Evaluation of Native Entomopathogenic Nematode Isolates for the Management of Selected Insect Pests in Nigeria

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Egypt. J. Agronematol., Vol. 20, No.2, PP. 167- 176 (2021)
DOI: 10.21608/EJAJ.2021.195928
Evaluation of Native Entomopathogenic Nematode Isolates for the
Management of Selected Insect Pests in Nigeria
A. O.* Cole-O. Y. and Claudius Alabi ;A. Y. Ottun
Dept. Crop Protection Environmental Biology, University of Ibadan, Ibadan, Nigeria
* Corresponding author email: b.claudiuscole@gmail.com; bi_cole@yahoo.com
Received: 10 February 2020 Revised: 21 April 2020 Accepted: 29 April 2020
ABSTRACT
Native entomopathogenic nematodes (EPNs) were evaluated for the management of
three selected pests, Sesamia calamistis, Spodoptera frugiperda and Rhynchophorus
ferrugineus in laboratory bioassays. The EPNs were isolated from soils from various
locations within Ibadan, Nigeria using the greater wax moth (Galleria mellonella) as
insect bait. A two-factor laboratory experimental assay was laid out in a completely
randomized design and replicated three times. The EPN suspension was applied on
insect larvae with distilled water as control. Number of days to mortality, percentage
mortality of insect larvae, and total EPN population of infective juveniles (IJs)
recovered from larval cadavers were assessed and reproductive factor (RF) determined.
Data were analyzed using analysis of variance, and means were separated using
Tukey’s Studentized Range Test at P<0.05. The number of days to mortality for
inoculated larvae of S. calamistis, S. frugiperda, R. ferrugineus and G. mellonella were
6.43, 3.57, 8.80 and 6.20, respectively. The EPN population achieved percentage
mortality of 82.0%, 54.0%, 60.0% and 84.0%, respectively for G. mellonella, S.
calamistis, S. frugiperda and R. ferrugineus larvae respectively. Mean numbers of EPN
IJs recovered from cadaver was in the order; R ferrugineus (9,407.0), >G. mellonella
(5075.08), >S. frugiperda (3957.23), >S. calamistis (742.31). From the results, EPN
had the greatest fecundity in G. mellonella and R. ferrugineus showing higher ability to
be recycled. This study reveals the potential of native EPNs as a biocontrol agent of
insect pests and emphasizes the need for a more environment-friendly and sustainable
approach to insect pest management.
Keywords: Biocontrol agents, insect mortality, Heterorhabditis sp., Sesamia calamistis
Spodoptera frugiperda, Rhynchophorus ferrugineus
INTRODUCTION
Cultivation of plants for the production of food and fibre, needed for man’s survival,
is an age-long practice. However, plants and their products are attacked by defoliators,
sap suckers, gall formers, stem borers, pod or fruit borers, and seed eaters either in the
early stage or during the later stages of development (Rao et al., 2000; Maina et al.,
2018). Insects attack and damage grains, fibers, fruits, vegetables, ornamentals and
forestry resulting in yield losses that contribute to food insecurity. The damage that
insects inflict on cultivated plants is highly varied, and includes direct feeding on
leaves, fiber, grain and fruits, facilitating the entry of pathogens or vectoring pathogens

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that cause plant diseases (Coates et al., 2015; Dhaliwal et al., 2015; Singh and Kaur,
2018). The most relevant stem boring species associated with maize production in
Nigeria are the lepidopterous moths.
The African pink stem borer, Sesamia calamistis Hampson (Lepidoptera:
Noctuidae), is one of the noxious noctuid pests of maize and most other plants of the
family Poaceae (Abate et al., 2000; Okweche et al., 2015). Damage to maize varies with
locations/regions within sub-Saharan Africa depending on the population of this stem
borer. Crop losses and grain yield reduction may result from the damage caused to
growing points leading to loss of stands (dead heart), large number of exit holes,
damage to leaves (windowpane damage), stem tunnelling, and direct damage to ear
shank and ears resulting in reduced cob yield (Okweche et al., 2015; Ramanujam et al.,
2017). Yield losses due to lepidopterous borers in Africa vary greatly between 0% to
100%, among ecological zones, regions, and seasons. Other reports suggest a potential
yield loss of 20% to 90% due to stem borers on cereals and sugarcane in Sub-Saharan
Africa (Assefa and Dlamini, 2018).
