
==== Front
Heliyon
Heliyon
Heliyon
2405-8440
Elsevier

S2405-8440(24)12396-1
10.1016/j.heliyon.2024.e36365
e36365
Review Article
Soil inhabiting bacto-helmith complex in insect pest management: Current research and future challenges
Tomar Preety a
Thakur Neelam a
Singh Sangram b
Kumar Sanjeev c
Rustagi Sarvesh d
Rai Ashutosh Kumar e
Shreaz Sheikh f
Yadav Neelam gh
Rai Pankaj Kumar i
Yadav Ajar Nath ajarbiotech@gmail.com
j⁎
a Department of Zoology, Akal College of Basic Sciences, Eternal University, Baru Sahib, Sirmour-173101, Himachal Pradesh, India
b Department of Biochemistry, Dr. Ram Manohar Lohia Avadh University, Ayodhya, Uttar Pradesh, India
c Faculty of Agricultural Sciences, GLA University, Mathura, Uttar Pradesh, India
d Depratment of Food Technology, School of Applied and Life Sciences, Uttaranchal University, Dehradun, Uttarakhand, India
e Department of Biochemistry, College of Medicine, Imam Abdulrahman Bin Faisal University, Dammam, Kingdom of Saudi Arabia
f Desert Agriculture and Ecosystem Department, Environment and Life Sciences Research Center, Kuwait Institute for Scientific Research, P. O. Box 24885, 13109, Safat, Kuwait
g Centre of Research Impact and Outcome, Chitkara University, Rajpura-140401, Punjab, India
h Chitkara Center for Research and Development, Chitkara University, Himachal Pradesh-174103, India
i Department of Biotechnology, Invertis University, Bareilly, Uttar Pradesh, India
j Department of Genetics, Plant Breeding and Biotechnology, Dr. Khem Singh Gill Akal College of Agriculture, Eternal University, Baru Sahib, Sirmaur-173101, Himachal Pradesh, India
⁎ Corresponding author. ajarbiotech@gmail.com
15 8 2024
30 8 2024
15 8 2024
10 16 e363654 11 2023
14 8 2024
14 8 2024
© 2024 The Authors. Published by Elsevier Ltd.
2024

https://creativecommons.org/licenses/by/4.0/ This is an open access article under the CC BY license (http://creativecommons.org/licenses/by/4.0/).
Pesticides have health consequences for humans, living organisms, and ecosystems. Research on biological management, with a primary focus on entomopathogens, has been accelerated by the rise in issues such as pesticide residue, soil degradation, and pest resistance. Entomopathogenic nematodes (EPNs) are among the most frequently used and commercialised biopesticides. However, they are restricted in their infectivity, persistence, storage, and cost of production. The nematodes, along with their endosymbiotic bacteria, combine to form a nemato-bacterial complex. This complex is responsible for causing mortality in insect pests due to the production of insecticidal compounds. The adaptation of EPNs is an eco-friendly method, economical, and safer for the environment as well as non-target organisms. Moreover, it's a better alternative to synthetic chemical pesticides, as it can be helpful in overcoming pest resistance and resurgence issues. Application of nematode juveniles is a cost-effective method, but the necessity of refrigeration and transportation may enhance their cost. This review emphasised the diversity of entomopathogenic nematodes and their endosymbiotic bacteria, the exploration of the biocontrol potential of insect pests by under-utilisation of nematodes, the development of nematode-based formulations, and the discussion of critical issues and required research in the future.

Highlights

• Portrayal of the distribution of entomopathogenic nematodes and their bacteria.

• Biocontrol potential of entomopathogenic nematodes against insect pests.

• Description of the nematode-based bioformulations available on the market.

Keywords

Biocontrol
Entomopathogens
Formulations
Infectivity
Pesticides
==== Body
pmc1 Introduction

Crop productivity for human consumption is a menace because of the prevalence and outbreak of insect pests in the agricultural industry. According to Food and Agricultural Organisation (FAO) statistics, an average 40 % loss in crop yield is due to the attack of insect pests, which account for $290 billion of the global economy [1]. The losses in crop productivity caused by these insect pests can be extensive and reduced by adapting crop protection measures [2]. Although several chemical-based insecticides and pesticides have been employed to manage the insect pest's populations, These pesticides enhanced economic potential, increased food productivity, and killed insects [3].

Pesticides possess a broad spectrum of compounds, such as antifungal, insecticidal, rodenticides, herbicides, nematicides, and mollusc-killing compounds, along with some plant growth regulators. Pesticides have grievous health consequences for humans, living organisms, and ecosystems. Large numbers of pest insects have developed resistance to the insecticides [4]. Immense evidence is available that indicates that synthetic chemicals possess potential hazards to humans and non-target organisms and cause other ill effects on the environment [5]. So an alternative economic and eco-friendly method was required to overcome the hazards of insecticides and pesticides. Applications of biocontrol agents such as entomopathogenic nematodes (EPNs) are the best alternative method to conquer all these problems.

Entomopathogenic nematodes, or insect-killing worms, are free-living organisms ubiquitous in ecosystems. These worms are mainly known to parasitize insects. Various kinds of interactions were seen among nematodes and insects, including phoretic, symbiotic, commensalistic, facultative, and obligatory interactions [6]. Soil offered admirable circumstances for EPNs survival [7]. The order Rhabditida has two main genus of EPNs, namely Steinernema under the family Steinernematidae and Heterorhabditis under the family Heterorhabditidae, with a total of 117 described species of EPNs. A total of 100 species in the genus Steinernema and 17 species from the genus Heterorhabditis have been reported worldwide [8]. The first EPN, Aplectana kraussei, was identified by Steiner in 1923. Genus Heterorhabditis was described by Poinar Jr [9]. In the 1980s, the biocontrol potential of EPNs was realised by scientists, which became the turning point of nematode-related research [10].

A complex way in which nematodes behave has been observed due to stimuli (chemical or physical) [11]. Five such habitual approaches were exhibited by nematodes: nictation, ambushing, jumping, cruising, and coalescences of ambush with cruising [12]. These behavioural approaches help the nematodes search for their host. Cruising behaviour is shown by highly mobile nematodes (Heteriorhabditis bacteriophora Poinar) that locate their host via their response to the chemical cues in the soil [13]. Ambusher behavioural approach exhibited by ambulatory nematodes that remains steady and attached to the passing host [12]. The ambusher sometimes lifts itself, standing over its tail and the entire body hanging like a loop. This is a kind of nictation [14]. Steinernema feltiae (Filipjev) exhibits a combination of both ambushers and cruisers. They are able to parasitize both immovable and movable hosts. Steinernema scapterisci (Nguyen and Smart) showed jumping actions where they attached to the insect body and migrated through phoresis [15].

The EPNs life cycle consists of three developmental stages: eggs, juveniles (04) and adults (male and female, or hermaphrodite female). Most of the life stages of EPNs were found feeding, growing, and reproducing inside the dead host. Only the “dauer juveniles (DJs), also aliased as the third juvenile form (IJ3), rm (IJ3), were found free-living in the environment or soil [16]. This third phase of juveniles (IJs) is regarded as the recovery stage and is actually responsible for invading and parasitizing the host insect [17]. As they perceive the host, the worm exuviates its cuticular lining to reveal the body openings [18]. They puncture the insect body through the natural interstice [19], including several entry routes such as anterior openings (mouth), posterior openings (anus), and the body surface opening (spiracle), along with the cuticular lining [20].

The nematodes showed a mutualistic alliance with the bacterial species that inhabit the alimentary canal of these nematodes. They combine to form a nemato-bacterial complex. The endosymbionts of the genus Steinernema contain the genus Xenorhabdus, and the nematode Heterorhabditis showed a mutual interrelation with the genus Photorhabdus [21]. The endosymbiotic bacteria of the genus Xenorhabdus and Photorhabdus are the members of family enterobacteriaceae under the order Enterobacterales and class Gammaproteobacteria [20]. The bacterial symbionts employed nematodes as a vector to get entry inside insects and devoted some part of their life cycle to the nematodes [22]. They effectively invade and parasitize the larval stages of soil inhabiting insect communities [23]. In the case of Steinernema spp., the bacterial cells were harboured in the special intestinal vesicles of the nematode, while in Heterorhabditis, the bacterial cells reside in the gut region and can be found scattered in the digestive tract of the nematode [24]. In Heterorhabditis spp., endosymbionts were regurgitated, while in Steinernema spp., they were expelled through the anal region [25].

The bacterial cells proliferated inside the insect haemocoel and, within 24–48 h, killed its host insect [26]. The bacteria released antimicrobial compounds that have immunosuppressive factors and a plethora that eroded with the host's tissues and created a nutritional medium for the nematode's development and reproduction [27]. In the case of Heterorhabditis nematodes hermaphrodites (hermaphroditic), females were produced in the first generation, but in the next generation, males and females (amphimictic) were developed. Steinernema species always produced amphimictic generations (males and females) [28]. The adults mate and lay eggs that hatch, and again the juveniles moulted and developed into adults. The cycle persisted for up to three generations until the nutrients were exhausted [29]. Due to an exhausting nutrient supply, the IJ3 colonised the endosymbionts inside their body, ripped the empty host cadaver, and quested for the new host [30].

EPNs of the genera Steinernema and Heterorhabditis have been effectively utilised as bio-agents in numerous insect management programs. One of the reasons for EPN utilisation as a bioagent is that 90 % or more insect pests inhabit soil to complete their life cycle. So they have been efficiently applied against ground-inhabited as well as folivore insects [31]. Due to their virulence, motility, quick and adaptive nature, and good sense of chemoception, EPNs have been effectively utilised in agricultural systems [32]. Additionally, they are eco-friendly, conservative, and free of harm [[33], [34], [35]]. Along with all the idiosyncrasy, the culturing of EPNs in vivo and in vitro is quite easy [36]. Even the applications of these EPNs are facile and can be showered via knapsack sprayer [37]. They are well-established biocontrol agents (BCA) and have been successfully employed in agricultural insect pest management strategies [38]. The attention paid to the biocontrol potential of EPNs began in the late 1930s. Although EPNs were familiar since the 17th century [39]. Immense studies on EPNs were accomplished in the 19th and 20th centuries (Table 1). This review emphasised the distribution of entomopathogenic nematodes and their endosymbiotic bacteria, the exploration of the biocontrol potential of insect pests by under-utilisation of nematodes, the development of nematode-based formulations, and the discussion of critical issues and required research in the future.Table 1 Global distribution of described species of entomopathogenic nematodes.

