Amandeep Kaur
David Kihoro Sirengo
Pratibha Karki2
Thomas O. Powers
Amanda M. V. Brown- 1Department of Biological Sciences, Texas Tech University, Lubbock, TX, United States
- 2Department of Plant Pathology, University of Nebraska-Lincoln, Lincoln, NE, United States
Entomopathogenic nematodes (EPNs) of the genera Heterorhabditis and Steinernema are increasingly recognized as potent biological control agents due to their ability to infect and kill diverse insect pest taxa through a symbiotic partnership with insect-pathogenic bacteria. Over the last decades, substantial progress has been made in improving EPN field performance through advances in formulation and application methods, use of biodegradable polymers and nanocarriers, and elucidation of stress tolerance mechanisms. However, despite their proven efficacy, large-scale commercialization of EPNs remains limited by high production costs, formulation instability, and environmental constraints. While numerous reviews have separately addressed EPN biology, mass production, or field application independently, a critical and integrative synthesis linking molecular mechanisms, and formulation strategies remains lacking. This review synthesizes current understanding of EPN biology with emphasis on molecular mechanisms governing host localization, invasion, and immune suppression, as well as their biotic ecological interactions within soil environments. We also discuss advances in stress tolerance mechanisms, innovations in formulation, and outline future research priorities needed to develop ecologically resilient EPN-based biocontrol products. As agriculture shifts toward more regenerative and environmentally sustainable systems, a comprehensive understanding of EPN biology, full ecological potential of EPN-bacteria partnerships holds promise not only for effective pest suppression but also for advancing fundamental understanding of host-microbe interactions and ecosystem resilience.
1 Introduction
The global population is projected to increase significantly by 2050 (United Nations, 2017), warranting a substantial increase in crop productivity through sustainable intensification and the adoption of innovative pest management strategies to ensure global food security. In this context, as agriculture shifts toward ecologically informed approaches, biological control has emerged as a cornerstone of environmentally sound, sustainable pest management. This paradigm shift has paved the way for the exploitation of naturally occurring beneficial microorganisms as alternatives and complements to synthetic chemical pesticides. As a result, the adoption of entomopathogenic nematodes (EPNs) as biocontrol agents against major agricultural insect pests is increasing across diverse cropping systems worldwide (Modic et al., 2020; Fallet et al., 2024; Polat et al., 2024; Ramakuwela et al., 2025). The successes of EPNs as biological control agents have been well demonstrated in various cropping systems against different insect taxa. In particular, EPNs have been effective in controlling stem flea beetle (Psylliodes chrysocephala) in winter oilseed rape (Godina et al., 2023), Pumpkin fruit fly (Bactrocera tau) in Chinese cucumber (Liu et al., 2025), false codling moth (Thaumatotibia leucotreta) in citrus (Moore et al., 2024), and codling moth (Cydia pomonella) in apple (Odendaal et al., 2016) among others. Moreover, compatibility of EPN with synthetic insecticides has been evaluated primarily through invitro bioassays, revealing species- and formulation-specific responses (Krishnayyaand and Grewal, 2002; Srivastava et al., 2011; Sabino PH de et al., 2014; Mahmoud et al., 2016; Wu et al., 2017; Dias S da et al., 2024). However, chemical compatibility with formulation components and application strategies needs to be tested under realistic agroecosystem conditions.
Beyond their practical applications, EPNs also serve as model organisms for advancing fundamental research on microbial mutualism and ecological interactions (Campos-Herrera et al., 2012; Tarasco et al., 2023; Stock et al., 2025). Notably, among EPN taxa, the infective juveniles (IJs) of Heterorhabditis and Steinernema, which maintain obligate symbioses with gammaproteobacteria of the genera Photorhabdus and Xenorhabdus, respectively, are the most intensively studied due to their effectiveness in biocontrol and relevance in both research and commercial applications. Within this highly specialized partnership, the nematode provides physical protection for its bacterial symbiont against environmental stresses such as temperature changes, moisture loss, and harmful microbes, and then releases the symbiont into the insect hemocoel upon infection (Stilwell et al., 2018). Once inside the host, these symbionts produce secondary metabolites and various toxins (Eleftherianos et al., 2007; Hinchliffe et al., 2010; Tobias et al., 2017) that suppress immune defenses and kill the host through septicemia, thereby creating a conducive environment for the nematode’s feeding and reproduction in the cadaver, facilitating later reacquisition of the bacteria before nematodes emerge as new IJs (Tarasco et al., 2023; Glazer et al., 2025). The insecticidal, antimicrobial and immunomodulatory roles of these secondary metabolites have been comprehensively reviewed elsewhere (Bode, 2009; Stock et al., 2017; Liao et al., 2025). Their application, however, is limited by host dependence and unresolved toxicity concerns, underscoring the need for systematic evaluation of stability, safety, and delivery strategies. Traditionally, this partnership has been viewed as species-specific, with each nematode species maintaining a single coevolved symbiont. Experimental evidence shows that these partnerships are tightly governed by partner recognition and fidelity and that pairing nematodes with non-cognates reduces fitness, virulence, and bacterial carriage (McMullen et al., 2017). Nonetheless, recent studies revealed the coexistence of other bacterial species with the primary symbiont. These secondary or facultative associates may enhance infective juvenile (IJ) insecticidal activity and expand the nematode’s host range (Pervez et al., 2020; Maher et al., 2021; Sugiyama and Hasegawa, 2025).
In addition to causing insect mortality, application of EPN in soil has been shown to modulate plant antioxidant defenses by influencing the activities of key enzymes such as peroxidases and catalase (Jagdale et al., 2009). For instance, the chemical cues such as ascarosides and 2,3-butanediol released by EPN-symbiont complex trigger oxidative bursts in root tissues, leading to activation of salicylic acid and jasmonic acid signaling pathways (Jagdale et al., 2009; Manohar et al., 2020). These hormonal cascades stimulate the expression of defense-related genes like PR-1 (Helms et al., 2019; Wang et al., 2025) and enhance enzymatic scavenging of reactive oxygen species through increased peroxidase and catalase activity. These multitrophic interactions reflect the ecological versatility of EPNs and their growing importance in integrated pest management (IPM) frameworks. Their effectiveness can be further augmented through co-application with other biocontrol agents, such as entomopathogenic fungi, where synergistic interactions frequently result in enhanced pest suppression and increased crop productivity (Sáenz-Aponte et al., 2020; Půža and Tarasco, 2023). The remarkable successes of EPNs across diverse cropping systems demonstrate their potential as reliable biological control agents and alternatives to chemical pesticides. However, realizing their full potential requires overcoming challenges related to formulation, shelf life, field persistence, regulatory policies, and large-scale delivery systems (Blanco-Pérez et al., 2025). Several comprehensive reviews have examined EPNs with emphasis on EPN biology, mass production, application, field efficacy and challenges (Shapiro-Ilan and Dolinski, 2015; Koppenhöfer et al., 2020; Abd-Elgawad, 2023; Tarasco et al., 2023). However, most existing reviews treat EPN biology, symbiosis, formulation strategies as largely independent topics. This review aims to bridge this gap by evaluating how molecular mechanisms underlying host immune suppression, stress tolerance affects field performance and commercialization.
2 Molecular mechanisms of EPNs on insect infection
2.1 Host localization, foraging strategies, and invasion
Successful infection by EPNs relies on host localization mediated by chemical, physical, and other environmental cues (Figure 1A) (O’Halloran and Burnell, 2003; Ali et al., 2010; Hallem et al., 2011; Filgueiras et al., 2016; Laznik and Trdan, 2016; Baiocchi et al., 2017; Laznik et al., 2020; Zhang et al., 2021). Among the important chemical cues are the paired BAG sensory neurons that detect olfactory signals such as CO2, a byproduct of insect respiration. In Heterorhabditis bacteriophora, BAG neuron ablation abolishes chemotaxis toward the greater wax moth, Galleria mellonella, confirming the essential role of BAGs in host localization (Hallem et al., 2011). BAG neurons also contribute to nictation-triggering circuits in ambusher EPN species like Steinernema carpocapsae (Hallem et al., 2011). In addition to insect-derived signals, EPNs also respond to herbivore-induced plant volatiles (HIPVs) through chemosensory neurons (Rasmann et al., 2005; Degenhardt et al., 2009). Root-derived compounds such as carvone have been shown to activate S. carpocapsae, suggesting that plant volatiles can trigger specific chemosensory responses in these nematodes (Bai et al., 2024). Although the underlying neuronal circuits in EPNs remain poorly characterized, evidence from the model nematode, Caenorhabditis elegans, suggests that attraction and repulsion behaviors are mediated by complex interneuronal networks (Hart and Chao, 2010; Guillermin et al., 2017).
Figure 1. Schematic representation of (A) host-finding and (B) infection by entomopathogenic nematodes. Figure was created using BioRender.
While these sensory mechanisms drive host detection, the specific cues used for host localization can vary depending on the EPN’s foraging strategy. Cruiser EPN species (e.g., H. bacteriophora) track long-range chemical, vibrational, and electrical signals, while ambushers (e.g., S. carpocapsae) use nictation to intercept mobile hosts (Lortkipanidze et al., 2016; Grunseich et al., 2021). Electrical fields generated by insect movement or root injury can elicit species-specific directional responses (Rasmann et al., 2005; Grunseich et al., 2021). For instance, the cruiser Steinernema glaseri moves toward higher electrical potentials, while the ambusher S. carpocapsae is attracted to lower potentials (Shapiro-Ilan et al., 2009; Shapiro-Ilan et al., 2012). Seismic cues produced by insect movement help detect hosts, particularly when chemical gradients are disrupted in organic-rich soils (Torr et al., 2004).