The fall armyworm (FAW), Spodoptera frugiperda (J.E. Smith) (Lepidoptera:
Noctuidae), is a key pest of maize (Zea mays L.) and many other crops. It is native to
tropical and subtropical regions of the western hemisphere from the United States of
America to Argentina. Spodoptera frugiperda was reported for the first time in 2016 in
the African continent within the following countries; Nigeria, Sao Tomé, Benin and
Togo, causing significant damage to maize hitherto (Deole and Paul, 2018). This
generalist insect pest attacks many crops but exhibits a preference for members of the
Poaceae family such as maize and sorghum. Larvae usually consume a large amount of
foliage (defoliation) and sometimes destroy the growing point of the plant causing dead
heart. The destruction of leaves, stems and/or flowers of the plant by the larval instars
occurs through feeding of all instar stages. In maize, armyworm larvae damage leaves
causing pinhole to shot holes and progresses into different sizes of lesions on leaves
during the vegetative stage. At tasseling and silking before maturity, they may damage
ears and kernels (Midega et al., 2018).
Late instar larvae can act as cutworms by entirely sectioning the stem base of maize
seedlings (Goergen et al., 2016; Midega et al., 2018). Late instars of fall armyworm
larvae consume large amounts of leaf tissue resulting in a ragged appearance of the
leaves leaving a mass of moist brown frass. Maize damage evaluation in Ethiopia
revealed that fall armyworm caused up to 30% loss at the late whorl stage even with
untimely pest control (Assefa and Ayalew, 2019). When late instars act as seedling
cutworms, maize losses can reach up to 100% (Fatoretto et al., 2017). Older larvae
burrow into maize tassels and feed into the ears, causing extensive damage.
Consequently, the impact of FAW is greatly felt in the economy of concerned
regions/countries.
Red palm weevil (RPW), Rhynchophorus ferrugineus (Coleoptera: Curculionidae),
is a globally obnoxious tissue borer of palm trees. This polyphagous palm insect pest
damage stem and core tissues of oil palm in West Africa (Mehdi et al., 2018). The
damage is completely caused by the grubs which feed internally by boring the tissues
of the stem and eventually kills the palm tree.
Entomopathogenic nematodes (Rhabditida: Heterorhabditidae and
Steinernematidae) have the potential for biological control of insect pests as conferred
by the symbiotic complex formed with certain bacteria (Kaya et al., 2006; El-Sadawy
et al., 2020). They possess a unique combination of attributes that make them a
promising alternative for pest control (Divya and Sankar, 2009; Abd-Elgawad, 2019).
The uniqueness of entomopathogenic nematodes can be further buttressed by their mass

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production mechanism from tissues (cadaver) of a wide variety of susceptible insects.
These insect-parasitizing nematodes kill the insects by serving as vectors of bacteria
which help to achieve quick kill of the target insect pests and thus have high potential
in pest management (Grewal et al., 2001; El-Sadawy et al., 2020).
Steinernematid and Heterorhabditid families infect several insect pest species, yet
pose no known negative effect on the environment. They can be mass-produced,
formulated, and commercially utilized as bio-control agents against insect pests (Lacey
et al., 2001; El-Sadawy et al., 2020). Entomopathogenic nematodes have the potential
to reproduce in soil environments and are capable of maintaining an efficacious
population density in soil for at least one additional season after their application has
been enhanced by habitat manipulation (Riga et al., 2001; Alramadan and Mamay,
2019). In this study, the potential of EPNs was evaluated as bio-control agent against
Sesamia calamistis, Spodoptera frugiperda and Rhynchophorus ferrugineus.
