nematode damage
        Page 75
        Page 76
        Page 77
        Page 78
        Page 79
        Page 80
        Page 81
        Page 82
        Page 83
    Back Cover
        Page 84
Full Text

Practical plant nematology:

A field and laboratory guide

D.L. Coyne, J.M. Nicol and B. Claudius-Cole

M.'B.M ,r,, 1, Rf5seaJc 4 &NowisVN nh Afi



This guide has been produced by the International Institute of Tropical Agriculture (IITA) and
the International Maize and Wheat Improvement Center (CIMMYT) as part of the strategy of
the Systemwide Program on Integrated Pest Management (SP-IPM) to improve the quality and
usefulness of pest management research. IITA, CIMMYT and the SP-IPM are supported by the
Consultative Group on International Agricultural Research (CGIAR; www.cgiar.org). Funding
for production of the guide was provided by the Technical Centre for Agricultural and Rural
Cooperation (CTA).

The SP-IPM is a global partnership programme whose task is to draw together the IPM efforts
of the international agricultural research centers and their partners and to focus these efforts
more clearly on the needs of resource-poor farmers in the developing world. The programme
tackles those areas where research promises to provide solutions to pressing problems in
sustainable agricultural development but where impact has so far been limited usually due to
fragmentation of efforts among different organizations or in different regions of the world, or
due to inadequate links between researchers and farmers. The SP-IPM expects to achieve rapid
progress by alleviating such constraints, breaking down barriers to information exchange, filling
research gaps where necessary and developing effective models of researcher-extensionist-
farmer partnerships to promote adoption of IPM technologies.

IITA is an Africa-based international research-for-development organization, established in
1967, and governed by a board of trustees. IITA's vision is to be one of Africa's leading research
partners in finding solutions for hunger and poverty. It has more than 100 international
scientists based in various IITA stations across Africa. This network of scientists is dedicated
to the development of technologies that reduce producer and consumer risk, increase local
production, and generate wealth.

CIMMYT's mandate is to act as a catalyst and leader in a global maize and wheat innovation
network that serves the poor in developing countries. Drawing on strong science and effective
partnerships, CIMMYT creates, shares, and uses knowledge and technology to increase food
security, improve the productivity and profitability of farming systems, and sustain natural

CTA's tasks are to develop and provide services that improve access to information for
agricultural and rural development, and to strengthen the capacity of African, Caribbean and
Pacific (ACP) countries to produce, acquire, exchange and utilise information in this area.

Practical plant nematology:
A field and laboratory guide

D.L. Coyne, J.M. Nicol and B. Claudius-Cole


Rmipm IITN
Rawrh to NN~Muh Al;,m


The authors are grateful to the SP-IPM and CTA for their generous financial support to produce
this guide. SP-IPM core donors who contributed to the development and production of this guide
are the Governments of Norway, Switzerland and Italy.

Dr BraimaJames (SP-IPM Coordinator) provided the impetus and continued support to preparing
this guide that we hope satisfies the demands of our partners and proves useful in increasing
awareness of and reducing nematological problems. We thank Braima for his continued support
in this.

Dr John Bridge's guidance and numerous contributions during the development of this guide are
particularly appreciated.

Thanks to Luma Al-Banna, John Bridge, Roger Cook, Don Dickson, Jon Eisenback, Georg
Georgen, Dieter Heinicke, Sean Kelly, Sandra Mack, Mariette Marais, Alex McDonald, Hans
Meerman, Edward Oyekanmi, Richard Plowright, Roger Rivoal, Richard Sikora, Paul Speijer,
Hugh Wallwork, Esther van den Berg and Vivien Vanstone for use of photographic images.
Thanks also to R. Esser for use of his figure, and to Graham Stirling for use of figures and related
reference material for the appendices.

Editing assistance from Mr Duncan Scudamore and Ms ElifSahin is gratefully acknowledged.

ISBN 978-131-294-7

2007 International Institute of Tropical Agriculture

All rights reserved. The publisher encourages fair use of this material provided proper citation
is made. No reproduction, copy or transmission of this report may be made without written
permission of the publisher.

Correct citation:
Coyne, D.L., Nicol, J.M. and Claudius-Cole, B. 2007. Practicalplant nematology: afield and
laboratory guide. SP-IPM Secretariat, International Institute of Tropical Agriculture (IITA),
Cotonou, Benin.

A PDF version of this document is also available on the CIMMYT, IITA and SP-IPM


Preface v

Introduction I

Basic biology of plant parasitic nematodes 3
Appearance and structure 3
Life cycle 3
Nematode types 3
Migratory endoparasites 7
Sedentary endoparasites 8
Ectoparasites 9

Symptoms of nematode damage I I
Above-ground symptoms I I
Symptoms caused by aerial nematodes I I
Symptoms caused by root nematodes I I
Below-ground symptoms 14
Root galls 14
Root galls versus root nodules 18
Abbreviated root systems 19
Root and tuber lesions 19
Root rot and tuber rot 21
Cracking 23
Cysts or'pearly root' 24

Sampling 25
Sampling tools 25
Number of samples 25
Sampling pattern 26
Time of sampling 26
Taking soil samples 27
Taking root samples 28
Taking samples from above-ground plant tissue 28
Care of samples 28

Nematode extraction 31
Choosing an extraction method 31
Preparation of samples 32
Labeling 32
Extraction tray method 34
Root maceration method 40
Sieving method 42
Incubation method 48

Practical plant nematology

Direct examination of plant tissue 51

Handling, fixing and staining nematodes 53
Handling nematodes 53
Picking nematodes 53
Sending nematodes for identification 55
Nematode identification services 57
Killing nematodes 57
Fixing nematodes 58
Killing and fixing in one step 59
Preserving sedentary nematodes in root or tuber tissue 59
Staining 60
Meloidogyne egg masses 60

Estimation of nematode density 61
Counting nematodes 61

Damage analysis 63
Scoring of nematode symptoms on plants 63

References and further reading 65

Appendix I. Examples of nematode genera and 67
species known to be important crop pests worldwide

Appendix 2. Basic identification of nematodes 72

Appendix 3. Score sheets for measuring 75
nematode damage
Root-knot gall (Meloidogyne spp.) scoring on cassava 75
Root-knot gall scoring on carrot 77
Root-knot gall scoring on lettuce 78
Lesion scoring for banana roots 79
Diagrammatic root-knot scoring chart 81
Cyst damage scoring sheet for wheat 82


This guide is a simple, easy-to-follow reference for assessing plant parasitic nematode
problems. It provides clear instructions, with many illustrations, on procedures for
collecting and processing samples for nematode assessment, as well as information
on accessing further identification and diagnosis support. The manual is aimed
at technicians, field workers, extension agents and others with an interest in crop
production and crop protection, particularly in those parts of the world where access
to expert help and advanced facilities is limited. It has been produced in response to
frequent demand from colleagues for a guide to aid diagnosis of nematode problems.
This guide will hopefully simplify some aspects ofnematology, and help to lessen
the mystery surrounding this crop production problem.

It is sometimes said that nematodes are viewed as a crop production problem only
when a nematologist is present, and that without the nematologist there would be
no nematode problem. Paradoxically, the unspecific symptoms of nematode damage
are often attributed to other causes, which may seem more likely or more obvious.
The reality is that a number of constraints often combine to reduce crop production,
and it is necessary to quantify all of the main constraints, including nematodes.
Keeping the nematode threat in perspective, in relation to other pests and diseases,
is a challenge, but one that will benefit enormously from better quantification of the
nematode problem through improved field and laboratory procedures.

Plant parasitic nematodes are ever-present and are incidental with plant growth and
crop production. They are significant constraints to sustainable agriculture and can
be difficult to control. Determining the importance of individual nematode species,
nematode communities and nematodes in combination with other problems is not
a simple task at the best of times, but is more difficult in tropical than in temperate
climates. Species previously not known to cause crop damage are continually being
discovered, particularly as agriculture changes to suit changing needs, and new crops
are introduced. Introduction of, or improvements in, nematological techniques and
diagnostics can lead to identification of nematodes as the cause of a problem which
had been present for many years, but through lack of local expertise had not been
properly diagnosed.

Much remains to be learned about nematodes and the damage they cause to crops.
There is, for example, a lack of reliable data on the relationship between nematode
numbers and yield for many different crops and types of nematodes. In many less
developed countries much basic information simply does not exist. It is therefore
important that even small developments and knowledge gains are recorded for future
use, through publications of the relevant networks and societies, or regional or
international journals.

Practical plant nematology

We hope that this guide will contribute to improving pest and disease management,
particularly where nematological expertise is scarce, such as in the less developed
countries of the world. An initial step to nematode management is establishing
their presence through collection and relation with symptoms, and with expert help
accurately identifying the species involved. This guide aims to support that initial step.

Danny Coyne IITA Nematologist, Uganda.

Julie Nicol CIMMYT Int. Nematologist, Turkey

Biodun Claudius-Cole Lecturer, Department of Crop Protection and
Environmental Biology, University of Ibadan, Nigeria.


Nematodes are a diverse group of worm-like animals. They are found in virtually
every environment, both as parasites and as free-living organisms. They are generally
minute, but some species can reach several meters in length. This guide focuses
specifically on plant parasitic nematodes, which are very small or microscopic, can
cause significant damage to crops, and are extremely widespread (Appendix 1).

Because nematodes are difficult or impossible to see in the field, and their symptoms
are often non-specific, the damage they inflict is often attributed to other, more
visible causes. Farmers and researchers alike often underestimate their effects. A
general assessment is that plant parasitic nematodes reduce agricultural production
by approximately 11% globally (Agrios, 2005), reducing production by millions of
tonnes every year.

The amount of damage nematodes cause depends on a wide range of factors, such
as their population density, the virulence of the species or strain, and the resistance
(ability of the plant to reduce the population of the nematode) or tolerance (ability
of the plant to yield despite nematode attack) of the host plant. Other factors also
contribute to a lesser extent, including climate, water availability, soil conditions,
soil fertility, and the presence of other pests and diseases. However, although we have
some knowledge on the nematode-crop relationship and influencing factors, much
remains to be learned. Damage thresholds for nematodes on various crops in various
parts of the world, for example, are often unknown, and the threat nematodes pose
often requires an educated guess.

Once nematodes are identified as significantly contributing to crop damage,
management options can then be decided. These depend on the nematodes involved,
the crop and cropping system, and local circumstances. If the species is identified
some specific interventions can be considered, for example, using resistant crops
or varieties. Otherwise more general options or combinations of options, such as
rotation, chemical application, biological control and sanitation practices, may be

The aims of this guide are to assist the reader to:
* Appreciate basic nematode biology
* Recognize the different groups and feeding behaviors of nematodes
* Identify damage symptoms
* Collect nematode samples
* Process the samples
* Dispatch nematodes and samples for more precise identification.

Practical plant nematology

On seeing suppressed growth/decreased production
in crop plants

Stage I: Look for and assess symptoms of nematode damage

Stage 2: Collect soil and plant tissue samples
Stage 2: Collect soil and plant tissue samples

Stage 3: Extract nematodes from samples

Stage 4: Identify nematodes

Stage 5: Nematode density assessment

Stage 6: Nematode damage analysis

Stage 7: Management decision

Figure 1. The stages in nematode assessment and management.

Figure 1 shows the stages in nematode assessment and management. This guide
provides information on carrying out stages 1-3. Stage 4 may also be attempted,
but assistance from a specialist nematologist will probably be needed, and will
certainly be required for the final stages. The References and Further Reading section
(page 65) includes some useful publications to take nematology tasks beyond those
described in this guide.

Basic biology of plant parasitic nematodes

Appearance and structure
Plant parasitic nematodes are mostly thread-like worms ranging from 0.25 mm to
>1.0 mm long, with some up to 4.0 mm. Although most taper toward the head
and tail, they come in a variety of shapes and sizes (Fig. 2). Females of some species
lose their worm-like shape as they mature, becoming enlarged and pear-, lemon- or
kidney-shaped or spherical as adults.
Like all animals, nematodes have circulatory, respiratory and digestive systems
(Fig. 3). Plant parasitic nematodes differ from nematodes that feed on bacteria and
fungi in that they have a specialized feeding structure, the spear or stylet (Fig. 3).
This is used to inject enzymes into plant cells and tissues and then to extract the
contents, in a similar way that aphids feed on plants.