Table 1Country	Nematode Species	Bait insect/Host organism	Habitat	References	
Argentina	Steinernema ritteri	Galleria mellonella	Soil	[105]	
Steinernema rarum	G. mellonella	Soil	[106]	
Australia	Heterorhabditis bacteriophora	Heliothis punctigera	Soil	[107]	
Steinernema spp.	G. mellonella	Soil	[44]	
Brazil	Heterorhabditis amazonensis	G. mellonella	Soil	[108]	
Steinernema brazilense	G. mellonella	Soil	[109]	
Benin	Steinernema kandii	G. mellonella	Eucalyptus	[110]	
Cameroon	Steinernema cameroonense	G. mellonella	Soil	[111]	
Steinernema nyetense	G. mellonella	Soil	[111]	
Chile	Steinernema australe	G. mellonella	Soil	[112]	
Steinernema unicornum	G. mellonella	Soil	[113]	
Heterorhabditis atacamensis	G. mellonella	Soil	[114]	
China	Steinernema caudatum	G. mellonella	Soil	[115]	
Steinernema longicaudum	G. mellonella	Soil	[116]	
Steinernema ceratophorum	G. mellonella	Soil	[117]	
Steinernema guangdongense	G. mellonella	Eucalypt forest	[118]	
Steinernema beddingi	G. mellonella	Soil	[119]	
Steinernema akhursti	G. mellonella	Soil	[119]	
Steinernema aciari	G. mellonella	Coastal area soil	[120]	
Steinernema hebeiense	G. mellonella	Sandy soil	[121]	
Steinernema sichuanense	G. mellonella	Roadside soil	[122]	
Steinernema leizhouense	G. mellonella	Soil	[123]	
Steinernema cholashanense	G. mellonella	Soil	[124]	
Steinernema xueshanense	G. mellonella	Dry meadows soil	[125]	
Steinernema pui	G. mellonella	Rubber plantations	[126]	
Steinernema xinbinense	G. mellonella	Soil	[127]	
Steinernema tielingense	G. mellonella	Shrubs	[127]	
Steinernema changbaiense	G. mellonella	Soil	[128]	
Heterorhabditis beicherriana	G. mellonella	Cherry orchard	[129]	
Colombia	Steinernema colombiense	G. mellonella	Coffee plantations	[130]	
Costa Rica	Steinernema puntauvense	G. mellonella	Soils	[131]	
Steinernema costaricense	G. mellonella	volcanic soils	[131]	
Cuba	Steinernema cubanum	G. mellonella	Citrus plantation soil	[132]	
Czech
Republic	Steinernema poinari	G. mellonella	Sandy soil	[133]	
Steinernema weiseri	G. mellonella	Fruit orchards and crop fields	[134]	
Czechoslovakia	Steinernema carpocapsae	Cydia pomonella	–	[135]	
Denmark	Steinernema affine	Phyla febrilis	Preserved specimens	[136]	
Steinernema feltiae	Feltia segetum (Agrotis segetum)	Preserved specimens	[137]	
Egypt	Heterorhabditis taysearae	G. mellonella	Hibiscus micranthus	[138]	
Ethiopia	Steinernema yirgalemense	G. mellonella	Field soil	[139]	
Steinernema ethiopiense	G. mellonella	Soil	[140]	
France	Steinernema boemarei	G. mellonella	Soil	[141]	
Germany	Steinernema kraussei	Cephaleia abietis (L.)	-	[142]	
Steinernema silvaticum	G. mellonella	Woodland soil	[143]	
Steinernema schliemanni	Osmoderma ceremita	Soil	[144]	
Indonesia	Steinernema hermaphroditum	G. mellonella	Different vegetation	[145]	
Iran	Steinernema arasbaranense	G. mellonella	Arasbaran forest	[146]	
Italy	Steinernema apuliae	G. mellonella	Soil	[147]	
Steinernema ichnusae	G. mellonella	Sandy soil of coastal region	[148]	
Steinernema vulcanicum	G. mellonella	Chestnut wood	[149]	
India	Heterorhabditis indica	Scirpophaga excerptalis	Sugarcane field soil	[150]	
Heterorhabditis casmirica	G. mellonella	Agricultural soils	[151]	
Steinernema shori	G. mellonella	Soil	[152]	
Steinernema anantnagense	G. mellonella	Agricultural fields soils	[153]	
Ireland	Heterorhabditis downesi	G. mellonella	Grassland soil	[154]	
Japan	Steinernema kushidai	G. mellonella	Soil from five different climates	[155]	
Steinernema litorale	G. mellonella	Pine forest of coastal region	[156]	
Steinernema ashiuense	G. mellonella	Grassland soil	[157]	
Korea	Steinernema monticolum	Agrotis segetum; Agrotis ipsilon,	Forest, Chestnut	[158]	
Kenya	Steinernema karii	G. mellonella	Soil	[159]	
Mexico	Heterorhabditis mexicana	G. mellonella	Soil	[160]	
Steinernema ralatorei	G. mellonella	Soil	[161]	
Heterorhabditis zacatecana	G. mellonella	Maize fields	[162]	
Nepal	Steinernema lamjungense	G. mellonella	Soil	[163]	
Steinernema surkhetense	G. mellonella	Soil	[164]	
Steinernema nepalense	G. mellonella	Soil	[164]	
New Zealand	Heterorhabditis zealandica	Heteronychus arator	–	[165]	
Oregon					
Pakistan	Steinernema pakistanense	G. mellonella	Soil	[166]	
Steinernema asiaticum	G. mellonella	Soil	[167]	
Steinernema bifurcatum	G. mellonella	Rose plant	[168]	
Steinernema balochiense	G. mellonella	Psidium guajava	[169]	
Poland	Steinernema sandneri	G. mellonella	Soil	[170]	
Puerto Rico	Steinernema puertoricense	G. mellonella	Coconut plantations	[171]	
Russia	Steinernema arenarium	–	Preserved specimens	[137]	
Republic of Rwanda	Heterorhabditis ruandica	G. mellonella	Cropland soil	[162]	
Steinernema africanum	G. mellonella	Agricultural soils	[172]	
Spain	Steinernema riojaense	G. mellonella	Soil from Grapevine field	[173]	
South Africa	Steinernema citrae	G. mellonella, T. molitor	Soil from citrus orchard	[174]	
Steinernema innovationi	G. mellonella	Grain field	[175]	
Steinernema beitlechemi	G. mellonella	Grassland soil	[176]	
Steinernema biddulphi	G. mellonella	Maize field	[177]	
Steinernema fabii	G. mellonella	Soil from Black wattle plantation	[178]	
Steinernema jeffreyense	G. mellonella	Guava tree	[179]	
Steinernema khoisanae	Whitegrub and black vine weevil	Soil from fruit orchards	[180]	
Steinernema litchii	T. molitor	Soil from litchi orchards	[181]	
Steinernema nguyeni	G. mellonella	Soil from Olea africana tree	[182]	
Steinernema sacchari	Eldana saccharina, G. mellonella	Soil from sugarcane field	[183]	
Steinernema tophus	G. mellonella	Vineyard soil	[184]	
Steinernema bertusi	G. mellonella	Soil from wattle plantation	[185]	
Steinernema batswanae	G. mellonella	Sub-tropical woodland area	[186]	
Heterorhabditis safricana	G. mellonella	Soil from peach orchard	[187]	
Heterorhabditis noenieputensis	G. mellonella	Soil from fig tree	[188]	
Undescribed spp.	G. mellonella	Citrus orchard	[189]	
Serbia	Steinernema bicornutum	G. mellonella	Soil	[190]	
Sultanate of Oman	Steinernema abbasi	G. mellonella	Sandy soil	[191]	
Taiwan	Steinernema taiwanensis	Spodoptera litura	Grassland soil	[192]	
Turkey	Steinernema anatoliense	G. mellonella	Grassland soil	[193]	
Tanzania	Steinernema pwaniensis	G. mellonella	Loamy sand soil	[194]	
Thailand	Steinernema siamkayai	G. mellonella	Soil from sweet tamarind orchard	[195]	
Steinernema minutum	-	Soil	[196]	
USA	Steinernema glaseri	Popillia japonica	Soil	[197]	
Steinernema riobrave	Helicoverpa zea	Soil from corn field	[198]	
Steinernema scarabaei	Anomala orientalis
Popillia japonica	Turfgrass	[199]	
Steinernema texanum	G. mellonella	Soil from mixed weedy vegetation	[200]	
Steinernema intermedium	G. mellonella	–	[201]	
Steinernema spp.	G. mellonella	Soil	[202]	
Steinernema jollieti	G. mellonella	–	[203]	
Heterorhabditis georgiana	G. mellonella	Soil from pecan orchard	[204]	
Steinernema tbilisiensis	G. mellonella	Soil from deciduous forest	[205]	
Steinernema borjomiense	Oryctes nasicornis, G. mellonella	-	[206]	
Heterorhabditis megidis	Popillia japonica	Soil	[207]	
Heterorhabditis marelatus	G. mellonella	Soil	[208]	
Steinernema neocurtillae	Neocurtilla hexadactylla	Field soil	[209]	
Steinernema diaprepesi	Diaprepes abbreviates, G. mellonella	Field soil	[210]	
Heterorhabditis floridensis	G. mellonella	Citrus plantations	[211]	
Steinernema phyllophagae	Phyllophaga sp.	Oak	[212]	
Steinernema khuongi	G. mellonella	Citrus tree	[213]	
Steinernema oregonense	G. mellonella	Soil	[214]	
Uruguay	Steinernema scapterisci	Scapteriscus vicinus	–	[215]	
Vietnam	Steinernema robustispiculum	G. mellonella	Woodland	[216]	
Steinernema tami	G. mellonella	Cat Tien forest	[217]	
Steinernema sangi	G. mellonella	Soil	[218]	
Steinernema thanhi	G. mellonella	beach soil	[219]	
Steinernema loci	G. mellonella	beach soil	[219]	
Steinernema sasonense	G. mellonella	Forest soil	[220]	
Steinernema cumgarense	G. mellonella	Forest soil	[220]	
Steinernema backanense	G. mellonella	Forest soil	[220]	
Steinernema eapokense	G. mellonella	Forest soil	[220]	
Steinernema huense	G. mellonella	Forest soil	[221]	
Heterorhabditis baujardi	G. mellonella	Forest soil	[222]	
Venezuela	Steinernema papillatum	G. mellonella	Grassland	[223]	
Steinernema goweni	G. mellonella	Fallow field	[224]	

2 Nemato-bacterial complex

The nematode-associated endosymbiotic bacteria Achromobacter nematophilus were first described by Poinar Jr. and Thomas [40] from infective juveniles of Steinernema carpocapsae (Weiser). They illustrated the position of this endosymbiont inside the nematode juvenile using microscopic observations, including a compound, as well as electronic microscopy techniques [41]. He also explained the contribution of this bacterium to the life cycle of the nematode (Fig. 1). He also demonstrated the key role of this bacterium in the host's death. Thomas and Poinar assigned a new genus to this bacterium, which was designated as genus Xenorhabdus on the basis of its identifying features [42]. The endosymbiotic bacterium was isolated from H. bacteriophora [43], and Thomas and Poinar named it Xenorhabditis luminescens because of its fluorescence ability [42]. Later, it was realised that the bacterium possessed different phenotypic and genotypic characters, so the newly isolated bacterium was placed in a separate genus designated as Photorhabdus [44].Fig. 1 Life cycle of entomopathogenic nematodes; IJ: Infective Juvenile; G: generation; M: Moulting; J: Juvenile form [292].

Fig. 1

Xenorhabdus bacteria were located inside the special vesicle inside the nematode intestines [45], while Photorhabdus bacteria were disseminated in the digestive tract, including the fore and midgut regions of Heterorhabditis juveniles [46]. All Steinernema showed mutualistic symbiotic associations with the genus Xenorhabdus, and Heterorhabditis species showed symbiosis with the genus Photorhabdus [47]. The symbiosis mechanism between EPNs and their endosymbiotic bacteria was a chief turning point for the utilisation of EPNs as bioagents. Based on their phenotype, these bacteria are Gram-negative, non-spore-forming, facultative anaerobes, and rod-like in appearance [48]. These bacteria are well known for their phenotypic dissimilarities and exhibit a mechanism of phase variation. The bacteria exhibit digenetic forms that show phase variation in their life forms [49]. Phase I was the metabolic phase with primary cells that imparted a red colour, or sometimes brick red colour, to the colony with characteristic bioluminescence properties. The phase II bacterial cells showed glistening, whitish colonies that exhibit non-bioluminescence characters. Earlier, it was reported that Photorhabdus luminescens (Thomas and Poinar) shows red-coloured colonies overlaid by the dark blue colour of the media [50]. The colonies of P. luminescens were observed to be circular, smooth, glistering, convex, and had white margins with a shiny appearance [51].

Phase-I is denoted as the primary phase variant found inside the infective juvenile stage of the nematode. It is more virulent and binds with specific dyes. It produced a wide range of metabolic compounds [52]. Phase II, or the secondary variant, is produced when resources (low oxygen and osmolarity levels) are exhausted [53] and the organism is less virulent. Phase II cells have low antibiotic production, untraceable bioluminescence, less pigmentation, and lower virulence than phase I cells [54]. Both phases help in the growth, development, and reproduction of the nematodes [55]. Smigielski et al. reported that phase I cells enhanced nematode growth and development during bacterial-nematode interactions [53]. Eckstein and Heermann also described the phenotypic switching among the primary and secondary cells of P. luminescens [56]. Tu et al. reported the two phases in the life cycle of Photorhabdus species that exhibit different secondary metabolite compositions in every phase [57].

3 Biocontrol attributes of entomopathogenic nematodes

Entomopathogenic nematodes showed biocontrol potential against a wide range of insect pests and contributed to the maintenance of insect populations naturally in the environment. In conventional, conservation, and augmentative biological control programmes, EPNs are an emerging and effective choice for managing this insect [58]. More than 200 insect species from various orders within the class Insecta are infected by nematodes, which have a vast host range [59]. EPNs have been successfully used in agricultural systems because of their virulence, motility, quickness, adaptability, and good sense of chemoception [32]. Because of their minimal negative impact on nontarget organisms, using native biocontrol agents to target a particular pest has been adopted. The process of pathogenicity is based on the characteristic interaction between the nematode bacteria and the host insect. It is influenced by insect resistance and by virulence factors of the bacteria and of the nematode acting separately or together to overcome the defence system [54]. EPNs are progressively utilised as biocontrol agents for insect pests (including humoral and cellular defences) [60].After infection, they kill the target host in 24–48 h. They successfully infiltrate and parasitize the larval stages of insect groups that live in soil [23]. The fact that 90 % or more insect pests need to live their entire life cycle in the soil is one of the reasons EPN is used as a bio-agent. Thus, they have been effectively used to combat both folivorous and ground-dwelling insects [31] (Table 2).Table 2 Biocontrol potential of entomopathogenic nematodes against insect pests.

Table 2Nematode species	Target insect	Order	Location	Reference	
S. scapterisci	Scapteriscus vicinus	Orthoptera	South America	[225]	
S.carpocapsae and H. megidis	Otiorhynchus sulcatus	Lepidoptera	Berkshire, USA	[226]	
S. abbasi, S. carpocapsae and H. indica	A. ipsilon	Lepidoptera	Bengaluru, India	[227]	
S. monticola, S. carpocapsae and H. bacteriophora	S. litura	Lepidoptera	Korea	[228]	
Heterorhabditis sp., GAU EPN 16 and S. riobrave GAU EPN 3	Helicoverpa armigera	Lepidoptera	Gujarat, India	[229]	
S. scarabaei	Asian garden beetle Maladera castanea	Coleoptera	New Brunswick	[230]	
S. riobrave	A. ipsilon	Lepidoptera	Gujarat, India	[231]	
H. mexicana, S. carpocapsae, H. bacteriophora	A. ipsilon	Lepidoptera	USA	[232]	
S. abbasi, S. feltiae, S. carpocapsae, S. glaseri, S. riborave, H. bacteriophora and H. indica	S. litura	Lepidoptera	Kerela, India	[233]	
S. carpocapsae and S. riobrave	Synanthedon exitiosa	Lepidoptera	Byron, USA	[234]	
H. indica and S. glaseri	H. armigera	Lepidoptera	Coimbatore, India	[235]	
H. megidis	Otiorhynchus sulcatus	Lepidoptera	Ireland and Norway	[236]	
S. carpocapsae	P. brassicae	Lepidoptera	Georgia	[237]	
H. zealandica	Cydia pomonella	Lepidoptera	Stellenbosch University, South Africa	[238]	
Steinernema spp. and Heterorhabditis spp.	Synanthedon Pictipes	Lepidoptera	Columbus, Ohio	[239]	
H. indica	Plutella xylostella, H. armigera, Leucinodes orbonalis, Earias vittella, S. litura, Pieris brassicae, Spilosoma obliqua, and Holotrichia consanguinea,	Lepidoptera and Coleoptera	Indian Agricultural Research Institute, New Delhi, India	[240]	
H. bacteriophora S. feltiae isolate IRA24 and S. carpocapsae	H. armigera	Lepidoptera	North-western Iran	[241]	
H. indica, S. feltiae, H. bacteriophora H. Georgiana, S. riobrave and S. carpocapsae	Corythucha ciliata and Stethobaris nemesis	Hemiptera and Coleoptera	Florida,USA	[242]	
H. bacteriophora	A. ipsilon	Lepidoptera	Jammu and Kashmir, India	[243]	
S. kraussei	A. segetum	Lepidoptera	Turkey	[244]	
S. carpocapsae and H. indica	S. litura	Lepidoptera	Bengaluru, India	[245]	
S. carpocapsae	Mamestra brassicae	Lepidoptera	Rumbeke-Beitem, Belgium	[246]	
S.carpocapsae, S. feltiae, H.bacteriophora, H. megidis and H. indica	Herpetogramma phaeopteralis	Lepidoptera	Florida, USA	[247]	
S. feltiae, S. carpocapsae, H. bacteriophora	Thrips tabaci	Thysanoptera	India	[248]	
H. bacteriophora	P. brassicae	Lepidoptera	Kashmir, India	[249]	
S. masoodi	H. armigera	Lepidoptera	Varanasi, India	[250]	
S. abbasi, S. masoodi, S. seemae, H. indica and H. bacteriophora	H. armigera	Lepidoptera	Tarai region of Uttar Pradesh, India	[251]	
H. bacteriophora and S. carpocapsae	A. segetum	Lepidoptera	Iran	[252]	
S. riobrave	Helicoverpa zea	Lepidoptera	Georgia, USA	[253]	
S.carpocapsae, S. feltiae and H.bacteriophora	Leptinotarsa decemlineata	Coleoptera	Czech Republic	[254]	
S. carpocapsae	S. litura	Lepidoptera	Udaipur, India	[255]	
Heterorhabditis sp. (HSG isolate), Heterorhabditis sp. (HKM isolate) and H. bacteriophora (HRJ isolate)	H. armigera, S. litura and P. xylostella	Lepidoptera	Himachal Pradesh, India	[256]	
S. abbasi, S. siamkayai and H. indica	H. armigera	Lepidoptera	Uttar Pradesh, India	[257]	
H. bacteriophora and S. carpocapsae, S. feltiae	Thrips tabaci	Thysanoptera	Giza, Egypt	[258]	
H. indica and S. glaseri	A. ipsilon	Lepidoptera	Coimbatore, Tamil Nadu, India	[259]	
H. bacteriophora and S. feltiae	P. brassicae	Lepidoptera	Iran	[260]	
S. glaseri and H. bacteriophora	S. litura	Lepidoptera	Faisalabad, Pakistan	[261]	
S. glaseri	Armyworms and bollworms	Lepidoptera	Faisalabad, Pakistan	[262]	
H. bacteriophora strain FLH-4-H and H. indica strain 216-H, S. carpocapsae strain E76-S, S. feltiae strain E76-S	A. ipsilon	Lepidoptera	Cappadocia Region, Central Turkey	[263]	
S. carpocapsae	Pieris rapae	Lepidoptera	Giza, Egypt	[264]	
H. indica, H. bacteriophora strain HRJ, Heterorhabditis sp. HSG, Heterorhabditis sp. strain HKM and Heterorhabditis sp. strain HSG	A. segetum	Lepidoptera	Palampur, Himachal Pradesh, India	[265]	
H. noenieputensis and S. yirgalemense	Ceratitis capitata	Diptera	South Africa	[266]	
H. bacteriophora	S. litura, C. cephalonica, Bombyx mori, G. mellonella and Brahmina coriacea	Lepidoptera and Coleoptera	Nauni, Solan, Himachal Pradesh, India	[267]	
H. indica	S. litura	Lepidoptera	Junagadh, India	[268]	
S. carpocapsae and H. indica	A. segetum	Lepidoptera	Shimla, Himachal Pradesh, India	[269]	
Heterorhabditis spp.	H. armigera	Lepidoptera	North West Himalayas	[270]	
H. indica	H. armigera and S. litura	Lepidoptera	Nagpur, Maharashtra, India	[271]	
H. bacteriophora, H. baujardi, H. indica, H. zealandica, H. noenieputensis and S. jeffreyense, S. yirgalemense	Holocacista capensis	Lepidoptera	Western Cape, South Africa	[272]	
S. yirgalemense	Frankliniella occidentalis	Thysanoptera	South Africa	[273]	
S. carpocapsae and H. bacteriophora	Heliothis virescens	Lepidoptera	South Africa	[274]	
S. litura to H. indica, H. bacteriophora, S. longicaudum and S. carpocapsae	S. litura	Lepidoptera	Daegu, Republic of Korea	[275]	
H. pakistanensis	P. brassicae	Lepidoptera	Srinagar, Jammu and Kashmir, India	[276]	
H. bacteriophora and S. glaseri	P. brassicae	Lepidoptera	Faisalabad, Pakistan	[277]	
S. feltiae and H. bacteriophora	P. brassicae	Lepidoptera	Himachal Pradesh, India	[278]	
H. bacteriophora and S. aciari	Odontotermes obesus and A. ipsilon	Blattodea and Lepidoptera	Assam, India	[279]	
H. bacteriophora	A. ipsilon	Lepidoptera	Assam, India	[280]	
H. amazonensis strain MC01	H. armigera	Lepidoptera	Brazil	[281]	
EPNs strains viz. NBS1, GNAF1 and MAF2	S. litura	Lepidoptera	Tamil Nadu, India	[282]	
H. bacteriophora	H. armigera, S. litura and A. segetum	Lepidoptera	Himachal Pradesh, India	[283]	
H. bacteriophora strain S26	P. brassicae	Lepidoptera	Himachal Pradesh, India	[35,284]	
H. bacteriophora	S. litura	Lepidoptera	Himachal Pradesh, India	[285]	
Five mid-himalyan isolates of H. bacteriophora	S. litura	Lepidoptera	Himachal Pradesh, India	[286]	
H. bacteriophora	H. armigera, S. litura, A. segetum and Mythimna separata	Lepidoptera	Himachal Pradesh, India	[287]	
S. carpocapsae and H. bacteriophora	Rhynchophorus ferrugineus	Coleoptera	Pakistan	[288]	
H. indica	Spodoptera frugiperda	Lepidoptera	Maharashtra, India	[289]	
H. bacteriophora	S. litura	Lepidoptera	Himachal Pradesh, India	[290]	
S. carpocapsae	P. brassicae	Lepidoptera	Himachal Pradesh, India	[291]	