Following localization, host invasion is a critical and often challenging step in the infection process. EPNs utilize multiple routes to enter their hosts, and penetration routes vary among insect species. The choice of penetration pathway is both species- and host-specific (Batalla-Carrera et al., 2014; Kallali et al., 2024). Typically, EPNs invade through natural openings such as the mouth, spiracles, or anus, but some species can also directly penetrate the insect cuticle (Dowds and Peters, 2002).
2.2 Overcoming the insect immune system
While penetration marks the beginning of an infection process, the survival of EPNs within their hosts depends on their ability to bypass host immunity through coordinated interactions with their symbiotic bacteria (Figure 1B) (Ciche and Ensign, 2003; ffrench-Constant et al., 2007). Upon invasion, IJs of H. bacteriophora use their anterior tooth to pierce host tissues while secreting proteolytic enzymes that degrade structural barriers. On the other hand, Steinernema species degrade midgut epithelia with enzymes before they can enter the hemocoel (Ciche and Ensign, 2003; Balasubramanian et al., 2010). Immediately after penetration into the hemocoel, the IJs shed their second-stage cuticle, exposing a dynamic surface coat that masks immunogenic epitopes and releases small immunomodulatory molecules (Wang and Gaugler, 1999), an early immune evasion strategy that suppresses host recognition and prepares the hemocoel for bacterial release and establishment.
Once in the hemocoel, IJs release effector molecules via their excretory-secretory glands, which target both humoral and cellular immunity. Among these effectors, some of the most important examples include proteolytic enzymes that suppress the prophenoloxidase (pro-PO) cascade, therefore blocking melanization and nodulation. Similarly, a Kunitz-type protease inhibitor (Sc-KU-4) disrupts hemocyte encapsulation and aggregation (Balasubramanian et al., 2010; Chang et al., 2019). As a result, transient immune paralysis is induced, halting the host defense response and enabling the establishment of IJs. This process creates favorable conditions for the subsequent release and proliferation of the EPN’s symbiotic bacteria within the host (Lu et al., 2017). Once released from the nematode gut, the symbionts reinforce and extend host suppression through multiple molecular pathways. For instance, the symbiotic bacterium Xenorhabdus nematophila inhibits Toll and IMD (immune deficiency) signaling by suppressing eicosanoid biosynthesis, thereby halting the production of antimicrobial peptides (AMPs) such as cecropins and lysozymes (Park and Kim, 2000; Hwang et al., 2013).
Moreover, extracellular proteases from Xenorhabdus degrade pre-existing AMPs, hence enhancing humoral suppression (ffrench-Constant et al., 2007; Balasubramanian et al., 2010). Similarly, the symbionts Photorhabdus luminescens and X. nematophila secrete rhabduscin. This phenoloxidase inhibitor suppresses melanization and Mcf-family toxins and induces death of hemocytes, the primary immune cells in the hemolymph of invertebrates responsible for defense against pathogens, foreign entities, and gut epithelial cells (Dowling et al., 2004; Crawford et al., 2012). Serralysin-type proteases such as PrtA further degrade immune-related proteins, thereby enhancing systemic infection (Felföldi et al., 2009). These events later lead to a collapse of humoral and cellular defenses, resulting in rapid septicemia in the target host (Forst et al., 1997; da Silva et al., 2020).
Interestingly, several studies have demonstrated the pathogenicity of axenic EPNs under laboratory conditions. For instance, Ehlers et al. (1997) investigated the pathogenicity of the Steinernema feltiae-Xenorhabdus bovienii complex to the marsh crane fly, Tipula oleracea, and G. mellonella larvae by injecting dauer IJs using monoxenic nematodes (with bacteria), axenic nematodes (bacteria-free), and the symbiotic bacteria alone. Axenic IJs were less pathogenic than monoxenic IJs in both insects, indicating that bacterial symbionts significantly enhance virulence. Similarly, when axenic cultures of G. mellonella larvae were treated with axenic IJs of H. bacteriophora and S. carpocapsae, all larvae were killed by S. carpocapsae, whereas H. bacteriophora failed to cause mortality despite a few nematodes being recovered from the hemocoel (Han and Ehlers, 2000). Although these results represent partial pathogenic success, the nematode-bacterium association remains essential for complete pathogenicity and successful reproduction (Bisch et al., 2015). These results support the potential of EPNs as candidate model systems for studying the molecular mechanisms of parasitism and insect immune defense.
3 Stress tolerance mechanisms and environmental persistence challenges
Despite advances in the biology, development, and deployment of EPNs, their integration into IPM programs is constrained by interrelated challenges ranging from formulation and application to environmental limitations. The following sections highlight the stress tolerance and environmental persistence mechanisms, and the challenges of formulation and application constraints that affect EPN-based biocontrol.
3.1 Stress tolerance and environmental persistence
EPNs encounter both abiotic and biotic stresses in the environment. Among these, extreme temperatures, desiccation, and UV radiation represent key abiotic challenges, while microbial antagonists constitute major biotic threats. The field efficacy of EPNs as biological control agents largely depends on their ability to withstand diverse biotic and abiotic stresses. The environmental persistence is further determined by stress tolerance mechanisms that may be species- or strain-specific as well as by interactions with soil type and the availability of suitable hosts. IJs acclimate to thermal stress by accumulating trehalose (Grewal and Jagdale, 2002; Jagdale and Grewal, 2003; Grewal et al., 2006) and by reducing metabolic activity (Brown and Gaugler, 1996; Andalo et al., 2011; Ali and Wharton, 2013). Although IJs possess mechanisms to lower their lipid reserves, species-specific differences exist in how metabolic adjustments occur. For instance, Heterorhabditis megidis reduces production of proteins involved in metabolism and protein synthesis, whereas H. carpocapsae downregulates proteins involved in intermediary metabolism and oxidative phosphorylation (Jagdale and Grewal, 2003; Andalo et al., 2011).
Under severe dehydration, IJs enter a state of anhydrobiosis, and osmoregulant molecules such as, heat-stable, water-stress-related protein (DESC47), and heat-shock protein (HSP40) are synthesized (Solomon et al., 2000; Gal et al., 2003; Somvanshi et al., 2008). In S. feltiae, desiccation induced a two-fold increase in trehalose content (Solomon et al., 1999). Furthermore, the induction of casein kinase (CK2) has been observed, which is hypothesized to participate in the transcriptional activation of nucleosome-assembly protein (NAP-1) (Gal et al., 2003). While UV tolerance mechanism in EPNs is unclear, the general assumption is that UV tolerance mechanisms parallel those described in C. elegans. In C. elegans, UV light is detected and avoided through four sensory neurons (ASJ, ASK, AWB, and ASH) with the neurotransmitter glutamate receptors (glc-3, mgl-1, mgl-2) expressed in ASH and ASK. These neurons play key roles in mediating this response (Bargmann, 2006; Ozawa et al., 2022).
In addition to species- and strain-specific physiological stress responses, ecological adaptation plays a critical role in determining EPN persistence and field efficacy. Locally adapted beneficial microbes exhibit superior tolerance to changing weather and climatic conditions, enhanced dispersal, and form a more stable association with the native soil microbiota (Campos-Herrera et al., 2013). Native EPNs have been shown to exhibit strong virulence against pests for major crops and turf grasses (Yadav and Lalramliana, 2012; Sun et al., 2021; Thakur et al., 2022; Tumialis et al., 2023; Fallet et al., 2024; Koppenhöfer and Luiza Sousa, 2024) and show greater potential when integrated with chemical control strategies (Shields et al., 2018; Tomar and Thakur, 2022). In addition, the EPN-symbiont complex produces a myriad of natural compounds that include scavenger deterrent factors (SDFs) with antifungal, insecticidal, and antibacterial properties. These metabolites not only deter scavengers from colonizing insect cadavers but also help the EPNs and their symbiotic bacteria overcome host immune defenses, ensuring successful infection and reproduction (Furgani et al., 2008; Gulcu et al., 2012; Fang et al., 2014; Hazir et al., 2016; Tobias et al., 2017; Gulcu et al., 2018).
3.2 Formulation and application
Successful commercialization of EPNs depends on their formulation and handling. However, one major obstacle limiting their field performance arises during formulation (Chen and Glazer, 2005). To increase EPN shelf life and efficacy, EPN treatments have been formulated using several carriers such as liquid concentrates, alginate gels, water-dispersible granules, synthetic sponge, and vermiculite applied in aqueous suspension through agricultural sprayers or irrigation systems (Georgis, 1990; Grewal, 2000) (Table 1). Traditionally, early work on water-dispersible granules showed enhanced improvements in EPN longevity and ease of handling compared with aqueous formulations (Grewal, 2000). Nonetheless, most aqueous products require refrigeration to maintain IJ viability, which results in increased production and transport costs and limits use in regions without reliable cold storage (Chen and Glazer, 2005; Karimi et al., 2025).
Table 1. Entomopathogenic nematode (Steinernema and Heterorhabditis spp.) formulation types, mechanisms governing efficacy and limitations, and directions for future research.
