The Isolation, Identification, and Insecticidal Activities of Indigenous Entomopathogenic Nematodes (Steinernema carpocapsae) and Their Symbiotic Bacteria (Xenorhabdus nematophila) against the Larvae of Pieris brassicae

Abstract

The cabbage butterfly, Pieris brassicae Linnaeus (Lepidoptera: Pieridae), is an oligophagous and invasive insect pest of various economically important cole crops. Recently, there have been reports about an increase in the incidence and damaging activities of cabbage butterflies, signifying that the existing control methods fail to meet the grower’s expectations. Entomopathogenic nematodes (EPNs) and their endosymbiotic bacteria have immense potential for the control of a wide range of insect pests. In this investigation, the EPN species Steinernema carpocapsae and its associated bacterial species, Xenorhabdus nematophila, were isolated and identified through morphological and molecular techniques. The laboratory bioassay experiment was performed using S. carpocapsae and X. nematophila against the 3rd instar larvae of P. brassicae (25 ± 1 °C; RH = 60%). The efficacy of EPN suspension (30, 60, 90, 120, 150 IJs/mL) and bacterial suspension (1 × 10⁴, 2 × 10⁴, 3 × 10⁴, 4 × 10⁴, and 5 × 10⁴ CFU/mL) via contact and oral routes showed significant mortality among the larvae. Surprisingly, 100% insect mortality within 48 h was recorded in the bacterial inoculum 5 × 10⁴ CFU/mL. However, in the case of EPNs (S. carpocapsae), 150 IJs/mL caused the highest, 92%, larval mortality rate after 96 h. The results signify that both indigenous EPNs and their associated bacteria can provide efficient control against P. brassicae larvae and could effectively contribute to IPM programs. However, further analyses are required to authenticate their effectiveness in field conditions.

1. Introduction

Insects are phytophagous organisms, with >85 percent of species containing more than 9 lakh identified insects [1]. Numerous agricultural insect pests, along with forest encroaching species, contribute to losses of approximately 76.9 billion USD per year, and it is a need of the hour to control the effects of insect bio-invasion worldwide [2]. Currently, phytophagous insect invasion has intensified due to global warming projects [3] that might contribute to the expansion of pest populations, empirical exceedance in outbreaks, and geographical spreading of several species, resulting in huge economic impairments and a decline in food safety [4]. Among insect arthropods, lepidopteran insect pests are causing substantial damage to agricultural produce. Pieris brassicae (the cabbage butterfly) is a destructive polyphagous pest that is responsible for huge economic losses to the crucifers [5] and is one of the limiting factors in crop production worldwide. Along with the crucifers, they also feed upon plants of the Tropaeolaceae, Resedaceae, and Capparaceae families [6], causing over 50% losses in production every year [7].

The management of cabbage butterflies is challenging due to their wide host range, feeding habits, and high reproduction potential. A wide spectrum of chemical-based insecticides and pesticides have been employed by farmers worldwide to protect their crops from invasive damage. The pesticides used are quite expensive and unsafe. The consumption of pesticides not only decays soil prolificacy but also affects the intrinsic habitats of advantageous organisms and causes resistance among insect pests [8,9,10]. Additionally, education and consciousness concerning the risks associated with pesticide consumption, including pollution, contamination, resistance, pest revival, and effect on other non-target creatures has risen. The consequences raised by these chemical-based formulations have drawn attention to a better and safer alternative. Continuous research in this field is being carried out, and an alternative means of insect pest control without affecting natural entities has emerged as “Biological control”. The utilization of bio-agents, such as indigenous EPNs, is the best way to tackle insect pest problems.

The word “Entomopathogenic” is integrated from two different words: first entomon, related to insects, and pathogen, related to causing disease. Thus, EPNs are responsible for causing diseases in various insects [11]. Phoretic, communalistic, symbiotic, obligatory, and facultative interactions have been recorded among nematodes and insects [12]. The EPN order Rhabditida belongs to two families, i.e., Heterorhabditidae [13] and Steinernematidae [14]. Genera Steinernema (Steinernematidae) and Heterorhabditis (Heterorhabditidae) exhibit a total of 117 species of EPNs, with 100 species described from Steinernema and 17 species from the genus Heterorhabditis [15]. The nematodes showed mutualistic alliances with the bacterial species that inhabit their alimentary canals and are responsible for forming nemato-bacterial complexes [16]. The EPNs of genus Steinernema showed mutualistic association with the bacterial genus Xenorhabdus whereas Heterorhabditis showed mutual interrelation with the bacterial genus Photorhabdus, respectively [17,18,19]. They are well-established biocontrol agents (BCAs) that play an imperative role in virulence and have been successfully employed against agricultural insect pests in different management strategies [20,21,22]. Third-stage juveniles, or infective juveniles (IJs), or dauer juveniles, are actually responsible for invading and parasitizing the host insect [23]. The symbiotic bacteria of EPNs releasing toxic substances upon invasion into the hemolymph and multiply there [24].