4 Formulations developed from entomopathogenic nematodes

A considerable enhancement has been made in the past few years for the development of nematode-based formulations. In general, an EPN-based formulation contains an active ingredient, i.e., nematodes, a carrier that may include liquids, solids, and gels, as well as cadavers, and an additive that may be adsorbents, absorbents, surfactants, emulsifiers, humectants, thickeners, antimicrobials, dispersants, and also a protector that protects from UV-rays [61]. These components in the formulation usually enhance the EPNs survival and sustain the pathogenicity of the nematodes. Different types of nematode-based formulations are aqueous suspensions, synthetic sponges, wheat-based gluten matrices, gels (alginate), starch matrices, polyacrylamide gels, clay, charcoal, vermiculite, and powder forms, including water-dispersible granules (Table 3).Table 3 Biocontrol potential of entomopathogenic nematodes against insect pests.

Table 3Nematode species	Type of formulation	Product name	Target pest	Manufacturers	
S. carpocapsae
H.bacteriophora	Polyether-polyurethane moist sponge	NOBUG	Plant borers	The National Research Centre (NRC), Egypt	
H. bacteriophora	Wettable powder	Cryptonem	Thaumatotibia leucotreta, Apples/codling moth, fruit weevil, black vine weevil, sciarids	River Bioscience, South Africa	
Wettable powder	Grubcide	White grubs, Termites,	Sri Biotech, India	
Inert carrier or clay	Larvanem	Curculionidae, Otiorhynchus	Biocont Laboratory spol. s r. o., Czech Republic, Europe	
Inert carrier or clay	Nematop	Curculionidae, Otiorhynchus	e-nema GmbH, Europe	
Clay powder	Nemagreen	Chafer grubs, billbugs, Japanese beetle, Oriental beetle, Asiatic garden beetle	e-nema GmbH, Germany	
Clay powder	Nematop	White grubs, black vine weevil	e-nema GmbH, Germany	
Wettable powder	OPTINEM H	Otiorhynchus sulcatus and other soil beetles	Agrifutur Srl, Italy	
Wettable powder	H. bacteriophora	O. sulcatus, Balaninus nucum, C. splendana, C. fagiglandana, Diabrotica virgifera, virgifera	Andermatt Biocontrol AG
Swiss Confederation, Italy	
Wettable powder	NEMAPACK HB	O. sulcatus and other soil beetles	Bioplanet s.c.a., Italy	
Wettable powder	Nematop	O. sulcatus	Biogard Divisone CBC (Europe) Srl, Italy	
Wettable powder	Terranem	Phyllopertha horticola, Serica brunnea, Hoplia spp., Aphodius spp	Koppert Italia Srl, Italy	
Wettable powder	Larvanem	O. sulcatus	Koppert Italia Srl, Italy	
Wettable powder	Nemax H	O. sulcatus and D. virgifera	Serbios Srl, Italy	
Wettable powder Clay	B-GREEN	P. horticola	Biobest, Poland	
Wettable powder Clay	LARVANEM	O. sulcatus, P. horticola, Melolontha spp., A. solstitialis	Koppert, Poland	
H. indica	Wettable powder	Soldier and Armour	White grubs, Termites, Ash weevil, Sweet potato weevil	Multiplex, Ponalab, and Camson Biotech, India	
H. indica LPP30
H. baujardi LPP7	Infected cadavers	Infected cadavers of G. mellonella	Conotrachelus psidii	UENF (Universidade Estadual Norte Fluminense) (state university in Brazil	
H. megidis	Wettable powder Clay	Heterorhabditis-System	O. sulcatus	Biobest, Poland	
Wettable powder Clay	Nemasys HM	O. sulcatus	Syngenta Bioline Becker Underwood, Poland	
Wettable powder Clay	Exhibilinehm	O. sulcatus	Syngenta Bioline, Poland	
S. carpocapsae	Clay powder	Nemastar	Capnodis tenebrionis, leatherjackets, mole crickets and black cutworms	e-nema GmbH,
Germany	
Wettable powder	OPTIMEN C	C. pomonella, C. molesta, Euzophera bigella	Agrifutur Srl, Italy	
Wettable powder	S. carpocapsae	C. pomonella, Agrotis spp., Rhynchophorus ferrugineus, Tipula spp., Gryllotalpa gryllotalpa, Duponchelia fovealis, Paysandisia archon	Andermatt Biocontrol AG
Swiss Confederation, Italy	
Wettable powder	NEMAPACK SC
Palme	R. ferrugineus, P. archon	Bioplanet s.c.a., Italy	
Wettable powder	NEMAPACK SC	C. pomonella	Bioplanet s.c.a., Italy	
Wettable powder	Nemastar	R. ferrugineus, P. archon, Cydia spp., Synanthedon spp., Spodoptera spp., Agrotis spp., D. fovealis, Amphimallon solstitialis, Rhizotrogus aestivus, C. tenebrionis, G. grillotalpa, Tipula paludosa, T. oleracea	Biogard Divisone
CBC (Europe) Srl, Italy	
Wettable powder clay	Capsanem	Noctuidae, Pyralidae, Tipulidae, Coleoptera and G. gryllotalpa	Koppert, Poland	
Wettable powder clay	Exhibitline Sc	Agrostis spp., Tipula spp.	Syngenta Bioline, Poland	
Wettable powder clay	Nemabact	Western flower thrips, wireworm	OOO Biometodika, Russia	
S. pakistanense	Sponge	PakNema-1	American, spotted, pink bollworm and root-knot nematodes	NNRC, Pakistan	
S. maqbooli	Sponge	PakNema-2	Root-knot nematodes	NNRC, Pakistan	
S. bifurcatum	Sponge	PakNema-3	Pupal stages in soil and termite	NNRC, Pakistan	
S. balochiense	Sponge	PakNema-4	Fruit fly pupa	NNRC, Pakistan	
S. feltiae
S. feltiae	Inert carrier or clay	Entonem	Sciaridae – larvae	Biocont Laboratory spol. s r. o., Czech Republic, Europe	
Inert carrier or clay	Nemaplus	Sciaridae – larvae	e-nema GmbH, Europe	
Inert carrier or clay	Steinernema-System	Sciaridae – larvae	Biobest NV, Europe	
Clay powder	Nemaplus	Glasshouse sciarids	e-nema GmbH, Germany	
Clay powder	Nemapom	Cydia pomonella Synanthedon myopaeformis	e-nema GmbH, Germany	
Wettable powder	OPTIMEN F	Cydia pomonella	Agrifutur Srl (Italy)	
Wettable powder	S. feltiae	Sciaridae, C. pomonella, thrips	Andermatt Biocontrol AG (Swiss Confederation), Italy	
Wettable powder	NEMAPACK SF	Sciarids, phorids, Agromyza spp., muscids, nottuids,
Agrotis spp., Cossus spp., sesids	Bioplanet s.c.a., Italy	
Wettable powder	Nemaplus	Cydia spp.,Synanthedon spp., Frankliniella occidentalis, Bradysia spp., Acrolepiopsis assectella, Ephrytidae, Scatella stagnalis, Scatellatenui costa, Parahypopta caestrum, Lyriomiza spp., Tuta absoluta	Biogard Divisone CBC (Europe) Srl Italy	
Wettable powder	Entonem	Sciarids and thrips	Koppert Italia Srl, Italy	
Wettable powder	Nemapom	Cydia spp., Synanthedon spp. C. tenebrionis	Biogard Divisone CBC (Europe) Srl, Italy	
Wettable powder clay	SCIARID	F. occidentalis, Sciaridae, Diptera	Koppert, Poland	
Wettable powder clay	Nemasys F	F. occidentalis, Sciaridae, Diptera	Syngenta Bioline, Poland	
Wettable powder clay	Exhibitline Sf	F. occidentalis, T. tabaci	Syngenta Bioline, Poland	
Wettable powder clay	Entonem	Thrips tabaci, P. horticola	Koppert, Poland	
Wettable powder clay	Steinernema- System	F. occidentalis, Sciaridae, Diptera	Biobest, Poland	
Wettable powder clay	Entonem F	Western flower thrips, wireworm, Melolontha chafers	OOO Biometodika, Russia	
S. kraussei	Wettable powder clay	Exhibitline Srb and Nemasys L	O. sulcatus	Syngenta Bioline, Poland	
S. braziliense	Sponge	Bio Steinernema	Sphenophorus levis	Bio Controle, Brazil	

4.1 Storage in the form of aqueous suspension

The most common method adapted for EPN storage is the aqueous suspension form. The methods have also been employed for the transportation as well as applications of EPNs [62]. The storage of several species of EPNs at a lower temperature (4–15 °C) increased their survival rates [63]. The storage of Steinernema spp. and Heterorhabditis spp. at this temperature range has survivorships of 6–12 months and 3–6 months, respectively. However, in several species, low temperatures typically reduce EPN mobility and enzymatic activity, which severely impairs IJ function [64]. Although nematode application is a cost-effective method, the necessity of refrigeration and transportation may enhance their cost [[65], [66], [67]]. The nematode's survival is also influenced by other factors such as nematode concentration, high oxygen requirement, sedimentation, highly reduced temperature ranges for some species, and contamination by microbes [68]. It was reported that the optimum relative humidity was required by S. feltiae and H. bacteriophora (>90 %) for their survival, while S. carpocapsae could survive at lower relative humidity (74 %), as rapid desiccation is the major limiting factor in nematode survival [69]. The nematode's survival also depends on the pH and oxygen concentration. There have been reports that suggest that the increased pH and decreased oxygen concentrations affect the viability of S. carpocapsae and Steinernema glaseri (Ssteiner) [70]. Nematode mortality can result from microbial contamination, particularly that caused by fungi, which can lower the quantity of oxygen available and compete with the nematodes [71].

4.2 Storage in the form of infected cadaver

The IJs can be stored in the form, of infected cadavers. Galleria mellonella (Linnaeus) larvae are the most frequently employed for this purpose due to their excellent qualitative value and high IJ-production rates [72]. Because the cadaver defends the released IJs from damaging biotic and abiotic elements, they have more energy reserves, are better able to spread and infect the host, and can survive longer in the soil [73]. A mean of 14,59,205 and 18,98,512 IJs can be produced per gram of host by employing wax moth larvae [74]. This technique allows small-to medium-sized farmers to use infected insect cadavers for insect pest control [75]. The breaking and rupturing of the insect carcass during its transportation, storage, and shipment is a drawback of employing this formulation approach. This restriction can be addressed, though, by covering the insect cadaver with a protective substance, such as clay or starch [76].

4.3 Storage in the form of inert carriers

Nematode storage in the form of an inert carrier is the most suitable method to keep the nematode in small quantities under the refrigerator. The commonly used carriers are in the form of sponge storage and vermiculite. The sponge-based formulation contained a polyether-polyurethane sponge on which nematode suspension was applied at a rate of 500–1000 IJs/cm2. The water was also applied to wet the remaining part of the sponge, and the sponge was kept inside the zip-lock plastic bag that was placed under the refrigerator (5–10 °C). The EPN survivorship inside these bags is up to 3 months [65,71]. Another inert carrier was vermiculite, which was a much upgraded form of EPN storage as well as transportation.

In vermiculite, the concentrated nematode suspension is mixed with micronized vermiculite and then kept under the zip-lock plastic bag for storage. The vermiculite was much more stable and easy to apply [68]. The EPN formulation containing EPNs S. carpocapsae, S. feltiae, and S. longicaudum (Shen and Wang) upon mixing with humus and vermiculite showed 90 % survivability after 120 days at 5 °C [77]. Recently, S. feltiae was stored at different temperature ranges with the use of a polyacrylamide gel and vermiculite mixture. About 80 % viability was recorded in stored nematodes after 241 days at 15 °C, which is reduced to 233 days at 25 °C and about 30 days at 35 °C [78].

4.4 Storage in the form of gels

The gel-based formulation of EPNs through encapsulation was first introduced by Kaya and Nelsen [79]. The EPN encapsulations include calcium alginate gel that was prepared for the slower release of nematode juveniles. However, it was not so successful. Polyacrylamide was used by Bedding et al. [80] to contain partially desiccated EPNs, but this storage gel was very difficult to dissolve and has low nematode survivability [68]. Much perfection in EPN storage and viability was achieved by Georgis [81] when he used calcium alginate-based sheets dispersed on plastic screens. The macrogel-based matrix containing mono-glycerides and diglycerides from the vegetable oil was also utilised for the encapsulation of S. carpocapsae, which considerably enhanced the nematode viability [82]. The method was further improved with the addition of acrylamide and hydrogenated vegetable oil, which increased the S. carpocapsae survival rate by up to 80 % for 35 days [83]. The osmotic treatment was applied to the same material (acrylamide and hydrogenated vegetable oil) before the development of the encapsulated formulation. When the formulation of S. feltiae was developed from this material, it showed 99.8 % viability even after six months [62].