MATERIALS AND METHODS
The laboratory assay was laid out in a completely randomized design (CRD) with
three replications. The two treatments were insects’ larvae species (G. mellonella, S.
calamstis, S. frugiperda, R. ferrugineus), and entomopathogenic nematode inoculation
with non-inoculated (uninfected) control. Galleria mellonella late instar larvae were
used as a basis for comparing the three test insects.
Source of Insects and Entomopathogenic Nematodes
Galleria mellonella larvae were obtained from infested honeycombs, collected from
apiaries in the University of Ibadan. The larvae were maintained in Kilner jars half-
filled with crushed combs. Laboratory cultured larvae of Sesamia calamistis were
obtained from the International Institute of Tropical Agriculture (IITA), Ibadan. The
larvae were fed on artificial media in which they were maintained until used. A field
population of Spodoptera frugiperda was collected from maize farms at Ijaye farm
settlement and were fed with fresh maize leaves in Kilner jars until used. Larvae of
Rhynchophorus ferrugineus were collected from a palm farm at Moniya, Ibadan. The
larvae were kept in palm stem dust in a large aerated bucket from where they were
selected for use.
Isolation of Native Entomopathogenic Nematodes (EPNs) from collected soil
samples
Sample Collection
Entomopathogenic nematodes were directly baited with the late instars of the
greater was moth (Bedding and Akhurst, 1974), from soil samples obtained during the
rainy season (July and August) from locations within Ibadan. The locations include
uncultivated field; Agodi gardens (N 7°24'23.5368", E 3°54'12.9852"), Botanical
garden University of Ibadan (UI) (7°27'25.1"N 3°53'41.8"E), Cocoa Research Institute
of Nigeria (CRIN) (7°13'29.2"N 3°52'02.6"E), Forestry Research Institute of Nigeria
(FRIN) (7°23'32.4"N 3°51'37.6"E), International Institute of Tropical Agriculture
(IITA) (7°30'08.0"N 3°54'34.9"E), National Institute of Horticulture (NIHORT)
(7°24'11.9"N 3°50'57.6"E), Teak reserve UI (7°27'27.3"N 3°53'55.0"E), and
uncultivated field types; IITA, NIHORT, Teaching and Research Farm UI. Samples

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were obtained by random sampling of fields from three quadrats (Župunski et al., 2017)
and five samples per quadrat.
The EPNs were isolated from these soil samples using the larvae of the greater wax
moth (Galleria mellonella), obtained from the Apiary Unit of the Department, as bait.
G. mellonella larvae are most commonly used to rear nematodes in in-vivo protocol
because of their commercial availability and high susceptibility to most EPN species
Grewal et al. (2001).
Soil samples were sieved then moistened and 300 cm3 of soil was placed in a clear
rectangular plastic container (6 cm x 4.5 cm x 2 cm) into which 5 larvae of G. mellonella
were introduced. Soil samples were incubated in the dark and observed every 24 hrs to
obtain infected and dead larvae which were removed and observed using a zoom
stereomicroscope to confirm the cause of death as internally borne EPNs. Each dead G.
mellonella was then placed in a White trap set-up (White, 1927; Rahoo et al., 2017) to
collect infective juveniles of the nematodes. The native population was preliminarily
identified as Heterorhabitis sp. and is further undergoing species identification. The
identification was based on cadaver colour and morphological characteristics of
infective juveniles and adult males.
Multiplication of Entomopathogenic Nematodes’ Infective Juveniles
Galleria. mellonella larvae were infected with IJs of harvested EPN from soil
samples. The larval cadavers recovered after periodic inoculation yielded a mass
population of entomopathogenic nematodes through White-trap extraction, that served
as inoculum for the laboratory bioassay.