EPN-infected cadavers have been explored as natural carriers (Shapiro and Glazer, 1996; Shapiro-Ilan et al., 2001). The IJs emerging from infected hosts disperse efficiently through soil, are highly infective, and provide better pest control than nematodes applied in suspension (Shapiro and Lewis, 1999; Perez et al., 2003; Shapiro-Ilan et al., 2003). For instance, field trials using H. bacteriophora-infected cadavers effectively reduced infestations of the sweet potato weevil, Cylas formicarius, and the guava weevil, Conotrachelus psidii. The reliability of cadavers as a dispersal agent, however, is limited because they desiccate and rupture during production and storage. These challenges were addressed by formulating cadavers with protective materials that prevent sticking and mechanical damage (Shapiro-Ilan et al., 2003). However, the influence of cadaver age on stability and efficacy remains unclear.
Recent advancements concentrate on improving stability in ambient conditions. For example, Darsouei et al. (2025) demonstrated that several EPN species encapsulated in calcium-alginate beads survived up to three months at room temperature under laboratory conditions. Similarly, hydrogel and emulsion capsules were shown to improve persistence of S. feltiae against the yellow mealworm beetle, Tenebrio molitor, under laboratory and field conditions (Perier et al., 2025). Other studies have demonstrated that encapsulation in alginate gels and hydrophilic colloids also shields IJs from desiccation and UV stress (Kim et al., 2021). Additionally, carboxymethyl cellulose (CMC), a water-soluble cellulose derivative, has emerged as an effective substrate for EPN formulation, particularly for S. carpocapsae (Kary et al., 2021). Despite these advances, cost-effective commercialization remains a challenge (San-Blas, 2013; Ramakuwela et al., 2016). Even with successful formulations, effectiveness ultimately depends on proper handling and field application.
During application, IJs are introduced to stressors such as tank sedimentation (Schroer et al., 2005), nozzle shear or pressure damage (Moreira et al., 2013), and post-application desiccation mortality (Platt et al., 2018). Several adjuvants and surfactants have been formulated recently to improve efficiency. The addition of xanthan gum or potassium alginate resulted in a two-fold increase of insect mortality at 80% relative humidity (RH) and a five-fold increase at 60% RH, while the mixtures of 0.3% xanthan or alginate with 0.3% surfactants further improved efficacy for foliar application of S. carpocapsae against diamondback moth larvae (Schroer et al., 2005). Similarly, S. carpocapsae formulated in an alkyl polyglycoside polymeric surfactant reduced the tomato pinworm, Tuta absoluta, larvae populations within two weeks post-treatment (Kajuga et al., 2025). Nonetheless, the efficacies of these adjuvants or surfactants are not consistently reliable. Some, for instance, increase longevity and enhance host invasion, whereas others reduce infectivity (Beck et al., 2013).
4 Future directions and opportunities
Given the growing demand for safer food and environmentally sustainable pest management practices, the commercial potential of EPNs is expanding (Dara, 2019). However, unlike chemical pesticides, their commercialization is constrained by multiple factors (Koppenhöfer et al., 2020). Addressing these limitations requires a stronger focus on biological and ecological factors that influence EPNs’ performance. Systematic screening of native EPN isolates, coupled with formulation optimization could further enhance persistence while reducing reliance on protective encapsulants.
Beyond strain selection, interactions among EPNs, their bacterial symbionts, and soil microbial communities may influence EPN efficacy. Nevertheless, the underlying mechanisms remain poorly understood. Therefore, metagenomic profiling of soil microbial community in soils with contrasting EPN efficacies can identify microbial taxa that promote EPN success or suppress their activity and enable the design of synthetic microbial communities that can be co-inoculated with EPNs to prime the soil environment, ensuring higher success rate in non-native soils. On the other hand, limited efficacy or genetic diversity in natural strains can be addressed through genomic-assisted breeding and molecular genetic approaches, including mutagenesis, transgenesis, and targeted gene modification. Over the past decades, these approaches have been applied to enhance EPN stress tolerance and longevity (Shapiro et al., 1997; Hashmi et al., 1998; Vellai et al., 1999; Sumaya et al., 2018). Selective breeding, however, can introduce fitness trade-offs, as demonstrated by altered reproduction, dispersal, and storage stability in modified S. carpocapsae strains (Gaugler et al., 1990; Bal et al., 2014). Future efforts to develop host-specific EPNs may extend their efficacy beyond soil-dwelling insects to targeted pests across various cropping systems (Lacey and Georgis, 2012). The application of precise and advanced genetic engineering technologies such as CRISPR/Cas9 offers a pathway to precisely enhance EPN environment resilience through targeted modification.
Recent advancements in IJ recovery through biodegradable polymers and nanocarriers are emerging as promising tools for enhancing the EPN performance and tolerance under various field stress conditions. For instance, alginate-starch hydrogel increased S. feltiae persistence and infectivity across different water regimes (Perier et al., 2025), while titanium-dioxide-based Pickering emulsions maintained S. carpocapsae viability for up to 120 days (Kotliarevski et al., 2022). Future research should prioritize precision-based delivery systems that synchronize nematode release with pest phenology and prevailing microclimatic conditions compared to conventional aqueous sprays. EPN application via drip irrigation systems resulted in more uniform nematode distribution, improved persistence, and significantly greater control of onion thrips compared with standard foliar sprays (Metwally et al., 2025). Emerging AI-driven automatic and controlled treatment systems could further optimize site-specific deployment via drones or automated irrigation, targeting pest “hotspots” and reducing costs and environmental impact. To maximize field efficacy, EPNs should be applied during early morning or late evening to reduce UV exposure, in pre-moistened soils to enhance IJ motility, and using methods that minimize mechanical shear. When combined with improved formulations and targeted genetic enhancements, such synchronized, precision-based delivery can elevate EPNs from supplemental inputs to reliable, site-specific components of modern IPM programs.
5 Concluding remarks
In summary, EPNs offer a promising avenue for sustainable pest management, providing a biologically robust alternative or complement to synthetic chemical control. Continued innovation in formulation, strain selection, and application technologies, supported by insights from genomics, nanotechnology, and microbial ecology, will be essential to enhance their persistence and reliability under field conditions. Omics-based discovery of genes and regulatory pathways in EPNs can guide more precise, targeted improvements in strain performance. However, potential trade-offs between enhanced traits should be considered. Future innovations should also focus on ecological interactions and development of strategies that integrate EPNs with beneficial microbes to enhance EPN persistence, infectivity, and strain stability in field deployment.
Author contributions
AK: Conceptualization, Writing – original draft, Writing – review & editing. DS: Conceptualization, Writing – original draft, Writing – review & editing. PK: Writing – original draft, Writing – review & editing, Visualization. TP: Writing – review & editing. AB: Writing – review & editing.
Funding
The author(s) declared that financial support was not received for this work and/or its publication.
Conflict of interest
The author(s) declared that this work was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Generative AI statement
The author(s) declared that generative AI was not used in the creation of this manuscript.
Any alternative text (alt text) provided alongside figures in this article has been generated by Frontiers with the support of artificial intelligence and reasonable efforts have been made to ensure accuracy, including review by the authors wherever possible. If you identify any issues, please contact us.
Publisher’s note
All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.
References
Abd-Elgawad, M. M. M. (2023). Optimizing entomopathogenic nematode genetics and applications for the integrated management of horticultural pests. Horticulturae 9, 865. doi: 10.3390/horticulturae9080865
Ali, F. and Wharton, D. A. (2013). Cold tolerance abilities of two entomopathogenic nematodes, Steinernema feltiae and Heterorhabditis bacteriophora. Cryobiology 66, 24–29. doi: 10.1016/j.cryobiol.2012.10.004
Ali, J. G., Alborn, H. T., and Stelinski, L. L. (2010). Subterranean herbivore-induced volatiles released by citrus roots upon feeding by Diaprepes abbreviatus recruit entomopathogenic nematodes. J. Chem. Ecol. 36, 361–368. doi: 10.1007/s10886-010-9773-7
Andalo, V., Moino, A., Maximiniano, C., Campos, V. P., and Mendonca, L. A. (2011). Influence of temperature and duration of storage on the lipid reserves of entomopathogenic nematodes. Rev. Colomb. Entomol. 37, 203–209. doi: 10.25100/socolen.v37i2.9075
Ansari, M. A., Hussain, M. A., and Moens, M. (2009). Formulation and application of entomopathogenic nematode-infected cadavers for control of Hoplia philanthus in turfgrass. Pest Manag. Sci. 65, 367–374. doi: 10.1002/ps.1699
Aquino-Bolaños, T., Ruiz-Vega, J., Hernández, Y. D. O., and Castañeda, J. C. J. (2019). Survival of entomopathogenic nematodes in oil emulsions and control effectiveness on adult engorged ticks (Acari: ixodida). J. Nematol. 51, 1–10. doi: 10.21307/jofnem-2019-001
Bai, P. H., Yu, J. P., Hu, R. R., Fu, Q. W., Wu, H. C., Li, X. Y., et al. (2024). Behavioral and molecular response of the insect parasitic nematode Steinernema carpocapsae to plant volatiles. J. Invertebr. Pathol. 203, 108067. doi: 10.1016/j.jip.2024.108067
Baiocchi, T., Lee, G., Choe, D. H., and Dillman, A. R. (2017). Host seeking parasitic nematodes use specific odors to assess host resources. Sci. Rep. 7, 6270. doi: 10.1038/s41598-017-06620-2
Bal, H. K., Michel, A. P., and Grewal, P. S. (2014). Genetic selection of the ambush foraging entomopathogenic nematode, Steinernema carpocapsae for enhanced dispersal and its associated trade-offs. Evol. Ecol. 28, 923–939. doi: 10.1007/s10682-014-9706-y
Balasubramanian, N., Toubarro, D., and Simões, N. (2010). Biochemical study and in vitro insect immune suppression by a trypsin-like secreted protease from the nematode Steinernema carpocapsae. Parasit. Immunol. 32, 165–175. doi: 10.1111/j.1365-3024.2009.01172.x
Bargmann, C. I. (2006). “Chemosensation in C. elegans,” in WormBook: The online review of C. elegans biology. The C. elegans Research Community. Pasadena (CA). 1–29. doi: 10.1895/wormbook.1.123.1
Batalla-Carrera, L., Morton, A., Shapiro-Ilan, D., Strand, M. R., and García-del-Pino, F. (2014). Infectivity of Steinernema carpocapsae and S. feltiae to larvae and adults of the hazelnut weevil, Curculio nucum: Differential virulence and entry routes. J. Nematol. 46, 281–286.