The hydrogel, kaolinite, and calcium alginate-based capsule formulations, including those of H. bacteriophora and S. carpocapsae, showed an EPN viability of 50 % after 40 days [84]. Navon and his co-workers developed calcium alginate-based formulations using EPNs S. feltiae, H. bacteriophora, S. carpocapsae, and Steinernema riobravae (Cabanillas, Poinar, and Raulston) and applied them against lepidopteran insect pests such as Spodoptera littoralis (Boisduval) and Helicoverpa armigera (Hübner) [85]. The NemaGel formulation of Steinernema abbassi (Elawad, Ahmad, and Reid) can be viable for about 9 months with a survival rate of 89 % in the temperature range of 15 °C–39 °C. However, the nematode stored in alginate gel beads can be escaped out of these beads due to the ultra-softness of the beads [86,87]. This might also happen due to the increased temperature during storage [88].

4.5 Storage in the form of clay

The sandwich model introduced by Bedding [89] is used for partial desiccation and the elimination of extra moisture from the body surface of the nematodes. The nematode juveniles were blended with clay and stored in a sandwich form that consisted of two layers of clay, and in-between the nematode juveniles were placed. S. feltiae, Steinernema bibionis (Bovien), S. glaseri, and Heterorhabditis heliothidis (Khan, Brooks, and Hirschmann) were used to develop this sandwich formulation that contained hygroscopic attapulgite clay with 8 weeks of viability. Although the formulation was commercialised and sold, it was soon terminated due to poor shelf life, poor storage, poor solubility, clogging during application, and a lower nematode-clay ratio [68].

Several studies indicate that EPNs may also be stored in the infected cadaver [90]. There is much research evidence that the use of an infected cadaver was much superior to the use of an use of an aqueous storage suspension [91]. However, it was found that the infected cadaver's formulation may have several problems in transportation and applications that resulted in a decrease in effectiveness. Ansari and his co-workers coated the infected cadaver with a kaolin-starch mixture and evidenced the maximum nematode viability even after 1 year, which caused 90 % mortality in Hoplia philanthus (Fuesslin) [92]. The infected cadaver-based formulations of S. carpocapsae, S. feltiae, and H. bacteriophora were covered with starch and clay by Lacey et al. [93]. The coated cadavers were again laminated with anti-desiccants such as wood flour foam that showed significant mortality in Cydia pomonella (Linnaeus) moth larvae upon its application in mulch and in aqueous suspensions.

4.6 Storage in the form of granules

The granular formulations of nematodes in the form of pellets were developed by Capinera and Hibbard [94]. The pellets contained a mixture of multiple components, including corn oil, alfalfa meal, wheat bran, wheat flour, and water. The pellet formulation containing S. feltiae was applied against Melanoplus spp. in field conditions, resulting in 78.1 % mortality. Connick et al. [95] encapsulated S. carpocapsae under a wheat flour-based gluten matrix called “Pesta''. The nematode viability inside this Pesta was quite low (6 weeks), and the risk of contamination by bacteria and fungi was very high. However, the formulation was dried to overcome the contamination issues, but the viability of the nematode was also reduced. Additionally, the dried granules turned into hard pellets that did not dissolve; even a 0.2 % formaldehyde solution was added during the preparation of Pesta to reduce the stress of microbial contamination [96]. The granular formulations developed from diatomaceous earth, amorphous silica, hydroxyethylcellulose, pregelatinized starch, lignosulfonate, starch, pregelled attapulgite clay, and fumed hydrophobic silica that contained S. carpocapsae, Steinernema scapterisci (Nguyen and Smart), S. feltiae, and S. riobravis showed 90 % viability when stored for up to 6 weeks. The 19 formulations, including combinations of starches, flours, clays, etc., were tested by Shapiro-Ilan et al. [76] to store H. bacteriophora. They found the formulated cadavers were amenable to desiccation, more resistant, and did not stick together.

4.7 Storage in the form of activated charcoal

The nematode can be stored in the form of activated charcoal. Yukawa and Pitt [97] mixed the nematodes with activated charcoal powder, which acts as an absorbent material. But the method was costly, difficult to apply, and had poor nematode viability and storage.

4.8 Storage in the form of wettable powder

The dispersible granular formulation contained a combination of materials like silica, starches, clays, cellulose, and lignin, which were used to encapsulate nematode juveniles [98]. The recent research tactics focused on the enhancement of viability, storage efficiency, effectiveness, and increased field persistence [99]. The wettable powder formulations have the ability to suspend in water. Nagesh et al. [100] granted a patent on the development of a silicate-based wettable powder mixture that contained IJs of H. indica strain NBAII Hi1 and H. bacteriophora strain NBAII Hb5. The diatomaceous earth powder (present naturally in the environment in the form of sedimentary rocks) was also utilised for the nematode formulation [101]. Recently, Kagimu and Malan [88] used diatomaceous earth powder for the preparation of EPN formulations that contained Steinernema yirgalemense (Nguyen, Tesfamariam, Gozel, Gaugler and Adams), Steinernema jeffreyense (Malan, Knoetze and Tiedt), and H. bacteriophora with high viability at 14 °C–25 °C temperature ranges. Cortés-Martínez et al. [102] reported longer survivability of S. glaseri upon pelletization with diatomaceous earth powder. High efficacy was also recorded when applied against Phyllophaga vetula (Horn).

5 Applications and future outlook

Generally, the most familiar methodology for EPN application is in the form of aqueous suspension. The nematode juveniles stored in the form of wettable powder, vermiculite, were suspended in water and applied. The juveniles stored in polyether-polyurethane sponges were also squeezed directly and suspended in the water. The gel-based formulations were not suspended in water, so a sodium citrate solution was also added to suspend the alginate gels [81]. Another important aspect was the regular shaking of the suspension during its application, as the suspended nematodes settled at the base of the water. Proper oxygen supply and mixing are required for the consistent distribution of EPNs [68,103]. Shapiro-Ilan et al. [104] suggested the nematode application time be in the early morning and evening, when the temperature is colder, so that the sprayed nematodes do not get desiccated. The nematodes can be applied using mist blowers, hand sprayers, pressure sprayers, and electrostatic sprayers [103].

Nematode has been recognised as an excellent biocontrol agent against a wide range of insect pests. Both forms of insects, above ground and below ground, are highly susceptible to nematode infection. The biocontrol potential of EPNs has been studied not only in the laboratory but also in green house and field conditions, and the nematode-based products (formulations) have been commercialised throughout the world, including agricultural and horticultural practices. However, it is not accepted by the farmers to that extent. So few research needs in the future include: cost reduction in the nematode-based formation as these products are too expensive; easy availability in the market to bring profit to the farmers; production efficiency should be enhanced with longer shelf life and better stability; requirements of the professional market; improved formulation should be developed to increase the viability in extreme temperatures so that farmers can use it easily; and farmers should be educated on the adaptation of EPNs as biocontrol agents.

6 Conclusions

This review identified the major concerns about why EPNs should be used for insect pest management. EPNs are ubiquitous in nature and have been successfully utilised as biocontrol agents for the management of insect pests. The nematodes, along with their endosymbiotic bacteria, combine to form a nemato-bacterial complex. This complex is responsible for causing mortality in insect pests within 72 h of infection due to the production of insecticidal compounds. The adaptation of EPNs is an eco-friendly method, economical, and safer for the environment as well as non-target organisms. Moreover, it's a better alternative to synthetic chemical pesticides, as it can be helpful in overcoming pest resistance and resurgence issues. Application of nematode juveniles is a cost-effective method, but the necessity of refrigeration and transportation may enhance their cost. But there are some biotic and abiotic stresses that affect the bio-efficacy of the nematodes and result in poor insect management. The nematode-based formulations are, although easy to apply, difficult to keep for a longer period of time. The attack of microbial contaminants is the limiting factor in the storage of EPNs. Temperature, storage, low oxygen, and transportation also affected the EPN's viability. Other factors that predominantly reduced the usage of nematodes for insect pest management are: the nematode formulations are usually more costly than the chemical-based pesticides; the mass production process is long and the in vivo process requires equipment and labour; the insect population increased the costs; the preparation of formulations, storage, and transportation also made formulations expensive; the nematode suspension should be applied in the evening and early morning to avoid sunlight exposure; and the nematode application was followed by irrigation to maintain humidity and efficacy.

Ethical approval

Not applicable.

Consent to publish

Not applicable.

Data availability statement

Not applicable.

CRediT authorship contribution statement

Preety Tomar: Writing – original draft. Neelam Thakur: Writing – review & editing. Sangram Singh: Writing – review & editing. Sanjeev Kumar: Writing – review & editing. Sarvesh Rustagi: Writing – review & editing. Ashutosh Kumar Rai: Writing – review & editing. Sheikh Shreaz: Writing – review & editing. Neelam Yadav: Writing – review & editing. Pankaj Kumar Rai: Writing – review & editing. Ajar Nath Yadav: Writing – review & editing, Conceptualization.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Acknowledgements

The authors are thankful to Vice Chancellor, Eternal University, Baru Sahib for providing necessary facilities.
==== Refs
References