Laboratory Bioassay
Five larvae of each test insect were placed into Petri-plates (12 cm diameter) lined
with filter paper moistened with distilled water and each larva of G. mellonella, S.
calamistis, and S. frugiperda, was inoculated with 50 IJs of EPN suspension in 1 ml of
water. Due to the large size of R. ferrugineus, one larva was introduced singly on each
filter paper-lined Petri-dish and inoculated with 140 IJs of EPN suspension per larva.
The choice of IJ numbers used in the inoculation of each insect species was based on
the relative sizes of the treated insect larvae. Five samples represented a replicate and
three replications were made per treatment with EPN inoculation and the respective
control (without EPN). The Petri-dishes were covered and labelled accordingly.
Gallaria mellonella, was used as the standard insect for comparison with other test
insects since it is widely accepted that entomopathogenic nematodes successfully yield
high progeny per larval cadaver of G. mellonella (Grewal et al., 2001). The larvae in
the Petri dishes were incubated in the dark at room temperature (26 ± 2 ℃) to allow
larval infection. All dead larvae were transferred singly to a White trap for nematode
extraction. Water suspension of emerged EPN juveniles from white traps was harvested
every 2-3 days over five weeks. The trial was repeated once following the same
procedures.
Data Collection and Analysis
Data collected were number of days to mortality, percentage larval mortality, EPN
population per week and the reproductive factor (RF) [Total population (Pf) of EPN/
Initial population (Pi) of EPN]. All data were analyzed using Analysis of Variance
(ANOVA), SAS 9.3 software and means were separated using Tukey’s Studentized
Range Test at P<0.05.

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RESULTS AND DISCUSSION
By 10 days post-inoculation, no mortality was observed in all non- infected insect
larvae in the Petri-dishes with a few larvae pupating in case of S. calamistis while those
inoculated with EPN died within 3 – 9 days after inoculation (Fig. 1). The initial levels
of inoculum and body tissue of the insects’ larvae both play a complementary role in
influencing the incidence of early mortality (Krishna, 2005). This was confirmed from
the results obtained where relatively large-sized larvae of R. ferrugineus died later than
other tested insects, (G. mellonella, S. calamistis, S. frugiperda), with smaller larval
size.
The results show that EPN had a significant impact on the insect larva types by
achieving a percentage mortality of 56.00%, 60.00%, 82.00% and 84.00%, respectively
for S. calamistis, S. frugiperda, G. mellonella, and R. ferrugineus (Fig. 2). There were
no significant differences in percentage mortality among the inoculated larvae and also
among the control (non-infected larvae). Mortality of more than 60% was not possible
for S. frugiperda either due to cannibalism amongst insect larvae of the same
experimental unit or migration out of the enclosure. This observation is in line with the
report from Fatoretto et al. (2017) that the S. frugiperda larvae movement is density-
dependent and influenced by strong cannibalism.
Figure 1: Number of days to mortality of insect larvae infected with native populations
of EPNs. Bars represent Standard error
Some of the S. calamistis larvae pupated during the study, this may have accounted for
the lower mortality (56%) recorded for them. Similarly, Garcia-del-Pino et al. (2013)
reported no pupal mortality in their trial with tomato leaf miner. White trap extraction
from the pupae in this study, in addition, yielded no EPN. Yet the findings are different
from those of Kurtz et al. (2009) who observed mortality in the studied pupae. Studies
have primarily revealed the infective ability of entomopathogenic nematodes (EPN) in
the management of insect pests. They can achieve relatively quick kill of the target
insect pests upon inoculation with number of active sufficient level of infective
juveniles capable of initiating infection and thus leading to mortality of insect pests
(Grewal et al., 2001; Divya and Sankar, 2009). It is evident from this study, that the

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native EPNs isolated from soils were effective in causing death of all the test insects to
varying degrees.