Life cycle
The nematode life cycle is typically divided into six stages: the egg, four juvenile
stages and the adult (Fig. 4). The duration of any of these stages and of the
complete life cycle differs for different species, and also depending on factors such
as temperature, moisture and plant host. Under favorable conditions in the tropics
many species have relatively short life cycles, with several generations possible per
season. This can lead to rapid population build up from just one (if self-fertilizing)
or two nematodes.

Nematodes can survive unfavorable conditions, such as a dry season or a cold winter.
Different species survive best at different life stages, for example Heterodera species
survive best as eggs encapsulated within cysts, Ditylenchus species as fourth stage
juveniles, and Anguina species as second stage juveniles.

Nematode types
Plant parasitic nematodes can be separated into aerial parasites those feeding on
above-ground parts of plants and root and tuber parasites those feeding on
below-ground parts.

They can also be grouped by their feeding behavior and motility into three main
* Migratory endoparasites mobile nematodes that feed inside the plant root
* Sedentary endoparasites nematodes that, once they have reached a feeding site
inside the plant, cease to be mobile and feed from a fixed location.
* Ectoparasites nematodes that feed on the plant from the outside of the plant.

Practical plant nematology

Pratylenchus (worm-like/
vermiform) [JB]

Nacobbus (swollen/fusiform) [JB]

Helicotylenchus (worm-like
vermiform/spiral) [GG]

Ah iiiiiiiiiiii (swollen/fusiform) JB
Achlysiella (swollen/fusiform) [JB]

Discocriconemella (slightly


Tylenchulus (pear-shaped) [JB]

Rotylenchulus (kidney-shaped/
reniform) [JB]

Scutellonema (worm-like
vermiform/C-shaped) [GG]

Heterodera (lemon-shaped)

Criconematid (ribbed/ridged)

Meloidogyne (spherical/gourd-
shaped) [JB]

Hirschmanniella vermiformm/
long) [JB]

appearance on
outside of root

Ogma surface structure (flanged/flared ribs) [EvB]

Figure 2. Diverse shapes of nematodes, as seen under the microscope.
(Photographs by J.Bridge [JB], G.Goergen [GG] or E. van den Berg [EvB].)

Basic biology of plant parasitic nematodes



Typical nematode structure (courtesy R. Esser).



Male and female Scutellonema.

Figure 3. (Top) Diagram of nematode structure as observed through a microscope.
(Bottom) Example of male and female nematode (Scutellonema bradys) as seen
under the microscope with male spiculee) and female (vulva and ovary) reproductive
organs indicated. Note that not all nematode species have males. A protective bursa
is found around the spicule on males of some species, as seen here, but not all species.
(Photograph by H. Meerman.)

Practical plant nematology

3'd juvenile stage

2nd juvenile stage (invasive)
1st juvenile stage
(in egg)

Infected maize root

4th juvenile stage

Healthy maize root
Healthy maize root

Adult female

Adult male

Egg mass and adult female

Figure 4. Nematode life cycle. Meloidogye used as an example on maize, as observed through the microscope.
Not all stages are to the same scale. Migratory nematode lifecycles follow a similar course, but remain wormlike
and do not produce egg masses.
(Photographs by E. Oyekanmi.)


Basic biology of plant parasitic nematodes

Migratory endoparasites (Fig. 5)
All life stages of migratory endoparasitic nematodes are mobile except the egg. The
nematodes burrow through the plant from cell to cell, or may leave the plant tissue
in search of new feeding sites. Whilst feeding they commonly lay eggs both inside
the plant cortical tissue and also in soil surrounding the root tissue. Damaged cells
release toxins which kill neighboring cells, resulting in small spots or lesions of
necrotic tissue. Root rot fungi and bacteria are often associated with infestations
of migratory endoparasitic nematodes, which enter the plant tissues through areas
damaged by nematodes.

it a

Scutellonema bradys in yam.

root cortex

- central stele

Hirschmanniella in rice [JB].

Figure 5. Migratory endoparasitic nematode female and eggs stained red in root tissue.
(Photograph by J. Bridge [JB].)

Practical plant nematology

Sedentary endoparasites (Fig. 6)
Sedentary endoparasitic nematodes invade plant tissue usually as newly hatched
second-stage juveniles the 'infective' wormlike stage. They move through the soil
to locate host roots, and then through the plant tissue to find a feeding site. At the
feeding site the female develops, remaining permanently sited for the duration of her
life. As she develops, her body swells to a spherical, lemon, kidney, or ovoid form.
The nematode feeds on a relatively small number of cells, which are regulated by the
nematode with growth substances. Some groups (e.g. cyst and root-knot nematodes)
cause 'giant' feeding cells to form in the host plant.

The males remain wormlike, feeding on the surface of the root for a few days, during
which they may or may not fertilize the females before moving into the soil where
they die.

Female sedentary endoparasitic nematodes generally produce a large number of eggs,
which remain in their bodies (e.g. cyst nematodes Heterodera spp.) or accumulate
in egg masses (e.g. root-knot nematodes Meloidogyne spp.) attached to their
bodies. Some other nematodes are sedentary, but only semi-endoparasitic, such as
the reniform (Rotylenchulus spp.) and citrus (Tylenchulus semipenetrans) nematodes,
which become only partly embedded in the root tissue.

Cyst nematode (Heterodera spp.)
breaking open the root cortex of

Adult cereal cyst nematode
(Heterodera filipjevi).

Root-knot nematodes (Meloidogyne
spp.) bursting and protruding out
of maize root.

Reniform nematode
(Rotylenchulus spp.) with head
buried in root tissue (semi-
sedentary endoparasite) [JB].

Root-knot nematode
(Meloidogyne spp.) embedded
in gourd root with eggs
released outside the root [JB].

Root-knot nematode (Meloidogyne
spp.) embedded in sweet potato
root [JB].

Figure 6. Endoparasitic nematodes.
(Photographs by J. Bridge [JB].)

Basic biology of plant parasitic nematodes

Ectoparasites (Fig. 7)
Ectoparasitic nematodes feed externally, on the surface of the plant, usually on root
hairs or cortical tissue. They are often found in high densities, but do not always pose
a problem. However, they may cause serious damage if the plant is suffering from
other biotic or abiotic stresses (e.g. fungal attack or low water availability). Examples
ofectoparasitic nematodes are ring nematodes (Criconemoides spp.), spiral nematodes
(Helicotylenchus spp.) and the aerial rice white-tip nematode (Aphelenchoides besseyi).

It is well recognized that some ectoparasites transmit plant viruses, for example some
species of dagger nematodes (Xiphinema spp.), needle nematodes (Longidorus spp.)
and stunt nematodes (Trichodorus and Paratrichodorus spp.).

Aulosphora feeding on rice root with
close-up of feeding stylet penetrating
root tissue.

Discocriconemella with close-up of head
region [GG].

-- -.
Tylenchorhynchus feeding at the
root tip [JB].

Figure 7. Ectoparasitic nematodes.
(Photographs by G. Georgen [GG] and J. Bridge [JB].)

Symptoms of nematode damage

One of the major challenges in identifying nematodes as the causal agent of crop
damage is the fact that many of them do not produce highly diagnostic symptoms,
which are specific and easy to identify. The damage caused by nematodes is often
non-specific and easily confused with symptoms of other abiotic or biotic stresses.
For example, chlorosis may be due to nitrogen deficiency or may be caused by
nematodes; poor stands of growth similarly may be caused by poor soil fertility or
moisture stress, or may be due to nematodes.

It is therefore highly recommended to assess for nematodes when crops are suffering
yield loss and exhibiting any of the symptoms described below. Additional knowledge
on the crop, cropping history, and management practices, combined with information
in this guide, may also indicate the possible nematode(s) involved.

Symptoms of nematode damage are found both above and below ground.

Above-ground symptoms
Above-ground symptoms fall into two categories: those caused by aerial nematodes
attacking foliage and those caused by root nematodes attacking plant roots.

Symptoms caused by aerial nematodes (Fig. 8)
These are often specific symptoms associated with the nematode pest and therefore
may be diagnostic. They include:
* Gall formation, or abnormal swelling of seeds (e.g. Anguina) or leaves (e.g.
* Leaf stripe, bleaching and discoloration of leaves (especially in temperate
climates) (e.g. Aphelenchoides)
* Swollen, crinkled and disorganized tissue growth (e.g. Ditylenchus)
* Internal stem necrosis, signified with a red ring (Bursaphelenchus cocophilus)
* Inflorescence necrosis
* Chlorosis/browning of leaves (needles in pines) and eventual tree death
(Bursaphelenchus xylophilus).

Symptoms caused by root nematodes (Fig. 9)
Root nematodes almost always cause varying degrees of abnormal above-ground
growth, but these symptoms alone are generally not enough to diagnose a root
nematode problem. Most symptoms reflect or can be mistaken for other problems,
such as reduced water uptake or disturbed mineral absorption. They include:
* Chlorosis (yellowing) or other abnormal coloration of foliage
* Patchy, stunted growth
* Thin or sparse foliage
* Symptoms of water stress, such as wilting or leaf rolling

Practical plant nematology

\b\ I I I

Deformed heads of barley and
wheat due to seed gall nematode
Anguina tritici [RS].

Crinkling/twisting of White tip disease of rice caused
rice leaves by Ditylenchus by Aphelenchoides besseyi [JB].
angustus [JB].

Red ring symptoms (Bursaph-
elenchus cocophilus) in coconut
trunk [JB].

Ufra disease on rice caused by
Ditylenchus angustus [JB].

Bleaching and streaking discoloration
of enset (Musa) leaf caused by
Aphelenchoides sp. [PS].

Oats severely infected with stem nematode causing patchy growth,
stunting, chlorosis (left) and basal swelling (right) associated with
Ditylenchus dipsaci infection [HW].

M-.. 0 -Jm,- 1; I
Seed gall of wheat caused by Anguina
tritici (crushed seed showing emerging
eggs and juveniles) [JB].

Figure 8. Above-ground symptoms caused by aerial nematodes attacking foliage.
(Photographs by J. Bridge [JB], R. Sikora [RS], P. Speijer [PS] and H. Wallwork [HW].)

Symptoms of nematode damage

Chlorosis and stunted Patchy growth and chlorosis of lower Delayed flowering/maturity and
growth of rice plants leaves of wheat caused by Heterodera spp. suppressed growth in a patch of solanum
(left) caused by [HW]. potatoes affected by potato cyst nema-
Heterodera sacchari. todes [DH].

Thin and sparse foliage of irrigated Stunting/reduced height of
rice caused by Hirschmanniella spp. plantain (plants on left) caused by
Pratylenchus coffee.

Patchy distribution and reduced
tillering in wheat attacked by the
root lesion nematode (Pratylenchus
neglectus) [RR and RC].

Patchy distribution, stunted and
chlorotic maize plants affected by root-
knot nematode (Meloidogyne spp.)

Dieback of citrus caused by Toppling over of banana caused
Radopholus similis. by Radopholus similis.

Figure 9. Above-ground symptoms caused by root nematodes attacking plant roots.
(Photographs by R. Cook [RC], D. Heinicke [DH], A. McDonald [AM], R. Rivoal [RR] and H. Wallwork [HW].)

Practical plant nematology

* Die-back of perennial or woody plants with little or no new foliage
* Reduced fruit and seed size
* Low yields.