1 FAO Climate change fans spread of pests and threatens plants and crops 2021 new FAO study
2 Oerke E.-C. Crop losses to pests J. Agric. Sci. 144 2006 31 43
3 He D.-c. Zhan J.-s. Xie L.-h. Problems, challenges and future of plant disease management: from an ecological point of view J. Integr. Agric. 15 2016 705 715
4 Pimentel D. Pesticides and pest control Peshin R. Dhawan A.K. Integrated Pest Management: Innovation-Development Process vol. 1 2009 Springer Dordrecht 83 87
5 Forget G. Balancing the need for pesticides with the risk to human health Forget G. Goodman T. De Villiers A. Impact of Pesticide Use on Health in Developing Countries: Proceedings of a Symposium Held in Ottawa, Canada, 17-20 Sept. 1990 1993 IDRC Ottawa, ON, CA
6 Z. MracekUse of entomoparasitic nematodes (EPANs) in biological control Upadhyay R.K. Advances in Microbial Control of Insect Pests 2003 Springer 235 264
7 Akhurst R.J. Xenorhabdus nematophilus subsp. poinarii: its interaction with insect pathogenic nematodes Syst. Appl. Microbiol. 8 1986 142 147
8 Bhat A.H. Chaubey A.K. Askary T.H. Global distribution of entomopathogenic nematodes, Steinernema and Heterorhabditis, Egypt J. Biol. Pest Co. 30 2020 1 15
9 Poinar G.O. Entomogenous Nematodes: a Manual and Host List of Insect-Nematode Associations 1975 Brill Archive 1 312
10 Ehlers R.-U. Mass production of entomopathogenic nematodes for plant protection Appl. Microbiol. Biotechnol. 56 2001 623 633 11601608
11 Gaugler R. Bilgrami A.L. Nematode Behaviour 2004 CABI 1 419
12 Burr A.J. Robinson A.F. Locomotion behaviour Gaugler R. Bilgrami A.L. Nematode Behaviour CABI 2004 25 62
13 Lewis E. Clarke D. Nematode parasites and entomopathogens Vega F.E. Kaya H.K. Insect Pathology 2012 Academic Press Amsterdam, The Netherlands 395 424
14 Campbell J.F. Kaya H.K. How and why a parasitic nematode jumps Nature 397 1999 485 486
15 Downes M.J. Griffin C.T. Dispersal behaviour and transmission strategies of the entomopathogenic nematodes Heterorhabditis and Steinernema Biocontrol Sci. Technol. 6 1996 347 356
16 Poinar G.O. Jr. Grewal P. History of entomopathogenic nematology J. Nematol. 44 2012 153 23482453
17 Kepenekci I. Hazir S. Lewis E.E. Evaluation of entomopathogenic nematodes and the supernatants of the in vitro culture medium of their mutualistic bacteria for the control of the root‐knot nematodes Meloidogyne incognita and M. arenaria Pest Manag. Sci. 72 2016 327 334 25721911
18 Griffin C. Boemare N. Lewis E. Biology and behaviour Grewal P.S. Ehlers R.U. Shapiro-Ilan D.I. Nematodes as Biocontrol Agents 2005 47 64
19 Devi G. Nath D. Entomopathogenic nematodes: a tool in biocontrol of insect pests of vegetables-A review Agric. Rev. 38 2017 137 144
20 Koppenhöfer A.M. Grewal P.S. Fuzy E.M. Differences in penetration routes and establishment rates of four entomopathogenic nematode species into four white grub species J. Invertebr. Pathol. 94 2007 184 195 17156793
21 Sajnaga E. Kazimierczak W. Evolution and taxonomy of nematode-associated entomopathogenic bacteria of the genera Xenorhabdus and Photorhabdus: an overview Symbiosis 80 2020 1 13
22 Stock S.P. Diversity, biology and evolutionary relationships Campos-Herrera R. Nematode Pathogenesis of Insects and Other Pests: Ecology and Applied Technologies for Sustainable Plant and Crop Protection 2015 Springer 3 27
23 Gaugler R. Entomopathogenic Nematology 2002 10.1079/9780851995670.0000
24 Clarke D.J. The genetic basis of the symbiosis between Photorhabdus and its invertebrate hosts Adv. Appl. Microbiol. 88 2014 1 29 24767424
25 Chaston J.M. Murfin K.E. Heath‐Heckman E.A. Goodrich‐Blair H. Previously unrecognized stages of species‐specific colonization in the mutualism between Xenorhabdus bacteria and Steinernema nematodes Cell Microbiol. 15 2013 1545 1559 23480552
26 Chaston J.M. Suen G. Tucker S.L. Andersen A.W. Bhasin A. Bode E. Bode H.B. Brachmann A.O. Cowles C.E. Cowles K.N. The entomopathogenic bacterial endosymbionts Xenorhabdus and Photorhabdus: convergent lifestyles from divergent genomes PLoS One 6 2011 e27909
27 Brivio M.F. Toscano A. De Pasquale S.M. De Lerma Barbaro A. Giovannardi S. Finzi G. Mastore M. Surface protein components from entomopathogenic nematodes and their symbiotic bacteria: effects on immune responses of the greater wax moth, Galleria mellonella (Lepidoptera: Pyralidae) Pest Manag. Sci. 74 2018 2089 2099
28 Poinar G.O. Jr. Taxonomy and biology of Steinernematidae and Heterorhabditidae Entomopathogenic nematodes in biological control 54 1990 23 74
29 Kaya H.K. Gaugler R. Entomopathogenic nematodes Annu. Rev. Entomol. 38 1993 181 206
30 Grewal P. Georgis R. Entomopathogenic nematodes Hall F.R. Menn J.J. Biopesticides: Use and Delivery 1999 271 299
31 Laznik Z. Vidrih M. Trdan S. The effects of different fungicides on the viability of entomopathogenic nematodes Steinernema feltiae (Filipjev), S. carpocapsae Weiser, and Heterorhabditis downesi Stock, Griffin & Burnell (Nematoda: Rhabditida) under laboratory conditions Chil. J. Agric. Res. 72 2012 62
32 Mahmoud M. Biology and use of entomopathogenic nematodes in insect pests biocontrol a generic view 49 2017 85 105
33 Sikandar A. Yuan R. Lian X. Zhen M. Zhao P. Li F. Lu X. Wang Y. Entomopathogenic nematodes as bioinsecticides–a Appl. Ecol. Environ. Res. 19 2021 2459 2476
34 Thakur N. Kaur S. Kaur T. Tomar P. Devi R. Thakur S. Tyagi N. Thakur R. Mehta D.K. A.N. Yadav Organic agriculture for agro-environmental sustainability Soni R. Suyal D.C. Yadav A.N. Goel R. Trends of Applied Microbiology for Sustainable Economy 2022 Elsevier 699 735
35 Tomar P. Thakur N. Isolation and evaluation of Heterorhabditis bacteriophora strain-S26 as biocontrol agents against Pieris brassicae L. under laboratory conditions Indian J. Nematol. 52 2022 49 58
36 Devi G. Mass production of entomopathogenic nematodes-A Review Int. J. Environ. Agric. Biotechnol 3 2018 1032 1043
37 Husin T.O.B. Port G.R. Efficacy of entomopathogenic nematodes against Tuta absoluta Biol. Control 160 2021 104699
38 Georgis R. Koppenhöfer A. Lacey L. Bélair G. Duncan L. Grewal P. Samish M. Tan L. Torr P. Van Tol R. Successes and failures in the use of parasitic nematodes for pest control Biol. Control 38 2006 103 123
39 Nguyen K. Smart G. Jr Taxonomy of insect parasitic nematodes Chen Z.X. Chen S.Y. Dickson D.W. Nematology: Advances and Perspectives. Volume 2: Nematode Management and Utilization 2004 CABI Publishing Wallingford UK 795 878
40 Poinar G.O. JR. Thomas G.M. new bacterium A. Achromobacter nematophilus sp. nov.(Achromobacteriaceae: eubacteriales) associated with a nematode Int. J. Syst. Evol. Microbiol. 15 1965 249 252
41 Poinar G.O. Thomas G.M. Significance of Achromobacter nematophilus Poinar and Thomas (Achromobacteraceae: Eubacteriales) in the development of the nematode, DD-136 (Neoaplectana sp. Steinernematidae) Parasitology 56 1966 385 390 4960247
42 Thomas G.M. Poinar JR G.O. Xenorhabdus gen. nov., a genus of entomopathogenic, nematophilic bacteria of the family Enterobacteriaceae Int. J. Syst. Evol. Microbiol. 29 1979 352 360
43 Poinar G.O. Lane R. Thomas G. Biology and redescription of Pheromermis pachysoma (v. Linstow) n. gen., n. comb.(Nematoda: mermithidae), a parasite of yellow jackets (Hymenoptera: vespidae) Nematologica 22 1976 360 370a
44 Boemare N. Akhurst R. Mourant R. DNA relatedness between Xenorhabdus spp.(Enterobacteriaceae), symbiotic bacteria of entomopathogenic nematodes, and a proposal to transfer Xenorhabdus luminescens to a new genus Photorhabdus gen. nov, Int. J. Sys. Evol. Microbiol. 43 1993 249 255
45 Bird A. Akhurst R. The nature of the intestinal vesicle in nematodes of the family Steinernematidae Int. J. Parasitol. 13 1983 599 606
46 Boemare N. Laumond C. Mauleon H. The entomopathogenic nematode-bacterium complex: biology, life cycle and vertebrate safety Biocontrol Sci. Technol. 6 1996 333 346
47 Boemare N. Akhurst R. The Genera Photorhabdus and Xenorhabdus, Prokaryotes vol. 6 2006 451 494
48 Imhoff J.F. Enterobacteriales Bergey's Manual® of Systematic Bacteriology 2005 Springer 587 850
49 Forst S. Nealson K. Molecular biology of the symbiotic-pathogenic bacteria Xenorhabdus spp. and Photorhabdus spp, Microbiol Rev. 60 1996 21 43
50 Sharad-Mohan A.-S. Gaur H. Successful management of mango mealy bug, Drosicha mangiferae by Photorhabdus luminescens, a symbiotic bacterium from entomopathogenic nematode Heterorhabditis indica Int. J. Nematol. 14 2003 195 198
51 Kumar K.V. Vendan K.T. Nagaraj S. Isolation and characterization of entomopathogenic symbiotic bacterium, Photorhabdus luminescens of Heterorhabditis indica from soils of five agro climatic zones of Karnataka Biosci. Biotech. Res. Asia 11 2014 129 139
52 Forst S. Dowds B. Boemare N. Stackebrandt E. Xenorhabdus and Photorhabdus spp.: bugs that kill bugs Ann. Rev Microbiol 51 1997 47 72 9343343
53 Smigielski A.J. Akhurst R.J. Boemare N.E. Phase variation in Xenorhabdusnematophilus and Photorhabdus luminescens: differences in respiratory activity and membrane energization Appl. Environ. Microbiol. 60 1994 120 125 16349145
54 Dowds B.C. Peters A. Virulence mechanisms Gaugler R. Entomopathogenic Nematology 2002 79 98
55 Volgyi A. Fodor A. Szentirmai A. Forst S. Phase variation in Xenorhabdus nematophilus Appl. Environ. Microbiol. 64 1998 1188 1193 16349534
56 Eckstein S. Heermann R. Regulation of phenotypic switching and heterogeneity in Photorhabdus luminescens cell populations J. Mol. Biol. 431 2019 4559 4568 31022406
57 Tu P.-W. Chiu J.-S. Lin C. Chien C.-C. Hsieh F.-C. Shih M.-C. Yang Y.-L. Evaluation of the antifungal activities of Photorhabdus akhurstii and its secondary metabolites against phytopathogenic Colletotrichum gloeosporioides J. Fungi 8 2022 403
58 Kumar D. Kumari P. Kamboj R. Kumar A. Banakar P. Kumar V. Entomopathogenic nematodes as potential and effective biocontrol agents against cutworms, Agrotis spp.: present and future scenario Egypt. J. Biol. Pest Co. 2 2022 42
59 Sharmila R. Subramanian S. Poornima K. Host range of entomopathogenic nematodes J. Entomol. Zool. Stud. 6 2018
60 Belien T. Entomopathogenic nematodes as biocontrol agents of insect pests in orchards CABI Reviews 2019 1 11
61 Grewal P. Anhydrobiotic potential and long-term storage of entomopathogenic nematodes (Rhabditida: Steinernematidae) Int. J. Parasitol. 30 2000 995 1000 10980289
62 Chen S. Glazer I. A novel method for long-term storage of the entomopathogenic nematode Steinernema feltiae at room temperature Biol. Control 32 2005 104 110
63 Hazir S. Kaya H.K. Stock S.P. Keskin N. Entomopathogenic nematodes (Steinernematidae and Heterorhabditidae) for biological control of soil pests Turk. J. Biol. 27 2003 181 202
64 Susurluk I.A. Influence of temperature on the vertical movement of the entomopathogenic nematodes Steinernema feltiae (TUR-S3) and Heterorhabditis bacteriophora (TUR-H2), and infectivity of the moving nematodes Nematology 10 2008 137 141
65 Grewal P.S. Formulation and application technology Gaugler R. Entomopathogenic Nematology 2002 CABI publishing Wallingford UK 265 287
66 Lacey L.A. Georgis R. Entomopathogenic nematodes for control of insect pests above and below ground with comments on commercial production J. Nematol. 44 2012 218 23482993
67 Shapiro-Ilan D.I. Cottrell T.E. Mizell R.F. III Horton D.L. Zaid A. Field suppression of the peachtree borer, Synanthedon exitiosa, using Steinernema carpocapsae: effects of irrigation, a sprayable gel and application method Biol. Control 82 2015 7 12
68 Grewal P. Peters A. Formulation and quality Grewal P.S. Ehlers R.U. Shapiro-Ilan D.I. Nematodes as Biocontrol Agents 2005 Cabi Publishing Wallingford UK 79 90
69 Ramakrishnan J. Salame L. Nasser A. Glazer I. Ment D. Survival and efficacy of entomopathogenic nematodes on exposed surfaces Sci. Rep. 12 2022 4629 35301390
70 Kung S.-P. Gaugler R. Kaya H.K. Influence of soil pH and oxygen on persistence of Steinernema spp J. Nematol. 22 1990 440 19287743
71 Grewal P.S. Formulations of entomopathogenic nematodes for storage and application Nematol. Res. 28 1998 68 74
72 Del Valle E.E. Dolinski C. Barreto E.L. Souza R.M. Samuels R.I. Efficacy of Heterorhabditis baujardi LPP7 (Nematoda: Rhabditida) applied in Galleria mellonella (Lepidoptera: Pyralidae) insect cadavers to Conotrachelus psidii,(Coleoptera: Curculionidae) larvae Biocontrol Sci. Technol. 18 2008 33 41
73 Shapiro-Ilan D.I. Lewis E.E. Son Y. Tedders W.L. Superior efficacy observed in entomopathogenic nematodes applied in infected-host cadavers compared with application in aqueous suspension, J. Invertebr. Pathol 83 2003 270 272
74 Van Zyl C. Malan A. Cost-effective culturing of Galleria mellonella and Tenebrio molitor and entomopathogenic nematode production in various hosts Afr. Entomol. 23 2015 361 375
75 de Oliveira Monteiro C.M. da Silva Matos R. Araújo L.X. Campos R. Bittencourt V.R.E.P. Dolinski C. Furlong J. de Azevedo Prata M.C. Entomopathogenic nematodes in insect cadaver formulations for the control of Rhipicephalus microplus (Acari: ixodidae) Vet. Parasitol. 203 2014 310 317 24836639
76 Shapiro-Ilan D.I. Lewis E.E. Behle R.W. McGuire M.R. Formulation of entomopathogenic nematode-infected cadavers J. Invertebr. Pathol. 78 2001 17 23 11500089
77 Guo S. Zhang S. Fang X. Liu Q. Gao J. Bilal M. Wang Y. Zhang X. Regulation of antimicrobial activity and xenocoumacins biosynthesis by pH in Xenorhabdus nematophila Microb. Cell Fact. 16 2017 1 14 28049473
78 Leite L.G. Shapiro-Ilan D.I. Hazir S. Survival of Steinernema feltiae in different formulation substrates: improved longevity in a mixture of gel and vermiculite Biol. Control 126 2018 192 197
79 Kaya H.K. Nelsen C. Encapsulation of steinernematid and heterorhabditid nematodes with calcium alginate: a new approach for insect control and other applications Environ. Entomol. 14 1985 572 574
80 Bedding R. Clark S. Lacey M. Butler K. Method and Apparatus for the Storage of Entomopathogenic Nematodes WIPO Patent No, WO 00/18887 2000
81 Georgis R. Formulation and application technology Gaugler R. Entomopathogenic Nematodes in Biological Control 1990 173 191
82 Chang F. Gehret M. Insecticide delivery system and attractant WIPO Patent No WO 91 1991 01736
