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Article

Distribution, Characterization, and Pathogenicity of Entomopathogenic Nematodes in Agricultural Crops in Amazcala, Querétaro

by
Gobinath Chandrakasan
1,*,
Mariana Beatriz Ávila López
2,3,
Markus Gastauer
4,
Genaro Martin Soto Zarazua
1,
Arantza Elena Sánchez Gutiérrez
1 and
Betsie Martinez Cano
1
1
Facultad de Ingeniería Campus Amazcala, Universidad Autónoma de Querétaro, Carr. Chichimequillas S/N Km 1, Amazcala, El Marqués 76265, Mexico
2
Comisión Intersecretarial de Bioseguridad de los Organismos Genéticamente Modificados (CIBIOGEM-SECIHTI), Av. Insurgentes Sur 1582, Ciudad de Mexico 03940, Mexico
3
Centro de Innovación para el Desarrollo Apícola Sustentable en Quintana Roo, Universidad Intercultural Maya de Quintana Roo, José María Morelos 77890, Mexico
4
Instituto Tecnológico Vale, R. Boaventura da Silva, 955, Bairro Nazaré, Belém CEP 66055-200, Pará, Brazil
*
Author to whom correspondence should be addressed.
Agriculture 2025, 15(15), 1603; https://doi.org/10.3390/agriculture15151603
Submission received: 31 May 2025 / Revised: 9 July 2025 / Accepted: 22 July 2025 / Published: 25 July 2025
(This article belongs to the Special Issue Advances in Biological Pest Control in Agroecosystems)
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Abstract

This study investigates the potential of entomopathogenic nematodes (EPNs) as biological control agents by exploring their occurrence and diversity in Amazcala, Querétaro. The aim was to characterise their distribution and evaluate their pathogenicity against insect pests. Soil samples were collected from various agricultural lands, followed by laboratory isolation and the molecular identification of EPN species. Morphological and genetic analyses confirmed the presence of several species with distinct pathogenic profiles. Pathogenicity assays using the larval stages of Galleria mellonella and Tenebrio molitor revealed that Heterorhabditis bacteriophora and Heterorhabditis atacamensis exhibited significant virulence, with Galleria mellonella being more susceptible. Among the 12 recovered EPN isolates, three strains—AMZX05 (Heterorhabditis atacamensis), AMZX10 (Heterorhabditis bacteriophora), and AMZX13 (Heterorhabditis atacamensis)—demonstrated particularly high pathogenic potential. These strains represent promising candidates for biological control and could contribute to sustainable integrated pest management (IPM) strategies. Further research is recommended to optimise their application across diverse agroecosystems.