The correct and precise recognition of EPNs is the initial and prime exigency to accomplish any biological control attribute. Additionally, they are eco-friendly, economic, and conservative in nature [25,26,27]. Keeping this in mind, a survey was conducted to explore EPN incidence in Himachal Pradesh and to isolate the bacteria associated with these EPNs in order to evaluate their virulence capacity towards cabbage butterfly larvae under laboratory conditions.

2. Materials and Methods

2.1. Survey and Sample Collection

Soil-inhabiting entomopathogenic nematodes are usually found naturally in all types of soils, but their incidence is high in undistributed soil. A survey study was conducted during 2018–2021, and soil samples were collected from the undisturbed land of forests and fruit orchards of district Shimla (altitude range between 1021 and 2276 m), Himachal Pradesh, as this region is known as the “fruit bowl” of India and is chiefly a horticulture state that produces and exports apples, peaches, plums, apricots, cherries, and pears to other states. The soil was sampled from the vicinity of plant roots at a depth of 15–30 cm with the help of a spade and hand shovel. The samples were brought into the experimental laboratory after proper labelling of details such as host plant name, locality, date, district, village, type of soil, and altitude [28]. The isolation of EPNs was done via the soil baiting technique [29]. For this, stones, detritus, and leaf litter were removed, and soil was filled into the plastic containers after proper labelling [30]. The soil baiting technique requires insect baits, so Galleria mellonella and Corcyra cephalonica were used as bait. In this technique, 5–10 last instar larvae of bait insects were kept in plastic containers with soil. The containers were searched for larval mortality after every 48 h up to 120 h. The dead insect cadavers were separated and removed regularly from the containers. These cadavers were investigated, and the existence of nematodes was explored using the white trap method [31]. The emerging nematodes were collected from the white trap and stored in the flask before being transferred to the roux bottle. The incidence of EPNs was assessed using the formula given by Norton [32].

2.2. In-Vivo Mass Multiplication of EPNs

The isolated nematodes were further mass multiplied under laboratory conditions using G. mellonella larvae, followed by the white trap methodology for nematode extraction. The isolated nematodes were stored in two forms: (i) directly as suspension in the roux bottle and (ii) in a sterilized synthetic sponge. They were stored at 12–15 °C.

2.3. Morphological Observations of Entomopathogenic Nematode

The isolated nematodes were examined live as well as after being sacrificed, fixed, and mounted. Nematodes were sacrificed and fixed with formalin (4%) fixative, and mounting was carried out using anhydrous glycerine [33]. A total of 10 nematode specimens from each life stage were collected randomly and observed for morphological analysis. The measurements were taken using a compound microscope (Leica DM750) equipped with Leica Application Suite (LAS) version 4.12. The morphological studies were carried out in accordance with the taxonomic keys [34], and the body ratio depiction was made on the basis of the formula given by de Man [35].

2.4. Isolation of Bacteria Associated with Entomopathogenic Nematode

The bacterium associated with the EPNs was also isolated following the methodology given by Thakur et al. [36]. To isolate bacterial endosymbionts from EPNs, 50–100 IJs were taken into an Eppendorf tube (1.5 mL). In this tube, 1 mL of 1% sodium hypochlorite solution was added for surface sterilization (4–6 min). The suspension was spun in the microcentrifuge for 5 min at 5000 rpm for the collection of sterilized IJs. The IJs pellet was resuspended in double distilled water and washed 3–5 times. After washing, the IJs were placed into a sterilized mortar and pestle and crushed properly. After crushing, a few drops of crushed nematodes were placed onto the Petri plate containing Nutrient Bromothymol Blue Tetrazolium Chloride Agar (NBTA) medium. The drops were spread over the plate with the help of a sterile spreader. The plates were then placed into the bacteriological incubator at 28 ± 1 °C for 24–48 h. Lawns of bacterial colonies appeared on the Petri plate after 48 h. A single colony of bacteria was chosen and re-streaking was carried out using a sterile loop until a pure uniform colony was achieved. The culture was stored under the nutrient broth (NB) media and used for DNA isolation as well as for the bioassay study.