83 Chang F. Gehret M. Stabilizer insect nematode compositions WIPO Patent No. WO WIPO Patent No WO 92 1992 10170
84 Hussein M.A. Abdel-Aty M. Formulation of two native entomopathogenic nematodes at room temperature J. Biopestic. 5 2012 23 27
85 Navon A. Keren S. Salame L. Glazer I. An edible-to-insects calcium alginate gel as a carrier for entomopathogenic nematodes Biocontrol Sci. Technol. 8 1998 429 437
86 Hiltpold I. Hibbard B.E. French B.W. Turlings T.C. Capsules containing entomopathogenic nematodes as a Trojan horse approach to control the western corn rootworm Plant Soil 358 2012 11 25
87 Kim J. Jaffuel G. Turlings T.C. Enhanced alginate capsule properties as a formulation of entomopathogenic nematodes BioControl 60 2015 527 535
88 Kagimu N. Malan A.P. Formulation of South African entomopathogenic nematodes using alginate beads and diatomaceous earth BioControl 64 2019 413 422
89 Bedding R.A. Storage of entomopathogenic nematodes Int. patent No. WO. 88 1988 (Google Patents)
90 Jansson R.K. Lecrone S.H. Gaugler R. Field efficacy and persistence of entomopathogenic nematodes (Rhabditida: Steinernematidae, Heterorhabditidae) for control of sweetpotato weevil (Coleoptera: apionidae) in southern Florida J. Econ. Entomol. 86 1993 1055 1063
91 Shapiro D.I. Lewis E.E. Comparison of entomopathogenic nematode infectivity from infected hosts versus aqueous suspension Environ. Entomol. 28 1999 907 911
92 Ansari M.A. Hussain M.A. Moens M. Formulation and application of entomopathogenic nematode‐infected cadavers for control of Hoplia philanthus in turfgrass Pest Manag. Sci. 65 2009 367 374 19165730
93 Lacey L.A. Shapiro-Ilan D.I. Glenn G.M. Post-application of anti-desiccant agents improves efficacy of entomopathogenic nematodes in formulated host cadavers or aqueous suspension against diapausing codling moth larvae (Lepidoptera: tortricidae) Biocontrol Sci. Technol. 20 2010 909 921
94 Capinera J. Hibbard B. Bait formulations of chemical and microbial insecticides for suppression of crop-feeding grasshoppers J. Agric. Entomol. 4 1987 337 344
95 Connick W. Jr. Nickle W. Vinyard B. Pesta“: new granular formulations for Steinernema carpocapsae J. Nematol. 25 1993 198 19279759
96 Connick W. Jr. Nickle W. Williams K. Vinyard B. Granular formulations of Steinernema carpocapsae (strain All)(Nematoda: Rhabditida) with improved shelf life J. Nematol. 26 1994 352 19279903
97 Yukawa T. Pitt J.M. Nematode Storage and Transport 1985 Google Patents
98 Silver S. Dunlop D. Grove D. WIPO Patent No. WO 95/0577 1995 World Intellectual Property Organization Geneva
99 Shapiro-Ilan D. Dolinski C. Entomopathogenic nematode application technology Campos-Herrera R. Nematode Pathogenesis of Insects and Other Pests, Ecology and Applied Technologies for Sustainable Plant and Crop Protection 2015 231 254
100 Nagesh M. Askary T. Balachander Manohar B.M. Arakalagud S. Rajan R. Strategies for making entomopathogenic nematodes cost-effective biocontrol agents Abd-Elgawad M.M. Askary T.H. Coupland J. Biocontrol Agents: Entomopathogenic and Slug Parasitic Nematodes 2017 CABI Wallingford UK 596 619
101 Kagimu N. Ferreira T. Malan A. The attributes of survival in the formulation of entomopathogenic nematodes utilised as insect biocontrol agents Afr. Entomol. 25 2017 275 291
102 Cortés-Martínez C. Ruiz-Vega J. Matadamas-Ortiz P. Lewis E. Aquino-Bolanos T. Navarro-Antonio J. Effect of moisture evaporation from diatomaceous earth pellets on storage stability of Steinernema glaseri Biocontrol Sci. Technol. 26 2016 305 319
103 Shapiro-Ilan D.I. Gouge D.H. Piggott S.J. Fife J.P. Application technology and environmental considerations for use of entomopathogenic nematodes in biological control Biol. Control 38 2006 124 133
104 Shapiro-Ilan D. Dolinski C. Entomopathogenic nematode application technology Campos-Herrera R. Nematode Pathogenesis of Insects and Other Pests 2015 Springer Cham 231 254
105 De Doucet M. Doucet M. Steinernema ritteri* N. Sp.(Nematoda: Steinernematidae) with a key to the species of the genus Nematologica 36 1990 257 265
106 De Doucet M. Bertolotti M. Giayetto A. Miranda M. Host range, specificity, and virulence of Steinernema feltiae, Steinernema rarum, and Heterorhabditis bacteriophora (Steinernematidae and Heterorhabditidae) from Argentina J. Invertebr. Pathol. 73 1999 237 242 10222175
107 Poinar G.O. Jr. Description and biology of a new insect parasitic Rhabditoid Heterorhabditis bacteriophora N. Gen., N. Sp.(Rhabditida; Heterorhabditidae N. Fam.) Nematologica 21 1975 463 470
108 Andaló V. Nguyen K.B. Moino A. Jr. Heterorhabditis amazonensis n. sp.(Rhabditida: Heterorhabditidae) from Amazonas, Brazil Nematology 8 2006 853 867
109 Nguyen K.B. Ginarte C.M.A. Leite L.G. dos Santos J.M. Harakava R. Steinernema brazilense n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from Mato Grosso, Brazil J. Invertebr. Pathol. 103 2010 8 20 19772861
110 Godjo A. Afouda L. Baimey H. Couvreur M. Zadji L. Houssou G. Bert W. Willems A. Decraemer W. Steinernema kandii n. sp. (Rhabditida: Steinernematidae), a new entomopathogenic nematode from northern Benin Nematology 21 2019 107 128
111 Kanga F.N. Trinh P.Q. Waeyenberge L. Spiridonov S.E. Hauser S. Moens M. Two new species of Steinernema Travassos, 1927 from the humid forest of southern Cameroon Russ. J. Nematol. 20 2012 15 36
112 Edgington S. Buddie A. Tymo L. France A. Merino L. Hunt D. Steinernema unicornum sp. n.(Panagrolaimomorpha: Steinernematidae), a new entomopathogenic nematode from Tierra del Fuego, Chile J. Nematode Morphol. Syst. 12 2009 113 131
113 Edgington S. Buddie A.G. Tymo L. Hunt D.J. Nguyen K.B. France A.I. Merino L.M. Moore D. Steinernema australe n. sp.(Panagrolaimomorpha: Steinernematidae), a new entomopathogenic nematode from Isla Magdalena, Chile Nematology 11 2009 699 717
114 Edgington S. Buddie A. Moore D. France A. Merino L. Hunt D. Heterorhabditis atacamensis n. sp.(Nematoda: Heterorhabditidae), a new entomopathogenic nematode from the Atacama Desert, Chile J. Helminthol. 85 2011 381 394 21087534
115 Zai-fu X. Guo-han W. Xiao-feng L. A new species of the genus Steinernema (Rhabditida: Steinernematidae) Zool. Res. 12 1991 17 20
116 Shen C. Wang G. Description of an entomopathogenic nematode, Steinernema longicaudum sp. nov. and its application Proceedings of the XIX International Congress of Entomology 1992 Beijing China 220 231
117 Jian H. Reid A. Hunt D. Steinernema ceratophorum n. sp.(Nematoda: Steinernematidae), a new entomopathogenic nematode from north-east China Syst. Parasitol. 37 1997 115 125
118 Qiu L. Fang Y. Zhou Y. Pang Y. Nguyen K.B. Steinernema guangdongense sp. n.(Nematoda: Steinernematidae), a new entomopathogenic nematode from southern China with a note on S. serratum (nomen nudum) Zootaxa 704 2004 1 20
119 Qiu L. Hu X. Zhou Y. Mei S. Nguyen K. Pang Y. Steinernema akhursti sp. n.(Nematoda: Steinernematidae) from Yunnan, China J. Invertebr. Pathol. 90 2005 151 160 16289544
120 Qiu L. Yan X. Zhou Y. Nguyen K. Pang Y. Steinernema aciari sp. n.(Nematoda: Steinernematidae), a new entomopathogenic nematode from Guangdong, China J. Invertebr. Pathol. 88 2005 58 69 15707870
121 Chen S. Li X. Yan A. Spiridonov S.E. Moens M. A new entomopathogenic nematode, Steinernema hebeiense sp. n.(Rhabditida: Steinernematidae), from North China Nematology 8 2006 563 574
122 Mrácek Z. Nguyen K.B. Tailliez P. Boemare N.l. Chen S. Steinernema sichuanense n. sp.(Rhabditida, Steinernematidae), a new species of entomopathogenic nematode from the province of Sichuan, east Tibetan Mts China, J. Invertebr. Pathol. 93 2006 157 169 16934830
123 Nguyen K.B. Qiu L. Zhou Y. Pang Y. Steinernema leizhouense sp. n.(Nematoda: Steinernematidae), a new entomopathogenic nematode from southern China Russ. J. Nematol. 14 2006 101 118
124 Nguyen K.B. Půža V. Mráček Z. Steinernema cholashanense n. sp.(Rhabditida, Steinernematidae) a new species of entomopathogenic nematode from the province of Sichuan, Chola Shan Mountains, China J. Invertebr. Pathol. 97 2008 251 264 17983627
125 Mráček Z. Qi-Zhi L. Nguyen K.B. Steinernema xueshanense n. sp.(Rhabditida, Steinernematidae), a new species of entomopathogenic nematode from the province of Yunnan, southeast Tibetan Mts China, J. Invertebr. Pathol. 102 2009 69 78 19460386
126 Qiu L. Zhao J. Wu Z. Lv Z. Pang Y. Steinernema pui sp. n.(Rhabditida, Steinernematidae), a new entomopathogenic nematode from Yunnan, China Zootaxa 2767 2011 1 13
127 Ma J. Chen S. Li X. Han R. Khatri-Chhetri H.B. De Clercq P. Moens M. A new entomopathogenic nematode, Steinernema tielingense n. sp.(Rhabditida: Steinernematidae), from north China Nematology 14 2012 321 338
128 Ma J. Chen S. De Clercq P. Moens M. Han R. Steinernema changbaiense sp. n.(Rhabditida: Steinernematidae), a new species of entomopathogenic nematodes from Northeast China Russ. J. Nematol. 20 2012
129 Xing-Yue L. Qi-Zhi L. Nermuť J. Půža V. Mráček Z. Heterorhabditis beicherriana n. sp.(Nematoda: Heterorhabditidae), a new entomopathogenic nematode from the Shunyi district of Beijing, China Zootaxa 3569 2012 25 40
130 López-Núñez J.C. Plichta K. Góngora-Botero C.E. Stock S.P. A new entomopathogenic nematode, Steinernema colombiense n. sp.(Nematoda: Steinernematidae), from Colombia Nematology 10 2008 561 574
131 Uribe-Lorío L. Mora M. Stock S.P. Steinernema costaricense n. sp. and S. puntauvense n. sp.(Rhabditida: Steinernematidae), two new entomopathogenic nematodes from Costa Rica Sys. Parasitol. 68 2007 167 182
132 Mráček Z. Hernández E.A. Boëmare N.E. Steinernema cubana sp. n.(Nematoda: Rhabditida: Steinernematidae) and the preliminary characterization of its associated bacterium J. Invertebr. Pathol. 64 1994 123 129
133 Mráček Z. Půža V. Nermut J. Steinernema poinari sp. n.(Nematoda: Steinernematidae) a new entomopathogenic nematode from the Czech Republic Zootaxa 3760 2014 336 350 24870085
134 Mráček Z. Bečvář S. Kindlmann P. Jersáková J. Habitat preference for entomopathogenic nematodes, their insect hosts and new faunistic records for the Czech Republic Biol. Control 34 2005 27 37
135 Weiser J. Neoaplectana carpocapsae n. sp.(Anguillulata, Steinernematinae), novy cizopasník housenek obalece jablecného, Carpocapsa pomonella L Vestnik Ceskoslovenske Spolecnosti Zoologicke 19 1955 44 52
136 Bovien P. Some types of association between nematodes and insects Vidensk. Meddelelser fra Dan. Naturhistorisk Foren. 101 1937 1 114
137 Wouts W. Mráček Z. Gerdin S. Bedding R. Steiner Neoaplectana 1929 a junior synonym of Steinernema Travassos 1927 (Nematoda; Rhabditida), Sys. Parasitol. 4 1982 147 154
138 Shamseldean M. Abou El-Sooud A. Abd-Elgawadoo M. Saleh M. Identification of a new Heterorhabditis species from Egypt, Heterorhabditis taysearae n. sp. (Rhabditida: Heterorhabditidae). Egypt J. Biol. Pest Co. 6 1996 129 138
139 Nguyen K.B. Tesfamariam M. Gozel U. Gaugler R. Adams B.J. Steinernemayirgalemense n. sp.(Rhabditida: Steinernematidae) from Ethiopia Nematology 6 2004 839 856
140 Tamiru T. Waeyenberge L. Hailu T. Ehlers R.-U. Půža V. Mráček Z. Steinernemaethiopiense sp. n.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from Ethiopia Nematology 14 2012 741 757
141 Lee M.-M. Sicard M. Skeie M. Stock S.P. Steinernema boemarei n. sp.(Nematoda: Steinernematidae), a new entomopathogenic nematode from southern France, Sys Parasitol. 72 2009 127 141
142 Steiner G. Aplectana kraussëi n. sp., eine in der Blattwespe Lyda sp. parasitierende Nematodenform, nebst. Bemerkungen über das Seitenorgan der parasitischen Nematoden G. Fischer 1923
143 Sturhan D. Spiridonov S. Mráček Z. Steinernema silvaticum sp. n.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from Europe Nematology 7 2005 227 241
144 Spiridonov S.E. Waeyenberge L. Moens M. Steinernema schliemanni sp. n.(Steinernematidae; Rhabditida)–a new species of steinernematids of the ‘monticolum’group from Europe Russ. J. Nematol. 18 2010 175 190
145 Stock S.P. Griffin C.T. Chaerani R. Morphological and molecular characterisation of Steinernema hermaphroditum n. sp.(Nematoda: Steinernematidae), an entomopathogenic nematode from Indonesia, and its phylogenetic relationships with other members of the genus Nematology 6 2004 401 412
146 Nikdel M. Niknam G. Morphological and molecular characterization of a new isolate of entomopathogenic nematode Steinernema feltiae (Filipjev) (Rhabditida: Steinernematidae) from the Arasbaran forests Iran, J. Asia-Pac. Biodivers. 8 2015 144 151
147 Triggiani O. Mrácek Z. Reid A. Steinernema apuliae sp. n.(Rhabditida: Steinernematidae): a new entomopathogenic nematode from southern Italy Zootaxa 460 2004 1 12
148 Tarasco E. Mráček Z. Nguyen K.B. Triggiani O. Steinernema ichnusae sp. n.(Nematoda: Steinernematidae) a new entomopathogenic nematode from Sardinia Island (Italy) J. Invertebr. Pathol. 99 2008 173 185 18556017
149 Clausi M. Longo A. Rappazzo G. Tarasco E. Vinciguerra M.T. Steinernema vulcanicum n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode species from Sicily (Italy) Nematology 13 2011 409 423
150 Poinar G. Jr. Karunakar G. David H. Heterorhabditis indicus n. sp.(Rhabditida: nematoda) from India: separation of Heterorhabditis spp. by infective juveniles, Fundam Appl. Nematol. 15 1992 467 472
151 Bhat A.H. Machado R.A. Abolafia J. Ruiz-Cuenca A.N. Askary T.H. Ameen F. Dass W.M. Taxonomic and molecular characterization of a new entomopathogenic nematode species, Heterorhabditis casmirica n. sp., and whole genome sequencing of its associated bacterial symbiont Parasit. Vectors 16 2023 383 37880744
152 Soni S. Patil J. Linga V. Mhatre P. Gowda M. Ganguli J. Půža V. Steinernema shori n. sp., a new entomopathogenic nematode (Nematoda: Steinernematidae) from India J. Helminthol. 97 2023 e72 37681419
153 Bhat A.H. Machado R.A. Abolafia J. Askary T.H. Půža V. Ruiz-Cuenca A.N. Rana A. Sayed S. Al-Shuraym L.A. Multigene sequence-based and phenotypic characterization reveals the occurrence of a novel entomopathogenic nematode species, n. sp J. Nematol. 55 2023 1 31
154 Stock S.P. Griffin C.T. Burnell A.M. Morphological characterisation of three isolates of Heterorhabditis Poinar, 1976 from theIrish group’(Nematoda: Rhabditida: Heterorhabditidae) and additional evidence supporting their recognition as a distinct species, H. downesi n. sp, Sys Parasitol. 51 2002 95 106
