GHENT UNIVERSITY.
FACULTY OF SCIENCE.
DEPARTMENT OF BIOLOGY.
Academic Year: 2015-2016
Host plant status of different green manure plants for Pratylenchus
penetrans and Meloidogyne chitwoodi
Alexander Mbiro
Promoter and supervisor: Prof. Dr. Ir. Wim Wesemael
Thesis submitted to obtain the degree of Master of Science in Nematology
Host plant status of different green manure plants for
Pratylenchus penetrans and Meloidogyne chitwoodi
Alexander MBIRO1, Wim .M.L.WESEMAEL1,2
1Ghent University, Department of Biology, K.L. Ledeganckstraat 35, 9000, Gent, Belgium
2Institute for Agricultural and Fisheries Research (ILVO), Burgemeester Van Gansberghelaan
96, 9820 Merelbeke, Belgium
Declaration
Submitting in this thesis, I declare that this work has never been submitted either in a whole or
part to this or any other institution of higher learning for any other degree and is, except where
otherwise stated, the original work of the author.
Abstract
Nine different green manure crops (different species or cultivars) were evaluated for their
potential use in management of Pratylenchus penetrans and Meloidogyne chitwoodi. Firstly, a
resistance screening test for each cultivar was carried out in small yellow tubes filled with soil
and inoculated with 100 P. penetrans (juveniles and adults) or M. chitwoodi (second-stage
juveniles). Eight weeks after inoculation, each cultivar was assessed for its resistance or
susceptibility to P. penetrans and M. chitwoodi. Based on the reproductive factor, bird’s-foot
trefoil cv. Franco, English ryegrass cv. Meltador, arugula cv. Nemat and fodder radish line
RsV79/80 were resistant to P. penetrans. For M. chitwoodi, Alfalfa cv. Alpha, bird’s-foot trefoil
cv. Lotar, bird’s-foot trefoil cv. Bull and fodder radish line RsV79/80 showed less than one egg
mass per root system being formed eight weeks after inoculation. These cultivars showed a high
level of resistance to M. chitwoodi multiplication. Secondly, a host evaluation pot test was
carried out for five green manure plants either singly or a mixture of cultivars, each inoculated
with 500 P. penetrans (juveniles and adults) or M. chitwoodi (second-stage juveniles). Each
cultivar or mixture was harvested 8 weeks after inoculation. Nematodes were extracted from
both roots and soil to assess the final nematode population. Fodder radish line RsV79/80, arugula
cv. Nemat and arugula- fodder radish mixture were non to poor hosts to both P. penetrans and
M. chitwoodi. Based on our results selected green manure crops or mixtures of green manure
crops can be used to control both P. penetrans and M. chitwoodi.
Key words: Plant-parasitic nematodes, resistance, susceptibility, reproductive factor.
INTRODUCTION
Plant-parasitic nematodes (PPN) do feed, reproduce on living plants and are capable of active
migration in the rhizosphere, on aerial plant parts and inside the plant especially the root (Dong
& Zhang, 2006). Decraemer et al. (2013), estimates that there are over 4000 species of plantparasitic nematodes described to date and they cause an important constraint on the agricultural
crop production globally. Economic loss caused by plant-parasitic nematodes was estimated to
be at $US80 billion per year by end of 2010 (Nicol et al., 2011). This figure is most likely to be
an underestimation, as most agricultural farmers in tropics are unaware of even the existence of
nematodes due to their microscopic nature, the atypical symptoms caused and their synergistic
association with other pathogens (De Waele & Elsen, 2007; Jones et al., 2013).
On a worldwide basis, root-knot nematode (Meloidogyne spp.), cyst nematode (Heterodera spp.
and Globodera spp.) and root lesion nematode (Pratylenchus spp.) are the first three in their
respective order of the ten most important and common genera of plant-parasitic nematodes
(Jones et al., 2013). The sedentary endoparasitic nematodes (Globodera, Heterodera,
Meloidogyne) (Back et al., 2002), semi-endoparasitic nematode (Rotylenchulus) and migratory
endoparasitic nematodes (Pratylenchus, Ditylenchus, Bursaphelenchus, Aphelenchoides and
Anguina) are the genera most commonly reported to be involved in disease complexes with
fungal and bacterial pathogens (Back et al., 2002; Moens & Perry, 2009).
As of October 2015, a total of 101 root-knot nematode species (Meloidogyne spp.) have been
described (Wesemael pers.comm). The most economically important species of Meloidogyne in
cooler climates are; M. naasi, M. hapla, M. chitwoodi and M. fallax, while M. arenaria, M.
javanica and M. incognita are the most common species in warmer conditions of southern
Europe (Moens & Perry, 2009; Wesemael et al., 2011). Meloidogyne chitwoodi and M. fallax are
the two most important species in Europe because they are quarantine pests (De Waele & Elsen,
2007; Wesemael et al., 2011).
Meloidogyne chitwoodi can parasitize a wide range of host plants which can be classified as
good hosts, maintenance hosts, poor hosts or non-hosts depending on host suitability for
nematode reproduction (Ferris et al., 1993). The classification may also vary with the nematode
ability to adapt to a particular environmental condition and the management system used.
1
Some of M. chitwoodi hosts are crop plants of economic importance, green manure plants (Cherr
et al., 2006) and common weed species (Kutywayo & Been, 2006). The common excellent crop
hosts include; potato (Solanum tuberosum), carrot (Daucus carota) and tomato (Solanum
lycopersicum) (Ferris et al., 1993). Barley (Hordeum vulgare), maize (Zea mays), oats (Avena
sativa), sugarbeet (Beta vulgaris var. saccharifera), wheat (Triticum aestivum) are maintenance
hosts (Ferris et al., 1993) while poor to non-host plants include; amaranth, oilseed radish, oilseed
rape, and safflower (Ferris et al., 1993). As for the green manure crops; a number of cultivars of
oil radish are known to be maintenance hosts while buckwheat (Fagopyrum esculentum),
rapeseed (Brassica napus), sundangrass (Sorghum vulgare), horsebean (Canavalia ensiformis),
velvetbean (Mucuna deeringina) castor (Ricinus communis), showy crotalaria (Crotalaria
spectabilis), joint-vetch Aeschynomene Americana), marigolds (Tagetes minuta and T. erecta),
sesame (Sesamum indicum cv. Paloma), barley (H. vulgare), are known to be in the range of poor
to non-host plants (Al-Rehiayani & Hafez, 1998). However, studies indicate that rapeseed as
green manure crop significantly reduces potato damage caused by M. chitwoodi (Mojtahedi et
al., 1993)
Genus Pratylenchus differ from root-knot nematode (RKN) in that they enter and leave root
tissues during their life cycle, move actively through soil and penetrate the root tissues for
feeding and reproduction (Esteves et al., 2015). Reduced growth, occasional yellowing of the
foliage and severe necrosis in roots and tubers are the major symptoms associated with the
nematodes of Pratylenchus (Castillo & Vovlas, 2007). With over 70 species of Pratylenchus
(Duncan et al., 2013), the most important species in agriculture are; P. crenatus, P. neglectus, P.
penetrans, P. thornei, P. brachyurus, P. coffeae (Jones et al., 2013). As potato is a good host of
Pratylenchus, P. penetrans was the most abundant species followed by P. neglectus, P. crenatus
and lastly P. thornei in Portugal (Esteves et al., 2015).
The migratory endoparasitic nematode P. penetrans has a wide host range, with over 350 host
plant species recorded (Mizukubo & Adachi, 1997; Moens & Perry, 2009; Duncan et al., 2013).