2.5. Molecular Characterization of Entomopathogenic Nematode and Bacteria

Molecular characterization was carried out in the molecular laboratory at ZSI, HARC-Solan. The genomic DNA from the nematodes and bacteria was procured using the Qiazen’s Dneasy^® blood and tissue kit-based method. Before proceeding to DNA isolation, 500 nematodes juveniles were stored in ethanol inside the Eppendorf tube (2 mL) and centrifuged at 12,000 rpm for 3 min. The supernatant was discarded, and the pellet was kept for drying, which was then transferred into the deep freezer (−20 °C). The nematode pellet was crushed using a micropestle inside the tube. The DNA was isolated, and the quantification of the extracted DNA was carried out using 0.8% agarose gel electrophoresis. The DNA quality was confirmed by visualizing the gel under the gel documentation unit (Alpha-Imager). The amplification of the isolated nematodes DNA was carried out, and the ITS region of the rDNA was amplified using the forward primer (18S: 5′-TTGATTACGTCCCTGCCCTTT-3′) and the reverse primer (26S: 5′-TTTCACTCGCCGTTACTAAGG-3′).

After that, the isolation of DNA from bacteria was carried out, and bacterial colonies were inoculated into 2 mL microcentrifuge tubes containing nutrient broth media and incubated for 24 h prior to extraction. The tubes were centrifuged, and the bacterial pallet was used for the isolation of DNA. The amplification of 16S rRNA was carried using pA/pH primers: forward primer pA (5′-AGAGTTTGATCCTGGCTCAG-3′) and reverse primer pH (5′-AAGGAGGTGATCCAGCCGCA-3′).

For the PCR amplification of nematode DNA, 25 μL reaction tube was prepared that contained genomic DNA 2.0 μL, Taq buffer (10X) 4.0 μL, MgCl₂ (25 mM) 1.0 μL, dNTPs (1 mM) 1.0 μL, primer forward 1.0 μL, primer reverse 1.0 μL, Taq polymerase 1.0 μL and Mili Q water 14 μL. The amplifications were accomplished in a gradient thermal cycler machine (Applied Biosys VeritiPro Thermal Cycle). The first step consisted of pre-heating the thermal cycler machine to 95 °C, followed by denaturation for 3 min at 94 °C. The third step involved annealing for 1 min at 55 °C and then for an extension of 1 min 30 s at 72 °C. The process was repeated for up to five cycles and was again followed by 30 s of denaturation at 94 °C; double annealing at 55 °C for 3 min; extension at 72 °C for 1 min; and, finally, extension of 5 min at 72 °C for up to 35 cycles to ensure the full length of amplified fragments. The amplified product was stored at 4 °C, which was later transferred to −20 °C.

Similarly, in bacterial DNA amplification, a 25 μL reaction tube was prepared and contained genomic DNA 2.0 μL, Taq buffer (10X) 4.0 μL, MgCl₂ (25 mM) 1.0 μL, dNTPs (1 mM) 1.0 μL, primer forward 1.0 μL, primer reverse 1.0 μL, Taq polymerase 1.0 μL, and Mili Q water 14 μL. The optimized PCR conditions include initial denaturation at 95° for 2 min, followed by denaturation at 94 °C for 30 s, followed by annealing at 54° for 30 s, then extension at 72 °C for 2 min, and final extension at 72 °C for 10 min for up to 35 cycles, and then storage at 4 °C.

The products after the PCR reactions, using primers, were observed on 1.2% agarose gels and their sizes were determined by comparison with the DNA ladder (100 bp). The amplified PCR products of the nematode genome were sent to the Eurofins Analytical Services India Pvt. Ltd. (Bengaluru, India) for sequencing. After sequencing, a phylogenetic tree of the obtained sequences was constructed, and the reverse sequence was antisense reversed. Using these sequences, 10 corresponding or similar sequences were downloaded from GenBank, NCBI. Multiple sequence alignment (MSA) of all the sequences was carried out using ClustalW (with a gap opening penalty for multiple alignments of 12 and an extension of 10) using MEGA v 11.0. A phylogenetic tree was constructed using maximum likelihood in bootstrapping 100, using the Tamura 3-parameter model. Gaps and missing data were treated as complete deletions [37]. The partial sequences were submitted to the National Centre for Biotechnology Information (NCBI) and accession numbers were assigned.