Figure 2: Mortality (%) of insect larvae infected with EPN compared to non-infected
larvae (control) 10 days after inoculation. Bars with the same letters are not
significantly different using Tukeys’ studentized multiple range test at p≤0.05)
Figure 3. Weekly nematode harvest from insect cadavers over a five-week period after
inoculation. Values of nematode numbers are square-root transformed; Bars represent
standard error of means for each nematode per week
Generally, the number of nematodes harvested from the dead insect larvae declined
throughout observation period (Fig. 3). By three weeks after inoculation no nematodes
were recoverable from dead larvae of both G. mellonella and S. calamistis however,
larvae were obtained in R. ferrugineus at five weeks after inoculation (Fig 3). This
demonstrates the ability of the nematodes to be available for reinfection of more larvae
in the community for up to three, four and five weeks respectively, in S. calamistis, S.
frugiperda and R. ferrugineus, but with majority of the nematode being dispersed at
one week after inoculation. Smits (1996) observed persistence of EPN for 2 to 6 weeks
after field application. Furthermore, the findings of Ebssa and Koppenhöfer (2011)
demonstrate that EPNs can persist to effectively manage the black cut worm in turf
grass on golf fields. Extended duration of IJs ability to infect insect host is a very
important trait in the biological control of pests.
With respect to EPN production of IJs from insect cadavers, R. ferrugineus had a
significantly higher population of EPN 9407.00, > G. mellonella 5075.08, > S.

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frugiperda 3957.23, > S. calamistis 742.31 (Table 1). There was no significant
difference between the total EPN population generated from cadavers of both G.
mellonella and S. frugiperda. However, G. mellonella had total EPN numbers that
differed significantly from each of the EPNs from individual cadavers of S. calamistis
and R. ferrugineus, with R. ferrugineus cadavers having the highest EPN yield (Table
1). Larvae in the non EPN control dishes did not yield harvestable EPN through to the
end of the extraction process.
Table 1: Total entomopathogenic nematodes from insects’ larval cadavers and
reproductive factor (RF) of entomopathogenic nematodes in larval types
Insect species Total EPN Population Reproductive factor
Infected Control Infected Control
Galleria mellonella 71.43(5075.08)b 0.71 (0.00)a* 101.50a 0.00a*
Sesamia. calamistis 38.99(742.31)c 0.71 (0.00)a* 14.85c 0.00a*
Sposoptera frugiperda 62.14(3957.23)b 0.71 (0.00)a* 79.59a 0.00a*
Rhynchophorus ferrugineus 143.68(9407.00)a 0.71 (0.00)a* 67.24a 0.00a*
Mean values with the same letters within the column are not significantly different according to Tukey’s
Studentized Range Test at P > 0.05. Values are transformed means with actual mean values in
parenthesis. * denotes significant difference between treatments in a row for each parameter.
Reproductive factor = total population divided by initial population.
The initial numbers of IJs used to infect the test insects can influence the final yield
of emerged IJs from the larval cadavers (Krishna, 2005), as observed with the
significantly lower EPN yield from standard insect, G. mellonella inoculated with a
lower level of inoculum, in contrast with the maximum EPN yield from R. ferrugineus.
At the same time, Sesamia calamistis yielded low EPN population upon inoculation
with a relatively low level of inoculum, the reason could be attributed to the small size
of the larvae in addition to the older stage of instars based on the appearance of pupae
shortly after inoculation. The highest populations of EPNs were recovered from
infected R. ferrugineus because EPNs continued to recycle in them due to their larger
size than other tested insect larvae. These findings imply that density of the initial
inoculum, nature of host, age of larval instar and size of larvae can influence the yield
of infective juveniles from the cadavers of infected larvae. This impacts on the ability
of the EPN population to be sustained in the system in order to enhance the longevity
of the management effect of the EPNs on the infected insect pests.
The greatest reproductive factor (RF), a measure of fecundity, of EPN was found in
G. mellonella, followed by S. frugiperda, and R. ferrugineus, (Table 1). S. calamistis,
displayed the lowest (P ≤ 0.05) level EPN reproductive factor in dead larvae when
compared to the EPN reproductive factor in all other insects’ larval cadavers.