Among them are cultivated crops, fruits, vegetables, green manure crops and numerous weeds.
Thus this makes the species difficult to manage with crop rotation (Jensen, 1953; Townshend &
Davidson, 1960; Manuel et al., 1980). The common cultivated crops attacked by P. penetrans
are; apple, cherry, citrus, roses, tomato, potato, corn, sugarbeet, ornamentals like Narcissus spp.
2
(Slootweg, 1956; Duncan et al., 2013). The seemingly good host green manure crops includes;
kura clover (Trifolium ambiguum), alsike clover (Trifolium hybridum), white clover (Trifolium
repens), oat (Avena sativa), and rye (Secale cereale) (Thies et al., 1995). On the other hand, P.
penetrans does not reproduce well on some green manure plants and they are termed as poor or
non-hosts. Among them are; pearl millet (Pennisetum glaucum), tall fescue (Festuca
arundinacea), perennial ryegrass (Lolium perenne), forage sorghum (Sorghum bicolor),
sudangrass (Sorghum sudanense), sweetclover (Melilotus alba), crownvetch (Coronilla varia)
and MNGRN-16alfalfa (Medicago sativa) (Thies et al., 1995).
Many different control strategies are being applied in agriculture and these include: chemical,
physical, cultural, genetic (resistance) and biological control (Nicol & Rivoal, 2008).
Specifically, effective control of root-knot and lesion nematodes commonly calls for the
integrated pest management approach, including the use of crop rotations with non-host plants,
the use of resistant cultivars if available, fallow, organic amendments (Haydock et al., 2013;
Viaene et al., 2013; Kruger et al., 2015). The overall aim of using these management strategies is
to decrease the nematode population densities below damage thresholds before the next primary
host crop is cultivated (Nicol & Rivoal, 2008).
Green manure crops are crops of economic importance to the soil and crop productivity. They
have been in existence in traditional agriculture for many decades but large scale agricultural
systems did not entirely adopt their use due to efficient and cost effective use of fertilizers and
pesticides that have been readily available on the market (Viaene et al., 2013). The primary
benefits of green manure crops are to; 1) protect the soil from erosion; 2) increase soil nutrients;
3) improve soil properties such as water-holding capacity; and 4) provide an energy source for
microbes, contributing to soil activity and biodiversity (Cherr et al., 2006; Ortiz et al., 2015).
Secondarily, green manure crops are used and applied in agricultural fields in the control of
soilborne pathogens and their mechanisms of action vary with species (Ortiz et al., 2015). Green
manure treatments may play a role in disease management by changing the Streptomycete
communities in soils, leading to pathogen suppression (Wiggins & Kinkel, 2005) or resulting in
bacterial communities that may induce plant systemic resistance (Cohen et al., 2005).
Some green manure crops act as non-host or poor host to the pathogen, produce allelochemicals
that are toxic or aggressive toward pathogens or stimulate antagonists of plant parasitic
3
nematodes (Hooks et al., 2010; Ortiz et al., 2015). A good manure crop grown for the
management of plant-parasitic nematodes is characterized by having non-host or poor host status
for the target nematode and/ or being suppressive to the population growth in the soil (Viaene &
Abawi, 1998).
Additionally, though different manure crops have been proved to be a ready source of
nematicidal compounds to suppress a number of plant-parasitic nematodes, some are good hosts
as well. Within a single green manure crop species, its cultivars may vary in their suppressive
effect to a single particular nematode species. In spite of all management strategies applied, P.
penetrans and M. chitwoodi continue to be a big threat to agricultural sector in Europe as far as
crop yield is concerned. Therefore, this study screened for resistance of different green manure
plant cultivars for P. penetrans and M. chitwoodi. In addition, the study evaluated the
reproductive potentials of P. pentrans and M. chitwoodi on different green manure cultivars
planted either singly or as a mixture.
MATERIALS AND METHODS
Nematode culture
Root-knot nematode, M. chitwoodi and root-lesion nematode, P. penetrans were used throughout
the study. The population of M. chitwoodi originated from a field in Belgium and was
maintained as a pure culture at Institute for Agricultural and Fisheries Research (ILVO) on
tomato raised under greenhouse conditions (18 - 23°C, with 16:8 hours of light and darkness
respectively), in a 16 cm diameter pot size with 2 litre volume of soil. The nematodes were mass
cultured on potato tubers (Solanum tuberosum cv. Bintje) in closed containers.
Prior to planting, the potato tubers were thoroughly washed in tap water to remove soil particles
and thereafter disinfected with a 5% NaOCl solution for a maximum of 4 minutes. After
disinfection, the tubers were rinsed with tap water to remove the disinfectant (NaOCl). The
rinsed tubers were spread on soft tissue paper and left at room temperature with maximum light
for about three weeks to sprout. 200 g of sterilized white river sand was placed in each closed
plastic container (10 cm diameter and 0.5 litre volume) and 30 ml of tap water was added. One
sprouted potato tuber was placed in each closed container with sprouting roots in the soil. The
closed containers were kept in the dark for 2 weeks to allow establishment of roots. After root
4
establishment, each closed container was singly inoculated with 2000 second-stage juveniles (J2)
of M. chitwoodi. The inoculated closed containers were stored in an incubator at 20-22°C in the
dark room for 10-14 weeks to allow nematode reproduction. After reproduction, potato roots
were chopped, placed on Baermann funnel to extract nematodes (Baermann, 1917) and freshly
hatched J2 which were used for inoculation were collected after 24 hours.
The P. penetrans population originated from a maize field in Belgium and was maintained and
mass cultured as pure culture on carrot discs at ILVO. Unblemished carrots with a cylindrical
shape and fresh leaves were selected. All the working tools plus laminar flow were disinfected
with 70% ethanol. The carrots were thoroughly cleaned with distilled sterile water before
peeling. With the help of forceps, the carrots were peeled using a peeling knife, by first dipping
in 70% ethanol for a few seconds and flamed over spirit lamp. The peeling knife was
continuously moistened with 70% ethanol between peelings to avoid contamination. The peeled
carrots were cut into small discs of about 1 cm thickness with a 3-4 cm diameter using the
sterilized knife. Using the sterilized forceps one carrot disc was placed into sterile disposable
petri dishes of 5 cm diameter and sealed with parafilm. The carrot discs were then kept in the
incubator at 21°C for 3 weeks.
The petri dishes with the prepared carrot discs were removed from the incubator and placed on
the laminar flow bench. With a micropipette, a solution of nematode inoculum containing about
30 infective juveniles and few males were inoculated on top of the discs and petri dishes were
sealed with parafilm. The inoculated discs were placed in a dark container and later transferred to
an incubator maintained at a temperature of 21°C for a period of 10 weeks. Callus formation
(whitish matter) on the surface of the carrot discs was observed, indicating formation of healthy
cultures during incubation. After 10 weeks, infective juveniles and adults were extracted from
infected carrot discs with brown coloring using a modified Baermann funnel technique under a
mistifier after 24 hours. The freshly hatched mixtures of juveniles and adults were used for
inoculation of different green manure plants for the two tests.
Green manure crops and cultivars
A combination of eleven cultivars was used in this study. Eight cultivars of green manure plants
and one candivar (fodder radish, Raphanus sativus, line RsV79/80) were included in the study.
Two non-green manure plants; maize and tomato were used as positive controls in both
5
resistance screening and host evaluation tests of P. penetrans and M. chitwoodi respectively
(Table 1). Meanwhile a fallow was used as a negative control for host evaluation test for each
nematode species.