Other symptoms that may suggest root nematode infection are:
* Failure to respond normally to fertilizers
* A tendency to react to water stress more rapidly than healthy plants, and slow
recovery from wilting
* Little or no new foliage development at the onset of a new growing season
* Severe weed problems (higher density of weeds), due to the nematode-infected
plant being less able to compete with weeds
* Greater disease incidence, because of suppressed resistance of nematode-infected

Below-ground symptoms
These are due to root nematodes, and may be specific enough to allow diagnosis of
the root nematode problem. Uprooting of plants or excavation of roots is needed to
observe symptoms. Symptoms include:
* Galling
* Shortened, stubby or abbreviated roots
* Root lesions
* Root or tuber necrosis, rotting or death
* Root or tuber cracking
* Cysts or 'pearly' root
* Deformed roots
* Altered root architecture.

Root galls
Root galls are caused mostly by the root-knot nematodes (Meloidogyne spp.),
although other nematodes such as Nacobbus aberrans may also cause galling (Fig. 10).
Feeding by some nematodes, such as Xiphinema spp., may result in swellings or less
defined galls, often at the root tips.

The galls can vary considerably depending on the Meloidogyne species, the crop and
cultivar, and whether occurring on roots or on tubers (Fig. 11). Typical appearance
* Small individual bead-like swellings
* Massive clumps of fleshy tissue stuck together
* Swollen root tips
* Irregular swellings along the root
* Hook-shaped root tips
* No form of visible root swelling other than a raised surface where the nematode
is embedded.

Symptoms of nematode damage

Meloidogyne spp. knobbling of
cassava roots.

Meloidogyne graminicola on rice causing stunted and galled
seedling roots [RP] and characteristic hooked root tips [JB].

Massive clumping and galling of root tissue in vegetables caused by
Meloidogyne spp.

Swollen maize root tips
caused by Meloidogyne spp.

Large woody galling on tree roots
caused by Meloidogyne spp.

Figure 10 continued overleaf

Practical plant nematology

Deformed banana root system with swollen roots (left) and cross section of banana
root which has raised root surface with embedded female Meloidogyne spp. nematodes
(highlighted in split root right).

Bead-like galls on lettuce caused by Meloidogyne spp.

Deformation, raised galls and root termination of maize roots caused by Meloidogyne spp.

Galling of potato roots caused by
Nacobbus aberrans [JB].

Figure 10. Root galls and other symptoms of root-knot nematode (Meloidogyne spp.) and Nacobbus sp.
(Photographs by J. Bridge [JB] and R. Plowright [RP].)

I -- ~s.e

Symptoms of nematode damage

It is a common misconception that plants of the family Graminaceae (grasses and
cereals) are not affected by root-knot nematodes. In fact, these species are readily
affected but galling is often less visible. Small galls, however, can usually be easily
observed (Fig. 10).

Tuber galls on yam (Discorea spp.).

Crenulated raised yam tuber surface with cross-
section revealing female nematodes embedded
in the tissue.

Galling on potato.

Galls on beetroot [RS].

Galls on cassava roots.

Galls on carrot.

Figure 11. Galls on storage organs caused by root-knot nematode (Meloidogyne spp.).
(Photograph by R. Sikora [RS].)


Practical plant nematology

Root galls versus root nodules (Fig. 12)
Other root swellings can be caused by beneficial nitrogen-fixing Rhizobium bacteria.
These are called nodules, and are commonly seen on the roots of legume crops. They
are distinguished from root galls by their contents, and by differences in how they
are attached to the root. Fresh nodules will have a milky pink to brown liquid inside

Root-knot nematode galls on cassava and lettuce (two pictures left) compared with beneficial nitrogen-fixing root
nodules (group on right) on soybean, bean and groundnut.

Galling-like root symptoms caused by
cabbage clubroot disease on cabbage.

Figure 12. Root galls caused by nematodes, compared with other gall-like symptoms.

Symptoms of nematode damage

them, whereas galls tend to be composed of a gelatinous clear creamy solid nature,
which is sometimes tough. Nodules are attached loosely to the root and can be easily
removed; root-knot galls originate from within the root, and removing them is more
difficult and will tear the root cortex.

Some root diseases such as the fungus clubroot (Plasmodiophora brassicae) can result
in deformed root systems, which can resemble root-knot damage (Fig. 12).

Abbreviated root systems
Nematode activity can also cause a shortening of the roots, so that the root mass is
greatly reduced or has a stubby appearance (Fig. 13).

Stunted and galled rice seedling
roots caused by Meloidogyne
graminicola [RP].

Severely abbreviated root system of olive
tree saplings resulting from root lesion
nematode (Pratylenchus spp.) infection.

Stubby root symptoms on maize, caused by Paratrichodorus minor [DD].

Figure 13. Root shortening.
(Photographs by D. Dickson [DD] and R. Plowright [RP].)

Root and tuber lesions
Roots and tubers may show areas of dead tissue (necrosis) as a result of nematode
activity (Figs 14 and 15). As the nematodes feed and migrate within the roots they
destroy plant cells and also disrupt cellular functions causing the tissue to die.

Practical plant nematology

Heavily lesioned strawberry roots
infected with root lesion nematode
(Pratylenchus vulnus) [JB].


Symptoms of root lesion
nematode (Pratylenchus
',., ... on wheat roots.

Maize roots infected with
K Pratylenchus lesion nematodes
(right) compared with uninfected
roots (left).

Banana roots showing extended
root lesioning (necrosis) caused
by Radopholus similis.

Figure 14. Lesion symptoms on roots.
(Photograph by J. Bridge [JB].)

Internal lesions and necrosis exposed
below the surface of yam due to the yam
nematode (Scutellonema bradys).

Lesions on sweet potato caused by
Scutellonema bradys.

Lesions on sweet
potato caused by
Paratrichodorus (middle
tuber) [MM].

Internal lesions on sweet
potato due to root-knot
Inematode (Meloidogyne
incognita) [JB].

Figure 15. Necrosis and lesions on storage organs.
(Photographs by J. Bridge [JB] and M. Marais [MM].)

Symptoms of nematode damage

Root rot and tuber rot
Nematodes alone can cause rotting of roots and tubers through extensive burrowing
leading to substantial necrosis and tissue and root death (Figs 16 and 17). The
banana burrowing nematode Radopholus similis, the root lesion nematodes
Pratylenchus spp., yam nematode Scutellonema bradys, and Hirschmanniella miticausa
on taro are examples. Frequently, secondary fungal and bacterial infections also
develop and contribute to rotting (Fig. 17).

Rot, surface cracking (left) and termination (right) of banana roots caused by a
combination of Radopholus similis, Helicotylenchus multicinctus and Meloidogyne spp.

Necrosis and reduction of sweet potato roots due to
Scutellonema bradys.

Figure 16. Root rot caused by nematode infection.

Practical plant nematology

Sub-surface necrosis on strips of cassava
storage root, caused by infection with root-
knot nematode (Meloidogyne spp.), compared
to clean strip on left.

Tuber rot of sweet potato infected
with root-knot nematode
(Meloidogyne spp.) [JB].

Internal rotting in yam tuber caused by fungal infection,
probably entering as a result of Scutellonema bradys damage to
the cortex.

Miti miti disease on
cocoyams caused by
Hirschmanniella miticausa

Dry rot of yams caused by the yam nematode
(Scutellonema bradys).

Figure 17. Tuber rot caused by nematode infection.
(Photographs by J. Bridge [JB].)

Symptoms of nematode damage

Roots and tubers sometimes develop a cracked surface following nematode infection
(Fig. 18). This symptom may be blamed on water or nutrient stress during growth.
Cracking is often seen in sweet potato tubers, caused by the reniform nematode
(Rotylenchulus reniformis). The yam nematode can cause the same symptom on yam.

Flaking and cracking of yam tuber surface (left) caused by Scutellonema

Surface cracking of potato tuber
caused by Scutellonema bradys.

Cracking of sweet potato tuber by Rotylenchulus spp.

Figure 18. Tuber cracking symptoms.

Practical plant nematology

Cysts or'pearly root' (Fig. 19)
Cyst nematodes (e.g. Heterodera and Globodera spp.) can often be observed on the
roots of their hosts without magnification, if the soil is gently tapped off and the
observer looks very carefully. The young adult females are visible as tiny white beads,
giving a pearly appearance when many are present. As the females mature, the cysts,
which can contain hundreds of eggs, harden and turn brown or black.

Dark brown cyst and white female
Heterodera oryzicola embedded in rice
Sroot tissue [RP].

White females

White females of the Heterodera avenue
cereal cyst nematode, each the size of
a pin-head, causing knotting of wheat
roots (left) before they mature to a
brown cyst, shown on the right spilling
eggs [HW].

Potato roots with white female Globodera rostochiensis
attached along its surface giving it an appearance of
white beads attached or being 'pearly' [JB].

Figure 19. Cyst or 'pearly root'.
(Photographs by J. Bridge [JB], R. Plowright [RP] and H. Wallwork [HW].)


Having observed symptoms that indicate possible or likely nematode infestation,
the next stage is to collect samples from the affected plants and from the soil around
the roots. These are then taken to the laboratory for analysis, to determine what
nematodes are present and possibly their density.

The following field characteristics have implications for the sampling method, and
should therefore be considered at this stage:
* Aggregated distribution of nematodes due to host root system and the seasonal
behavior of the nematode
* Crop type and history
* Areas planted to different varieties
* Soil moisture
* Soil compaction
* Soil type
* Temperature and seasonal changes.

Sampling tools
Useful tools for sampling, some of which are shown in Fig. 20, include a spade, a
hand trowel, a screwdriver, a soil auger corerr), knives (for cutting roots), scissors,
polythene sample bags, tags. Also marker pens for labeling the sample bag and a
pencil and notebook for recording information. The soil auger or corer should have
a blade 20-30 cm in length and 20-25 mm diameter, and can be either a complete
cylinder or a half cylinder. Half cylinders help in removing soil from the corer.

Figure 20. An assortment of tools for sampling nematodes.

Number of samples
Take enough samples to ensure they are representative of the situation in the field.
The greater the number ofsub-samples/cores combined for each field sample, the
more accurate the assessment will be. A balance between available time and resources
is, however, necessary.

Practical plant nematology

The sampling procedure and number of samples taken should allow for nematode
variation or aggregation. From an area of 0.5 to 1 hectare, take a minimum of 10
core sub-samples, and even as many as 50. Combine these to make one composite
sample to represent the field area sampled. Bulking of samples in this way helps to
preserve them by maintaining the temperature and moisture of samples.

Sampling pattern
Nematodes are rarely distributed evenly in a field, and samples should therefore be
collected from several areas within the field. Collect separate samples from both the
poor growth areas and an area of relative good growth, where this is obvious, for
comparison. Maintain a consistent sampling style and pattern during surveys and
experiments to enable meaningful comparisons between fields, plots, treatments, etc.

Sampling patterns can be random or systematic (Fig. 21). Random sampling does not
accommodate the patchy nature of nematode distribution, and is only representative
if the sampling area is small. Systematic sampling is a more structured way to remove
samples as it takes into consideration the nature of the field and nematode distribution.

a b c d
0 O .i O O
S. ..... 0 o o
0 0 0

0 0 ................... 0 0 0 0
S0 0 ..0.0 0
0 0 0 0
O O ... .. ................ 0 0 O O
0 0O

Figure 21. Sampling patterns for nematodes. (a) Random sampling; (b-d) systematic sampling.

Time of sampling
The optimum time for sampling varies between crops and is related to the growth
stage of the crop and the objective of the sampling (predictive or diagnostic).
Predictive sampling (not the focus of this guide) is often done early in the season,
such as at or just before planting, or at the end of the previous cropping season to
determine the number of nematodes (density).

Many nematode species increase to high levels during the growing season and reduce
during the off (dry) season; this is easier to see in annual crops than in perennial and
tree crops. Samples should therefore ideally be collected in the middle of the season
and/or at final harvest for diagnostic purposes. Perennials can be sampled during the
active growing period such as during the rainy/growing season to identify the problem.


Taking soil samples
As a rule, avoid sampling very wet or very dry soil. However, where crops normally
grow in, for example, swamp (e.g. paddy rice) or arid conditions (e.g. sisal), these
should be sampled to represent these conditions.