Meanwhile, Galleria ranked in the category of highest EPN yielding larvae
(significantly high reproductive factor) in the short-term period of extraction of
harvestable nematode IJs. This conforms with the report of Grewal et al. (2001);

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solidifying its use as the standard larvae for EPN inoculum production. It being not
significantly different from the fecundity of EPN in S. frugiperda and R. ferrugineus
implies that these two insect hosts can encourage the persistence of the pathogenic
nematodes in the environment.
Entomopathogenic nematodes possess a unique combination of attributes that make
them a promising alternative to synthetic chemical insecticides for pest control (Divya
and Sankar, 2009; Abd-Elgawad, 2019). These nematodes have exceptional potential
for biological control of insects as conferred by the symbiotic complex formed with
bacteria of the genus Xenorhabdus associated with the Steinernema species and
Photorhabdus with the Heterorhabditis species (Grewal et al., 2001; Kaya et al., 20006;
Ardpairin et al., 2020) to achieve their successful parasitism and quick-kill effect on
their insect host. This ability was evidenced in the high mortality observed on the test
insects. The uniqueness of entomopathogenic nematodes can be further buttressed by
their mass production mechanism from tissues (cadaver) of susceptible insects. The
infective juveniles of these EPNs are motile, virulent with high reproductive potential
and possess the ability to disperse from the dead host to infect new healthy ones
(Krishna, 2005; Ramakuwela, 2014; Assefa and Ayalew, 2019). This explains the rapid
multiplication of the offspring (infective juveniles) in the parasitized larval cadavers
and their subsequent release from the cadaver’s dead tissues.
From the present research, it can be concluded that the native/indigenous Nigerian
population of entomopathogenic nematodes are potentially effective as a bio-control
agent against lepidopterous insect pests Sesamia calamistis and Spodoptera frugiperda
insect larvae and a coleopteran destructive palm pest Rhynchoporus ferrugineus larvae.
Production of several generations of EPNs in insect cadaver is possible, when there is
sufficient food (large-tissue cadaver), as observed for R. ferrugineus, to nourish the
developing infective juveniles. However, further studies should be conducted to
evaluate the efficacy of entomopathogenic nematodes on insect pests of different crops
under screen house and field conditions.
REFERENCES
Abate, T., Van Huis, A. and Ampofo, J.K.O. (2000). Pest management strategies in
traditional agriculture: An African perspective. Ann. Rev. Entomol. 45: 631–
659.
Abd-Elgawad, M.M.M. (2019). Towards optimization of entomopathogenic nematodes
for more service in the biological control of insect pests. Egypt J. Biol. Pest
Co. 29: 77 https://doi.org/10.1186/s41938-019-0181-1.
Alramadan, Y. and Mamay, M. (2019). The importance of entomopathogenic nematode
and their role in the control of agricultural pests. 1st international
GOBEKLITEPE Agriculture Congress (IGAC) November 25-27, 2019. 275-
282.
Ardpairin, J., Muangpat, P., Sonpom, S., Dumidae, A., Subkrasae, C., Tandhavanant,
S., Thanwisai, A. and Vitta, A. (2020). A survey of entomopathogenic
nematodes and their symbiotic bacteria in agricultural areas of northern
Thailand. J. Helminthol. 94, e192, 1-11. https://doi.org/10.1017/
S0022149X20000735.
Assefa, F. and Ayalew, D. (2019) Status and control measures of fall armyworm
(Spodoptera frugiperda) infestations in maize fields in Ethiopia: A review.

Evaluation of Native Entomopathogenic Nematode Isolates….. 175
Egypt. J. Agronematol., Vol. 20, No.2 (2021)
Cogent Food Agric. 5:1, 1641902. https: //doi. org/ 10.1080/ 23311932. 2019.
1641902.