Table 1: Plant common names, scientific names seed source and 1000 grain weight.
1000 grain
Common name
Scientific name
Cultivar
weight (g)
Seed source
Red clover
Trifolium pretense
Lemmon
2
ILVO, Belgium
White clover
Trifolium repens
Melital
1
ILVO, Belgium
Bird’s-foot trefoil
Lotus corniculatus
Lotar
1.3
Oseva Uni, Czech Republic
Feldsaaten Freudenburger,
Bird’s-foot trefoil
Lotus corniculatus
Bull
1.5
Germany
Bird’s-foot trefoil
Lotus corniculatus
Franco
1.1
Italy
English ryegrass
Lolium multiflorum
Meltador
3
ILVO, Belgium
Alfalfa
Medicago sativa
Alpha
2
Barenbrug, The Netherlands
Arugula
Eruca sativa
Nemat
1.6
Alliance, Belgium
Fodder radish
Raphanus sativus
RsV79/80
16.4
ILVO, Belgium
Maize
Zea mays
LG3220
32.6
Limagrain, Belgium
Tomato
Solanum lycopersicum
Marmande
1.4
AVEVE, Belgium
Resistance screening test
Resistance screening of each cultivar was carried out in small yellow tubes (RLC4 type) of 3 cm
diameter, and 16 cm height with a surface area of 7x10-4m2 made by Stuewe and Sons, USA.
One seed from each cultivar was placed in a plastic yellow tube containing soil sterilized at
100°C for 16hours. The soil comprised of 74% sand, 14% sandy loam, 6% clay, 5% loam, 1%
organic matter content and a neutral pH.
After seed germination and root establishment, each tube was inoculated with 100 J2 of M.
chitwoodi or 100 P. penetrans (a mixture of juveniles and adults). The plants were watered daily,
grown in the greenhouse at a temperature range of 18-23°C and received 16:8 hours of light and
darkness respectively. The experiment was terminated 8 weeks after nematode inoculation.
6
Plant harvest during resistance screening test
For M. chitwoodi, roots were thoroughly washed clean and separated from the soil. The roots
were then dipped in a 1 litre solution of 0.15 g phloxine B for 15 minutes to stain the gelatinous
matrix of egg masses produced by the female M. chitwoodi on the roots. After staining, root
systems were rinsed in tap water and the number of egg masses per root system were observed
and counted using a binocular microscope and thereafter quantified. The soils were not
considered and hence it was discarded after washing the roots.
For P. penetrans, roots were thoroughly washed, chopped in small pieces of about 2 cm long and
macerated in a laboratory blender (Waring commercial) for one minute. The blended roots were
added to the beaker containing soil suspension. The root soil mixture was subjected to an
automated zonal centrifuge technique (Hendrickx, 1995).
The automated zonal centrifuge technique works based on the principle of density differences;
root and soil samples are first diluted to 1 litre and half of the dilution is taken up by the machine
for extraction. The extracted sample of 500 ml is subjected to; 1), MgSO4 at a density of 1.20
kg.m-3 which plays a role in separating particles with lower and higher specific gravity than its
own specific gravity. 2), water so that nematodes are retained at the interface with the MgSO4
solution and 3) kaolin suspension which is added at the end of the centrifugation cycle to the
rotor to avoid soil particles, root and other debris from mixing with the nematode suspension
when the centrifugation process stops. At the end of the centrifugation, a supernatant of clean
water and MgSO4 containing the nematodes is collected in a small beaker of 150ml via the
hollow shaft of the rotor (figure 1). The rotor and the tubes are cleaned automatically after each
sample to avoid contamination of samples (Wander et al., 2007).
The final population (Pf) of P. penetrans for each sample was obtained by counting nematodes in
the whole 40 ml of the supernatant and multiplying by a factor of 2. These nematode final
population (Pf) included eggs, juveniles and adults from both organic (root) and mineral (soil)
fractions.
Host status evaluation test
The evaluation of host status of individual or plant mixtures was carried out in pots of 16 cm
diameter, 15.5 cm height with a 2 litre volume of soil and surface area of 0.02 m2. The number of
seeds planted in the pot was determined depending on the surface area of the pot, the seed
7
density of the plant/cultivar and whether planted individually or as a mixture. Table 2 shows the
details of the plant/cultivar in relation to seed density. White clover, red clover, English ryegrass,
fodder radish and arugula were singly planted in pots. A fallow for each nematode species was
also set up without any green manure crop planted. Two mixtures were set up for each nematode
species and these included the clover- English ryegrass mixture, and arugula- fodder radish
mixture. Seeds from each cultivar(s) were sown in pots containing sterilized soil with soil
properties as mentioned above.
Table 2: crop seed density and number of seeds per pot.
seed
Plant
density/hectare
seed density/pot
(cultivar)
(g/10000 m2)
(g/0.02 m2)
Seeding
No of seeds/pot
status
Red clover
20000
0.04
20
Single
White clover
10000
0.02
20
Single
English ryegrass
30000
0.06
20
Single
Arugula
8000
0.016
10
Single
Fodder radish
40000
0.08
6
Single
Red-white clover-
7000,
0.014, 0.006,0.04
7, 6, 13
Mixture
ryegrass
3000,20000
Arugula-fodder
radish
Mixture
8000, 40000
0.016, 0.08
10, 6
After seed germination and root establishment, each pot was inoculated with an initial population
(Pi) of 500 J2 of M. chitwoodi or 500 P. penetrans (a mixture of juveniles and adults). The plants
were watered daily, grown in the greenhouse at a temperature range of 18-23°C, received 16:8
hours of light and darkness respectively. The experiment was terminated 8 weeks after nematode
inoculation. The experimental setup was a randomized complete block design with five
replicates per plant/mixture for each nematode species.
8
Nematode extraction during host status test
The organic (root) and mineral (soil) fractions were extracted separately. For both M. chitwoodi
and P. penetrans roots were thoroughly washed clean, fresh root weight taken, chopped in small
pieces of 2 cm long, homogenized and a subsample of 5 g was macerated in a laboratory blender
(Waring commercial) for one minute. The subsample of blended roots was sieved and added in a
plastic beaker. After homogenization of 2000 cm3 soil, a subsample of 200 cm3 of soil was
sieved and added in a 1 litre plastic beaker. The root and soil subsamples separately were
subjected to an automated zonal centrifuge technique (Hendrickx, 1995) as described above.
After automated zonal centrifugation, the collected nematode suspension was left to settle down
for at least 3 hours and thereafter the supernatant was removed with a hand controlled vacuum
pump machine (Vacuum brand BVC 21 NT VARIO) to reduce the volume of the nematode
suspension to about 40ml for easy counting and quantification. The nematode population of both
M. chitwoodi and P. penetrans for each sample was obtained by counting all the nematodes
(eggs, juveniles and adults) in 40 ml of the supernatant.
As for the root subsample, the machine only extracts 500 ml out 1000 ml, therefore to obtain the
nematodes in whole subsample it was multiplied by a factor of 2. If the root system weight was
more than 5 g, the nematode final population (Pf) was extrapolated by multiplying the nematodes
in the subsample with actual root weight of the bulk sample divided by the subsample of the root.
For the soil subsample, the machine only extracts 100 cm3 out 200 cm3, therefore to obtain the
nematodes final population (Pf) in whole bulk sample it was multiplied by a factor of 20.
9
Figure 1: A schematic representation of automated zonal centrifuge machine (courtesy of
Wim Wesemael).