Divide fields larger than 1 hectare into 1 hectare (10,000 m2) plots and sample these
plots separately. Take 10 to 50 sub-samples (cores) and combine them to make a
composite sample that weighs 1-2 kg. Remove the soil sub-sample from the root
zone using a trowel, auger, corer, spade or similar implement that is suitable for the
crop being sampled.

Carefully bag, label and seal samples (Fig. 22).

Make sure to take
samples between rows.

Once the corer
is full, carefully
remove it from the

The sample should
represent a cross section
of the soil from the
surface (0 cm) to around
20-30 cm below.
2 -0


Samples can also be taken using a
trowel or other suitable implement
if a corer is not available.

Place the corer over a With a strong blunt Place samples in sturdy Or even easier, just label
large flat and sturdy box instrument, scrape all the bags with a tie at the top. the bag on the outside
(preferably plastic). contents of the corer into Label clearly with a card with a permanent thick
the box. Make sure you written with pencil (not marker.
thoroughly shake out pen as it smudges).
any excess soil before
taking another sample.

Figure 22. Taking and bagging samples.

Practical plant nematology

Taking root samples
Roots can be collected at the same time and from the same locations as for soil,
and in general should be combined in the same sample bag, so that the soil helps
to preserve the roots.

Generally, depending on the crop, 25-100 g of roots per total sample is sufficient,
but a lower weight may be collected for finer roots such as from rice, and a higher
weight for thick, heavy roots such as from banana or trees. Where both fine and
heavy roots occur, as on crops such as banana, it is suggested to sample these

Avoid sampling dead plants or those in advanced stages of senescence, as nematodes
will often have migrated from these to other food sources. For small crop plants, the
whole root system of a plant can be used for each sub-sample. Lift the plants and
their roots from the soil using a spade or trowel, so that a sizeable proportion of the
root system is unearthed intact, and taking care not to break off the roots and leave
them in the ground. After tapping soil free, randomly remove roots with a knife or

Taking samples from above-ground plant tissue
Leaf, stem, seed or other aerial material should be collected where symptoms are
present and nematodes suspected. Again, it is important to select from a number
of affected plants, which should be compared with non-affected plant tissue from
different plants.

Care of samples
Collect samples in strong plastic bags, and label them clearly and systematically.
Plastic labels marked with a water-resistant permanent marker or pencil can be
placed in the sample bag (Fig. 22), or alternatively write directly on the plastic bag
with a permanent marker pen the sample number or reference. Paper labels are best
attached to the outside of the bag with wire or twine. If using paper labels, use a
pencil, not a pen (which will run or smudge when wet). But remember, paper labels
deteriorate quickly during wet conditions.

Record, where possible:
* The crop and cultivar
* The sampling date
* The farmer
* The location (and GPS coordinates if possible)
* A reference number (or plot) if within an experimental trial
* The previous crop(s).


Nematodes are very sensitive and
perishable, and it is very important that
appropriate care is taken to keep them in
good condition. Samples should NOT be
left in direct sunlight or in a closed vehicle
in the sun. They should also not be left for
long periods before processing.

After collection, samples should be
placed in a coolbox (insulated container;
Fig. 23), or packed in strong cardboard
boxes and placed in a shaded area where
conditions are cool. If unable to process
immediately, samples can be stored in
a refrigerator (approx. 10oC) for up to
2 weeks. Nematode survival decreases
with time however, and nematodes from
relatively hot environments can suffer Figure 23. Insulated cool box for
chilling injury. sample storage.

Nematode extraction

The next stage is to extract nematodes from the samples. This should be done as soon
after collection as possible as samples deteriorate over time.

There are four basic extraction techniques, which are covered in this guide:
* Extraction tray method
* Root or leaf maceration method
* Sieving method
* Incubation method.

Three further methods elutriation, the Fenwick can and centrifugal flotation
- require specialist equipment and are not described in this guide. Details on these
methods can be found in publications listed in the References and Further Reading
section (page 65). It is also possible in some cases to examine nematodes directly
from plant tissue specimens.

Choosing an extraction method
The choice of which method to use depends on the conditions and materials
available, the sample type, and also the type of nematodes present. Some methods
of extraction are more useful for specific types of nematode, while others are more
general. This guide provides details of the most straightforward methods, including
the extraction tray method, which is useful in the most basic conditions, provides a
reasonable assessment of nematodes from soil, roots, seeds or plant tissue, and can be
easily replicated.

Table 1 shows which extraction methods are suitable for which types of nematodes
(sedentary or migratory) in soil or root/foliar samples.

Table 1. The suitability of extraction methods for different nematode types and samples.

Soil sample Root/foliar sample
Sedentary Migratory Sedentary Migratory
nematodes nematodes nematodes nematodes
Extraction tray (p. 34) X X

Sieving (p. 42) X X

Root/leaf maceration (p. 40) X X

Incubation (p. 48) X X

Practical plant nematology

If the target nematode is known such as in research experimental plots it may
be possible to precisely identify which material should be sampled and processed for
the nematodes (soil, root, tuber, leaf, etc.). If you are not sure though, both roots
and soil should be used for extraction with both sedentary and migratory nematodes

Preparation of samples
Dry soil should be properly mixed before sub-sampling for nematode extraction.
Break up clumps and remove stones, roots and debris. Pass (dry) soil through a
coarse sieve with holes of approx. 1-2 mm (Fig. 26, step 1) into a suitable container
and then mix thoroughly. Remove a sub-sample using a beaker or container of
known volume (Fig. 26, step 2); 100 ml soil is commonly used. From each bulked
field sample two x 100 ml soil samples should be processed and the mean taken.

Wet soil, such as from a rice paddy, needs to be removed from the field sample
using small balls or clumps from various parts of the sample or from each root base,
and measured using displacement of water to get samples of the same size (Fig. 29,
steps 1 and 2). Fill the beaker to a set (marked) volume (e.g. 200 ml) and raise the
volume to the required marked amount using soil clumps, e.g. raising 200 to 300 ml
measures 100 ml of soil. Then use the sieving method to extract nematodes.

Roots should be separated from soil and any soil adhering removed by gently tapping
off, or rinsing gently under a tap or in a container of water. Dab the roots dry with
paper towel, and then chop and extract according to the chosen extraction method.

It is important to use clear, correct and consistent labeling for samples (Fig. 24).
Label containers with either a waterproof marker or chinagraph pencil, or use labels
that can be moved along with each stage of the extraction. Ensure all samples are
correctly labeled at all times.

Nematode extraction

Labeling samples with a paper Labeling a sample bag with a
tag and pencil. permanent black marker.

Labeling an extraction tray with both an accompanying plastic tag
and also number on tray with a permanent marker or chinagraph.

Labeling a collection cup with
a black permanent text or a
chinagraph pencil.

Waterproof chinagraph pencil.

Figure 24. Materials for and examples of labeling.

Practical plant nematology

Extraction tray method
This method (or variations of it) is sometimes also called the modified Baermann
technique, the pie-pan method, or the Whitehead tray method.

* Specialist equipment is not required
* It is easy to adapt to basic circumstances using locally available materials
* It extracts a wide variety of mobile nematodes
* It is a simple technique.

* Large and slow moving nematodes are not extracted very well
* The extractions can sometimes be quite dirty (especially if the clay content of the
soil is high) and therefore difficult to count
* The proportion of nematodes extracted can vary with temperature, causing
potential variation in results between samples extracted at different times
* Maximum recovery takes 3-4 days.

* A basket (or domestic sieve) made with coarse mesh (Fig. 25)
* A dish/tray/plate, slightly larger than the basket
* Tissue paper
* Beakers or containers to wash the extraction into
* Wash bottles
* Waterproof pen
* Knife/scissors
* Weighing scales
* Large bench space.

Most items can be purchased ready-made or easily constructed from readily available
materials (Fig. 25). Funnels can be held in a rack or stand with rubber tubing
attached to the bottom and sealed with a clip (Fig. 25, top) or insect mesh attached
to an -10 cm section of -15 cm diameter plastic piping can be used to construct the
sieve (Fig. 25, middle). It is very important that the mesh and sieve base are raised
slightly (-2 mm) from the bottom of the dish/plate using, for example, three or four
small 'feet' glued to the base of the sieve (Fig. 25, bottom). If this is not done, the
nematodes cannot easily migrate into the water.

It is very important to ensure good, consistent labeling of all containers used for
each sample, as it is very easy to make mistakes. Root and soil extractions should be
labeled separately.

Nematode extraction

Example of the funnel method of extraction with rubber tube
clipped and labeled at the base. Nematodes will be concentrated
just above the clip.

Example of different simple types of sieves and trays. The orange sample on the left uses a
home-made cross section of PVC piping with mosquito mesh attached to the bottom. It is
very important that the sieve does not sit flat on the bottom of the tray but is slightly raised.

Home-made PVC piping and net sieve with plastic 'feet'
which ensures the nematodes can migrate easily into the

Figure 25. Different ways to extract the nematodes from the sample.

Practical plant nematology

For soil samples:

Remove roots from sample and place in a separate dish. Label.
Using a coarse sieve, remove stones and debris from soil and break up soil lumps
(Fig. 26, step 1).
In a plastic container (basin, bucket) thoroughly mix the soil sample.
Remove a measure of soil (e.g. 100 ml) (Fig. 26, step 2).
Place tissue paper (milk filter, paper napkin etc.) in the plastic sieve/basket (placed on a plastic plate)
ensuring that the base of the sieve is fully covered by the tissue (Fig. 26, step 3). Label.
Place the soil measure on the tissue in the sieve. It is important that the soil remains on the tissue paper
- spillover results in dirty extractions (Fig. 26, step 4).
Add water to the extraction plates (Fig. 26, step 5). Take care to gently pour water into the plate (dish) and
not onto the tissue paper or soil (between the edge of the mesh and the side of the tray). Add a set volume to
each dish to wet but not cover the soil or root tissue, ensuring there is sufficient not to dry out. More water is
needed for soil samples than root material. Add more later if necessary.
Leave (preferably in the dark) undisturbed for a set period (48 hours if possible) (Fig. 26, step 6) adding
more water if it is likely to dry out. Nematodes from the soil or plant tissue will move through the tissue
paper into the water below, resting on the tray/plate.
After the extraction period, drain excess water from the sieve and the soil into the extraction (Fig. 26, step 7).
Remove the sieve and dispose of plant tissue/soil.
Pour the water from the plate into a labeled beaker (or cup), using a water bottle to rinse the plate (Fig. 26,
steps 8 and 9). Leave samples to settle (Fig. 26, step 10).
For counting the nematodes in the extraction, reduce the volume of water by gently pouring off or siphoning
the excess (taking care not to lose nematodes and sediment), or by passing the extract through a very small
aperture sieve (e.g. 20-30 pm) (Fig. 26, step 11). Wash the nematodes off the small aperture sieve into a
beaker (or tube) for counting, or for preserving, if sending away or counting later (Fig. 26, steps 12 and 13).

2. Measure out a standard
sized sub-sample by 3. Line a sieve with tissue paper
volume, e.g. 100 ml. and place it on a plastic plate.
1. Coarsely sieve the sample to
remove debris and lumps.

Figure 26 continued opposite

4. Place soil on the tissue 5. Carefully pour water into th
ensuring that the soil remains tray making sure you pour the
on the tissue and does not to water down the gap between th
spill over the edges. tray and the sieve.

7. Carefully drain and remove 8. Pour the water containing
the sieve from the tray and dis- the nematodes into a labeled
card the tissue paper and soil. beaker/cup.

10. Leave samples to settle for a few hours or overnight. us

12. The sample can be
stored in a tube if not
observing immediately.

Nematode extraction

e 6. Store extract from samples over 2 days,
constantly checking that the samples
ie remain wet and do not dry out due to

ig 9. Thoroughly rinse the tray into the
ed beaker.

SReduce the suspension by decanting or
ing a small aperture (i.e. 28 pm) sieve and
llect in a beaker ready for assessing nematodes.

13. If sending away for assessment, the
nematodes can be removed from the bottom of
the large tube with a pipette, after settling, for
storage/dispatch in small tube.