Assefa, Y. and Dlamini, T. (2018). Evaluation of stem borer species diversity and
identification of the fall armyworm strain in cereal crops and sugarcane fields
of Swaziland. UNESWA 50-50 Commemoration Journal 1: 1-6.
Bedding, R.A. and Akhurst, R.J. (1974). A simple technique for the detection of insect
parasitic nematodes in soil. Nematologica, 21: 109-110.
Coates, B.S., Poelchau, M., Childers, C., Evans, J.D., Handler, A., Guerrero, F., Skoda,
S., Hopper, K., Wintermantel, W.M., Ling, K., Hunter, W.B., Oppert, B.S.,
Pérez De León, A.A., Hackett, K. and Shoemaker, D. (2015). Arthropod
genomics research in the United States Department of Agriculture-Agricultural
Research Service: current impacts and future prospects. Trends Entomol. 11: 1–
27.
Deole, S. and Paul, N. (2018). First report of fall army worm, Spodoptera frugiperda
(J.E. Smith), their nature of damage and biology on maize crop at Raipur,
Chhattisgarh. J. Entomol. Zool. Stud. 6(6): 219-221.
Dhaliwal, G.S., Jindal, V. and Mohindru, B. (2015). Crop losses due to insect pests:
global and Indian scenario. Indian J. Entomol. 77(2): 165–168.
Divya, K. and Sankar, M. (2009). Entomopathogenic nematodes in pest management.
Review “Biological control of crops in India”. Indian J. Sci. Technol. 2(7): 53-
60.
Ebssa, L., and Koppenhöfer, A. M. (2011). Efficacy and persistence of
entomopathogenic nematodes for black cutworm control in turfgrass. Biocont.
Sci. Tech. 21(7): 779-796.
El-Sadawy, H.A., Ramadan, M.Y., Abdel Megeed, K.N., Ali, H.H., El Sattar, S.A. and
Elakabawy, L.M. (2020). Biological control of Phlebotomus papatasi larvae by
using entomopathogenic nematodes and its symbiotic bacterial toxins. Trop.
Biomed. 37(2): 288–302.
Fatoretto, J.C., Michel, A.P., Filho, M.C.S. and Silva, N. (2017). Adaptive potential of
Fall armyworm (Lepidoptera: Noctuidae) limits Bt trait durability in Brazil. J.
Integr. Pest Manag. 8(1): 1–10. doi: 10.1093/jipm/pmx011.
Garcia-del-Pino, F., Alabern, X., & Morton, A. (2013). Efficacy of soil treatments of
entomopathogenic nematodes against the larvae, pupae and adults of Tuta
absoluta and their interaction with the insecticides used against this
insect. BioControl. 58(6):723-731.
Goergen, G., Kumar, P.L., Sankung, S.B., Togola, A. and Tamò, M. (2016). First report
of outbreaks of the fall armyworm Spodoptera frugiperda (J.E. Smith)
(Lepidoptera, Noctuidae), a new alien invasive pest in West and Central Africa.
PLoS One 11 (10) e0165632. http://dx.doi.org/10.1371/journal. pone.0165632.
Grewal, P.S., De Nardo, E.A.B. and Aguillera, M.M. (2001). Entomopathogenic
nematodes: potential for exploration and use in South America. Neotrop.
Entomol. 30(2): 191-205.
Kaya, H.K., Aguillera, M.M., Alumai, A., Choo, H.Y., De la Torre, M., Fodor, A. and
Ganguly, S. (2006). Status of entomopathogenic nematodes and their symbiotic
bacteria from selected countries or regions of the world. Biol. Control 38: 134-
155.
Krishna, S.N. (2005). Studies on entomopathogenic nematodes. Department of
Entomology, Faculty of Agriculture, Acharya N.G. Ranga Agricultural
University, Tirupati. 517 502 (A.P.).

Ottun et al. 176
Egypt. J. Agronematol., Vol. 20, No.2 (2021)
Kurtz, B., Hiltpold, I., Turlings, T. C., Kuhlmann, U., and Toepfer, S. (2009).