Data analysis
The number of egg masses of M. chitwoodi for screening tests and final nematode population
(Pf) for P. penetrans in both tests and M. chitwoodi in host status test were subjected to analysis
of variance (ANOVA) using a software program R x64 3.2.3. Differences among treatment
means were compared using Fisher’s least significant differences (LSD) at P < 0.05 and data was
normalized by log transformation. Nematode reproductive factor for each species was equally
calculated (Rf = Pf/Pi).
Based on the Rf, the plant cultivars were classified under five different categories (Schomaker et
al., 2013) as follows.
Non-host = (Rf <0.15), Poor host = (Rf < 1.0 ≥ 0.15), Maintenance host = (Rf ≤ 2.0 ≥1.0), Good
host (Rf≤ 4.0 ≥2.0) and Excellent host (Rf >4.0)
10
RESULTS
Resistance screening tests
Pratylenchus penetrans
600
Mean nematode final populations (Pf)
a
500
b
400
b
b
300
cb
Pf
Pi
cb
200
cd
cd
100
d
100
d
0
BFT- Bull BFTLotar
BFTRed
Franco clover
White Ryegrass Alfalfa Arugula Fodder Maize
clover
radish
Figure 2: Mean population of P. penetrans from a combination of organic and mineral soil
fractions of different green manure plant cultivars extracted 8 weeks after inoculation with 100
(juveniles and adults) in yellow tubes. Error bars show standard error and letters indicate
significantly similar and dissimilar groups (n= 6 and P ≤ 0.05). Pf = Nematode final population,
Pi = Nematode initial inoculation, BFT = Bird’s-foot trefoil.
A non-green manure and susceptible control (maize cv. LG 3220) supported the highest
nematode reproduction with the highest reproductive factor of 4.5 to P. penetrans in comparison
to other green manure plants and the difference was significantly different (P< 0.05) from all
other green manure plants (Fig. 2). Nematode reproduction rates on alfalfa cv. Alpha, bird’s-foot
trefoil cv. Bull and red clover cv. Lemmon were not significantly different with reproductive
factors of 2.83, 2.56 and 2.55 respectively. Bird’s-foot trefoil cv. Lotar and white clover cv.
Melital showed reproductive factors of 2.20 and 1.55 respectively and were not significantly
different from each other though significantly different from the other cultivars (Fig. 2). The
final populations of P. penetrans on bird’s-foot trefoil cv. Franco and arugula cv. Nemat were
11
less than the initial population with reproductive factors of 0.93 and 0.89 respectively. Ryegrass
cv. Meltador and fodder radish line RsV79/80 showed the lowest reproductive factors of 0.3 and
0.24 respectively, with lowest final populations below the initial nematode population and
significantly different from the other crop hosts (Fig. 2).
Meloidogyne chitwoodi
Mean number of egg masses
30
a
25
20
15
b
10
c
5
c
c
c
c
c
BFTLotar
BFT-Bull
c
c
0
Alfalfa Ryegrass
Red
clover
White
clover
BFT- Arugula Fodder Tomato
Franco
radish
Figure 3: Mean number of egg masses observed on roots of different green manure cultivars 8
weeks after inoculation with 100 second-stage juveniles of M. chitwoodi. Error bars show
standard error and letters indicate significantly similar and dissimilar groups (n= 10 and P ≤
0.05), BFT = Bird’s-foot trefoil.
A non-green manure and susceptible control tomato cv. Marmande was observed to have the
highest mean number of egg masses per root system (20.7) and was significantly different from
all other green manure plants (P< 0.05) (Fig. 3). Generally all the green manure plants did not
show high nematode multiplication with the observation of less number of egg masses on the
12
root if not absent for some plant cultivars. Among the green manure plants White clover cv.
Melital had the highest mean number of egg masses (8.3). It was significantly different from the
rest of green manure plants and the susceptible control tomato cv. Marmande (P< 0.05).
Ryegrass cv. Meltador, arugula cv. Nemat, red clover cv. Lemmon and bird’s-foot trefoil cv.
Franco had mean number of egg masses greater than one but less than eight (3.1, 1.7, 1.6 and
1.6) respectively and they were not significantly different from each other. Alfalfa cv. Alpha,
bird’s-foot trefoil cv. (Bull and Lotar) and fodder radish line RsV79/80 had a mean number of
egg masses less than one and they were not significantly different from each other. Above all no
single egg mass was observed on the roots of fodder radish line RsV79/80 (Fig 3).
13
Host evaluation tests
P. penetrans
Table 3: The number of eggs, juveniles and adults in organic and mineral fraction and their
respective reproductive factor (Rf) on different green manure plant cultivars extracted 8 weeks
after inoculation with 500 P. penetrans (mixture of juvenile and adult stages). A susceptible
maize control and a fallow were subjected to the same inoculation.
Plants
Root weight Nematode population after 8 weeks
(g)
Organic
Mineral
Final
fraction
proportion
Populations
g-1
fresh 2000 g-1 soil
Rf
Host status
(Pf)
root
Arugula
3.65±0.27
6.0
236±116.10 de
258±110.97 e
0.52
Poor
Fodder radish
3.54±0.78
9.3
184±60.66 de
217.2±65.52 e
0.43
Poor
ESFR Mix
2.30±0.20
5.9
176±43.36 de
189.6±35.65 e
0.38
Poor
Ryegrass
14.23±1.44
7.2
256±63.88 de
358±81.895 e
0.72
Poor
Red clover
4.32±1.27
52.6
612±114.54 bc
839.2±145.74 bc
1.68
Maintenance
White clover
3.70±0.85
77.9
920±367.97 ab
1177.2±375.88 b
2.35
Good
Clover Mix
13.0±2.46
15.6
444±84.14 cd
647.2±167.47 cd
1.29
Maintenance
76±38.47 e
76.0±38.47 e
0.15
940±167.33 a
2632.4±391.97 a
5.26
Fallow
Maize
29.47±3.66
57.47
Excellent
Letters indicate significantly similar and dissimilar groups (n = 5 and P ≤ 0.05). Reproductive
factor (Rf) = final nematode population at harvest (Pf)/initial nematode inoculation (Pi). Values
are actual means with standard errors (±), ESFR = Arugula-fodder radish mix.
14
3500
a
Mean nematode final populations (Pf)
3000
2500
2000
Organic
proportion
Mineral
proportion
Pi
b
1500
cb
1000
500
cd
ed
ed
e
e
e
0
Arugula
Fodder
radish
ESFR Mix Ryegrass Red clover White Clover Mix Fallow
clover
Maize
Figure 4: Mean population of P. penetrans (organic and mineral soil fraction) of different green
manure plant cultivars extracted 8 weeks after inoculation with 500 juveniles and adults. Error
bars show standard error and letters indicate significantly similar and dissimilar groups (n = 5
and P ≤ 0.05). Pf = Nematode final population, Pi = Nematode initial inoculation, ESFR =
Arugula-fodder radish mix.
Inoculation of the different green manure crops with P. penetrans resulted in varying final
populations (Fig. 4). Higher nematode numbers were recovered from the soil than in the roots
with the exception of susceptible maize cv. LG3220, where more nematode numbers were
recorded in the root fraction. Nematode populations did not correspond to root weights of green
manure cultivars. More nematode numbers were recovered in plant cultivars with a relative small
root system (low root weight), with the highest numbers recovered from roots of white clover
and red clover respectively (Table.3). Nematode reproduction was highest on susceptible control
maize (Rf 5.25) followed by white clover and red clover, (Rf 2.35 and 1.68) respectively.
Nematode population recovered from clover- English ryegrass mixture was slightly higher than
the initial population (Rf 1.29). However, nematode populations recovered from ryegrass cv.