Beck, B., Brusselman, E., Nuyttens, D., Moens, M., Pollet, S., Temmerman, F., et al. (2013). Improving foliar applications of entomopathogenic nematodes by selecting adjuvants and spray nozzles. Biocontrol Sci. Technol. 23, 507–520. doi: 10.1080/09583157.2013.777692
Bedding, R. A. (1988). Storage of insecticidal nematodes. World Patent No. WO 88/08668. Geneva, CH: World Intellectual Property Organization.
Bedding, R. A. and Butler, K. L. (1994). Method for the storage of entomopathogenic nematodes. WIPO Patent No. WO 94/05150. Geneva, CH: World Intellectual Property Organization.
Bedding, R. A., Clark, S. D., Lacey, M. J., and Butler, K. L. (2000). Method and apparatus for the storage of entomopathogenic nematodes. WIPO Patent No. WO 00/18887. Geneva, CH: World Intellectual Property Organization.
Bisch, G., Pagès, S., McMullen, J. G., Stock, S. P., Duvic, B., Givaudan, A., et al. (2015). Xenorhabdus bovienii CS03, the bacterial symbiont of the entomopathogenic nematode Steinernema weiseri, is a non-virulent strain against Lepidopteran insects. J. Invertebr. Pathol. 124, 15–22. doi: 10.1016/j.jip.2014.10.002
Blanco-Pérez, R., San-Blas, E., Rivera, M. J., and Campos-Herrera, R. (2025). Population ecology of entomopathogenic nematodes: Bridging past insights and future applications for sustainable agriculture. J. Invertebr. Pathol. 211, 108313. doi: 10.1016/j.jip.2025.108313
Bode, H. B. (2009). Entomopathogenic bacteria as a source of secondary metabolites. Curr. Opin. Chem. Biol. 13, 224–230. doi: 10.1016/j.cbpa.2009.02.037
Brown, I. M. and Gaugler, R. (1996). Cold tolerance of Steinernematid and Heterorhabditid nematodes. J. Therm. Biol. 21, 115–121. doi: 10.1016/0306-4565(95)00033-X
Campos-Herrera, R., El-Borai, F. E., and Duncan, L. W. (2012). Wide interguild relationships among entomopathogenic and free-living nematodes in soil as measured by real time qPCR. J. Invertebr. Pathol. 111, 126–135. doi: 10.1016/j.jip.2012.07.006
Campos-Herrera, R., Pathak, E., El-Borai, F. E., Schumann, A., Abd-Elgawad, M. M. M., and Duncan, L. W. (2013). New citriculture system suppresses native and augmented entomopathogenic nematodes. Biol. Control. 66, 183–194. doi: 10.1016/j.biocontrol.2013.05.009
Chang, D. Z., Serra, L., Lu, D., Mortazavi, A., and Dillman, A. R. (2019). A core set of venom proteins is released by entomopathogenic nematodes in the genus Steinernema. PloS Pathog. 15, e1007626. doi: 10.1371/journal.ppat.1007626
Chang, F. N. and Gehret, M. J. (1992). Insecticide delivery system and attractant. U.S. Patent No. 5141744. Washington, DC: U.S. Patent and Trademark Office.
Chang, F. N. and Gehret, M. J. (1995). Stabilized insect nematode compositions. U.S. Patent No. 5401506. Washington, DC: U.S. Patent and Trademark Office.
Chen, S. and Glazer, I. (2005). A novel method for long-term storage of the entomopathogenic nematode Steinernema feltiae at room temperature. Biol. Control. 32, 104–110. doi: 10.1016/j.biocontrol.2004.08.006
Ciche, T. A. and Ensign, J. C. (2003). For the insect pathogen Photorhabdus luminescens, which end of a nematode is out? Appl. Environ. Microbiol. 69, 1890–1897. doi: 10.1128/AEM.69.4.1890-1897.2003
Connick, W. J., Nickle, W. R., and Vinyard, B. T. (1993). Pesta”: New granular formulations for Steinernema carpocapsae. J. Nematol 25, 198–203.
Connick, W., Nickle, W. R., Williams, K., and Vinyard, B. (1994). Granular formulations of Steinernema carpocapsae (strain all) (Nematoda: Rhabditida) with improved shelf life. J. Nematol. 26, 352–359.
Cortés-Martínez, C. I., Ruiz-Vega, J., Matadamas-Ortiz, P. T., Lewis, E. E., Aquino-Bolaños, T., and Navarro-Antonio, J. (2016). Effect of moisture evaporation from diatomaceous earth pellets on storage stability of Steinernema glaseri. Biocontrol Sci. Technol. 26, 305–319. doi: 10.1080/09583157.2015.1104650
Crawford, J. M., Portmann, C., Zhang, X., Roeffaers, M. B. J., and Clardy, J. (2012). Small molecule perimeter defense in entomopathogenic bacteria. Proc. Natl. Acad. Sci. 109, 10821–10826. doi: 10.1073/pnas.1201160109
Dara, S. K. (2019). The new integrated pest management paradigm for the modern age. J. Integr. Pest Manag., 10(1). doi: 10.1093/jipm/pmz010
Darsouei, R., Karimi, J., and Stelinski, L. L. (2025). Properties of enhanced calcium-alginate beads as a formulation for disseminating the entomopathogenic nematodes Heterorhabditis bacteriophora, Steinernema carpocapase, and Steinernema feltiae. J. Nematol 57, 20250020. doi: 10.2478/jofnem-2025-0020
da Silva, W. J., Pilz-Júnior, H. L., Heermann, R., and da Silva, O. S. (2020). The great potential of entomopathogenic bacteria Xenorhabdus and Photorhabdus for mosquito control: A review. Parasit. Vectors. 13, 376. doi: 10.1186/s13071-020-04236-6
Degenhardt, J., Hiltpold, I., Köllner, T. G., Frey, M., Gierl, A., Gershenzon, J., et al. (2009). Restoring a maize root signal that attracts insect-killing nematodes to control a major pest. Proc. Natl. Acad. Sci. 106, 13213–13218. doi: 10.1073/pnas.0906365106
Dias S da, C., de Brida, A. L., Jean-Baptiste, M. C., Leite, L. G., Ovruski, S. M., Lee, J. C., et al. (2024). Compatibility of entomopathogenic nematodes with chemical insecticides for the control of Drosophila suzukii (Diptera: Drosophilidae). Plants 13, 632. doi: 10.3390/plants13050632
Dowds, B. C. and Peters, A. (2002). "Virulence mechanisms." in Entomopathogenic Nematology. Ed. R. Gaugler (CABI, Wallingford), 79–98. doi: 10.1079/9780851995670.0079
Dowling, A. J., Daborn, P. J., Waterfield, N. R., Wang, P., Streuli, C. H., and Ffrench-Constant, R. H. (2004). The insecticidal toxin Makes caterpillars floppy (Mcf) promotes apoptosis in mammalian cells: An insecticidal toxin promoting apoptosis. Cell Microbiol. 6, 345–353. doi: 10.1046/j.1462-5822.2003.00357.x
Ehlers, R. U., Wulff, A., and Peters, A. (1997). Pathogenicity of axenic Steinernema feltiae, Xenorhabdus bovienii, and the bacto–helminthic complex to larvae of Tipula oleracea (Diptera) and Galleria mellonella (Lepidoptera). J. Invertebr. Pathol. 69, 212–217. doi: 10.1006/jipa.1996.4647
Eleftherianos, I., Boundy, S., Joyce, S. A., Aslam, S., Marshall, J. W., Cox, R. J., et al. (2007). An antibiotic produced by an insect-pathogenic bacterium suppresses host defenses through phenoloxidase inhibition. Proc. Natl. Acad. Sci. 104, 2419–2424. doi: 10.1073/pnas.0610525104
Fallet, P., Bazagwira, D., Ruzzante, L., Ingabire, G., Levivier, S., Bustos-Segura, C., et al. (2024). Entomopathogenic nematodes as an effective and sustainable alternative to control the fall armyworm in Africa. PNAS. Nexus. 3, 122. doi: 10.1093/pnasnexus/pgae122
Fang, X., Zhang, M., Tang, Q., Wang, Y., and Zhang, X. (2014). Inhibitory effect of Xenorhabdus nematophila TB on plant pathogens Phytophthora capsici and Botrytis cinerea in vitro and in planta. Sci. Rep. 4, 4300. doi: 10.1038/srep04300
Felföldi, G., Marokházi, J., Képiró, M., and Venekei, I. (2009). Identification of natural target proteins indicates functions of a serralysin-type metalloprotease, PrtA, in anti-immune mechanisms. Appl. Environ. Microbiol. 75, 3120–3126. doi: 10.1128/AEM.02271-08
ffrench-Constant, R. H., Dowling, A., and Waterfield, N. R. (2007). Insecticidal toxins from Photorhabdus bacteria and their potential use in agriculture. Toxicon 49, 436–451. doi: 10.1016/j.toxicon.2006.11.019
Filgueiras, C. C., Willett, D. S., Junior, A. M., Pareja, M., Borai, F. E., Dickson, D. W., et al. (2016). Stimulation of the salicylic acid pathway aboveground recruits entomopathogenic nematodes belowground. PloS One 11, e0154712. doi: 10.1371/journal.pone.0154712
Forst, S., Dowds, B., Boemare, N., and Stackebrandt, E. (1997). Xenorhabdus and Photorhabdus spp.: Bugs that kill bugs. Annu. Rev. Microbiol. 51, 47–72. doi: 10.1146/annurev.micro.51.1.47
Furgani, G., Böszörményi, E., Fodor, A., Máthé-Fodor, A., Forst, S., Hogan, J. S., et al. (2008). Xenorhabdus antibiotics: A comparative analysis and potential utility for controlling mastitis caused by bacteria. J. Appl. Microbiol. 104, 745–758. doi: 10.1111/j.1365-2672.2007.03613.x
Gal, T. Z., Glazer, I., and Koltai, H. (2003). Differential gene expression during desiccation stress in the insect-killing nematode Steinernema feltiae IS-6. J. Parasitol. 89, 761–766. doi: 10.1645/GE-3105
Gaugler, R., Campbell, J. F., and McGuire, T. R. (1990). Fitness of a genetically improved entomopathogenic nematode. J. Invertebr. Pathol. 56, 106–116. doi: 10.1016/0022-2011(90)90151-U
Georgis, R. (1990). “Formulation and application technology,” in Entomopathogenic Nematodes in Biological Control, Eds. Gaugler, R. and Kaya, H. K. (CRC Press, Boca Raton, FL), 173–194.