Figure 26. Extraction tray method for soil samples.

Practical plant nematology

For root samples:
Roots can sometimes be divided into separate categories, such as larger tough roots
and finer feeder roots. It is useful to extract nematodes separately from each category,
as the root tissue texture varies and the type of nematodes invading may also vary, as
well as densities of the same nematode. Extraction efficiencies may also vary, with the
nematodes exiting slower from a larger root.

Gently tap soil off the roots/tubers or rinse under a tap and then gently dab dry
with tissue paper. Peel tubers carefully with a knife or kitchen peeler to just below
the surface.

Chop the roots (or tuber peel) finely with a knife or scissors and place in a labeled
dish (Fig. 27, step 1). Mix all chopped root material thoroughly.

Remove and weigh a sub-sample (e.g. 5 g) of chopped root material using
measuring scales (Fig. 27, step 2).

Place weighed sub-sample on the tissue paper in the labeled sieve/basket (Fig. 27,
step 3).

Follow the rest of the procedure for soil extraction above (Fig. 26, steps 5-13)


Use the root maceration method for tough plant tissue (Fig. 28, pages 40-41), then
follow the remaining procedure for tray extraction (Fig. 26, steps 5-13)

If sending nematodes away for identification or counting, preserve using the
protocol on pages 57-60.

Nematode extraction

1. Chop roots and/or tuber peel and place in labeled dish.

2. Weigh out root sub-samples.

3. Place root sub-sample in sieves for extraction.

Then follow steps 5-13 in Figure 26.

Figure 27. Extraction tray method for root samples.

Practical plant nematology

Root maceration method
This method is also known as root maceration followed by incubation in an
extraction tray.

* Does not require specialized equipment.

* The amount of time spent macerating is critical it must be sufficient to allow
nematodes to easily move out from plant tissue, but damage must be minimized.

* Beakers
* Scissors/knife
* Water bottle
* Waterproof pen
* Weighing scales
* Domestic blender


Cut roots or tuber peel into small pieces (Fig. 28, steps 1 and 2). Mix root tissue

Weigh a sub-sample (Fig. 28, step 3). Place it in an electric blender with just
enough water to cover the blades.

Blend fine roots and tuber peel for two 5 second bursts and tougher roots for
two 10 second bursts, waiting for the suspension to settle briefly between the two
blendings (Fig. 28, step 4).

Pour the blended suspension of roots and water into a beaker, rinsing out the
blender container of all debris, using a water bottle if possible (Fig. 28, step 5).

Gently pour the suspension of roots and water onto tissue paper (Fig. 28, step 6)
and then follow the extraction tray method (Fig 26, step 5 onwards).

The roots can be stained before maceration to improve visibility of the nematode
after extraction (see page 60).


Figure 28. Root maceration method.

Nematode extraction

1. After rinsing the tubers, peel them thinly with a knife or peeler.

2. Chop the peelings or roots coarsely with a pair of scissors or a knife.

3. Weigh out sub-samples. 4. Macerate roots/peel using a blender.

5. Pour blended suspension into a labeled beaker 6. Gently pour the sample onto the tissue paper
and rinse out blender into the beaker. in the sieve as for extraction tray method.

Then follow steps 5-13 in Figure 26.


Practical plant nematology

Sieving method
Good extraction of all types of nematodes
Good for extraction of large and slow moving soil nematodes
Suitable for extracting nematodes from wet soil
Useful for cyst extraction from soil.

Nematodes may settle out with soil particles unless soil is well dispersed
Nematodes can be easily damaged.
Requires slightly more specialized equipment.

For soil motile nematodes:
Beakers and buckets
Waterproof pen
Brass 20 cm diameter (Endecotts' or Retsch') sieves: 2 mm, 90 pm (or 53 pm)
and 38 pm
Extraction tray equipment

For sedentary cyst (e.g. Heterodera) recovery:
As for soil motile nematodes, plus
Brass 20 cm diameter (Endecotts' or Retsch') sieves: 2 mm, 250 pm, 150 pm
Filter paper (or paper towel/tissue)

If it is not known whether sedentary nematodes are present then all equipment
should be used.
For soil motile nematodes:

Fill a bucket with about 6 liters of water. Mark a water line on the inside of the bucket with a waterproof pen
for consistent water volume between samples (Fig. 29, step 3).
Place a sub-sample of sieved and mixed dry soil, or of wet soil measured by displacing water in a beaker (Fig.
29, steps 1 and 2) into the bucket (Fig. 29, step 4).
Mix the water thoroughly using your hand (Fig. 29, step 5). Allow larger particles to settle for 30 seconds
(Fig. 29, step 6).
Slowly pour off the upper 34 of the water through the nested sieves: use a 2 mm sieve to catch debris for disposal,
or just 90 pm and 38 pm ones to catch nematodes if there is no debris (Fig. 29, step 7). This requires two people.
Take great care to ensure that water does not escape over the sides of the nested sieves (in between the stacked
sieves) when pouring the bucket of water through, as nematodes will be lost in the escaping water. Pour slowly
and tap the underside of the bottom sieve gently, if necessary, to help water flow through the the sieves.
Refill the bucket to the marked line (Fig. 29, step 8) and repeat the process once or twice (Fig. 29, steps 5- 7).
Wash off the debris from the 90 and 38 pm sieves into a labeled beaker, ensuring that sieves are properly
cleared (Fig. 29, steps 9-11) by washing gently from behind.
Leave beakers for 2-3 hours for nematodes to settle to the bottom. If necessary gently pour off and discard
excess water.

Nematode extraction

S Water line

1. Measure water in
a beaker to a known
volume, e.g. 200 ml.

4. Pour pre-measured soil
sub-sample into the water.

2. Measure soil by adding clumps
from each bulk sample and displac-
ing water to marked volume.

5. Mix thoroughly.

3. Measure a set volume of
water using a pre-marked line
on the inside of a bucket.

6. Leave the soil to settle for
30 seconds.

7. Slowly pour three-quarters
of the water through the
nested sieves (such as 90 and
38 [tm size) with the 90 [tm
sieve on the top.

8. Refill bucket and
repeat steps 5, 6 and 7.

9. Condense the extract debris by 10. Ensure the sieves are back- 11. Gently wash debris from the
gently rinsing the sieves thoroughly washed properly and all debris 90 )tm and the 38 tm sieve into
with a hose, mainly from the back. and nematodes are collected a labeled beaker.
from the sieve surface at the
bottom point of the sieve.

Figure 29. Extraction of soil nematodes using the sieving method.

Practical plant nematology

The sievings can then be processed further via the extraction tray method (Fig. 26,
steps 3-13, page 37).

For recovery of sedentary cysts:

Air-dry the soil sample before using for extraction (Fig. 30, step 1).

Fill a bucket with about 6 liters of water, and mark the water line on the inside of
the bucket with a waterproof pen (Fig. 30, step 2).

Place the measured soil sub-sample in the bucket (Fig. 30, step 3).

Mix the water thoroughly using your hand, then allow soil particles to settle for
60 seconds. Cysts should float (Fig. 30, steps 4 and 5).

Slowly pour off the top 12 of water through the nested sieves: 2 mm to catch debris
for disposal, and 250 pm and 150 pm to trap cysts (Fig. 30, step 6).

Wash off the debris from the 250 pm and 150 pm sieves into a labeled beaker
(Fig. 30, steps 7 and 8).

Refill the bucket to the marked line and repeat the process (steps 4-8) at least once,
collecting all debris for each sample into the same beaker. Repeat this process as
much as necessary until you are satisfied that no cysts remain in the bucket.

Prepare and label a paper lining (filter paper, milk filter, paper towel etc.) for a
funnel (i.e. in a cone shape) held in a stand or beaker (Fig. 30, step 10).

Pour the wash-off (sievings) in the beaker through the filter in the funnel (Fig. 30,
steps 11 and 12). Allow water to drain through.

Carefully remove filter papers from the funnel and place in a moistened tray to
await direct observation under the microscope (Fig. 30, step 13). Viewing can be
done by gently opening the filter paper and spreading the contents across the filter
paper, followed by viewing under stereomicroscope.


Allow filter papers to dry in the funnel, for removal and storage for observation,
picking or counting at a later time (Fig. 30, step 14).

Nematode extraction

SWater line


1. Air dry the soil in an open dish.

2. Fill the bucket with water to a marked line.

3. Pour the pre-measured dry soil
sub-sample into the water.

4. Mix thoroughly.

5. Allow the soil to settle
for 60 seconds.

6. Slowly pour half of the water
through the 2 mm, 250 and 150
tm sieves.

7. Rinse down each of the 2 catchment sieves, backwashing where

Figure 30 continued overleaf

Practical plant nematology

8. Using a wash-bottle, wash the debris
from both the 250 and 150 tm sieves
into beakers for collection.

9. Refill the bucket with water
to the marked line and repeat
steps 4-8.

10. Fold a circular piece of 11. Pour the extraction 12. Leave the liquid to drain
filter/tissue paper into quarters, through the filter paper, through the filter paper.
then open it as a cone and
place in the funnel which is
held in a beaker or stand.

13. Place filter paper with cyst extract
debris into a dish with water in the
bottom to keep moist for immediate
cyst assessment.

14. Leave extracts to dry in the funnel
to assess cysts at a later date.

Figure 30. Extraction of cyst nematodes from soil using the sieving method.

Nematode extraction

Cysts from air-dried samples will float to the surface of the bucket, but if conditions
do not allow air drying, cysts can be extracted from fresh soil samples. Many cysts
should still float, but fresh, heavier cysts may not, and agitating the water and
reducing the settling time may be needed, before decanting the water/suspension
through sieves.

Cysts can also be picked from the sieved, dried samples directly under the
stereomicroscope, using a fine paintbrush. Determining the cyst density in soil is
useful, but it is also necessary to assess the egg number, by first crushing cysts to
release eggs (see References and Further Reading).

Practical plant nematology

Incubation method
This method is also called root incubation in plastic bags or jars.

* Does not require specialized equipment.

* Extraction efficiency may be relatively poor
* Nematodes are often in poor condition due to lack of oxygen.

Plastic bags, jars, conical flask or similar vessel.


Cut/chop tissue finely and mix together (Fig. 31, step 1).
Weigh out a sub-sample/sample.
Place in a closed labeled plastic bag or covered container (not metal) holding a
small quantity (10-20 ml) of water (Fig. 31, step 2).
Nematodes hatch from eggs or migrate from root tissue into water over a period of
2 to 7 days.
Remove water regularly (e.g. daily) from the container, which will help nematode
survival, and bulk in a labeled beaker, using the same beaker for each sample
(Fig. 31, step 3).
Replace water after each decanting, cover and leave, repeating the process over 2-7
days (Fig. 31, step 4).
Reduce or concentrate the suspension/extract for each sample and observe directly
or store for later use following steps 11-13 in Fig. 26.

Nematode extraction

1. Chop the roots and weigh the sub-

2. Place weighed sub-sample of roots in jar, conical flask or plastic
bag with water and leave for 2-7 days. Take care not to fully seal the
container, but loosely cover.

3. Each day, shake/swirl the
container and gently pour the
suspension into a beaker, leaving the
plant material in the container.

4. Replenish with fresh water after
pouring off suspension.

5. Concentrate the suspension
and collect nematodes for further
assessment, e.g. using a small
aperture sieve, or leave the beaker
to settle and pour off the excess.
Then follow Fig 26, steps 12 and 13.

Figure 31. Incubation method of extraction.

Direct examination of plant tissue

Infected plant tissue can be examined for nematodes under a dissecting microscope,
for example to assess that nematodes are present before sending material for expert
identification. The adult sedentary females, which are embedded inside roots (see
Figs 10 and 11) can also be teased out of the tissue and used for identification
purposes. Where samples are being sent elsewhere for species identification,
the (galled) root material itself needs to be sent to the taxonomist, preserved in
lactoglycerol solution. Direct observation is also useful for assessing and observing
foliar tissue and seed-infecting nematodes (see Fig. 8), and for picking out of
individual and specific nematodes to prepare slide collections etc.