15
Meltador, arugula cv. Nemat, radish line RsV79/80 and arugula- fodder radish (ESFR) mixture
were less than the initial populations (Rf 0.72, 0.52, 0.43 and 0.38 respectively). Least root
weight was observed on arugula- fodder radish (ESFFR) mixture and this corresponded with the
minimum number of nematodes recovered from the root fraction in comparison to other green
manure plants. Nematode populations were lowest on fallow treatment with a reproductive factor
of 0.15. There were significant differences in total nematode numbers recovered from the
different groups of green manure plant cultivars as indicated in Fig. 4 (letters indicating
significantly similar and dissimilar groups with P ≤ 0.05).
16
M. chitwoodi
Table 4: The mean number of eggs, juveniles and adults in organic and mineral fraction and their
respective reproductive factor (Rf) on different green manure plant cultivars extracted 8 weeks
after inoculation with 500 second-stage juveniles of M. chitwoodi. A negative control (fallow)
was subjected to the same inoculation.
Plant
Root weight Nematode population after 8 weeks
(g)
Organic
Mineral
Final
fraction
proportion
populations
g-1
fresh 2000 g-1 soil
Rf
Host status
(Pf)
root
Red clover
3.64±0.56
100.3
448±180.33 c
813.2±163.65 c
1.63
Maintenance
White clover
3.33±1.29
201.8
1312±263.29 a
1984±347.69 b
3.97
Good
Ryegrass
11.09±1.76
121.1
824±12.81 b
2167.6±396.91 b
4.33
Good
Clover mix
11.81±1.06
160.1
852±136.09 b
2743.2±332.22 a
5.49
Good
Fodder radish
3.02±0.22
0
236±38.47 cd
236±38.47 d
0.47
Non
Arugula
2.29±0.71
6.8
312±71.55 cd
327.6±64.92 d
0.65
Poor
ESFR mix
2.64±0.51
3.9
104±81.73 d
114.4±83.51 d
0.23
Poor
124±26.08 d
124±26.07 d
0.25
Fallow
Letters indicate significantly similar and dissimilar groups (n = 5 and P ≤ 0.05). Reproductive
factor (Rf) = final nematode population at harvest (Pf)/initial nematode inoculation (Pi). Values
are actual means with standard errors (±). Pf = Nematode final population, Pi = Nematode initial
inoculation, ESFR = Arugula-fodder radish mix.
17
3500
a
3000
b
Mean nematode final populations (Pf)
2500
b
2000
Organic
proportion
Mineral
proportion
1500
Pi
c
1000
500
d
d
d
d
ESFR Mix
Fallow
0
Red clover
White
clover
Ryegrass Clover Mix Fodder
radish
Arugula
Figure 5: Mean population of M. chitwoodi from a combination of organic and mineral soil
fraction of different green manure plant cultivars extracted 8 weeks after inoculation with 500
second-stage juveniles. Error bars show standard error and letters indicate significantly similar
and dissimilar groups (n = 5 and P ≤ 0.05). Pi = initial nematode inoculation, ESFR = Arugulafodder radish mix.
The clover-ryegrass mixture supported the highest nematode reproduction with the highest
reproductive factor of 5.49 and significantly different from all the green manure plants and a
fallow (P < 0.05). Ryegrass cv. Meltador and white clover cv. Melital equally had relatively high
reproductive factors of 4.33 and 3.97 respectively in comparison to other hosts and the recovered
18
final populations were not significantly different from each other (P< 0.05) (Fig 5). Arugula cv.
Nemat, fodder radish line RsV79/80, arugula- fodder radish (ESFR) mixture and fallow did not
support nematode reproduction yielding final populations lower than the initial populations (Rf
0.65, 0.47, 0.23 and of 0.25) respectively The least number of nematodes was recovered from
arugula- fodder radish (ESFR) mixture though there were no significant differences with the
individual arugula cv. Nemat and fodder radish line RsV79/80 (Fig. 5).
DISCUSSION
Resistance screening of different green manure plants
This study presents results based on cultivar of a particular plant species. Alfalfa cv. Alpha had a
reproductive factor of 2.83. However, these findings are not in agreement with findings of a
greenhouse study carried out in Ontario, Canada which showed that the final population of P.
penetrans on alfalfa cv. Saranac was less than the initial population (Townshend & Potter, 1976).
This could be explained based on the fact that there could be differences in the level of resistance
between the two cultivars. Different Alfalfa cultivars are mostly like to influence nematode
invasion and multiplication differently.
The bird’s-foot trefoil cv. Bull, Lotar and Franco had reproductive factors of 2.56, 2.20 and 0.93
respectively. Therefore, bird’s-foot trefoil cv. Bull and Lotar are susceptible to P. penetrans
while bird’s-foot trefoil cv. Franco has a certain level of resistance to P. penetrans reproduction
similar to bird’s-foot trefoil cv. Empire which equally yielded a final population less than the
initial inoculation population in an experiment of Townshend and Potter (1976). Resistance to
nematode could be as a result of unknown plant proteins or unsuitable host cell for nematode
reproduction (Gheysen et al., 2006). This is the first report on screening of these three cultivars
for P. penetrans in temperate region. Therefore, pot greenhouse and field tests should be carried
out to assert the host status.
In many research studies, root gall index was used as factor for resistance screening of different
plants for the invasion of Melodoigyne species. Presence or absence of galls may not be
correlated with any Melodoigyne spp. reproduction (Al-Rehiayani & Hafez, 1998) as root galling
depends on physiological reaction of a particular plant (Wesemael, pers.comm). It is on this basis
the study decided to use egg mass numbers to assert invasion and reproduction of M. chitwoodi
on different green manure plant cultivars. However, present screening criterion of using number
19
of egg masses is backed up with root galling information which is available in many previous
studies.
Susceptible control and a non-green manure plant (tomato cv. Marmade) had the highest number
of egg masses from all the green manure plants and this is in agreement with the findings of
Kutywayo and Been (2006), where similarly a high number of egg masses and galls on the roots
were observed. Arugula cv. Nemat had a low mean number of egg masses compared to the
susceptible control tomato (cv. marmade). These findings substantiates the results of
Melakeberhan et al. (2006) who recorded more galling on tomato cv. Rutgers than arugula cv.
Nemat after inoculation with Meloidogyne hapla under greenhouse conditions. Few egg masses
were recovered from alfalafa cv. Alpha and this is in agreement with previous findings of
(Mermans, 2015) where fewer numbers of egg masses on alfalfa cv. Alpha under similar
greenhouse conditions were equally observed. Thus these two recent studies indicate that alfalfa
cv. Alpha has some level of resistance to M. chitwoodi multiplication. However, Griffin and
Rumbaugh (1996) reported that alfalfa cv. 1 and alfalfa cv. 2 had 60-80 % of the root tissues
galled hence contributing to reproductive factors of 12 and 10 respectively. This indicates that
not all alfalfa cultivars are resistant to the nematode and care should be taken when choosing a
cultivar for nematode management.
Among the bird’s-foot trefoil plants, high mean number of egg masses were observed on cv.
Franco and lowest on cv. Lotar. Griffin and Rumbaugh (1996) also reported the presence of galls
on bird’s-foot trefoil in greenhouse experiment with a gall index of 2.5 (60-80% roots galled)
and reproductive factor of 4.0, thus further highlighting its susceptibility to M. chitwoodi.