1. Introduction

The growing concern over the environmental and health impacts caused by chemical pesticides has fuelled a global shift toward sustainable and eco-friendly pest control methods [1]. In this context, biological control agents, particularly entomopathogenic nematodes (EPNs), have emerged as a promising alternative [2,3]). EPNs, which are parasitic to insects, offer an effective and environmentally benign solution for managing insect pests across various ecosystems [4]. EPNs (Heterorhabditidae and Steinernematidae) inhabit the soil and infect different types of insects, including the larvae of Lepidoptera, Coleoptera, and Diptera, as well as immature and adult Orthoptera [5,6,7].
Species from the genera Heterorhabditis and Steinernema have gained particular attention for their ability to invade and propagate within insect hosts. Specifically, Heterorhabditis bacteriophora and Heterorhabditis atacamensis have demonstrated broad-spectrum pathogenicity, making them suitable candidates for pest control in agriculture, horticulture, and forestry [8]. They associate symbiotically with Photorhabdus bacteria, which release toxins to kill the host. These bacteria, once released into the insect hemocoel, cause rapid septicaemia, leading to host death in 24–72 h [9]. Heterorhabditis bacteriophora is widely distributed and known for its efficacy in controlling soil-dwelling pests such as root weevils, while H. atacamensis, a more recently described species, has shown promising results in arid environments [10,11].
EPNs have been isolated from all inhabited continents and many islands and occur in a range of ecologically diverse habitats from cultivated fields to deserts [12,13]. In Mexico, the isolation of native strains of Heterorhabditis and Steinernema, another major EPN genus, from local soils has been performed to enhance biological control efforts in specific agricultural zones. These crops face significant pest pressure from soil-dwelling and foliar insect pests, making EPNs a valuable tool for integrated pest management (IPM) strategies [14,15]. Studies on EPN ecology, mass production, and effectiveness in pest management have also been performed in various crops in Mexico [16]. However, the limiting factor in the use of these nematodes as biopesticides is the high mortality of infective juveniles (IJs) occurring within the first few days after application because of abiotic factors, such as high temperature, soil type, low moisture, UV light, and dehydration [17]. Although various EPN species have been reported worldwide, their natural occurrence, distribution, and pathogenic performance can vary considerably depending on local environmental conditions and agroecosystems. In Mexico, studies on native EPN populations remain limited, especially in regions such as Amazcala, Querétaro, where agricultural activity is prominent and sustainable pest control solutions are increasingly needed.
The use of model insect species such as (Pyralidae, Lepidoptera) Galleria mellonella (greater wax moth) and Tenebrio molitor (Tenebrionidae—Coleoptera: yellow mealworm) in research provides valuable insights into EPN efficacy. Both G. mellonella and T. molitor larvae are important agricultural and beekeeping pests that, due to their susceptibility to EPNs, make them ideal organisms to evaluate the pathogenic potential of species of Heterorhabditis spp. Given the rising demand for safer and more sustainable pest control methods, research on the isolation, characterisation, and pathogenicity of EPNs, such as H. bacteriophora and H. atacamensis, has become increasingly important [18]. This study aimed to explore the diversity and distribution of native EPNs in agricultural soils of Amazcala, and to assess their pathogenicity against two model insect hosts: Galleria mellonella and Tenebrio molitor. We hypothesised that native EPN isolates would exhibit species-specific differences in virulence and that some strains would demonstrate high pathogenic potential suitable for biocontrol applications.

2. Materials and Methods

2.1. Geography of Study Area and Sample Collection

Sampling was carried out at the Autonomous University of Queretaro, Natural Sciences Campus Amazcala, Queretaro, Mexico, which is a rich, fertile agricultural area in which different annual crops, including alfalfa, maize, sorghum, and avena, are grown. Amazcala is a town in the Queretaro state in Mexico. The town is located at latitude: 20°42′12.20″ N and longitude: −100°15′54.40″ W. The predominant climate is high-altitude subtropical, temperate semidry in 80% of the municipality, and the remaining 20% has a temperate humid climate. Soil samples were collected from various agricultural areas across 50 acres to monitor EPN persistence and abundance, with a minimum sampling distance of about 50 m between two samples. A total of 160 soil samples, 40 per soil type and crop, were collected from February 2023 to September 2024 (10-day intervals). The geographical location of the study area is shown on the map, and the physicochemical characteristics of the soil were analysed using samples collected at approximately 50 m intervals (Figure 1). Each soil sample (approximately 1 kg) was collected at a depth of 2–20 cm, placed in Zip-lock or plastic bags (labelled) to prevent water loss, transported to the laboratory, and stored at 12–15 °C until processing [19]. The abundance and distribution of EPNs were determined in different soil types: sandy soil, loamy soil, silt soil, sandy clay. For each soil sample, temperature, soil type, pH, and electrical conductivity (EC) were recorded. Specifically, soil pH and EC were measured using a soil-to-distilled water suspension at a 1:2.5 ratio (w/v). For each sample, 10 g of air-dried, sieved soil was mixed with 25 mL of distilled water and stirred thoroughly. The suspension was allowed to settle for 30 min. pH was measured using a calibrated digital pH meter (Oakton® pH 700 Benchtop pH Meter, Cole-Parmer, LLC—Vernon Hills, USA), and EC (Orion Lab Star EC112 conductivity bench meter, Thermo fisher scientific, CDMX, Mexico) was measured using a conductivity metre.

2.1.1. Rearing of G. mellonella and T. molitor (Model Host Insects)

Greater wax moth (G. mellonella) larvae were used as a model insect to isolate EPNs and evaluate their pathogenicity [20]. T. molitor was used as another model insect to evaluate pathogenicity. Wheat bran was purchased from the nearby market and sterilised in an autoclave at 121 °C for 30 min to remove microorganisms. For wheat bran to be suitable for Tenebrio larvae consumption, it was sieved through a mesh sieve (size 15), after being sterilised. The mealworm colony was cultured at 28 ± 5 °C, with a relative humidity of 70–75%, and a 10/14 h (light/dark) photoperiod in the growth chamber.