2.6. Evaluation of EPNs Infectivity

Rearing of Pieris brassicae was carried out in the laboratory following the methodology given by Tomar et al. [38]. A petri plate bioassay experiment was performed to evaluate the infectivity of S. carpocapsae and X. nematophila against the 3rd instar larvae of P. brassicae under laboratory conditions. Two bioassay experiments were performed separately, in which one experiment contained EPNs suspension and another contained bacterial suspension. The nematodes were used at concentrations of 30, 60, 90, 120, 150 IJs/mL as well as a control. The bacterial suspension was applied at a rate of 1 × 10⁴, 2 × 10⁴, 3 × 10⁴, 4 × 10⁴, and 5 × 10⁴ CFU/mL with absolute control. Healthy, laboratory-reared 3rd instar larvae of P. brassicae were kept in the petri plates at a rate of 10 larvae per petri plate along with food (cabbage leaves). Each treatment was replicated five times, and in the control, only 1–2 mL of distilled water was applied. The Petri dish was incubated at 26 ± 1 °C. Insect mortality was checked after 24, 48, and 96 h of bacterial inoculation, and the data were recorded daily. The experiment was conducted in a completely randomized design. The data recorded on the insect mortality under laboratory conditions were subjected to statistical analysis, and the significance of the results was determined. The corrected percent mortality was analyzed using Abbott’s formula [39]. The analysis of variance (ANOVA) was evaluated (three factorial analysis) using arcsine transformation on software (Op Stat) developed by HAU, Haryana. A Tukey’s post hoc test was executed for pairwise comparisons between the EPNs and bacterial treatments.

3. Results

3.1. Entomopathogenic Nematode Occurrence in (Surveyed) Soils

For the soil sampling survey, a total of 31 soil samples were gathered from Chhupari, Rampur, Badiyara, Shimla, Fagu, and Jabbal and processed using the soil baiting technique with G. mellonella and C. cephalonica larvae (

Table 1. Distribution and frequency of occurrence of entomopathogenic nematodes.

Location/ Villages	Vegetations	Total No. of Samples	Samples Having EPNs	Samples without EPNs	Frequency of Occurrence (%)
Chhupari	Peach, Apple, Pear, Cucumber and Plum	05	01	04	25
Rampur	Apple and Persimmon	04	-	04	-
Badiyara	Peach and Apple	06	01	05	16.67
Shimla	Apricot, Plum, Cherry, Apple, Pear, Peach	05	-	05	-
Fagu	Apple	05	01	04	25
Jabbal	Pear, Apple and Apricot	06	02	04	33.33

The isolated nematodes were identified on the basis of their morphological observations based on the species-specific diagnostic keys.

). Out of these 31 samples, only 5 were observed positive for EPNs, which included 4 Heterorhabditis and 1 Steinernema species based upon the infected cadaver appearance. The overall frequency of Steinernema sp. occurrence was 25% from an apple orchard in the Fagu region. Furthermore, the isolated Steinernema sp. strain EUPT-R2 was mass multiplied under laboratory conditions using G. mellonella and C. cephalonica larvae. The average infective juveniles (IJs) produced by G. mellonella were 66,036 (55,120–67,154) and C. cephalonica 48,320 (42,532–50,795) IJs/larva.

3.2. Morphological and Molecular Characterization of EPNs

The morphometrics of infective juveniles, adult males, and adult females corresponded with Steinernema carpocapsae, Weiser. The 10 individual infective juveniles, female and male, were observed morphologically. The IJs were much narrower, 508.00–598.00 μm in length, 23.64–26.58 μm in width, and the esophagus and intestinal region were collapsed. The nerve ring distance from the anterior end was 72.00–86.00 μm, the mouth was closed, the tail was pointed, and the lateral line fields were clearly visible. The excretory pore distance from the anterior end was 30.00–35.00 μm, the pharynx distance from the anterior end was 111.50–123.00 μm, and the tail length was 48.00–54.00 μm.