For direct observation of plant material:

Wash the plant tissue under a gentle stream of water, or place in a bowl of water for
a few minutes, to remove soil and debris, taking care not to dislodge ectoparasitic
nematodes feeding or attached on the outside of roots.
Cut the plant tissue into -2 cm pieces with a pair of sharp scissors or a knife.
Place the plant tissue into an open Petri dish that has water in the base (Fig. 32).
For immediate observation tease open the tissue with the aid of mounted needles
and forceps to release the nematodes from the plant tissue. This is suitable for
sedentary endoparasites (Fig. 32).
If the plant tissue contains migratory nematodes it may be useful to leave in a Petri
dish overnight or longer even. Nematodes will migrate out of the tissue into the
Nematodes can then be picked under the stereomicroscope for identification or
preserved (and stained) and/or sent for further identification.

Sheaths teased apart. Seed coats broken to allow nematodes
free movement.

Figure 32. Direct examination of plant material in water.

Handling, fixing and staining nematodes

Various techniques aid the handling and identification of nematodes. These are
described in this section.

Handling nematodes
Due to their microscopic size, nematodes can be difficult to handle, particularly for
beginners. It is nearly always necessary to handle them in a fluid medium, usually
water. Using a dissecting microscope rather than a compound microscope also
helps. Individual nematodes need to be selected when establishing pure cultures
or preparing identification slides and therefore need to be 'picked'. If handling
nematodes from pure cultures, batches of specimens can be transferred with the use
of glass pipettes (see Fig. 35). By narrowing the aperture of glass pipettes using heat
from a Bunsen burner, even individual nematodes can be handled and transferred.

Picking nematodes
To look at nematodes closely for identification, it is often necessary to individually
'pick' the nematodes from the extraction suspension and place them on a glass
slide (Fig. 33). This can be difficult, but gets easier with practice. Nematodes are
translucent (see-through) and may be difficult to see; under-stage lighting and a
stereoscopic microscope help (Fig. 34). If picking nematodes from plant tissue or
root surface, top lighting can also be beneficial.

Figure 33. Picking nematodes using a bamboo splinter on a dissection microscope.

Practical plant nematology

Various instruments can be used for picking, for example a fine insect pin, a
bamboo splinter, an eyelash or bristle glued to the end of a mounted needle, a
sharpened toothpick, or feather spine (Fig. 35).
Pour or use a pipette to place some of the nematode suspension (or infected plant
tissue) into a Petri dish, counting dish or glass block. Keep the suspension shallow.
Place on a stereomicroscope using the lowest convenient magnification.
Gently swirl the suspension to move nematodes to the center of the dish.
Locate a nematode and gently lift the nematode to the surface of the water with the
picking tool.
Adjust microscope focus to keep the nematode in view whilst picking the nematode
out of the water solution.
Holding the picking instrument under the nematode lift or very gently 'flick' the
nematode out of the water. The nematode should be 'hanging' on the tip of the
picking instrument.
Gently place the tip of the pick into a drop of water on a slide, in a glass block or
other vessel containing some water.
To view nematode(s) on a slide on a compound microscope, it is first useful to
'relax' them by heating briefly on a hotplate (not too hot).

Place glass beads or cover slip splinters at edges of the water droplet and place a
cover slip over the droplet.
View under compound microscope.

Using a dissection stereo-microscope with Using a compound stereo-microscope with
understage lighting. understage lighting.
Figure 34. Microscopes used for picking.

Handling, fixing and staining nematodes

Figure 35. A sample of potential nematode picking tools: a bamboo splint; the tip
of a pipette thinned using heat; dissection needle with an eyelash glued to the tip.

Sending nematodes for identification
If, after sampling and extracting, the nematodes can be immediately identified
to genus level and counted then this will quickly provide an indication of which
parasitic nematode groups are present and whether they are at potentially damaging
levels or can be associated with crop damage. However if the expertise to do this is
not available, or the species involved need to be established, nematode samples will
need to be sent away for identification by a specialist taxonomist.

Before sending, samples usually need to be killed (see next section) and preserved,
especially if being sent out of the country. Nematodes can be sent live if pre-arranged
with the destination lab or if sent within the same country. Sometimes soil and/
or plant tissue can also be sent, however it is essential to respect the quarantine
regulations for a given country.

Nematodes should be collected in a small vial or tube (Fig. 36) and packed carefully
in insulated containers for transportation to the laboratory where they will be
identified. The vials should be clearly labeled with a code/number. The codes should
be recorded on duplicate sheets (one to accompany specimens and one to remain)
with all the details of the sample and kept until the results of the identification are

Practical plant nematology

A range of sample bottles for storing or sending samples.

Small microtubes useful for sending samples away for identification.

Using a glass block for preserving nematodes
by pipetting in drops of formalin.

Transferring nematodes into a microtube for storage
or sending.

Figure 36. Storing nematode samples for transporting.

Handling, fixing and staining nematodes

Nematode identification services
A number of taxonomists based at various centers around the world are able
to precisely identify nematodes. However, very few offer a service for routine
identification, especially to species level. Some centers that do offer this service are:

Plant Disease and Diagnostic Services
CABI Bioscience UK
Bakeham Lane
Surrey TW20 9TY, UK
Tel: +44 (0)1784 470111

P/Bag X134
Pretoria 0001, Republic of South Africa
Tel: +27 (0)12 356 9830

Department of Plant Protection
Faculty of Agriculture
University of Jordan
Amman 11942, Jordan
Tel: +962 (0)6 535 5000-3004

Central Science Laboratory
Sand Hutton
Y04 1LZ, UK
Tel: +44 (0)1904 462000

Before sending any samples for identification, it is essential to contact the centre
and establish their capacity to handle your samples and how best to preserve and
transport them.

General advice and training is offered by the nematology sections of both CIMMYT
and IITA upon contact. Information can also be sought from the nematology related
societies, which are listed on the inside back cover.

Killing nematodes
It is important to kill nematodes quickly, as each species assume a particular 'death
shape' when killed quickly which can help in identification. Nematodes are best
killed with gentle heat (55-65C), which retains the nematode body content. If
killed at too hot a temperature, body contents are cooked and denatured, causing
difficulty for identification. Nematodes can either be killed first and then fixed or
killed and fixed in the same process.

Practical plant nematology

A simple and efficient method for killing nematodes is to add an equal volume of
boiling water to the nematode suspension. If the whole extract is being sent, the
nematode suspension may need to be reduced in volume so that there is less than
half in the sample tube or vial. It may be easier to kill and fix the nematodes in
larger tubes then place a reduced volume into smaller tubes (Fig. 36), or remove
nematodes from the bottom (Fig. 26, step 13) with a pipette and place in the tubes
for transportation.

Nematodes can also be killed by holding the tube containing a small volume of
nematode suspension in near-boiling water for 1-2 minutes, but this can take a long
time for a large number of samples. It can also be cumbersome and care is needed to
ensure sample bottles remain upright and do not topple over into the water, losing
the samples.

Fixing nematodes
The simplest method for fixing or preserving samples is to pipette a few drops of
formaldehyde (formalin) into recently heat-killed samples. Two or three drops into a
7 ml sample bottle is sufficient (Fig. 37); larger sample bottles will require more. This
is a quick and easy method, which will prevent samples from deteriorating during
transit and storage before identification, however it does not provide good quality
specimens for long-term preservation and can also cause difficulty for identification,
especially if not examined immediately.

Take great care with formaldehyde as it is dangerous to health.

Pipetting formalin into a nematode suspension in a 7 ml
tube to preserve the sample.

Staining plant tissue with acid fuchsin in lactoglycerol
solution on a hot plate.

Fig 37. Fixing and staining nematodes.

Handling, fixing and staining nematodes

Killing and fixing in one step

Heat the fixative to near boiling in a test tube or beaker by immersing in boiling
Pour an equal volume of hot fixative to that of the nematode suspension into the
nematode suspension (i.e. 2 ml of hot fixative into 2 ml of suspension = 4 ml).


Collect the nematodes in a glass block in a small drop of water and add 2-3 ml of
hot fixative with a pipette.

The most suitable fixatives to use are:

Triethanolamine 2 ml
Formalin (40% formaldehyde) 7 ml
Distilled water 91 ml
The fixative remains stable for a long time and nematode appearance remains lifelike
because the specimens do not dry out.

FA 4:1
Formalin (40% formaldehyde) 10 ml
Glacial acetic acid (proponic acid) 1 ml
Distilled water 89 ml
In FA 4:1 nematodes maintain their structure though they may become discolored
after some time.

Formalin glycerol
Formalin (40% formaldeyde) 10 ml
Glycerol 1 ml
Distilled water 89 ml

This has the advantage of keeping the nematode from drying out even if the vials
are not properly sealed. Again take great care with all these fixatives as they are
dangerous to health.

Preserving sedentary nematodes in root or tuber tissue
The females of sedentary nematodes are required for species identification. Therefore
the plant tissue containing the nematodes, such as galled roots, needs to be preserved
and sent for examination. Placing a small sub-sample of infected plant tissue into
a sample bottle containing lactophenol or lactoglycerol can be sufficient. Staining
before preserving can help identification.

Practical plant nematology

Lactophenol can be purchased ready made, or made by mixing equal volumes of
glycerol, lactic acid and distilled water (lactoglycerol) and dissolving a small amount
(1%) of phenol into it (lactophenol). Phenol is very toxic however, so it is usually
best to just use lactoglycerol, although this does not preserve samples for long periods.

Observation of nematodes embedded in plant tissue can be made easier by using
appropriate stains, which stain the nematodes while plant tissue remains relatively
clear (e.g. see Fig. 5). Thick or bulky roots should be sliced thinly before staining to
ensure transmission of sufficient light after clearing.

Stain in lactoglycerol + 0.1% cotton blue or 0.05-0.1% acid fuchsin, then destain
in a beaker containing a solution of equal volumes of glycerol and distilled water + a
few drops of lactic acid. Destaining is most effective if done over several days.

Gently wash plant material free of soil and other debris, and dry gently by dabbing
with paper towels.
Cut or slice thick or wide roots or tuber into small lengths.
Place in muslin cloth, tie up the corners with a piece of cotton string, and label
clearly with labels attached to each separate muslin 'bag'.
Bring stain solution, using a glass beaker on a hot plate, to near boiling.
Place muslin bags into boiling stain solution and leave for approximately
3 minutes, depending on root thickness. Use a deep beaker, approx. half full of
stain solution, as it will froth up when plant tissue is added (Fig. 37).
Remove the muslin bags and rinse in running water.
Place the muslin bags in the clearing solution and leave overnight or longer.
Examine under the microscope. Placing roots side-by-side on a microscope slide
and gently squashing them using another slide placed on top enables the stained
nematodes to be seen more clearly. Nematodes will be stained red with acid fuchsin
or blue with cotton blue.

Meloidogyne egg masses
Phloxine B stains the gelatinous matrix that surrounds Meloidogyne eggs, increasing
the visibility of egg masses and enabling a rapid count of adult female nematodes/egg
masses present. The eggs also remain viable after staining. The solution is made by
adding 15 mg (a very small sprinkle) of Phloxine B to 1 liter of water.

Place the rinsed roots in a tray or dish (preferably white) containing Phloxine B
solution and leave for 15-20 min. Count stained (blue) egg masses.

Estimation of nematode density

Once nematodes have been extracted from soil or plant tissue, they must be first
identified and then quantified. This enables assessment of their association with, or
their potential to cause, damage.

This guide does not provide an identification guide to plant parasitic nematodes; for
these see the Further Reading section on page 65. (A very basic description of the
most common plant parasitic nematodes is given in Appendix 2.) Nematode density
estimation should only be attempted if nematodes have been definitely identified.
If the capacity and skills to do this are not available, samples should be sent to the
experts (see page 57).