Comparative susceptibility of larval instars and pupae of the western corn
rootworm to infection by three entomopathogenic nematodes. Biocontrol 54(2),
255-262.
Lacey, L.A., Frutos, R., Kaya, H. K. and Vail, P. (2001). Insect pathogens as biological
control agents: Do they have a future? Biol. Control 21: 230–248.
Maina, U.M., Galadima, I.B., Gambo, F.M. and Zakaria, D. (2018). A review on the
use of entomopathogenic fungi in the management of insect pests of field crops.
J. Entomol. Zool. Stud. 6(1): 27-32 E-ISSN: 2320-7078 P-ISSN: 2349-6800
JEZS.
Mehdi, M.Z., Wakil, W., Jan, H., Raza, M.M., Shah, Q. and Zia-ul-Haq, M. (2018).
Evaluation of entomopathogenic nematode and fungi alone and their
combination against red palm weevil, Rhynchophorus ferrugineus(Olivier). J.
Entomol. Zool. Stud. 6(2): 2038-2042.
Midega, C.A.O., Pittchar, J.O., Pickett, J.A., Hailu, G.W. and Khan, Z.R. (2018). A
climate-adapted push-pull system effectively controls fall armyworm,
Spodoptera frugiperda (J E Smith), in maize in East Africa. Crop Prot. 105: 10–
15 https://doi.org/10.1016/j.cropro.2017.11.003.
Okweche, S.I., Osai, E.E. and Umoetok, S.B.A. (2015). Maize borer damage in
Nigeria’s Guinea Savanna: Timing of planting overrides effects of insecticides
treatments. Revista Colombiana de Entomología 41(2): 170-175.
Rahoo, A.M., Mukhtar, T., Gowen, S.R., Rahoo, R.K. and Abro, S.I. (2017).
Reproductive potential and host searching ability of entomopathogenic
nematode, Steinernema feltiae. Pakistan J. Zool. 49(1): 229-234. DOI:
10.17582/journal. pjz/2017. 49.1.229.234.
Ramakuwela, T., (2014). Development of a new entomopathogenic nematode species,
Steinernema innovationi: Biological characterization and mass production.
Plant Pathology, School of Agricultural, Earth and Environmental Sciences,
College of Agriculture, Engineering and Science, University of KwaZulu-Natal,
Pietermaritzburg, South Africa.
Ramanujam, B., Japur, K., Poornesha, B., Shylesha, A.N. and Rangeshwaran, R.
(2017). Field evaluation of endophytic entomopathogenic fungi against maize
stem borer (Chilopartellus) (Crambidae: Lepidoptera). Indian J. Agric. Sci.
87(8): 1099–1103.
Rao, M.R., Singh, M.P. and Day, R. (2000). Insect pest problems in tropical
agroforestry systems: Contributory factors and strategies for management.
Agrofor. Syst. 50: 243–277.
Riga, E., Whistlecraft, J. and Potter, J. (2001). Potential of controlling insect pests of
corn using entomopathogenic nematodes. Can. J. Plant Sci. 81(4): 783-787
Singh, B. and Kaur, A. (2018). Control of insect pests in crop plants and stored food
grains using plant saponins: A review. LWT - Food Sci. and Tech. 87: 93-101.
https://doi.org/10.1016/j.lwt.2017.08.077.
Smits,P.H.(1996).Post-application persistence of entomopathogenic nematodes.
Biocont. Sci. Tech. 6 (3): 379-388.
White, G.F. (1927). A method for obtaining infective juveniles from cultures. Science,
66: 302-303. http://dx.doi.org/10.1126/science.66.1709.302-a.
Župunski, V., Jevtić, R., Jokić, V. S., Župunski, L., Lalošević, M., Ćirić, M. and Ćurčić,
Ž. (2017). Sampling Error in Relation to Cyst Nematode Population Density
Estimation in Small Field Plots. J. Nematol. 49(2): 150–155.