Females (n = 10) were somewhat larger in size, 3210.00–4060.00 μm in length and 183.00–211.00 μm, with a smooth cuticle and slightly rounded head, united lips, and a collapsed stoma. Pharynx distance from the anterior end was 180.00–196.00 μm, with a Rhabditoid esophagus. Anterior-to-basal bulb nerve ring was present surrounding the isthmus, and reflexed amphidelphic gonads and the vulval region showed protuberance. The tail was conical with a spiny tip and was 42.00–49.00 μm in length. Males (n = 10) are anteriorly similar to females, with a truncated, slightly rounded head, smooth cuticle, 1080.00–1658.00 μm in length, and 78.00–113.00 μm in width. The nerve ring distance from the anterior end was 93.00–124.00 μm, with a partially collapsed stoma, an absent collar, and the presence of a small amphid. The excretory pore distance from the anterior end was 48.00–72.00 μm, and the pharynx distance from the anterior end was 138.00–164.00 μm. The testis was reflexed, and the paired spicule was slightly curved and symmetrical. Spicule has a wider head, tapered gubernaculums, and a tail bearing genital papillae. The tail was 28.00–34.00 μmwithmucron and no bursa (Figure 1).

The results obtained through amplified PCR products showed a band size of 850 bp in the agarose gel electrophoresis run with a 100 bp ladder. The blast comparison based upon the ITS region amplification showed that the Steinernema sp. strain EUPT-R2 exhibited 99.38% sequence similarity and 99% query cover with S. carpocapsae (MK977607). The sequence was submitted to the gene bank with accession number OP295355. The calculated patterns of nucleotide substitution in the isolated taxon were T = 37.8; C = 16.3; A = 23.6; and G = 22.3 (Figure 2).

3.3. Identification and Molecular Characterization of the Endosymbiotic Bacteria

Initially, the identification of bacteria was carried out based on morphological characteristics. The bacterial colony appeared maroon on the blue agar, and, after Gram staining, it was found to be Gram-negative, rod-shaped, a facultative anaerobe, and motile. The genomic DNA of a bacterium was isolated and amplified using 16S and 18S gene sequencing. The blast comparison of 16S gene amplification in EUDPTB-R2 attributes 99.35% sequence similarity and 100% query cover with X. nematophila (MW619913). The sequence was submitted to the gene bank with accession number OQ842895. The calculated patterns of nucleotide substitution in the isolated taxon were T = 18.6; C = 22.1; A = 24.7; and G = 34.6. Based on gene amplification, the phylogenetic tree was constructed using the neighbor-joining method (Figure 3).

3.4. Entomopathogenic Capacity of Isolated EPN Species and Associated Bacterium

During the present investigation, the entomopathogenicity of EPNs (30, 60, 90, 120, and 150 IJs/mL) and their associated bacteria (1 × 10⁴, 2 × 10⁴, 3 × 10⁴, 4 × 10⁴, and 5 × 10⁴ CFU/mL) was evaluated against the 3rd instar larvae of P. brassicae. A bioassay experiment showed the highest mortality (92%) was recorded at the highest nematode inoculum concentration (150 IJs/mL) after 96 h of infection (Figure 4). Considerable variations have been observed in insect mortality at various inoculums that vary from 45, 58, 64, 82 to 92% after 96 h of nematode inoculation (F =13.647; DF = 5; p < 0.05). A significant increase in larval mortality rate was recorded as exposure time increased.

The EPN-associated bacterium X. nematophila also caused significant mortality among insect larvae. The highest concentration of bacterial suspension (5 × 10⁴ CFU/mL) resulted in 100% larval mortality within 48 h. A noticeable difference in the mortality rate was observed after the application of different concentrations of bacterial suspension. The mortality ranges vary from 94, 96, 100 to 100% after 96 h of bacterial inoculation (F = 258.01; DF = 5; p < 0.05). Amongst both of these treatments, applications of different concentrations of X. nematophila caused rapid infectivity, followed by insect mortality (Figure 5).

4. Discussion

The isolation and identification of infinitesimal EPNs is quite difficult and thus necessitates greater attention during these processes. The isolation of EPNs and mass multiplication using the insect baiting technique has been mentioned by many researchers. During the present investigation, EPN S. carpocapsae was isolated and mass multiplied using C. cephalonica and G. mellonella larvae, showing that the average infective juveniles (IJs) produced by G. mellonella were 66,036 (55,120–67,154) and C. cephalonica 48,320 (42,532–50,795) IJs/larva. The results were also supported by an earlier study, where 54,347, 44,697, and 36,860 IJs of S. carpocapsae were recorded on large, medium, and small-sized larvae of C. cephalonica [40]. The highest 71,532.0 IJs/larva were recovered during the in vivo mass multiplication of S. carpocapsae on G. mellonella larvae [41].