155 Yoshida M. Reid A.P. Briscoe B.R. Hominick W.M. Survey of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) in Japan, fundam Appl. Nematol. 21 1998 185 198
156 Yoshida M. Steinernema litorale n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from Japan Nematology 6 2004 819 838
157 Phan L.K. Takemoto S. Futai K. Steinernema ashiuense sp. n.(Nematoda: Steinernematidae), a new entomopathogenic nematode from Japan Nematology 8 2006 681 690
158 Choo H.Y. Kaya H.K. Stock S.P. Isolation of entomopathogenic nematodes (Steinernematidae and Heterorhabditidae) from korea, nematol Res. 25 1995 44 51
159 Waturu C. Hunt D. Reid A. Steinernema karii sp. n.(Nematoda: Steinernematidae), a new entomopathogenic nematode from Kenya Int. J. Nematol. 7 1997 68 75
160 Nguyen K. Sharpiro-Ilan D. Stuart R. McCoy C. James R. Adams B. Heterorhabditis mexicana n. sp.(Rhabditida: Heterorhabditidae) from Tamaulipas, Mexico, and morphological studies of the bursa of Heterorhabditis spp Nematology 6 2004 231 244
161 Grifaldo-Alcantara P.F. Alatorre-Rosas R. Segura-León O. Hernandez-Rosas F. Steinernema ralatorei n. sp. 1 isolated from sugarcane areas at veracruz, Mexico, southwest Entomol 42 2017 171 190
162 Machado R.A. Bhat A.H. Abolafia J. Muller A. Bruno P. Fallet P. Arce C.C. Turlings T.C. Bernal J.S. Kajuga J. Multi-locus phylogenetic analyses uncover species boundaries and reveal the occurrence of two new entomopathogenic nematode species, Heterorhabditis ruandica n. sp. and Heterorhabditis zacatecana n. sp J. Nematol. 53 2021 1 42 34179819
163 Khatri-Chhetri H.B. Waeyenberge L. Spiridonov S. Manandhar H.K. Moens M. Steinernema lamjungense n. sp.(Rhabditida: Steinernematidae), a new species of entomopathogenic nematode from Lamjung district, Nepal Nematology 13 2011 589 605
164 Khatri-Chhetri H.B. Waeyenberge L. Spiridonov S. Manandhar H.K. Moens M. Two new species of Steinernema Travassos, 1927 with short infective juveniles from Nepal Russ. J. Nematol. 19 2011 53 74
165 Poinar G.O. Jr. Taxonomy and biology of Steinernematidae and Heterorhabditidae Gaugler R. Entomopathogenic Nematodes in Biological Control vol. 54 1990 CRC Press
166 Shahina F. Anis M. Reid A. Rowe J. Maqbool M. Steinernema pakistanense sp. n.(Rhabditida: Steinernematidae) from Pakistan Int. J. Nematol. 11 2001 124 133
167 Anis M. Shahina F. Reid A. Rowe J. Steinernema asiaticum sp. n.(Rhabditida: Steinernematidae) from Pakistan Int. J. Nematol. 12 2002 1 12
168 Fayyaz S. Yan X. Qiu L. Han R. Gulsher M. Khanum T.A. Javed S. A new entomopathogenic nematode, Steinernema bifurcatum n. sp.(Rhabditida: Steinernematidae) from Punjab, Pakistan Nematology 16 2014 821 836
169 Fayyaz S. Khanum T.A. Ali S. Solangi G.S. Gulsher M. Javed S. Steinernema balochiense n. sp.(Rhabditida: Steinernematidae) a new entomopathogenic nematode from Pakistan Zootaxa 3904 2015 387 402 25660789
170 Lis M. Sajnaga E. Skowronek M. Wiater A. Rachwał K. Kazimierczak W. Steinernema sandneri n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from Poland J. Nematol. 53 2021 1 24 34179819
171 Román J. Figueroa W. Steinernema puertoricensis n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from Puerto Rico J. Agric. University Puerto Rico 78 1994 167 175
172 Machado R.A. Bhat A.H. Abolafia J. Shokoohi E. Fallet P. Turlings T.C. Tarasco E. Půža V. Kajuga J. Yan X. Steinernema africanum n. sp.(Rhabditida, Steinernematidae), a new entomopathogenic nematode species isolated in the Republic of Rwanda J. Nematol. 54 2022 1 28
173 Půža V. Campos-Herrera R. Blanco-Pérez R. Jakubíková H. Vicente-Díez I. Nermuť J. Steinernema riojaense n. sp., a new entomopathogenic nematode (Nematoda: Steinernematidae) from Spain Nematology 22 2020 825 841
174 Malan A.P. Knoetze R. Moore S.D. Isolation and identification of entomopathogenic nematodes from citrus orchards in South Africa and their biocontrol potential against false codling moth J. Invertebr. Pathol. 108 2011 115 125 21839086
175 Çimen H. Lee M.-M. Hatting J. Hazir S. Stock S.P. Steinernema innovationi n. sp.(Panagrolaimomorpha: Steinernematidae), a new entomopathogenic nematode species from South Africa J. Helminthol. 89 2015 415 427 24698548
176 Cimen H. Půža V. Nermuť J. Hatting J. Ramakuwela T. Faktorova L. Hazir S. Steinernema beitlechemi n. sp., a new entomopathogenic nematode (Nematoda: Steinernematidae) from South Africa Nematology 18 2016 439 453
177 Cimen H. Půža V. NermuŤ J. Hatting J. Ramakuwela T. Hazir S. Steinernema biddulphi n. sp., a new entomopathogenic nematode (Nematoda: Steinernematidae) from South Africa J. Nematol. 48 2016 148 27765988
178 Abate B.A. Malan A.P. Tiedt L.R. Wingfield M.J. Slippers B. Hurley B.P. Steinernema fabii n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from South Africa Nematology 18 2016 235 255
179 Malan A. Knoetze R. Tiedt L. Steinernema jeffreyense n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from South Africa J. Helminthol. 90 2016 262 278 25758326
180 Hatting J. Stock S.P. Hazir S. Diversity and distribution of entomopathogenic nematodes (Steinernematidae, Heterorhabditidae) in South Africa J. Invertebr. Pathol. 102 2009 120 128 19615373
181 Steyn W.P. Knoetze R. Tiedt L.R. Malan A.P. Steinernema litchii n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from South Africa Nematology 19 2017 1157 1177
182 Malan A.P. Knoetze R. Tiedt L.R. Steinernema nguyeni n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from South Africa Nematology 18 2016 571 590
183 Nthenga I. Knoetze R. Berry S. Tiedt L.R. Malan A.P. Steinernema sacchari n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from South Africa Nematology 16 2014 475 494
184 Cimen H. Lee M.M. Hatting J. Hazir S. Stock S.P. Steinernema tophus sp. n.(Nematoda: Steinernematidae), a new entomopathogenic nematode from South Africa Zootaxa 3821 2014 337 353 24989748
185 Katumanyane A. Malan A.P. Tiedt L.R. Hurley B.P. Steinernema bertusi n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from South Africa Nematology 22 2020 343 360
186 Didiza L. Lephoto T.E. Gray V.M. Morphological and molecular phylogenetic description of Steinernema batswanae n. sp.(Rhabditida: Steinernematidae): a new species of an entomopathogenic nematode from South Africa Arch. Phytopathol. Plant Protect. 2021 1 14
187 Malan A.P. Nguyen K.B. De Waal J.Y. Tiedt L. Heterorhabditis safricana n. sp.(Rhabditida: Heterorhabditidae), a new entomopathogenic nematode from South Africa Nematology 10 2008 381 396
188 Malan A.P. Knoetze R. Tiedt L. Heterorhabditis noenieputensis n. sp.(Rhabditida: Heterorhabditidae), a new entomopathogenic nematode from South Africa J. Helminthol. 88 2014 139 151 23232041
189 Ferreira T. Van Reenen C.A. Endo A. Spröer C. Malan A.P. Dicks L.M. Description of Xenorhabdus khoisanae sp. nov., the symbiont of the entomopathogenic nematode Steinernema khoisanae Int. J. Syst. Evol. Microbiol. 63 2013 3220 3224 23456807
190 Tallosi B. Peters A. R-U E. Steinernema bicornutum sp. n.(Rhabditida: Steinernematidae) from Vojvodina, Yugoslavia Russ. J. Nematol. 3 1995 71 80
191 Elawad S. Ahmad W. Reid A. Steinernema abbasi Sp. n.(Nematoda: Steinernematidae) from the Sultanate of Oman, Fundam vol. 20 1997 Appl. Nematol. 435 442
192 Tseng C.-T. Hou R.F. Tang L.-C. Steinernema taiwanensis n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from Taiwan Zootaxa 4434 2018 466 480 30313174
193 Hazir S. Stock S.P. Keskin N. A new entomopathogenic nematode, Steinernema anatoliense n. sp.(Rhabditida: Steinernematidae), from Turkey, Sys Parasitol. 55 2003 211 220
194 Půža V. Nermut J. Mráček Z. Gengler S. Haukeland S. Steinernema pwaniensis n. sp., a new entomopathogenic nematode (Nematoda: Steinernematidae) from Tanzania J. Helminthol. 91 2017 20 34 26765335
195 Stock S. Steinernema siamkayai n. sp.(Rhabditida: Steinernematidae), an entomopathogenic nematode from Thailand, Sys Parasitol. 41 1998 105 113
196 Maneesakorn P. Grewal P. Chandrapatya A. Steinernema minutum sp. nov.(Rhabditida: Steinernematidae): a new entomopathogenic nematode from Thailand Int. J. Nematol. 20 2010 27 42
197 Steiner G. glaseri Neoaplectana ng, n. sp.(Oxyuridae), a new nemic parasite of the Japanese beetle (Popillia japonica Newm.), J. Washington Acad Sci. 19 1929 436 440
198 Cabanillas H. Poinar G. Jr. Raulston J. Steinernema riobravis n. sp.(Rhabditida: Steinernematidae) from Texas, Fundam Appl. Nematol. 17 1994 123 131
199 Stock S.P. Koppenhöfer A. Steinernema scarabaei n. sp.(Rhabditida: Steinernematidae), a natural pathogen of scarab beetle larvae (Coleoptera: scarabaeidae) from New Jersey, USA Nematology 5 2003 191 204
200 Nguyen K.B. Stuart R.J. Andalo V. Gozel U. Rogers M.E. Steinernema texanum n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from Texas, USA Nematology 9 2007 379 396
201 Poinar G.O. Jr. Neoaplectana intermedia n. sp.(Steinernematidae: nematoda) from South Carolina Rev. Nematol. 8 1985 321 327
202 Fischer-Le Saux M. Mauléon H. Constant P. Brunel B. Boemare N. PCR-ribotyping of Xenorhabdus and Photorhabdus isolates from the Caribbean region in relation to the taxonomy and geographic distribution of their nematode hosts Appl. Environ. Microbiol. 64 1998 4246 4254 9797272
203 Spiridonov S. Krasomil-Osterfeld K. Moens M. Steinernema jollieti sp n.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from the American midwest Russ. J. Nematol. 12 2004 85 95
204 Nguyen K.B. Shapiro-Ilan D.I. Mbata G.N. Heterorhabditis georgiana n. sp.(Rhabditida: Heterorhabditidae) from Georgia, USA Nematology 10 2008 433 448
205 Gorgadze O. Lortkhipanidze M. Ogier J.-C. Tailliez P. Burjanadze M. [i] Steinernema tbilisiensis sp. n.[/i](Nematoda: Steinernematidae)—a New Species of Entomopathogenic Nematode from Georgia J. Agric. Sci.Technol. 13 2015
206 Gorgadze O. Fanelli E. Lortkhipanidze M. Troccoli A. Burjanadze M. Tarasco E. De Luca F. Steinernema borjomiense n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from Georgia Nematology 20 2018 653 669
207 Poinar G.O. Jr. Jackson T. Klein M. Heterorhabditis megidis sp. n.(Heterorhabditidae: Rhabditida), parasitic in the Japanese beetle, Popillia japonica (Scarabaeidae: Coleoptera) Proceed. Helminthol. Soc. Washington 54 1987 53 59
208 Liu J. Berry R.E. Heterorhabditis marelatusn. sp.(Rhabditida: Heterorhabditidae) from Oregon J. Invertebr. Pathol. 67 1996 48 54
209 Nguyen K.B. Smart G.C. Jr. Steinernema neocurtillis n. sp.(Rhabditida: Steinernematiclae) and a Key to Species of the Genus Steinernema J. Nematol. 24 1992 463 19283024
210 Nguyen K.B. Duncan L.W. Steinernema diaprepesi n. sp.(Rhabditida: Steinernematidae), a parasite of the citrus root weevil Diaprepes abbreviatus (L)(Coleoptera: Curculionidae) J. Nematol. 34 2002 159 19265926
211 Nguyen K.B. Gozel U. K_Ppenh_Fer H.S. Adams B.J. Heterorhabditis floridensis n. sp.(Rhabditida: Heterorhabditidae) from Florida Zootaxa 1177 2006 1–19-11–19
212 Nguyen K. Buss E. Steinernema phyllophagae n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from Florida, USA Nematology 13 2011 425 442
213 Stock S.P. Campos-Herrera R. El-Borai F. Duncan L. Steinernema khuongi n. sp.(Panagrolaimomorpha, Steinernematidae), a new entomopathogenic nematode species from Florida, USA J. Helminthol. 93 2019 226 241 29743130
214 Liu J. Berry R. Steinernema oregonensis n. sp.(Rhabditida: Steinernematidae) from Oregon, USA, Fundam Appl. Nematol. 19 1996 375 380
215 Nguyen K. Smart G. Jr. Steinernema scapterisci n. sp.(Rhabditida: Steinernematidae) J. Nematol. 22 1990 187 19287709
216 Phan L.K. Subbotin S.A. Waeyenberge L. Moens M. new entomopathogenic nematode A. Steinernema robustispiculum n. sp.(rhabditida: Steinernematidae), from chumomray national park in vietnam Syst. Parasitol. 60 2005 23 32 15791398
217 Pham V.L. Nguyen K. Reid A. Spiridonov S. Steinernema tami sp. n.(Rhabditida: Steinernematidae) from Cat Tien forest, Vietnam Russ. J. Nematol. 8 2000 33 43
218 Phan K.L. Nguyen N. Moens M. Steinernema sangi sp n.(Rabditida: Steinernematidae) from Vietnam Russ. J. Nematol. 9 2001 1 7
219 Phan K.L. Nguyen N.C. Moens M. Steinernema loci sp. n. and Steinernema thanhi sp. n.(Rhabditida: Steinernematidae) from Vietnam Nematology 3 2001 503 514
220 Phan K.L. Spiridonov S.E. Subbotin S.A. Moens M. Four new species of Steinernema Travassos, 1928 with short infective juveniles from Vietnam Russ. J. Nematol. 14 2006 11
221 Phan K.L. Mráček Z. Půža V. Nermut J. Jarošová A. Steinernema huense sp. n., a new entomopathogenic nematode (Nematoda: Steinernematidae) from Vietnam Nematology 16 2014 761 775
222 Phan K.L. Subbotin S.A. Nguyen N.C. Moens M. Heterorhabditis baujardi sp. n.(Rhabditida: Heterorhabditidae) from Vietnam and morphometric data for H. indica populations Nematology 5 2003 367 382
223 San-Blas E. Portillo E. Nermuť J. Půža V. Morales-Montero P. Steinernemapapillatum n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from Venezuela Nematology 17 2015 1081 1097
224 San-Blas E. Morales-Montero P. Portillo E. Nermuť J. Puza V. Steinernema goweni n. sp.(Rhabditida: Steinernematidae), a new entomopathogenic nematode from Zulia State, Venezuela Zootaxa 4067 2016 200 214 27395871
225 Parkman J. Smart G. Jr. Entomopathogenic nematodes, a case study: introduction of Steinernema scapterisci in Florida Biocontrol Sci. Technol. 6 1996 413 420
226 Kakouli‐Duarte T. Labuschagne L. Hague N.M. Biological control of the black vine weevil, Otiorhynchus sulcatus (Coleoptera: Curculionidae) with entomopathogenic nematodes (Nematoda: Rhabditida) Ann. Appl. Biol. 131 1997 11 27
227 Hussaini S. Singh S. Parthasarathy R. Shakeela V. Virulence of native entornopathogenic nematodes against black cutworms, Agrotis ipsilon (Hufnagel) and A. segtetum (Noctuidae: Lepidoptera) Indian J. Nematol. 30 2000 103 105
228 Park S.H. Yu Y.S. Park J.S. Choo H.Y. Bae S.D. Nam M.H. Biological control of tobacco cutworm, Spodoptera litura Fabricius with entomopathogenic nematodes Biotechnol. Bioproc. Eng. 6 2001 139 143