Glazer, I., Simões, N., Eleftherianos, I., Ramakrishnan, J., Ment, D., Toubarro, D., et al. (2025). Entomopathogenic nematodes: Survival, virulence and immunity. J. Invertebr. Pathol. 212, 108363. doi: 10.1016/j.jip.2025.108363
Godina, G., Vandenbossche, B., Schmidt, M., Sender, A., Tambe, A. H., Touceda-González, M., et al. (2023). Entomopathogenic nematodes for biological control of Psylliodes chrysocephala (Coleoptera: Chrysomelidae) in oilseed rape. J. Invertebr. Pathol. 197, 107894. doi: 10.1016/j.jip.2023.107894
Grewal, P. S. (1998). Formulations of entomopathogenic nematodes for storage and application. Nematol Res. Jpn. J. Nematol 28, 68–74. doi: 10.3725/jjn1993.28.supplement_68
Grewal, P. S. (2000). Enhanced ambient storage stability of an entomopathogenic nematode through anhydrobiosis. Pest Manag. Sci. 56, 401–406. doi: 10.1002/(SICI)1526-4998(200005)56:5<401::AID-PS137>3.0.CO;2-4
Grewal, P. (2002). “Formulation and application technology,” in Entomopathogenic Nematology. Ed. Gaugler, R. (CABI, Oxfordshire), 265–287.
Grewal, P. S., Bornstein-Forst, S., Burnell, A. M., Glazer, I., and Jagdale, G. B. (2006). Physiological, genetic, and molecular mechanisms of chemoreception, thermobiosis, and anhydrobiosis in entomopathogenic nematodes. Biol. Control. 38, 54–65. doi: 10.1016/j.biocontrol.2005.09.004
Grewal, P. S. and Jagdale, G. B. (2002). Enhanced trehalose accumulation and desiccation survival of entomopathogenic nematodes through cold preacclimation. Biocontrol Sci. Technol. 12, 533–545. doi: 10.1080/0958315021000016207
Grunseich, J. M., Aguirre, N. M., Thompson, M. N., Ali, J. G., and Helms, A. M. (2021). Chemical cues from entomopathogenic nematodes vary across three species with different foraging strategies, triggering different behavioral responses in prey and competitors. J. Chem. Ecol. 47, 822–833. doi: 10.1007/s10886-021-01304-8
Guillermin, M. L., Carrillo, M. A., and Hallem, E. A. (2017). A single set of interneurons drives opposite behaviors in C. elegans. Curr. Biol. CB. 27, 2630–2639.e6. doi: 10.1016/j.cub.2017.07.023
Gulcu, B., Hazir, S., and Kaya, H. K. (2012). Scavenger deterrent factor (SDF) from symbiotic bacteria of entomopathogenic nematodes. J. Invertebr. Pathol. 110, 326–333. doi: 10.1016/j.jip.2012.03.014
Gulcu, B., Hazir, S., Lewis, E. E., and Kaya, H. K. (2018). Evaluation of responses of different ant species (Formicidae) to the scavenger deterrent factor associated with the entomopathogenic nematode-bacterium complex. Eur. J. Entomol. 115, 312–317. doi: 10.14411/eje.2018.030
Gumus, A., Karagoz, M., Shapiro-Ilan, D., and Hazir, S. (2015). A novel approach to biocontrol: Release of live insect hosts pre-infected with entomopathogenic nematodes. J. Invertebr. Pathol. 130, 56–60. doi: 10.1016/j.jip.2015.07.002
Hallem, E. A., Dillman, A. R., Hong, A. V., Zhang, Y., Yano, J. M., DeMarco, S. F., et al. (2011). A sensory code for host seeking in parasitic nematodes. Curr. Biol. 21, 377–383. doi: 10.1016/j.cub.2011.01.048
Han, R. and Ehlers, R. U. (2000). Pathogenicity, development, and reproduction of Heterorhabditis bacteriophora and Steinernema carpocapsae under axenic in vivo conditions. J. Invertebr. Pathol. 75, 55–58. doi: 10.1006/jipa.1999.4900
Hart, A. C. and Chao, M. Y. (2010). "From odors to behaviors in Caenorhabditis elegans." in The Neurobiology of Olfaction. Ed. A. Menini (CRC Press, Boca Raton, FL), 1–35.