Counting nematodes
Extracted nematodes can be viewed and counted using a dissecting or a compound
microscope (Fig. 34); access to both is ideal. Good quality illumination (understage
lighting) is essential. A magnification of about 40x is usually suitable (i.e. a 4x
objective combined with a lOx eyepiece), but a compound microscope can also
be used (i.e. using the lOx objective), which is useful for nematodes that are
in poor condition or are hard to identify. Dissection microscopes allow greater
maneuverability and depth of focus, especially for dirty samples. Nematodes that
cannot be identified in the counting dish with the counting magnification or at
higher magnification using a compound microscope, should be manually picked
(Fig. 33) and mounted on a glass slide for identification using higher magnification
with a compound microscope.

Various forms of counting dishes exist, but basically a clear plastic dish is needed that
has a grid etched on the bottom (Fig. 38). This can easily be prepared by carefully
scratching lines on the underside of a small plastic Petri dish. An open rectangular
plastic dish with approximately 5 ml capacity is useful for general purposes; it also
makes it possible to move nematodes or debris in the dish, and to hand pick for
identification at higher magnification (Fig. 33). Sloping sides help minimize optical
distortion caused by the meniscus, while raised grids can help reduce the nematodes
moving between the grid lines.

Samples that have been reduced to 5 ml can be counted as a whole sample, but
if nematode density is high or the sample is dirty, a proportion aliquott) can be
counted, diluting with water as required. However, care must be taken to ensure
that a representative proportion of the total sample is counted which is achieved by
thoroughly mixing the sample before taking the aliquout.

Practical plant nematology

f ~, Extract nematodes from a known weight of plant tissue
tij or volume of soil using one of the previously described

Concentrate the extracted suspension to a precise
known volume in a measuring cylinder or graduated
tube (e.g. 10 ml).

Shake or stir the suspension immediately before remov-
.i ing aliquots.

Use a wide mouth pipette to remove aliquots, to
prevent blockage by debris. Pipette tips can be cut if
A sample of counting dishes. they are too narrow.

Carefully pipette aliquots into the counting dish,
avoiding splashing.

If only a few nematodes are present, count them in the
total suspension volume.

If nematode density is high, count the nematodes from
an aliquot (e.g. 1 or 2 ml). Dilution of the suspension
may be necessary to aid counting, for example dou-
bling the volume.
A sample of pipettes.
Count all the nematodes in the counting dish in a sys-
tematic way following the gridlines on the dish. Some-
times nematodes may float on the surface, but adding a
tiny spot of liquid soap overcomes this.

Use a tally counter (ideally a multiple tally counter;
Fig. 38) to count the various different nematodes
present, or score using the Roman tally system if no
tally counter is available.

Return the counted aliquot to the suspension after

Repeat using 2-3 aliquots per sample and then calcu-
late the mean for the combined aliquot score before
calculating the total nematode number per sample.

The mean number of nematodes calculated from the
aliquots should be multiplied by the total volume of
the suspension to calculate the total number in the
Multiple and single tally counter. plant tissue or soil that they were extracted from (e.g.
100 ml soil or 5 g root).
Figure 38. Counting tools.

Damage analysis

Scoring of nematode symptoms on plants
Nematode damage can be evaluated at the same time as field sampling for
nematodes. The amount of root damage is estimated visually (as a percentage) using
a scoring procedure (Appendix 3). Useful damage estimates can be made for root-
knot nematode damage in particular, but also for other nematode damage. The
damage score usually has a strong relationship with crop yield losses.

Scoring nematode damage provides a rapid indication of the damage at that time.
Where basic nematological equipment and expertise are lacking, it may be the only
means of assessment. Damage scoring can also be used to help identify resistance or
tolerance in varietal screening exercises.

The number of plants assessed can be one or two, up to 25 or more, depending
on the crop and area under assessment, and also whether the farmer wants to take
a low or a high risk approach to assessment. One person or at least as few people
as possible should carry out the scoring, for consistency. The use of score sheets to
regularly refer back to is advisable for the same reason.

Some judgment may be needed when assessing nematode damage, for example,
plants that have severe root-knot infection may have very few roots left to assess, with
galls having rotted away. Galling damage may therefore appear minimal, but in fact
the damage due to nematodes is high. It must also be remembered that the response
of different crops and crop varieties to a nematode species may vary, in particular to
root-knot nematodes. Different nematode species also cause different symptoms, for
example, infection by Meloidogyne hapla will often result in bead-like galling (as seen
on the lettuce in Appendix 3), while Meloidogyne incognita may cause more massive
galling and fused root flesh (Appendix 3).

The score sheets in Appendix 3 provide examples and a basis upon which to create
damage scoring for other crops and nematode damage circumstances. The sheets
mostly score damage on a scale from 1 to 5, which balances speed of assessment with
accuracy. If more time is available, scoring on a 1-10 scale will provide more accurate
damage estimates.

References and further reading

Agrios, G.N. (2005). Plant FP: ',-;. 5th edn. Academic Press, USA. 922 pp.

Bridge, J. and Page, S.L.J. (1980). Estimation of root-knot nematode infestation
levels on roots using a rating chart. Tropical Pest Management 26: 296-298.

Brown, R.H. and Kerry, B.R. (1987). Principles and Practice ofNematode Control in
Crops. Academic Press, Sydney. 447 pp.

Evans, D., Trudgill, D.L. and Webster, J.M. (1993). Plant Parasitic Nematodes in
Temperate Agriculture. CAB International, Wallingford. 648 pp.

Luc, M., Sikora, R.A. and Bridge, J. (2005). Plant Parasitic Nematodes in Subtropical
and TropicalAgriculture, 2nd edn. CAB International, Wallingford. 871 pp.

Mai, WE and Mullin, PG. (1996). Plant Parasitic Nematodes. A Pictorial Key to
Genera, 5th edn. Comstock, London and Cornell University, Ithaca. 276 pp.

Moens, M. and Perry, R. (2006). Plant Nematology. CAB International, Wallingford.
447 pp.

Southey, J.E (1986). Laboratory Methods for Work with Plant and Soil Nematodes. Ref.
Book 402. Ministry of Agriculture, Fisheries and Food. Commercial Colour Press,
London. 202 pp.

Speijer, PR. and De Waele, D. (1997). Screening ofMusa Germplasmfor Resistance and
Tolerance to Nematodes. INIBAP Technical Guidelines 1. International Network for the
Improvement of Banana and Plantain, Montpellier. 47 pp.

Stirling, G.R., Nicol, J. and Reay, F. (1999). Advisory Services for Nematode Pests
- Operational Guidelines. Rural Industries Research and Development Corporation.
RIRDC Publication No. 99/41. 111 pp.

Waller, J.M., Lennd, J.M. and Waller, S.J. (2002). Plant Pathologists Handbook, 3rd
edn. CAB International, Wallingford. 516 pp.

Wallwork, H. (2000). Cereal Root and Crown Diseases. Grains Research Development
Corporation Publications, Canberra, Australia. 58 pp.

Whitehead, A.G. (1997). Plant Nematode Control. CAB International, Wallingford.
400 pp.

Zuckerman, B.M., Mai, WF and Krusberg, L.R. (eds) (1990). PlantNematology
Laboratory Manual. University of Massachusetts Agricultural Experiment Station,
Massachusetts. 252 pp.

Appendix I.

Examples of nematode genera and species

known to be important crop pests worldwide

Nematodes and damage symptoms Main crops affected Distribution

Necrosis of roots
Achlysiella williamsi Sugarcane Australasia
Anguina Cereals and grasses Temperate: worldwide
Seed and leaf galls, distortion of leaves
A. tritici (ear cockle nematode) Temperate cereals, mainly Temperate: China, Eastern Europe,
wheat India, North Africa, West Asia
Poor root growth, chlorosis
Aphasmatylenchus straturatus Groundnuts Tropical: West Africa
Necrosis and distortion of leaves and
seeds, destruction of fungal mycelium
A. arachidis (groundnut testa nematode) Groundnut Tropical: West Africa
A. besseyi (rice white tip nematode) Rice Tropical: rice-growing areas worldwide
A. fragariae (strawberry crimp nematode) Strawberry Temperate: Europe, North America,
A. ritzemabosi (leaf nematode) Chrysanthemum Temperate: Europe, North and South
America, East and southern Africa,
A. composticola (mushroom nematode) Mushrooms Mushroom cultivation areas
Belonolaimus (sting nematodes)
Necrosis of roots, chlorosis, wilt
B. longicaudatus Sweet corn, vegetables, Subtropical: southeastern USA
groundnut, citrus, cotton
Chlorosis and tree death
B. xylophilus (pine wilt nematode) Pine Temperate: China, Korea, North
America, Taiwan, Turkey
B. cocophilus (red ring nematode) Coconut, oil palm South and Central America,
Necrosis (red ring) of stems and Caribbean
inflorescence, nut fall

Practical plant nematology

Nematodes and damage symptoms Main crops affected Distribution

Criconemella (ring nematodes) Temperate and tropical
Chlorosis, necrosis of roots and pods,
C. onoensis Rice Tropical: USA, West Africa, Central
and South America
C ornata Groundnut Subtropical: USA
C xenoplax Fruit trees Subtropical: USA
Ditylenchus (Stem/bulb nematodes)
Lesions of stems and leaves, distortion of
flowers and foliage, bulb and tuber rot
D. africanus (groundnut pod nematode) Groundnut Subtropical: southern Africa
D. angustus (ufra nematode) Rice Tropical: Bangladesh, India, Burma,
D. dipsaci Field beans, onions daffodils Europe, North and South America,
and other bulb crops, cereals Eastern Australia
D. myceliophagus Mushrooms Temperate: mushroom cultivating
areas worldwide
Helicotylenchus (spiral nematodes) Widespread on many crops Temperate and tropical: worldwide
Necrosis of roots but damage largely unknown
H. multicinctus Bananas and plantains Tropical/subtropical: banana growing
areas worldwide
Root destruction, chlorosis, twig dieback
H. mangiferae Fruit trees Subtropical: South Asia, Africa, South
and Central America, Caribbean
Heterodera (cyst nematodes)
Cysts on roots, poor root growth,
chlorosis, wilting
H. avenue (cereal cyst nematode) Small grain cereals (wheat, Global: Central West Asia and North
barley, oats) Africa, Northern Europe, China, India,
Australia, Pacific North West USA
H. cajani (pigeon pea cyst nematode) Pigeon pea India
H. ciceri (chickpea cyst nematode) Chickpea, lentil Mediterranean
H. filipjevi (cereal cyst nematode) Small grain cereals (wheat, Central West Asia, India, China,
barley, oats) Northern Europe
H. glycines (soybean cyst nematode) Soybean, beans Subtropical: North and South
America, Japan, China
H. latipons (cereal cyst nematode) Small grain cereals (wheat, West Asia
barley, oats)
H. mani Small grain cereals (wheat, West Asia
barley, oats)
H. oryzae (rice cyst nematode) Rice Tropical: India, Bangladesh
H. sacchari (sugarcane cyst nematode) Sugarcane, rice Tropical: West Africa, India
H. schachtii (sugarbeet cyst nematode) Beets, swedes and other Temperate/subtropics Europe, North
brassicas America, West and southern Africa,
H. zeae (maize cyst nematode) Maize Tropical: India

Appendix I. Examples of nematode genera and species

Nematodes and damage symptoms Main crops affected Distribution

Hirschmanniella (Rice root nematodes
and mitimiti nematode)
Root lesions and corm rot (mitimiti
H. gracilis Rice Tropical
H. imamuri Rice Tropical
H. oryzae Rice Tropical: West Africa, North and
South America, South and Southeast
H. spinicaudata Rice Tropical: Africa, North and South