The mean number of egg masses for red clover cv. Lemmon were significantly different from
white clover cv. Melital, (1.6 and 8.3) respectively. This study contradicts with Griffin and
Rumbaugh (1996) who reported that red clover is susceptible and white clover resistant to M.
chitwoodi in the greenhouse. This could also be explained in terms of cultivar differences were
different cultivars influence nematode reproduction differently. No egg masses were found on
the roots of fodder radish line RsV79/80, deviating from the many galls observed in a
greenhouse test of fodder radish cultivars (Melodie, Adagio 5a-3 and Trez) by Ferris et al.
(1993). In a greenhouse study, Teklu et al. (2014) noted no differences between fodder radish
varieties (Radical, Doublet, Contra, Anaconda, Defender and Terranova) based on root galling
20
index. They concluded that all cultivars were partially resistant to M. chitwoodi. However,
fodder radish line RsV79/80 is a newly bred line which might have been developed with an
important trait of resistance to the important PPN of Europe. Egg masses were observed on the
root system of ryegrass cv. Meltador. Cook et al. (1999), showed that some cultivars of ryegrass
were heavily galled when inoculated with M. naasi. This could be of the reason that M. naasi has
been documented to be a host of many monocotyledonous plants (grasses and cereals)
(Wesemael et al., 2011)
For easy resistance screening of plants/crops or cultivars for Melodoigyne species, plant breeders
and Nematologists should use number of egg masses per root criterion. Egg masses are formed
on a susceptible plant while galling might not be present or difficult to observe. Secondly the use
of small yellow tubes in a greenhouse condition is simple, convenient and cost effective way of
screening.
Host evaluation different green manure plants
Maize cv. LG3220 (susceptible control) was found to be an excellent host and this is in
agreement with Kutywayo and Been (2006), who reported on maize cv. Husar as an excellent
host with reproductive factor of 6.4. During the pot experiment, maize exhibited a dense root
system which is of an advantage to the P. penetrans in accessing food and rapid multiplication.
Generally, the host evaluation study showed that ryegrass cv. Meltador is a poor host to P.
penetrans, findings which are in agreement with resistance screening based on the low
reproductive factor of 0.33. Abawi and Ludwig (1995) reported the similar results on treatment
of the nematode with ryegrass cv. Pennant.
Red clover cv. Lemmon and white clover cv. Melital are reported a maintenance and good host
respectively in the present study. This contradicts with the conclusions of Abawi and Ludwig
(1995) classifying both of them as intermediate hosts. However, their reproductive factor scaling
was different and maintenance and good hosts in this study are regarded as intermediate hosts in
their study. In a greenhouse study carried out in Canada, Papadopoulos et al. (2002) evaluated 18
cultivars and breeding lines of red clover and their reaction to inoculation of P. penetrans, only
one cultivar (AC Kingston) was highly susceptible to P. penetrans, cultivar Florex had low
levels of being invaded and three breeding lines (CRS 15,CRS 5 and CRS 11) registered both
low levels of invasion and low multiplication of the nematodes in the root. A four year field
21
micro-plot study in Abbotsford (British Columbia) found out that white clover was more
susceptible to P. penetrans (Vrain et al., 1996). Fodder radish line RsV79/80 is reported as a
poor host to P. penetrans in the current study, though at a species level, Hoek, (pers. comm)
indicates that fodder radish is regarded as a good host. Additionally, fodder radish cv. (Melodie
and Trez) were reported maintenance hosts to P. neglectus (Al-Rehiayani & Hafez, 1998). In
addition, planted Raphanus sativus (irrespective of cultivar) reduced population levels of P.
neglectus below 60% prior to planting potato in the field (Al-Rehiayani et al., 1999). Arugula cv.
Nemat is reported a poor host in this study. It is believed to be a trap crop hence suppressing
certain nematodes which enter the root with help of allelochemicals. The cultivar Nemat is
known to reduce plant-parasitic nematode populations and can be included in a crop rotation
scheme for organic farming (Curto et al., 2005; Kruger et al., 2013).
The clover- English ryegrass mixture is reported a maintenance host with a reproductive factor of
1.29, which is less than the reproductive factor of red clover cv. Lemmon (1.68) and white clover
cv. Melital (2.35) but greater than that of ryegrass cv. Meltador (0.72). Ryegrass cv. Meltador
was observed having a dense root system which covered the whole pot. It is therefore assumed to
have a contributing influence in reduction of nematode final populations of clover- English
ryegrass mixture. Ryegrass extensive root system and high forage is supplemented by the
Nitrogen supply from the leguminous clovers (Goh & Bruce, 2005) which in turn is of an
advantage to nematode reduction. Arugula- fodder radish mixture is a poor host along with the
singly planted arugula cv. Nemat and fodder radish line RsV79/80 to P. penetrans. Thus
integrating a combination of these green manure crops could be an advisable strategy in
nematode management nematode program.
It is important to note that green manure plant species well known for control of one type of
nematode may show susceptibility to other PPN (Cherr et al., 2006). Within a plant species,
different cultivars may not control a particular nematode species. Therefore, a green manure
mixture of different plant species is ideal in managing PPN and promoting organic farming.
The decline in nematode population in both fallows of M. chitwoodi and P. penetrans is in
agreement with the findings of (Kutywayo & Been, 2006; Wesemael & Moens, 2008), though
the experimental periods differ in weeks. Townshend (1984) revealed that nematodes while in
anhydrobiotic state can still thrive for about 110 weeks provided the loss of moisture is gradual.
Townshend (1984) suggested that temperature is a great abiotic factor in the persistence of the
22
nematode, as he reported that P. penetrans while in moist soils can survive at -4: 40-70˚C for 13
and 9 weeks respectively. Other factors for nematode persistence in a fallow include soil
moisture content, soil type and soil properties, the physiological age of the nematode and their
lipid reserves (Kutywayo & Been, 2006). The infectivity of the nematode decreases after a
prolonged state without food (Karssen et al., 2013), followed by dying of the nematode (Nježić
et al., 2014).
Egg masses and subsequent reproduction were noted within the root system of the green manure
crops which supported nematode invasion while absence of egg masses reproduction could have
been the result of nematode failure to invade and establish feeding sites in the root cells of some
manure crops. In this study red clover cv. Lemmon and white clover cv. Melital are classified
maintenance host and good hosts respectively. The plant species though not based on cultivars
were susceptible to M. chitwoodi with the reproductive factor of 9 and 10 respectively (Griffin &
Rumbaugh, 1996). Fodder radish line RsV79/80 is reported a non-host. This is in consensus with
findings of Ferris et al. (1993), where 9 of the 10 cultivars tested were poor or non-host to M.
chitwoodi with a reproductive factor less than 0.3. Presently, fodder radish line RsV79/80 being
a non-host complies with field micro-plot findings of (Al-Rehiayani & Hafez, 1998) for the two
fodder radish cultivars (Melodie and Trez) each with a reproductive factor of 0.1. Further this
study findings contradicts with the greenhouse evaluation for the three fodder radish cv.
(Melodie, Trez and Adagio 5a-3) as maintenance hosts with reproductive factors of 2.9, 2.8 and
1.7 respectively (Al-Rehiayani & Hafez, 1998). Recently, fodder radish varieties Anaconda,
Contra, Defender, Doublet and Terranova, known to have partial resistance, were evaluated and
their relative susceptibility were 0.17, 0.10, 0.42, 0.32 and 0.14% respectively (Teklu et al.,
2014). The M. chitwoodi populations reduced by more than 98% but (Teklu et al., 2014) could
not regard the varieties as non-hosts because their final nematode populations were dependent on
the initial nematode inoculation. Further field testing of RsV79/80 is thus recommended.