Hashmi, S., Hashmi, G., Glazer, I., and Gaugler, R. (1998). Thermal response of Heterorhabditis bacteriophora transformed with the Caenorhabditis elegans hsp70 encoding gene. J. Exp. Zool. 281, 164–170. doi: 10.1002/(SICI)1097-010X(19980615)281:3<164::AID-JEZ2>3.0.CO;2-L
Hazir, S., Shapiro-Ilan, D. I., Bock, C. H., Hazir, C., Leite, L. G., and Hotchkiss, M. W. (2016). Relative potency of culture supernatants of Xenorhabdus and Photorhabdus spp. on growth of some fungal phytopathogens. Eur. J. Plant Pathol. 146, 369–381. doi: 10.1007/s10658-016-0923-9
Helms, A. M., Ray, S., Matulis, N. L., Kuzemchak, M. C., Grisales, W., Tooker, J. F., et al. (2019). Chemical cues linked to risk: Cues from below-ground natural enemies enhance plant defences and influence herbivore behaviour and performance. Funct. Ecol. 33, 798–808. doi: 10.1111/1365-2435.13297
Hiltpold, I., Hibbard, B. E., French, B. W., and Turlings, T. C. J. (2012). Capsules containing entomopathogenic nematodes as a Trojan horse approach to control the western corn rootworm. Plant Soil. 358, 11–25. doi: 10.1007/s11104-012-1253-0
Hinchliffe, S. J., Hares, M. C., Dowling, A. J., and Ffrench-Constant, R. H. (2010). Insecticidal toxins from the Photorhabdus and Xenorhabdus bacteria. Open Toxinol. J. 3, 83–100. doi: 10.2174/1875414701003010101
Hussein, M. and Abdel-Aty, M. (2012). Formulation of two native entomopathogenic nematodes at room temperature. J. Biopestic. 5, 23–27. doi: 10.57182/jbiopestic.5.0.23-27
Hwang, J., Park, Y., Kim, Y., Hwang, J., and Lee, D. (2013). An entomopathogenic bacterium, Xenorhabdus nematophila, suppresses expression of antimicrobial peptides controlled by Toll and Imd pathways by blocking eicosanoid biosynthesis. Arch. Insect Biochem. Physiol. 83, 151–169. doi: 10.1002/arch.21103
Jagdale, G. B. and Grewal, P. S. (2003). Acclimation of entomopathogenic nematodes to novel temperatures: Trehalose accumulation and the acquisition of thermotolerance. Int. J. Parasitol. 33, 145–152. doi: 10.1016/S0020-7519(02)00257-6
Jagdale, G. B., Kamoun, S., and Grewal, P. S. (2009). Entomopathogenic nematodes induce components of systemic resistance in plants: Biochemical and molecular evidence. Biol. Control. 51, 102–109. doi: 10.1016/j.biocontrol.2009.06.009
Kajuga, J. N., Waweru, B. W., Bazagwira, D., Ishimwe, P. M., Ndacyayisaba, S., Mukundiyabo, G. C., et al. (2025). Efficacy of foliar applications of entomopathogenic nematodes in the management of the invasive tomato leaf miner Phthorimaea absoluta compared to local practices under open-field conditions. Agronomy 15, 1417. doi: 10.3390/agronomy15061417
Kallali, N. S., Ouijja, A., Goura, K., Laasli, S. E., Kenfaoui, J., Benseddik, Y., et al. (2024). From soil to host: Discovering the tripartite interactions between entomopathogenic nematodes, symbiotic bacteria and insect pests and related challenges. J. Nat. Pestic. Res. 7, 100065. doi: 10.1016/j.napere.2023.100065
Karimi, S., Eivazian Kary, N., and Mohammadi, D. (2025). Optimization of carboxymethyl cellulose-based formulations for enhanced shelf life of an entomopathogenic nematode, Steinernema carpocapsae. Egypt. J. Biol. Pest Control. 35, 23. doi: 10.1186/s41938-025-00858-z
Kary, N. E., Chahardoli, S., Mohammadi, D., and Dillon, A. B. (2021). Efficacy of carboxymethyl cellulose as an inert water-soluble carrier for formulation of entomopathogenic nematodes, Heterorhabditis bacteriophora and Steinernema carpocapsae. Biol. Control. 160, 104690. doi: 10.1016/j.biocontrol.2021.104690
Kim, J., Hiltpold, I., Jaffuel, G., Sbaiti, I., Hibbard, B. E., and Turlings, T. C. J. (2021). Calcium-alginate beads as a formulation for the application of entomopathogenic nematodes to control rootworms. J. Pest Sci. 94, 1197–1208. doi: 10.1007/s10340-021-01349-4
Kim, J., Jaffuel, G., and Turlings, T. C. J. (2015). Enhanced alginate capsule properties as a formulation of entomopathogenic nematodes. BioControl 60, 527–535. doi: 10.1007/s10526-014-9638-z
Koppenhöfer, A. M. and Luiza Sousa, A. (2024). Long-term suppression of turfgrass insect pests with native persistent entomopathogenic nematodes. J. Invertebr. Pathol. 204, 108123. doi: 10.1016/j.jip.2024.108123
Koppenhöfer, A. M., Shapiro-Ilan, D. I., and Hiltpold, I. (2020). Entomopathogenic nematodes in sustainable food production. Front. Sustain. Food Syst. 4. doi: 10.3389/fsufs.2020.00125
Kotliarevski, L., Cohen, R., Ramakrishnan, J., Wu, S., Mani, K. A., Amar-Feldbaum, R., et al. (2022). Individual coating of entomopathogenic nematodes with titania (TiO2) nanoparticles based on oil-in-water pickering emulsion: a new formulation for biopesticides. J. Agric. Food Chem. 70, 13518–13527. doi: 10.1021/acs.jafc.2c04424
Krishnayyaand, P. V. and Grewal, P. S. (2002). Effect of neem and selected fungicides on viability and virulence of the entomopathogenic nematode Steinernema feltiae. Biocontrol Sci. Technol. 12, 259–266. doi: 10.1080/09583150210388
Lacey, L. A. and Georgis, R. (2012). Entomopathogenic nematodes for control of insect pests above and below ground with comments on commercial production. J. Nematol. 44, 218–225.
Laznik, Ž., Košir, I. J., Košmelj, K., Murovec, J., Jagodič, A., Trdan, S., et al. (2020). Effect of Cannabis sativa L. root, leaf and inflorescence ethanol extracts on the chemotrophic response of entomopathogenic nematodes. Plant Soil. 455, 367–379. doi: 10.1007/s11104-020-04693-z
Laznik, Ž. and Trdan, S. (2016). Attraction behaviors of entomopathogenic nematodes (Steinernematidae and Heterorhabditidae) to synthetic volatiles emitted by insect damaged potato tubers. J. Chem. Ecol. 42, 314–322. doi: 10.1007/s10886-016-0686-y
Leite, L. G., Shapiro-Ilan, D. I., and Hazir, S. (2018). Survival of Steinernema feltiae in different formulation substrates: Improved longevity in a mixture of gel and vermiculite. Biol. Control. 126, 192–197. doi: 10.1016/j.biocontrol.2018.05.013
Liao, C., Zhang, S., Fu, T., Jin, X., Xu, W., Liu, D., et al. (2025). Mutualistic bacteria of entomopathogenic nematodes as an insecticidal agent for sustainable agriculture. Chem. Biol. Technol. Agric. 12, 141. doi: 10.1186/s40538-025-00862-3
Liu, J., Li, W., Wang, C., Liang, H., Chu, Z., Wang, G., et al. (2025). Mixtures of entomopathogenic nematodes provided successful suppression of Bactrocera tau by stimulated dispersal, aggregation, and invasion. Biol. Control. 208, 105858. doi: 10.1016/j.biocontrol.2025.105858
Llácer, E., Martínez de Altube, M. M., and Jacas, J. A. (2009). Evaluation of the efficacy of Steinernema carpocapsae in a chitosan formulation against the red palm weevil, Rhynchophorus ferrugineus, in Phoenix canariensis. BioControl 54, 559–565. doi: 10.1007/s10526-008-9208-3
Lortkipanidze, M. A., Gorgadze, O. A., Kajaia, G., Gratiashvili, N. G., and Kuchava, M. A. (2016). Foraging behavior and virulence of some entomopathogenic nematodes. Ann. Agrar. Sci. 2, 99–103. doi: 10.1016/j.aasci.2016.05.009
Lu, D., Macchietto, M., Chang, D., Barros, M. M., Baldwin, J., Mortazavi, A., et al. (2017). Activated entomopathogenic nematode infective juveniles release lethal venom proteins. PloS Pathog. 13, e1006302. doi: 10.1007/s00248-020-01573-y
Maher, A. M. D., Asaiyah, M., Quinn, S., Burke, R., Wolff, H., Bode, H. B., et al. (2021). Competition and co-existence of two Photorhabdus symbionts with a nematode host. Microb. Ecol. 81, 223–239.
Mahmoud, M. F., Mahfouz, H. M., and Mohamed, K. M. (2016). Compatibility of entomopathogenic nematodes with neonicotinoids and Azadirachtin insecticides for controlling the black cutworm, Agrotis ipsilon (Hufnagel) in canola plants. IJRES 2, 11–18. doi: 10.20431/2454-9444.0201002
Manohar, M., Tenjo-Castano, F., Chen, S., Zhang, Y. K., Kumari, A., Williamson, V. M., et al. (2020). Plant metabolism of nematode pheromones mediates plant-nematode interactions. Nat. Commun. 11, 208. doi: 10.1038/s41467-019-14104-2
Matadamas-Ortiz, P. T., Ruiz-Vega, J., Vazquez-Feijoo, J. A., Cruz-Martínez, H., and Cortés-Martínez, C. I. (2014). Mechanical production of pellets for the application of entomopathogenic nematodes: Factors that determine survival time of Steinernema glaseri. Biocontrol Sci. Technol. 24, 145–157. doi: 10.1080/09583157.2013.852161
McMullen, J. G., Peterson, B. F., Forst, S., Blair, H. G., and Stock, S. P. (2017). Fitness costs of symbiont switching using entomopathogenic nematodes as a model. BMC Evol. Biol. 17, 100. doi: 10.1186/s12862-017-0939-6
Metwally, H. M. S., Saleh, M. M. E., and Abonaem, M. (2025). Formulation for foliar and soil application of entomopathogenic nematodes for controlling the onion thrips Thrips tabaci Lindeman (Thysanoptera: Thripidae). Egypt. J. Biol. Pest Control. 35, 4. doi: 10.1186/s41938-025-00841-8
Modic, Š., Žigon, P., Kolmanič, A., Trdan, S., and Razinger, J. (2020). Evaluation of the field efficacy of Heterorhabditis bacteriophora poinar (Rhabditida: Heterorhabditidae) and synthetic insecticides for the control of western corn rootworm larvae. Insects 11. doi: 10.3390/insects11030202