The taxonomic identification of genus level has been considered unstable due to overlapped morphological characteristics. With the advancement of molecular techniques, morphological observations can be accompanied with molecular observations that have been considered stable. The analysis of nucleotide sequences has also been effective for providing the phylogenetic relationship amongst the nematodes. On the basis of morphological and morphometrical analysis, the juveniles and adults of isolated EPNs resembled S. carpocapsae [42,43]. Earlier, H. bacteriophora, Steinernema sp. and S. feltiae were recovered from the apple orchards of Himachal Pradesh, India [44]. Steinernema glaseri, Steinernema thermophilum, and H. indica recovered from Meghalaya, India [45]. Based upon the amplification of the ITS region, the EPNs for Steinernema sp. strain EUPT-R2 attribute 99.38% sequence similarity with S. carpocapsae. Earlier, H. bacteriophora and S. carpocapsae were reported from Uttar Pradesh (India) using morphological and molecular analyses [46,47]. S. carpocapsae was identified on the basis of morphological and molecular characterization in Italy (Veneto) [48].

The nematodes genus Steinernema showed mutualistic alliance with the bacterial genus Xenorhabdus. The mutualism between both partners is highly specialized and both of them are mutually benefitted. Here, in this study, S. carpocapsae-associated bacteria, X. nematophila, was isolated and identified morphologically as well as molecularly. Similar observations were recorded by researchers in Cauca-Colombia [49]. S. carpocapsae (MK350941), S. monticolum (MK503674), and Rhabditis blumi (MN453373) were isolated and identified on the basis of 18S rDNA sequencing from the tropical and subtropical agro-ecosystems of Tamil Nadu, India [50]. The nematode-associated bacterial isolates X. nematophila were also identified morphologically, biochemically, and molecularly. Earlier, X. nematophilus from S. carpocapsae was isolated in Korea [51].

EPNs and their symbionts have been known for their virulence activities against insect pests since ancient times. During this investigation, a laboratory bioassay experiment was conducted for the estimation of the insecticidal potential of S. carpocapsae and X. nematophila against the 3rd instar larvae of P. brassicae. Amongst both of these treatments, applications of different concentrations of X. nematophila caused rapid infectivity followed by insect mortality. The observations of the current findings are supported by previous observations that showed 100% larval mortality in the early instars after 96 h [45,52,53,54]. It was also suggested that increased exposure time significantly increased larval mortality and vice versa [55]. In total, 72.08% and 85.38% larval mortality was recorded among 2nd and 4th instar larvae upon treatment with H. bacteriophora [56]. The mortality rate of 67.42 and 69.50% was recorded among the 2nd and 4th instars upon treatment with S. feltiae. High insect mortality was recorded among the 3rd and 4th instar larvae with the inoculum of 200 IJs, which is similar to the observations of this investigation [57]. In total, 100 percent mortality was recorded among all larval stages of the cabbage butterfly after exposure to S. glaseri and H. bacteriophora [58].

High mortality in P. brassicae larvae was recorded by several researchers upon treatment with native EPN isolates from Himachal Pradesh [56,59]. The virulence of Xenorhabdus sp. was also reported against mushroom mites [60]. EPNs and their associated bacteria were found to be highly effective in managing the 3rd and 4th larval instars of Pieris rapae and Pentodonal gerinus [61]. The Xenorhabdus and Photorhabdus bacteria (cell suspensions and cell-free supernatant) applied for the management of Agrotis ipsilon and good management potential was recorded [62]. The biocontrol potential of Steinernema sp. and their symbiontic bacteria, Xenorhabdus sp., was also found to be effective for the management of Ephestia cautella larvae [63].

5. Conclusions

Entomopathogens have been known to manage insect populations for years. Steinernema carpocapsae (OP295355) has been isolated from the apple (Malus domestica) orchards of the district of Shimla, Himachal Pradesh. The bacteria associated with this EPN species were recognized as X. nematophila (OQ842895). The laboratory bioassay experiments were performed using different concentrations of S. carpocapsae and X. nematophila against the 3rd instar larvae of P. brassicae. Among both of these treatments, applications of different concentrations of X. nematophila resulted in rapid infection and faster mortality when compared with EPNs. It can be concluded from the present investigation that the soil environment of the fruit orchards of Himachal Pradesh provides an excellent habitat to proliferate the entomopathogenic nematodes naturally in the environment. Further survey studies are required to explore nematode diversity and evaluate their biocontrol attributes in this region.