Ryegrass cv. Meltador is classified a good host in this current study with a reproductive factor
greater than 1.0. These findings are not in agreement with Hoek (pers. comm) that an English
ryegrass is a moderate host but whose crop damage is not clearly known. In a pot greenhouse
study Cook et al. (1999) confirmed a number of clones of ryegrass to be either resistant or
susceptible to M. naasi. Arugula cv. Nemat was identified as a poor host. This classification was
23
supported by the fewer number of nematodes in the root system and egg masses. The
significantly fewer nematode population densities of all stages extracted from arugula cv. Nemat
indicates that arugula hinders the development of all stages hence its capable of acting as trap
plant to manage M. hapla (Melakeberhan et al., 2006). Riga (2011) found a 99% reduction in M.
chitwoodi populations after planting potato as a follow-up crop in a greenhouse study. Riga
(2011) further narates that arugula on its own has not been able to control M. chitwoodi in the
field where lengthy season potatoes can support more than one generation of the nematode
species unlike in the green house. In South Africa, cv. Nemat, was reported a poor host and able
to inhibit M. javanica gall formation when applied as green manure in a glasshouse trial (Kruger
et al., 2015). The main allelochemical known for killing the nematodes is glucosinolates
particularly 4-methylthiobuthyl (Curto et al., 2005).
The current study presents the first report of clover- ryegrass mixture as a good host to M.
chitwoodi. It is reported that red clover cv. Lemmon, white clover cv. Melital and ryegrass cv.
Meltador are hosts to M. chitwoodi with reproductive factor of 1.63, 3.97 and 4.33 respectively.
This might have stimulated the high reproduction of the nematode in the mixture with a
reproductive factor of 5.49. It was observed that a good host ryegrass has dense root system
which covered the whole experimental pot, therefore its influence on the final nematode
reproduction of the mixture may be very fundamental than the two clovers. Obviously in the
field root development will be different and this might influence results. Therefore further field
testing is required. This initiative is in support of Peter and Rayns (2008) who recommended for
red clover-ryegrass, white clover/ryegrass or a complex containing several different cultivars of
several species of grasses and clovers. In promoting organic farming and management of M.
chitwoodi, this clover- English ryegrass mixture should not be used in the rotation. However,
clover- English ryegrass biofumigation investigation is recommended to further ascertain the
nematode levels especially in the field soils. Furthermore, the study presents the first report of
arugula- fodder radish mixture as a poor host with a reproductive factor less than that of the
individual plants, which seems logically true. The individual fodder radish and arugula are nonhost and poor host respectively, therefore the mixture is expected to reduce the nematode
reproduction. This would be the ideal green manure mixture to be used by farmers in
management of M. chitwoodi.
24
CONCLUSION AND RECOMMENDATION
Due to a policy to increase biodiversity farmers are financially rewarded if they use mixtures of green
manures. For farmers with P. penetrans and M. chitwoodi problems the options are limited and therefore
research on green manure mixtures is needed. Based on this study the classical clover-ryegrass mixtures
are not ideal but fodder radish (resistant cultivar) and arugula (cv. Nemat) can be successful.
Putting into consideration that different geographical areas have differing soil properties and
abiotic factors. It is essential first to select potential green manure plants that are adapted and
best fit into the local climatical crop rotation. It is recommended to carry out greenhouse or field
micro-plots for the various selected green manure plants to test for their host suitability to the
target pathogens before attempting larger scale field experience through farmers.
For a Belgian farmer willing to adopt organic farming, increase his or her crop yields and obtain
monetary incentives from the use of green manure mixtures provided by the European Union, it
should be a collaborative approach and advice from plant pathologists in different disciplines,
plant breeders and geneticists as well.
Acknowledgement
I do convey my appreciations to the Flemish government of Belgium (VLIR-UOS) for
sponsoring my master program. I take this moment to thank ILVO under the leadership of Prof
Nicole Viaene for the offer to work with the research team in the Nematology laboratories
especially Ms Nancy De Sutter. Many thanks to university of Ghent Nematology research unit
especially Prof wim Bert, Prof Decraemer, Ms Inge Dehennin and Emmanuelle De Bock for
their technical and administrative support during the whole period of my master program.
25
REFERENCES
Abawi, G. & Ludwig, J. (1995). Host efficiency of 16 cover crops to the lesion nematode.
Phytopathology 85, 1554.
Al-Rehiayani, S. & Hafez, S. (1998). Research: Host Status and Green Manure Effect of
Selected Crops on Meloidogyne chitwoodi Race 2 and Pratylenchus neglectus.
Nematropica 28, 213-230.
Al-Rehiayani, S., Hafez, S., Thornton, M. & Sundararaj, P. (1999). Investigation-Research:
Effects of Pratylenchus neglectus, Bacillus megaterium, and Oil Radish or Rapeseed
Green Manure on Reproductive Potential of Meloidogyne chitwoodi on Potato.
Nematropica 29, 37-49.
Back, M., Haydock, P. & Jenkinson, P. (2002). Disease complexes involving plant parasitic
nematodes and soilborne pathogens. Plant pathology 51, 683-697.
Baermann, G. (1917). Eine einfache methode zur auffindung von ankylostomum (Nematoden)
larven in erdproben. Geneeskunding Tijdschrift voor Nederlandsch-Indië 57, 131-137.
Castillo, P. & Vovlas, N. (2007). Pratylenchus (Nematoda: Pratylenchidae): diagnosis, biology,
pathogenicity and management. Brill, pp.
Cherr, C., Scholberg, J. & Mcsorley, R. (2006). Green manure approaches to crop production.
Agronomy Journal 98, 302-319.
Cohen, M., Yamasaki, H. & Mazzola, M. (2005). Brassica napus seed meal soil amendment
modifies microbial community structure, nitric oxide production and incidence of
Rhizoctonia root rot. Soil Biology and Biochemistry 37, 1215-1227.
Cook, R., Mizen, K. A. & Person-Dedryver, F. (1999). Resistance in ryegrasses, Lolium spp., to
three European populations of the root-knot nematode, Meloidogyne naasi. Nematology
1, 661-671.
Curto, G., Dallavalle, E. & Lazzeri, L. (2005). Life cycle duration of Meloidogyne incognita and
host status of Brassicaceae and Capparaceae selected for glucosinate content. Nematology
7, 203-212.
De Waele, D. & Elsen, A. (2007). Challenges in tropical plant nematology. Annu. Rev.
Phytopathol. 45, 457-485.
Decraemer, W., Hunt, D. J., Perry, R. & Moens, M. (2013). Structure and classification. Plant
nematology, 3-39.
26
Dong, L. Q. & Zhang, K. (2006). Microbial control of plant-parasitic nematodes: a five-party
interaction. Plant and Soil 288, 31-45.
Duncan, L., Moens, M. & Perry, R. (2013). Migratory endoparasitic nematodes. Plant
nematology, 144-178.
Esteves, I., Maleita, C. & Abrantes, I. (2015). Root-lesion and root-knot nematodes parasitizing
potato. European Journal of Plant Pathology 141, 397-406.
Ferris, H., Carlson, H., Viglierchio, D., Westerdahl, B., Wu, F., Anderson, C., Juurma, A. &
Kirby, D. (1993). Host status of selected crops to Meloidogyne chitwoodi. Journal of
Nematology 25, 849.
Gheysen, G., Jones, J., Perry, R. & Moens, M. (2006). Molecular aspects of plant-nematode
interactions. Plant nematology, 234-254.