Hoplolaimus (lance nematodes)
Necrosis of roots
H. columbus Cotton Tropical: USA, Egypt
H. seinhorsti Cotton, vegetables Tropical: Africa, South Asia, South
Longidorus (needle nematodes)
Root tip galling. Transmit viruses
L. elongatus Strawberry, sugarbeet Temperate: Europe, Canada
Meloidogyne (root-knot nematodes)
Galling of roots and tubers, chlorosis,
M. acronea Cotton, sorghum Tropical: southern Africa
M. africana Coffee Tropical: Africa
M. arenaria Groundnut Tropical: worldwide
M. ., : Wheat, barley and legumes Mediterranean countries including
Italy, France, Greece and Spain, West
Asia, Israel and Western Siberia
M. chitwoodi Potatoes, sugarbeet, cereals Temperate: North America, Mexico,
South Africa, Europe
M. coffeicola Coffee Tropical: South America
M. exigua Coffee Tropical: South America
M. graminicola Rice Tropical: South and Southeast Asia
M. hapla Pyrethrum, vegetables, Temperate/subtropical: worldwide
M. incognita Vegetables, cotton, tobacco, Tropical: worldwide
very wide host range
M. javanica Vegetables, cotton, tobacco, Tropical: worldwide
very wide host range
M. oryzae Rice Tropical: South Asia
M. mayaguensis Vegetables, papaya, wide Tropical: worldwide
host range
M. naasi Wheat, barley Northern Europe, New Zealand,
Chile, USA, Iran and former USSR

Practical plant nematology

Nematodes and damage symptoms Main crops affected Distribution

Nacobbus (false root-knot nematodes)
Root galling
N. aberrans Vegetables, potato, sugar Temperate/subtropical: South,
beet Central and North America, Europe

Paralongidorus (needle nematodes)
Root tip galling. Transmit viruses
P. australis Rice Subtropical: Australia
Paratrichodorus (stubby root nematode)
Shortened (stubby) blackened roots.
Transmit viruses
P. minor Vegetables Temperate/subtropical: Europe, USA
Pratylenchus (lesion nematodes)
Necrosis of roots, corms and tubers
P. brachyurus Groundnuts, pineapple, Tropical: worldwide
P. coffee Bananas, yams, coffee, Tropical: worldwide
citrus, spices, very wide host
P. goodeyi Bananas Subtropical: East and West Africa,
P. loosi Tea Subtropical: South Asia
P. neglectus Potatoes, vegetables, small Australia, West Asia, North Africa,
grained cereals (wheat, USA, Canada
barley, oats)
P penetrans Fruit and nut trees, Temperate: worldwide
vegetables, soft fruits, flower
P. thornei Small grained cereals (wheat, Australia, West Asia, North Africa,
barley, oats) Israel, Mexico, USA
P. zeae Maize, upland rice Tropical: South and Southeast Asia;

Radopholus (burrowing nematodes)
Necrosis of roots and tubers, rots, root
breakage, toppling
R. citri Citrus Tropical: Indonesia
R. similis Bananas, citrus, root and Tropical: worldwide
tubers, coconut, tea, black
pepper and other spices
R. nativus Cereal and grain legumes Temperate: Australia

Appendix I. Examples of nematode genera and species

Nematodes and damage symptoms Main crops affected Distribution

Rotylenchulus reniformm nematodes)
Poor root growth, chlorosis, stunting
R. parvus Pigeon pea, sweet potato Tropical: Africa
R. reniformis Pineapple, vegetables Tropical: worldwide
R. variabilis Sweet potato Tropical: Africa
Rotylenchus (spiral nematodes)
Unthrifty growth
R. robustus Vegetables, tree seedlings Temperate: Europe, North America,
Scutellonema (spiral nematodes) Mainly in the tropics and Africa
Dry rot of tubers, poor root growth
S. bradys (yam nematode) Yams, cassava Tropical: West Africa, Caribbean
S. cavenessi Groundnuts Tropical: West Africa
Trichodorus (stubby root nematodes)
Shortened (stubby) blackened roots.
Transmit viruses
T primitivus
T viruliferus Sugarbeet, potato Temperate/subtropical: Europe, North
Reduced root growth
T obscurus Coffee Tropical: Africa
Tylenchorhynchus (stunt nematodes)
Stunting of roots Cereals, vegetables Temperate and tropical
Poor root growth, slow decline of trees
T semipenetrans (citrus nematode) Citrus Subtropical/tropical: citrus-growing
areas worldwide
Xiphinema (Dagger nematodes)
Root tip galling. Transmit viruses
X americanum Trees, grape, Temperate/subtropical: worldwide
X diversicaudatum Roses, grape, soft fruits Temperate: Europe, North America,
Australia, New Zealand
X index Grape, fruit trees, rose Temperate: Europe, South America,
Mediterranean, southern Africa,
Eastern Australia

Adapted from Table 13.1 in J. Bridge and TD. Williams, Plant parasitic nematodes,
pp.140-162, in Waller et al. (2002) (courtesy of CABI Publishing).

Practical plant nematology

Appendix 2.

Basic identification of nematodes

This guide does not provide a taxonomic guide to plant parasitic nematodes; for this
see the References and Further Reading section. A very basic description of the most
common plant parasitic nematodes is however given below. Figure 39 shows the
visual differences between plant parasitic nematodes (head regions) and non-plant
parasitic nematodes, to aid differentiation.

Meloidogyne root-knot nematode
Adult female is pear shaped or spheroid with an elongated neck while males are
worm-like vermiformm).
Juveniles and females are endoparasitic causing galls.
Stylet is slender with basal knobs.
The eggs are laid in a gelatinous matrix.

Pratylenchus lesion nematode
Vermiform nematode.
Females with one ovary.
Relatively broad, flattened head and rounded tail.
Lip region is flat and the stylet is stout 14-19 pm long with massive basal knobs.
Causes lesions in plant roots.
All stages are infective and are migratory endoparasites.

Heterodera cyst nematode
Adult female lemon shaped forming cyst on maturity.
Female is semi-endoparasitic with only anterior portion inside the plant material.
Eggs are retained in the cyst but can additionally possess an egg mass.
Juvenile stylet is strong with prominent basal knobs.

Helicotylenchus spiral nematode
Lip region is high, rounded conical.
Nematode is usually coiled into a loose spiral or C shape.
Female with 2 ovaries with dorsally curved tail.
They are semi-endoparasites or ectoparasite and usually found in soil.

Scutellonema false spiral nematode
Basically the same description as Helicotylenchus, but:
The stylet is shorter and basal knobs are more pronounced.
The relaxed shape is straight or a slight C shape.
They are mostly ectoparasitic.

Xiphinema and Longidorus spear nematodes
Very long nematodes
The stylet is a long needle shaped structure without pronounced basal knobs.
The relaxed shape is usually straight.
They are ectoparasitic.

Appendix 2. Basic identification of nematodes

Plant parasitic nematodes:

Paratrichodorus [JB] Heterodera [JB]

Hemicriconemoides [JB]

Tylenchorhynchus [JB]

Hemicycliophora [JB]

Plectus [JB]

Scutellonema [JB]

Hoplolaimus [JB]

Meloidogyne J2 head [SM]

I "

Xiphinema [LA-B]

Filenchus [LA-B]

.. ......

Meloidogyne female head [LA-B]

Xiphinema [SM]

Helicotylenchus [SM] Longidorus anterior female head [LA-B]

Pratylenchus [SK]

Figure 39 continued overleaf

Practical plant nematology

Non-plant parasitic nematodes:

Mononchus [JB] Discolaimus [JB]

Figure 39. Comparison of various plant and non-plant parasitic nematode
head regions.
(Photographs by J. Bridge [JB], S. Mack [SM], L. Al-Banna [LA-B]
and S. Kelly [SK].)

Appendix 3.
Score sheets for measuring nematode damage

Root-knot gall (Meloidogyne spp.) scoring on cassava
Use a score combination of both roots and tubers when assessing mature harvested
plants or a score of roots only for assessment of roots removed from standing plants.

Cassava roots

1. No galls observed, feeder roots intact.

3. Numerous galls, about 50% of roots affected.

2. At least one gall observed.

4. Numerous galls, most roots affected.

5. Heavy galling on most roots,
with necrosis, and feeder roots
heavily affected or absent.

Practical plant nematology

Cassava plants

1. No galls observed, healthy feeder roots and tubers.

3. Galls obvious on roots, a few feeder roots and
tubers reduced in size.

2. At least one gall observed on roots.

4. Numerous galls, roots necrotic, and tubers reduced
in size.

5. Heavy galling on most roots, feeder roots largely
absent, and few tubers.

Appendix 3. Score sheets for measuring nematode damage

Root-knot gall scoring on carrot

4. Moderate galling.

OF S ti

5. Severe galling.

Practical plant nematology

Root-knot gall scoring on lettuce

1. No galling damage.

2. Slight galling. 3. Mild galling.

4. Moderate galling.

5. Severe galling.

Appendix 3. Score sheets for measuring nematode damage

Lesion scoring for banana roots
Adapted from Paul Speijer and Dirk De Waele (1997).

Practical plant nematology

Lesion scoring for Musa
Example of scoring five lengthwise sliced banana roots for root necrosis (%) of root
cortex surface showing necrosis caused by migratory lesion endoparasites (courtesy of
Paul Speijer and Dirk De Waele, 1997).

10 cm
% root necrosis
of each split root

% proportion of each split root
(5 x 20% 100)

: 2%



0% 5% 10% 15% 20%
% proportion of each split root
(5 x 20% = 100%)

Randomly select five funtional roots per sample (plant). Each one should be at least
10cm long. Slice each root lengthwise and discard one half. Score the other half of
the root for the percentage of root cortex showing necrosis. Each root contributes
20% of the whole sample so that when added up you get 100% for the 5 roots. So
if half the root exhibits necrosis then score it at 10%. If the root shows no necrosis
score it at 0% (see figure above). Once you've scored each root out of 20 then add up
the 5 scores to get the total percentage of necrosis across the whole sample.

Appendix 3. Score sheets for measuring nematode damage

Diagrammatic root-knot scoring chart
Courtesy of John Bridge and Sam Page (1980).

0 No knots on roots. 1 Few small knots, difficult
to find.

2 Small knots only but clearly
visible. Main roots clean.

5 50% of roots affected.
Knotting on some main roots.
Reduced root system.

8 All mainfroots, including tap
root, knotted. Few clean roots

3 Some larger knots visible.
Main roots clean.

6 Knotting on main roots.

9 All roots severely knotted.
Plant usually dying.

4 Larger knots predominate but
main roots clean.

7 Majority of main roots

10 All roots severely knotted. No
root system. Plant usually dead.

Practical plant nematology

Cyst damage scoring sheet for wheat
Adapted from A.D. Rovira in Brown and Kerry (1987).

1 No damage, clean.

2 Slight damage.

3 Mild damage. 4 Moderate damage.

5 Severe damage.

Useful networks and organizations

Afro-Asian Society of Nematologists

Australasian Association of Nematologists

Brazilian Nematological Society

Cereal Nematode Network

Chinese Society of Plant Nematologists

Nematology Initiative in East and Southern Africa

Egyptian Society of Agricultural Nematology

European Society of Nematologists

International Federation of Nematology Societies

Japanese Nematological Society

Nematological Society of India

Nematological Society of Southern Africa

Organization of Nematologists for Tropical America

Society for Invertebrate Pathology

Society of Nematologists

West and Central African Nematology Network

Photos: All photographs are by the authors unless otherwise credited.
Editing, design, layout andproofreading: Green Ink Publishing Services Ltd, UK (www.greenink.co.uk)
Printing: Pragati Offset Pvt. Ltd, India (www.pragati.com)

Plant parasitic nematodes are ever-present in farmers' fields, but the damage they cause is often attributed to other
pests and diseases or other crop problems. In developing countries in particular, where resources and facilities are
scarce, it is difficult to accurately identify and quantify the nematode problem.

This guide aims to help overcome this limitation by providing an easy-to-follow reference for assessing plant
parasitic nematode problems. It provides clear instructions, with many illustrations, on procedures for collecting
and processing samples for nematode assessment, as well as information on accessing further identification and
diagnosis support. The manual is aimed at technicians, field workers, extension agents and others with an interest
in crop production and crop protection, particularly in those parts of the world where access to expert help and
advanced facilities is limited. This guide will hopefully simplify some aspects of nematology, and help to lessen the
mystery surrounding this crop production problem.

ISBN 978-131-294-7

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