Moore, S. D., Ehlers, R. U., Manrakhan, A., Gilbert, M., Kirkman, W., Daneel, J. H., et al. (2024). Field-scale efficacy of entomopathogenic nematodes to control false codling moth, Thaumatotibia leucotreta (Lepidoptera: Tortricidae), in citrus orchards in South Africa. Crop Prot. 179, 106610. doi: 10.1016/j.cropro.2024.106610
Moreira, G. F., Batista, E. S. D. P., Campos, H. B. N., Lemos, R. E., and Ferreira, M. D. C. (2013). Spray nozzles, pressures, additives and stirring time on viability and pathogenicity of entomopathogenic nematodes (Nematoda: Rhabditida) for greenhouses. PloS One 8, e65759. doi: 10.1371/journal.pone.0065759
Navon, A., Keren, S., Salame, L., and Glazer, I. (1998). An edible-to-insects calcium alginate gel as a carrier for entomopathogenic nematodes. Biocontrol Sci. Technol. 8, 429–437. doi: 10.1080/09583159830225
Navon, A., Nagalakshmi, V. K., Levski, S., Salame, L., and Glazer, I. (2002). Effectiveness of entomopathogenic nematodes in an alginate gel formulation against Lepidopterous pests. Biocontrol Sci. Technol. 12, 737–746. doi: 10.1080/0958315021000039914
O’Halloran, D. M. and Burnell, A. M. (2003). An investigation of chemotaxis in the insect parasitic nematode Heterorhabditis bacteriophora. Parasitology 127, 375–385. doi: 10.1017/S0031182003003688
Odendaal, D., Addison, M. F., and Malan, A. P. (2016). Entomopathogenic nematodes for the control of the codling moth (Cydia pomonella L.) in field and laboratory trials. J. Helminthol. 90, 615–623. doi: 10.1017/S0022149X15000887
Ozawa, K., Shinkai, Y., Kako, K., Fukamizu, A., and Doi, M. (2022). The molecular and neural regulation of ultraviolet light phototaxis and its food-associated learning behavioral plasticity in C. elegans. Neurosci. Lett. 770, 136384. doi: 10.1016/j.neulet.2021.136384
Park, Y. and Kim, Y. (2000). Eicosanoids rescue Spodoptera exigua infected with Xenorhabdus nematophilus, the symbiotic bacteria to the entomopathogenic nematode Steinernema carpocapsae. J. Insect Physiol. 46, 1469–1476. doi: 10.1016/S0022-1910(00)00071-8
Perez, E. E., Lewis, E. E., and Shapiro-Ilan, D. I. (2003). Impact of the host cadaver on survival and infectivity of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) under desiccating conditions. J. Invertebr. Pathol. 82, 111–118. doi: 10.1016/S0022-2011(02)00204-5
Perier, J. D., Wu, S., Arthurs, S. P., Toews, M. D., and Shapiro-Ilan, D. I. (2025). Persistence of the entomopathogenic nematode Steinernema feltiae in a novel capsule formulation. Biol. Control. 200, 105684. doi: 10.1016/j.biocontrol.2024.105684
Pervez, R., Lone, S. A., and Pattnaik, S. (2020). Characterization of symbiotic and associated bacteria from entomopathogenic nematode Heterorhabditis sp. (nematode: Heterorhabditidae) isolated from India. Egypt. J. Biol. Pest Control. 30. doi: 10.1186/s41938-020-00343-9
Platt, T., Stokwe, N. F., and Malan, A. P. (2018). Foliar application of Steinernema yirgalemense to control Planococcus ficus: Assessing adjuvants to improve efficacy. South Afr. J. Enol. Vitic. 40. doi: 10.21548/40-1-2920
Polat, B., Cengiz, A., Koc, S., Kokten, S. K., Gultekin, Z. N., Caliskan, C., et al. (2024). Efficacy of Steinernema feltiae nematode against lesser mealworm (Alphitobius diaperinus) populations from poultry farms in Türkiye. Vet. Sci. 11, 567. doi: 10.3390/vetsci11110567
Půža, V. and Tarasco, E. (2023). Interactions between entomopathogenic fungi and entomopathogenic nematodes. Microorganisms 11, 163. doi: 10.3390/microorganisms11010163
Raja, R., Hazir, C., Gümüş, A., Asan, C., Karagöz, M., and Hazir, S. (2015). Efficacy of the entomopathogenic nematode Heterorhabditis bacteriophora using different application methods in the presence or absence of a natural enemy. Turk. J. Agric. For. 39, 277–285. doi: 10.3906/tar-1410-33
Ramakrishnan, J., Salame, L., Mani, K. A., Feldbaum, R., Karavani, E., Mechrez, G., et al. (2023). Increasing the survival and efficacy of entomopathogenic nematodes on exposed surfaces by Pickering emulsion formulations offers new venue for foliar pest management. J. Invertebr. Pathol. 199, 107938. doi: 10.1016/j.jip.2023.107938
Ramakuwela, T., Hatting, J., Laing, M. D., Hazir, S., and Thiebaut, N. (2016). In vitro solid-state production of Steinernema innovationi with cost analysis. Biocontrol Sci. Technol. 26, 792–808. doi: 10.1080/09583157.2016.1159284
Ramakuwela, T., Tarasco, E., Chavarría-Hernández, N., and Toepfer, S. (2025). Entomopathogenic nematodes: Commercial use and future perspectives. J. Invertebr. Pathol. 212, 108388. doi: 10.1016/j.jip.2025.108388
Rasmann, S., Köllner, T. G., Degenhardt, J., Hiltpold, I., Toepfer, S., Kuhlmann, U., et al. (2005). Recruitment of entomopathogenic nematodes by insect-damaged maize roots. Nature 434, 732–737. doi: 10.1038/nature03451
Sabino PH de, S., Sales, F. S., Guevara, E. J., Junior, A. M., and Filgueiras, C. C. (2014). Compatibility of entomopathogenic nematodes (Nematoda: Rhabditida) with insecticides used in the tomato crop. Nematoda 1, e03014. doi: 10.4322/nematoda.03014
Sáenz-Aponte, A., Correa-Cuadros, J. P., and Rodríguez-Bocanegra, M. X. (2020). Foliar application of entomopathogenic nematodes and fungi for the management of the diamondback moth in greenhouse and field. Biol. Control. 142, 104163. doi: 10.1016/j.biocontrol.2019.104163
San-Blas, E. (2013). Progress on entomopathogenic nematology research: A bibliometric study of the last three decades: 1980–2010. Biol. Control. 66, 102–124. doi: 10.1016/j.biocontrol.2013.04.002
Schroer, S., Yi, X., and Ehlers, R. U. (2005). Evaluation of adjuvants for foliar application of Steinernema carpocapsae against larvae of the diamondback moth (Plutella xylostella). Nematology 7, 37–44. doi: 10.1163/1568541054192126
Shapiro, D. I., Behle, R., McGuire, M. R., and Lewis, E. E. (2003). Formulated arthropod cadavers for pest suppression. U.S. Patent No. 6,524,601. Washington, DC: U.S. Patent and Trademark Office.
Shapiro, D. I. and Glazer, I. (1996). Comparison of entomopathogenic nematode dispersal from infected hosts versus aqueous suspension. Environ. Entomol. 25, 1455–1461. doi: 10.1093/ee/25.6.1455
Shapiro, D. I., Glazer, I., and Segal, D. (1997). Genetic improvement of heat tolerance in Heterorhabditis bacteriophora through hybridization. Biol. Control. 8, 153–159. doi: 10.1006/bcon.1996.0488
Shapiro, D. I. and Lewis, E. E. (1999). Comparison of entomopathogenic nematode infectivity from infected hosts versus aqueous suspension. Environ. Entomol. 28, 907–911. doi: 10.1093/ee/28.5.907
Shapiro-Ilan, D. I., Campbell, J. F., Lewis, E. E., Elkon, J. M., and Kim-Shapiro, D. B. (2009). Directional movement of Steinernematid nematodes in response to electrical current. J. Invertebr. Pathol. 100, 134–137. doi: 10.1016/j.jip.2008.11.001
Shapiro-Ilan, D. I., Cottrell, T. E., Mizell, R. F., Horton, D. L., Behle, R. W., and Dunlap, C. A. (2010a). Efficacy of Steinernema carpocapsae for control of the lesser peachtree borer, Synanthedon pictipes: Improved aboveground suppression with a novel gel application. Biol. Control. 54, 23–28. doi: 10.1016/j.biocontrol.2009.11.009
Shapiro-Ilan, D. and Dolinski, C. (2015). Entomopathogenic nematode application technology. Nematode. Pathog. Insects. Pests. Ecol. Appl. Technol. Sustain. Plant Crop Prot. 231–254. doi: 10.1007/978-3-319-18266-7_9
Shapiro-Ilan, D. I. and Goolsby, J. A. (2021). Evaluation of Barricade® to enhance survival of entomopathogenic nematodes on cowhide. J. Invertebr. Pathol. 184, 107592. doi: 10.1016/j.jip.2021.107592
Shapiro-Ilan, D. I., Lewis, E. E., Behle, R. W., and McGuire, M. R. (2001). Formulation of entomopathogenic nematode-infected cadavers. J. Invertebr. Pathol. 78, 17–23. doi: 10.1006/jipa.2001.5030
Shapiro-Ilan, D. I., Lewis, E. E., Campbell, J. F., and Kim-Shapiro, D. B. (2012). Directional movement of entomopathogenic nematodes in response to electrical field: Effects of species, magnitude of voltage, and infective juvenile age. J. Invertebr. Pathol. 109, 34–40. doi: 10.1016/j.jip.2011.09.004
Shapiro-Ilan, D. I., Lewis, E. E., Son, Y., and Tedders, W. L. (2003). Superior efficacy observed in entomopathogenic nematodes applied in infected-host cadavers compared with application in aqueous suspension. J. Invertebr. Pathol. Washington, DC: U.S. Patent and Trademark Office 83, 270–272. doi: 10.1016/S0022-2011(03)00101-0
Shapiro-Ilan, D. I., Morales-Ramos, J. A., Rojas, M. G., and Tedders, W. L. (2010b). Effects of a novel entomopathogenic nematode-infected host formulation on cadaver integrity, nematode yield, and suppression of Diaprepes abbreviatus and Aethina tumida. J. Invertebr. Pathol. 103, 103–108. doi: 10.1016/j.jip.2009.11.006
Shields, E. J., Testa, A. M., and O’Neil, W. J. (2018). Long-term persistence of native New York entomopathogenic nematode isolates across crop rotation. J. Econ. Entomol. 111, 2592–2598. doi: 10.1093/jee/toy258
Silver, S. C., Dunlop, D. B., and Grove, D. I. (1995). Granular formulation of entities with improved storage stability. WIPO Patent No. WO 95/0577. Geneva, CH: World Intellectual Property Organization.