Goh, K. M. & Bruce, G. E. (2005). Comparison of biomass production and biological nitrogen
fixation of multi-species pastures (mixed herb leys) with perennial ryegrass-white clover
pasture with and without irrigation in Canterbury, New Zealand. Agriculture, ecosystems
& environment 110, 230-240.
Griffin, G. & Rumbaugh, M. (1996). Host suitability of twelve Leguminosae species to
populations of Meloidogyne hapla and M. chitwoodi. Journal of Nematology 28, 400.
Haydock, P., Woods, S., Grove, I., Hare, M., Perry, R. & Moens, M. (2013). Chemical control of
nematodes. Plant nematology, 459-479.
Hendrickx, G. (1995). An automatic apparatus for extracting free-living nematode stages from
soil. Nematologica 41, 308.
Hooks, C. R., Wang, K.-H., Ploeg, A. & Mcsorley, R. (2010). Using marigold (Tagetes spp.) as a
cover crop to protect crops from plant-parasitic nematodes. Applied Soil Ecology 46, 307320.
Jensen, H. (1953). Experimental greenhouse host range studies of two root-lesion nematodes,
Pratylenchus vulnus and Pratylenchus penetrans. Plant Disease Reporter 37, 384-387.
Jones, J. T., Haegeman, A., Danchin, E. G., Gaur, H. S., Helder, J., Jones, M. G., Kikuchi, T.,
Manzanilla‐López, R., Palomares‐Rius, J. E. & Wesemael, W. M. (2013). Top 10
plant‐parasitic nematodes in molecular plant pathology. Molecular Plant Pathology 14,
946-961.
27
Karssen, G., Wesemael, W., Moens, M. & Perry, R. (2013). Root-knot nematodes. Plant
nematology, 73-108.
Kruger, D., Fourie, J. & Malan, A. (2015). Control Potential of Brassicaceae Cover Crops as
Green Manure and their Host Status for Meloidogyne javanica and Criconemoides
xenoplax. S. Afr. J. Enol. Vitic 36.
Kruger, D., Fourie, J. & Malan, A. P. (2013). Cover crops with biofumigation properties for the
suppression of plant-parasitic nematodes: A review.
Kutywayo, V. & Been, T. H. (2006). Host status of six major weeds to Meloidogyne chitwoodi
and Pratylenchus penetrans, including a preliminary field survey concerning other weeds.
Nematology 8, 647-657.
Manuel, J. S., Bendixen, L., Reynolds, D. & Reidel, R. (1980). Weeds as hosts of Pratylenchus.
Research Bulletin, Ohio Agricultural Research and Development Center.
Melakeberhan, H., Xu, A., Kravchenko, A., Mennan, S. & Riga, E. (2006). Potential use of
arugula (Eruca sativa L.) as a trap crop for Meloidogyne hapla. Nematology 8, 793-799.
Mizukubo, T. & Adachi, H. (1997). Effect of temperature on Pratylenchus penetrans
development. Journal of Nematology 29, 306.
Moens, M. & Perry, R. N. (2009). Migratory plant endoparasitic nematodes: a group rich in
contrasts and divergence. Annual review of phytopathology 47, 313-332.
Mojtahedi, H., Santo, G. & Ingham, R. (1993). Suppression of Meloidogyne chitwoodi with
sudangrass cultivars as green manure. Journal of Nematology 25, 303.
Nicol, J. M. & Rivoal, R. (2008). Global knowledge and its application for the integrated control
and management of nematodes on wheat. Integrated management and biocontrol of
vegetable and grain crops nematodes. Springer, pp. 251-294.
Nicol, J. M., Turner, S. J., Coyne, D., Den Nijs, L., Hockland, S. & Maafi, Z. T. (2011). Current
nematode threats to world agriculture. Genomics and molecular genetics of plantnematode interactions. Springer, pp. 21-43.
Nježić, B., De Sutter, N. & Moens, M. (2014). Interaction of Tagetes patula cv. Single Gold with
the life cycle of the plant-parasitic nematodes Meloidogyne chitwoodi and Pratylenchus
penetrans. Russian Journal of Nematology 22, 101-108.
28
Ortiz, A. M., Sipes, B. S., Miyasaka, S. C. & Arakaki, A. S. (2015). Green Manure Crops for
Management of Meloidogyne javanica and Pythium aphanidermatum. HortScience 50,
90-98.
Papadopoulos, Y. A., Kimpinski, J., Mcrae, K. B., Christie, B. R., Gallant, C. E. & Fillmore, S.
A. (2002). Invasion and reproduction by rootlesion nematode (Pratylenchus penetrans)
differs among selected lines of red clover (Trifolium pratense). Phytoprotection 83, 4145.
Peter, K. & Rayns, F. (2008). A Rosenfeld Green manures: A review conducted by HDRA as
part of HDC Project FV 299. An investigation into the adoption of green manures in both
organic and conventional rotations to aid nitrogen management and maintain soil
structure 37.
Riga, E. (2011). The effects of Brassica green manures on plant parasitic and free living
nematodes used in combination with reduced rates of synthetic nematicides. Journal of
Nematology 43, 119.
Schomaker, C., Been, T., Perry, R. & Moens, M. (2013). Plant growth and population dynamics.
Plant nematology, 301-330.
Slootweg, A. (1956). Rootrot of bulbs caused by Pratylenchus and Hoplolaimus spp.
Nematologica 1, 192-201.
Teklu, M. G., Schomaker, C. H. & Been, T. H. (2014). Relative susceptibilities of five fodder
radish varieties (Raphanus sativus var. Oleiformis) to Meloidogyne chitwoodi.
Nematology 16, 577-590.
Thies, J., Petersen, A. & Barnes, D. (1995). Host suitability of forage grasses and legumes for
root-lesion nematode Pratylenchus penetrans. Crop science 35, 1647-1651.
Townshend, J. (1984). Anhydrobiosis in Pratylenchus penetrans. Journal of Nematology 16, 282.
Townshend, J. & Davidson, T. (1960). Some weed hosts of Pratylenchus penetrans in Premier
strawberry plantations. Canadian Journal of Botany 38, 267-273.
Townshend, J. & Potter, J. (1976). Evaluation of forage legumes, grasses, and cereals as hosts of
forage nematodes. Nematologica 22, 196-201.
Viaene, N., Coyne, D., Davies, K., Perry, R. & Moens, M. (2013). Biological and cultural
management. Plant nematology, 383-410.
29
Viaene, N. M. & Abawi, G. S. (1998). Management of Meloidogyne hapla on lettuce in organic
soil with sudangrass as a cover crop. Plant Disease 82, 945-952.
Vrain, T., Deyoung, R., Hall, J. & Freyman, S. (1996). Cover crops resistant to root-lesion
nematodes in raspberry. HortScience 31, 1195-1198.
Wander, J., Van Den Berg, W., Van Den Boogert, P., Lamers, J., Van Leeuwen, G., Hendrickx,
G. & Bonants, P. (2007). A novel technique using the Hendrickx centrifuge for extracting
winter sporangia of Synchytrium endobioticum from soil. European Journal of Plant
Pathology 119, 165-174.
Wesemael, W. M. & Moens, M. (2008). Vertical distribution of the plant-parasitic nematode,
Meloidogyne chitwoodi, under field crops. European Journal of Plant Pathology 120,
249-257.
Wesemael, W. M., Viaene, N. & Moens, M. (2011). Root-knot nematodes (Meloidogyne spp.) in
Europe. Nematology 13, 3-16.
Wiggins, B. & Kinkel, L. (2005). Green manures and crop sequences influence potato diseases
and pathogen inhibitory activity of indigenous streptomycetes. Phytopathology 95, 178185.
30