229 Vyas R. Patel N. Yadav P. Ghelani Y. Patel D. Performance of entomopathogenic nematodes for management of gram pod borer Helicoverpa armigera Ann. Plant Protec. Sci. 11 2003 107 109
230 Koppenhöfer A.M. Fuzy E.M. Steinernema scarabaei for the control of white grubs Biol. Control 28 2003 47 59
231 Mathasoliya J. Maghodia A. Vyas R. Efficacy of Steinernema riobrave against Agrotis ipsilon hufnagel (Lepidoptera: noctuidae) on potato Indian J. Nematol. 34 2004 177 179
232 Shapiro-Ilan D.I. Stuart R.J. McCoy C.W. Characterization of biological control traits in the entomopathogenic nematode Heterorhabditis mexicana (MX4 strain) Biol. Control 32 2005 97 103
233 Shakeela V. Hussaini S. Susceptibility of tobacco cutworm, Spodptera litura (F), to some indigenous isolates of entomopathogenic nematodes J. Ecofriendly Agric. 1 2006 64 67
234 Cottrell T.E. Shapiro-Ilan D.I. Susceptibility of the peach tree borer, Synanthedon exitiosa, to Steinernema carpocapsae and Steinernema riobrave in laboratory and field trials J. Invertebr. Pathol. 92 2006 85 88 16707138
235 Banu J.G. Jothi B.D. Narkhedkar N. Susceptibility of different stages of cotton bollworm, Helicoverpa armigera (Lepidoptera: noctuidae) to entomopathogenic nematodes Int. J. Nematol. 17 2007 41
236 Haukeland S. Lola-Luz T. Efficacy of the entomopathogenic nematodes Steinernema kraussei and Heterorhabditis megidis against the black vine weevil Otiorhynchus sulcatus in open field-grown strawberry plants Agric. Forest Entomol. 12 2010 363 369
237 Kvinikhidze G. Lortkipanidze G.K.M. Kokhia M. Histological Study of the Effect of Nematode-Bacteriumi Complex on the Structure and Vital, Activity of Cabbage Moth vol. 8 2010 80 85
238 De Waal J.Y. Malan A.P. Addison M.F. Efficacy of entomopathogenic nematodes (Rhabditida: Heterorhabditidae and Steinernematidae) against codling moth, Cydia pomonella (Lepidoptera: tortricidae) in temperate regions Biocontrol Sci. Technol. 21 2011 1161 1176
239 Cottrell T. Shapiro-Ilan D. Horton D. Mizell R. III Laboratory virulence and orchard efficacy of entomopathogenic nematodes against the lesser peach tree borer (Lepidoptera: sesiidae) J. Econ. Entomol. 104 2011 47 53 21404838
240 Prasad C. Hussain M.A. Pal R. Prasad M. Virulence of nematode Heterorhabditis indica (Meerut strain) against lepidopteran and coleopteran pests Vegetos 25 2012 343 351
241 Kary N.E. Golizadeh A. Dastjerdi H.R. Mohammadi D. Afghahi S. Omrani M. Morshedloo M. Shirzad A. A laboratory study of susceptibility of Helicoverpaarmigera (Hübner) to three species of entomopathogenic nematodes Munis Entomol. Zool. 7 2012 372 379
242 Shapiro-Ilan D.I. Mizell R.F. Laboratory virulence of entomopathogenic nematodes to two ornamental plant pests, Corythucha ciliata (Hemiptera: tingidae) and Stethobaris nemesis (Coleoptera: Curculionidae) Fla. Entomol. 95 2012 922 927
243 Mantoo M. Zaki F. Waliullah M. Virulence of kashmir isolate of EPN Heterorhabditis bacteriophora (nematoda: Heterorhabditidae) against black cutworm (Agrotis ipsilon) SKUAST J. Res. 14 2012 67 72
244 Gokce C. Yilmaz H. Erbas Z. Demirbag Z. Demir I. First record of Steinernema kraussei (Rhabditida: Steinernematidae) from Turkey and its virulence against Agrotissegetum (Lepidoptera: noctuidae) J. Nematol. 45 2013 253 24379483
245 Holajjer P. Patil J.B. Harish G. Nataraja M. Savaliya S. Evaluation of entomopathogenic nematodes, Steinernema carpocapsae and Heterorhabditis indica for their virulence against Spodoptera litura Ann. Plant Protec. Sci. 22 2014 163 165
246 Beck B. Brusselman E. Nuyttens D. Moens M. Temmerman F. Pollet S. Van Weyenberg S. Spanoghe P. Improving the biocontrol potential of entomopathogenic nematodes against Mamestra brassicae: effect of spray application technique, adjuvants and an attractant Pest Manag. Sci. 70 2014 103 112 23512412
247 Tofangsazi N. Cherry R. Arthurs S. Efficacy of commercial formulations of entomopathogenic nematodes against tropical sod webworm, Herpetogrammaphaeopteralis (Lepidoptera: crambidae) J. Appl. Entomol. 138 2014 656 661
248 Kashkouli M. Khajehali J. Poorjavad N. Impact of entomopathogenic nematodes on Thrips tabaci Lindeman (Thysanoptera: thripidae) life stages in the laboratory and under semi-field conditions J. Biopestic. 7 2014 77
249 Mantoo M.A. Zaki F. Biological control of cabbage butterfly, Pieris brassicae, by a locally isolated entomopathogenic nematode, Heterorhabditis bacteriophora SKUASTK-EPN-Hr-1 in Kashmir SKUAST J. Res. 16 2014 66 70
250 Meena L. Raju S. Efficacy of insecticides against fruit borer, Helicoverpa armigera (Hubner) on tomato Indian J. Entomol. 77 2015 201 202
251 Singh S. Yadav A.K. Vardhan S. Tripathi C. Diversity analysis of entomopathogenic nematodes against Helicoverpa armigera (Hübner) from Tarai region of IGP, India Curr. Life Sci. 1 2015 15 23
252 Goudarzi M. Moosavi M.R. Asadi R. Effects of entomopathogenic nematodes, Heterorhabditis bacteriophora (Poinar) and Steinernema carpocapsae (Weiser), in biological control of Agrotis segetum (Denis & Schiffermüller)(Lepidoptera: noctuidae), Turk J. Entomol. 39 2015 239 250
253 Hazir S. Shapiro-Ilan D.I. Hazir C. Leite L.G. Cakmak I. Olson D. Multifaceted effects of host plants on entomopathogenic nematodes J. Invertebr. Pathol. 135 2016 53 59 26896698
254 Kepenekci İ. Atay T. Alkan M. Biological control potential of Turkish entomopathogenic nematodes against the Colorado potato beetle, Leptinotarsadecemlineata Biocontrol Sci. Technol. 26 2016 141 144
255 Yadav S. Patil J. Sharma H. Bio-efficacy of Steinernema carpocapsae against Spodoptera litura under laboratory condition J. Pure Appl. Biosci. 5 2017 165 172
256 Vashisth S. Chandel Y. Chandel R. Biological control potential of himalayan strains of entomopathogenic nematodes against Lepidopteran insects infesting cole crops under screen house conditions, Biopestic Int. 12 2016 53 59
257 Chaubey A. Impact of soil dwelling entomopathogenic nematodes, recovered from the soil of Uttar Pradesh (India), on cotton bollworm Helicoverpa armigera (Hübner)(Lepidoptera: noctuidae) Appl. Biol. Res. 19 2017 63 70
258 Hussein M.A. El-Mahdi I.F. Efficiency of three formulated entomopathogenic nematodes against onion thrips, Thrips tabaci under aquaculture system J. Biopestic. 12 2019 134 138
259 Radhakrishnan S. Shanmugam S. Ramasamy R. Bio control efficacy of entomopathogenic nematodes against black cutworms, Agrotis ipsilon (Hufnagel)(Noctuidae: Lepidoptera) in potato Chem. Sci. Rev. Lett. 6 2017 219 224
260 Abdolmaleki A. Rafiee Dastjerdi H. Tanha Maafi Z. Naseri B. Virulence of two entomopathogenic nematodes through their interaction with Beauveria bassiana and Bacillus thuringiensis against Pieris brassicae (Lepidoptera: pieridae) J. Crop Protec. 6 2017 287 299
261 Safdar H. Javed N. Khan S.A. Arshad M. Reproduction potential of entomopathogenic nematodes on armyworm (Spodoptera litura) Pak. J. Zool. 50 2018 771 774
262 Khan B. Javed N. Khan S.A. Rajput N.A. Abbas H. Jabbar A. Walait M. Akash Z. Evaluation of entomopathogenic nematode Steinernema glaseri an effective biological entity against pink bollworm and armyworm in laboratory Int. J. Biol. Biotechnol. 15 2018 301 306
263 Yuksel E. Canhilal R. Evaluation of local isolates of entomopathogenic nematodes for the management of black cutworm, Agrotis ipsilon Hufnagel (Lepidoptera: noctuidae) Egypt. J. Biol. Pest Co. 28 2018 1 7
264 Reda E. Sallam A. Ibrahim H. Eid S. Efficacy of entomopathogenic nematode, Steinernema carpocapsae and its interaction with Beauveria bassiana against Pieris rapae L.(Lepidoptera: pieridae) J. Plant Protec. Pathol. 9 2018 795 798
265 Vashisth S. Chandel Y. Chandel R. Biological control potential of North West Himalayan strains of heterorhabditid nematodes against the turnip moth, Agrotis segetum (Denis & Schiffermuller)(Lepidoptera: noctuidae) Egypt. J. Biol. Pest Control 28 2018 1 8
266 James M. Malan A.P. Addison P. Surveying and screening South African entomopathogenic nematodes for the control of the Mediterranean fruit fly, Ceratitis capitata (Wiedemann) Crop Protec 105 2018 41 48
267 Foundation G.M. The World's Healthiest Food 2010 https://archive.org/stream/TheWorldsHealthiestFoods/TheWorldsHealthiestFoods_djvu.txt
268 Kamaliya R. Jethva D. Kachhadiya N. Bhut J. Ahir V. Bio-efficacy of entomopathogenic nematode Heterorhabditis indica against Spodoptera litura (Fabricius) J. Pharmacogn. Phytochem. 8 2019 1563 1567
269 Kumari N. Dc K. C V. Virulence of Steinernima and Heterorhabditis against black cutworm, Agrotis segetum (Lepidoptera: noctuidae) in potato crop Int. J. Curr. Microbiol. App. Sci. 8 2019 2404 2409
270 Vashisth S. Chandel Y. Chandel R. Comparative efficacy of indigenous heterorhabditid nematodes from north western Himalaya and Heterorhabditis indica (Poinar, Karunakar & David) against the larvae of Helicoverpa armigera (Hubner) Int. J. Pest Manage. 65 2019 16 22
271 Gokte-Narkhedkar N. Bhanare K. Nawkarkar P. Chilliveri P. Fand B.B. Kranthi S. Parasitic potential of entomopathogenic nematode Heterorhabditis indica against two Lepidopteran insect pests of cotton, Helicoverpa armigera (Hubner) and Spodoptera litura (Fabricious) Phytoparasitica 47 2019 31 41
272 Steyn L. Addison P. Malan A. Potential of South African entomopathogenic nematodes to control the leaf miner, Holocacista capensis (Lepidoptera: heliozelidae) South African J. Enol. Vitic. 40 2019 1-1
273 Dlamini T. Allsopp E. Malan A. Application of Steinernema yirgalemense to control Frankliniella occidentalis (Thysanoptera: thripidae) on blueberries Crop Protec. 128 2020 105016
274 Gulzar A. Mukhtar T. Wright D.J. Effects of entomopathogenic nematodes Steinernema carpocapsae and Heterorhabditis bacteriophora on the fitness of a Vip3A resistant subpopulation of Heliothis virescens (Noctuidae: Lepidoptera) Bragantia 79 2020 281 292
275 Acharya R. Yu Y.-S. Shim J.-K. Lee K.-Y. Virulence of four entomopathogenic nematodes against the tobacco cutworm Spodoptera litura Fabricius Biol. Control 150 2020 104348
276 Askary T.H. Ahmad M.J. Efficacy of entomopathogenic nematodes against the cabbage butterfly (Pieris brassicae (L.)(Lepidoptera: pieridae) infesting cabbage under field conditions Egypt. J. Biol. Pest Control 30 2020 1 7
277 Abbas W. Javed N. Haq I.U. Ahmed S. Pathogenicity of entomopathogenic nematodes against cabbage butterfly (Pieris brassicae) Linnaeus (Lepidoptera: pieridae) in laboratory conditions Int. J. Tropic. Insect Sci 41 2021 525 531
278 Kasi I.K. Singh M. Waiba K.M. Monika S. Waseem M. Archie D. Gilhotra H. Bio-efficacy of entomopathogenic nematodes, Steinernema feltiae and Heterorhabditis bacteriophora against the Cabbage butterfly (Pieris brassicae [L.]) under laboratory conditions Egypt. J. Biol. Pest Control 31 2021 1 7
279 Bhairavi K.S. Bhattacharyya B. Devi G. Bhagawati S. Das P.P.G. Devi E.B. Manpoong N.S. Evaluation of two native entomopathogenic nematodes against Odontotermes obesus (Rambur)(Isoptera: termitidae) and Agrotis ipsilon (hufnagel)(Lepidoptera: noctuidae) Egypt. J. Biol. Pest Control 31 2021 1 8
280 Devi G. Saikia M. Bhagawati S. Bhattacharyya B. Bio-efficacy of entomopathogenic nematode, Heterorhabditis bacteriophora against cutworm, Agrotis ipsilon damaging potato under field condition J. Entomol. Zool. Stud. 9 2021 323 325
281 Andaló V. Faria Lsd Carvalho F.J. Assis Gad Santos V. Sm M. Ahr G. Entomopathogenic nematodes for the control of Helicoverpa armigera (Hübner) (Lepidoptera: noctuidae) pupae Arq. Inst. Biol. 88 2021 1 8
282 Lalitha K. Venkatesan S. Balamuralikrishnan B. Shivakumar M.S. Isolation and biocontrol efficacy of entomopathogenic nematodes Steinernema carpocapsae, Steinernema monticolum and Rhabditis blumi on lepidopteran pest Spodoptera litura Biocatal. Agric. Biotechnol. 39 2022 102291
283 Thakur N. Tomar P. Kaur S. Kumari P. Virulence of native entomopathogenic nematodes against major lepidopteran insect species of tomato (Solanum lycopersicum L.) J. Appl. Biol. Biotechnol. 10 2022 6 14
284 Tomar P. Thakur N. Sharma A. Infectivity of entomopathogenic nematode against the cabbage butterfly (Pieris brassicae L.) in polyhouse and in field condition Egypt. J. Biol. Pest Control 32 2022 1 7
285 Thakur N. Tomar P. Sharma S. Kaur S. Sharma S. Yadav A.N. Hesham A.E.-L. Synergistic effect of entomopathogens against Spodoptera litura (Fabricius) under laboratory and greenhouse conditions Egypt. J. Biol. Pest Control 32 2022 1 10
286 Tomar P. Thakur N. Biocidal potential of indigenous isolates of Entomopathogenic Nematodes (EPNs) against tobacco cutworm, Spodoptera litura Fabricius (Lepidoptera: noctuidae) Egypt. J. Biol. Pest Control 32 2022 1 10
287 Tomar P. Thakur N. Yadav A.N. Indigenous entomopathogenic nematode as biocontrol agents for insect pest management in hilly regions Plant Sci. Today 8 2021 51 59
288 Rehman G. Mamoon-ur-Rashid M. Evaluation of entomopathogenic nematodes against red palm weevil, Rhynchophorus ferrugineus (Olivier)(Coleoptera: Curculionidae) Insects 13 2022 733 36005358
289 Shinde S.P. Ingole D. Biradar V. Gokte-Narkhedkar N. Lavhe N. Thube S.H. Shah V. Prasad Y. Efficacy of native strains of entomopathogenic nematode, Heterorhabditis indica against the fall armyworm, Spodoptera frugiperda (JE Smith)(Lepidoptera: noctuidae) from India Egypt. J. Biol. Pest Control 32 2022 1 13
290 Thakur N. Tomar P. Kaur J. Kaur S. Sharma A. Jhamta S. Yadav A.N. Dhaliwal H.S. Thakur R. Thakur S. Eco-friendly management of Spodoptera litura (Lepidoptera: noctuidae) in tomato under polyhouse and field conditions using Heterorhabditis bacteriophora Poinar, their associated bacteria (Photorhabdus luminescens), and Bacillus thuringiensis var. kurstaki Egypt. J. Biol. Pest Control 33 2023 7
291 Tomar P. Thakur N. Sidhu A.K. Laskar B.A. Hashem A. Avila-Quezada G.D. Abd_Allah E.F. The isolation, identification, and insecticidal activities of indigenous entomopathogenic nematodes (Steinernema carpocapsae) and their symbiotic bacteria (Xenorhabdus nematophila) against the larvae of Pieris brassicae Horticulturae 9 2023 874
292 Tomar P. An Inventory of Entomopathogenic Nematodes and Their Bacterial Symbionts Based on Molecular Characterization in Himachal Pradesh 2022 Eternal University, Himachal Pradesh India 1 419 PhD Thesis