Solomon, A., Paperna, I., and Glazer, I. (1999). Desiccation survival of the entomopathogenic nematode Steinernema feltiae: Induction of anhydrobiosis. Nematology 1, 61–68. doi: 10.1163/156854199507983
Solomon, A., Salomon, R., Paperna, I., and Glazer, I. (2000). Desiccation stress of entomopathogenic nematodes induces the accumulation of a novel heat-stable protein. Parasitology 121, 409–416. doi: 10.1017/S0031182099006563
Somvanshi, V. S., Koltai, H., and Glazer, I. (2008). Expression of different desiccation-tolerance related genes in various species of entomopathogenic nematodes. Mol. Biochem. Parasitol. 158, 65–71. doi: 10.1016/j.molbiopara.2007.11.012
Srivastava, C. N., Mohan, L., Sharma, P., and Maurya, P. (2011). A review on prospectives of synergistic approach in insect pest management. J. Entomol. Res. 35, 255–266.
Stilwell, M. D., Cao, M., Goodrich-Blair, H., and Weibel, D. B. (2018). Studying the symbiotic bacterium Xenorhabdus nematophila in individual, living Steinernema carpocapsae nematodes using microfluidic systems. mSphere 3. doi: 10.1128/msphere.00530-17
Stock, S. P., Campos-Herrera, R., and Shapiro-Ilan, D. (2025). The first 100 years in the history of entomopathogenic nematodes. J. Invertebr. Pathol. 211, 108302. doi: 10.1016/j.jip.2025.108302
Stock, S. P., Kusakabe, A., and Orozco, R. A. (2017). Secondary metabolites produced by Heterorhabditis symbionts and their application in agriculture: what we know and what to do next. J. Nematol. 49, 373. doi: 10.21307/jofnem-2017-084
Strauch, O., Niemann, I., Neumann, A., Schmidt, A. J., Peters, A., and Ehlers, R. U. (2000). Storage and formulation of the entomopathogenic nematodes Heterorhabditis indica and H. bacteriophora. BioControl 45, 483–500. doi: 10.1023/A:1026528727365
Sugiyama, T. and Hasegawa, K. (2025). Synergistic actions of symbiotic bacteria modulate the insecticidal potency of entomopathogenic nematode Steinernema monticolum KHA701. Sci. Rep. 15. doi: 10.1038/s41598-025-06488-7
Sumaya, N. H., Gohil, R., Okolo, C., Addis, T., Doerfler, V., Ehlers, R. U., et al. (2018). Applying inbreeding, hybridization and mutagenesis to improve oxidative stress tolerance and longevity of the entomopathogenic nematode Heterorhabditis bacteriophora. J. Invertebr. Pathol. 151, 50–58. doi: 10.1016/j.jip.2017.11.001
Sun, B., Zhang, X., Song, L., Zheng, L., Wei, X., Gu, X., et al. (2021). Evaluation of indigenous entomopathogenic nematodes in Southwest China as potential biocontrol agents against Spodoptera litura (Lepidoptera: Noctuidae). J. Nematol. 53, e2021–e2083. doi: 10.21307/jofnem-2021-083
Tarasco, E., Fanelli, E., Salvemini, C., El-Khoury, Y., Troccoli, A., Vovlas, A., et al. (2023). Entomopathogenic nematodes and their symbiotic bacteria: From genes to field uses. Front. Insect Sci. 3, 1195254. doi: 10.3389/finsc.2023.1195254
Thakur, N., Tomar, P., Kaur, S., and Kumari, P. (2022). Virulence of native entomopathogenic nematodes against major lepidopteran insect species of tomato (Solanum lycopersicum L.). J. Appl. Biol. Biotechnol. 10, 6–14. doi: 10.7324/JABB.2022.10s102
Tobias, N. J., Wolff, H., Djahanschiri, B., Grundmann, F., Kronenwerth, M., Shi, Y. M., et al. (2017). Natural product diversity associated with the nematode symbionts Photorhabdus and Xenorhabdus. Nat. Microbiol. 12, 1676–1685. doi: 10.1038/s41564-017-0039-9
Tomar, P. and Thakur, N. (2022). Biocidal potential of indigenous isolates of entomopathogenic nematodes (EPNs) against tobacco cutworm, Spodoptera litura Fabricius (Lepidoptera: Noctuidae). Egypt. J. Biol. Pest Control. 32, 107. doi: 10.1186/s41938-022-00607-6
Torr, P., Heritage, S., and Wilson, M. J. (2004). Vibrations as a novel signal for host location by parasitic nematodes. Int. J. Parasitol. 34, 997–999. doi: 10.1016/j.ijpara.2004.05.003
Tumialis, D., Mazurkiewicz, A., Florczak, L., and Skrzecz, I. (2023). The potential of entomopathogenic nematodes of the genera Steinernema and Heterorhabditis for biological control of the pine lappet moth Dendrolimus pini L. (Lepidoptera: Lasiocampidae) in Scots pine stands. For. Int. J. For. Res. 96, 733–739. doi: 10.1093/forestry/cpad008
United Nations (2017). World population prospects: The 2017 revision (New York, USA: United Nations Department of Economic and Social Affairs), 24.
Valle, E. E. D., Dolinski, C., Barreto, E. L. S., and Souza, R. M. (2009). Effect of cadaver coatings on emergence and infectivity of the entomopathogenic nematode Heterorhabditis baujardi LPP7 (Rhabditida: Heterorhabditidae) and the removal of cadavers by ants. Biol. Control. 50, 21–24. doi: 10.1016/j.biocontrol.2009.01.007
Vellai, T., Molnár, A., Lakatos, L., Bánfalvi, Z., Fodor, A., and Sáringer, G. (1999). Transgenic nematodes carrying a cloned stress resistance gene from yeast. Surviv. Entomopathog. Nematodes. Off. Off. Publ. Eur. Commun. Luxemb., 105–119.
Wang, Y. and Gaugler, R. (1999). Steinernema glaseri surface coat protein suppresses the immune response of Popillia japonica (Coleoptera: Scarabaeidae) larvae. Biol. Control. 14, 45–50. doi: 10.1006/bcon.1998.0672
Wang, A., Tang, H., Sun, J., Wang, L., Rasmann, S., Ruan, W., et al. (2025). Entomopathogenic nematodes-killed insect cadavers in the rhizosphere activate plant direct and indirect defences aboveground. Plant Cell Environ. 48, 923–939. doi: 10.1111/pce.15193
Wu, H., Gong, Q., Fan, K., Sun, R., Xu, Y., and Zhang, K. (2017). Synergistic effect of entomopathogenic nematodes and thiamethoxam in controlling Bradysia odoriphaga Yang and Zhang (Diptera: Sciaridae). Biol. Control. 111, 53–60. doi: 10.1016/j.biocontrol.2017.05.006
Yadav, A. K. and Lalramliana (2012). Efficacy of indigenous entomopathogenic nematodes from Meghalaya, India against the larvae of taro leaf beetle, Aplosonyx chalybaeus (Hope). J. Parasit. Dis. 36, 149–154. doi: 10.1007/s12639-012-0139-7
Yukawa, T. and Pitt, J. M. (1985). Nematode storage and transport. WIPO Patent No. WO 85/03412. Geneva, CH: World Intellectual Property Organization.
Zhang, X., Li, L., Kesner, L., and Robert, C. A. M. (2021). Chemical host-seeking cues of entomopathogenic nematodes. Curr. Opin. Insect Sci. 44, 72–81. doi: 10.1016/j.cois.2021.03.011
Keywords: biological control, EPN, formulation, heterorhabditis, steinernema, stress tolerance mechanisms, sustainable agriculture, symbiotic bacteria
Citation: Kaur A, Sirengo DK, Karki P, Powers TO and Brown AMV (2026) Harnessing entomopathogenic nematodes for sustainable pest management: mechanisms, challenges, and innovations. Front. Plant Sci. 17:1755114. doi: 10.3389/fpls.2026.1755114
Received: 26 November 2025; Accepted: 12 January 2026; Revised: 10 January 2026;
Published: 29 January 2026.
Edited by:
Dun Wang, Northwest A&F University, ChinaReviewed by:
Rui Tang, Guangdong Academy of Science (CAS), ChinaArkadeb Mukhopadhyay, Central Island Agricultural Research Institute (ICAR), India
Copyright © 2026 Kaur, Sirengo, Karki, Powers and Brown. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.
*Correspondence: Amandeep Kaur, YW1hbmRrYXVAdHR1LmVkdQ==; David Kihoro Sirengo, ZHNpcmVuZ28yQGh1c2tlcnMudW5sLmVkdQ==