2.1.2. Insect Baiting Method

Under laboratory conditions, the collected soil samples were processed within one week using the insect-baiting method (Galleria trap). A 200 g subsample of soil was placed in a plastic box and baited with fifth instar larvae of G. mellonella [21]. The boxes were stored in a dark environment at 26 ± 2 °C to facilitate nematode infection. After five days, any dead larvae were collected and transferred to white traps to confirm the presence of EPNs and to harvest the emerging IJs [22]. The emergence of IJs from the cadavers typically occurred within 6 to 12 days, after which the nematodes settled in the water surrounding the white trap. To ensure the maximum recovery of EPNs from each soil sample, the baiting procedure was repeated three times with fresh G. mellonella larvae.

2.1.3. Survivability of Recovered Isolates at Different Storage Time Intervals at 20 °C

In this study, the survivability of the recovered isolates was evaluated at regular intervals over a period of 90 days following the method described by the Deol group [23]. The isolates, which consisted of IJs, were stored in distilled water containing 0.1% (v/v) formalin as a preservative, at a concentration of approximately 200–500 IJs/mL. The storage conditions were maintained in a controlled environment within tissue culture flasks at a constant temperature of 20 °C to minimise environmental fluctuations that could affect their longevity. At specified intervals, samples of approximately 100 IJs were extracted from each flask for survivability analysis. To determine whether the IJs remained viable using a stereoscopic microscope (Microscopios Nikon, LepsiPrisma, Microscopia, Monterry, Mexico), each one was probed with a fine needle. If an IJ exhibited movement or a response to the needle, it was classified as alive. In contrast, if the IJ did not react to the mechanical stimulation, it was deemed non-viable (dead). The total number of dead IJs was recorded, and the percentage of survivability was calculated for each sample by comparing the number of live individuals to the total number sampled.

2.1.4. Pathogenicity of Recovered EPNs over Time Against G. mellonella and T. molitor

Fourth-instar larvae of G. mellonella and T. molitor were collected and used to perform pathogenicity assays, as previously described by [24]. For the pathogenicity treatment, 10 larvae of G. mellonella and 20 g of sterile sand were placed in each experimental plate, and a specific volume of the EPN isolate suspension (25, 50, or 100 µL) containing 100 IJs/mL was added to each plate. This setup ensured that different EPN concentrations were tested against T. molitor, enabling an assessment of the dose-dependent virulence against the insect larvae. The treated plates were maintained at 25 °C ± 3 °C to mimic natural environmental conditions that favour EPN activity (triplicate was performed). After treatment, the larvae were monitored in 24 h intervals to observe the rate and extent of mortality. Observations were made every 24 h for a total period of 72 h, allowing for a time-based analysis of how quickly the nematodes can cause mortality in the larvae. Insect mortality was recorded at each time point, and the cumulative mortality was calculated at the end of the 72 h observation period. This helped explain both the rapidity and efficiency of the EPNs in killing the host insects. To ensure the reliability and reproducibility of the results, the entire experiment was repeated three times under the same conditions.

2.2. Scanning Electron Microscopy (SEM)

SEM was used to study the morphological features of first-generation and second-generation adults and IJs. Specimens were processed following [25]. After rinsing in Ringer’s solution, they were fixed in 3% glutaraldehyde buffered with sodium cacodylate at pH 7.2, post-fixed with osmium tetroxide, dehydrated through a graded ethanol series, and coated with a 200 nm gold layer. SEM imaging was conducted with the Hitachi SU8230 (Hitachi-shi, Japan) cold field emission (CFE) SEM/STEM microscope EVO-50.

2.3. Polymerase Chain Reaction (PCR) and Sequencing

For DNA extraction, two male specimens were cut into small pieces using a sterile scalpel to reduce the cell lysis time according to the protocol of the Quick-DNA™ Miniprep Plus Kit (Zymo Research-California, USA). The 28S gene of ribosomal DNA was amplified by PCR. For the PCR mixture, the following were added to each reaction: 12.5 µL of Green GoTaq Master Mix (Promega, Madison, WI, USA), 2 µL of primer, 8.5 µL of distilled water, and 2 µL of genomic DNA for a final volume of 25 µL. The primers used were 391F and 536R 5′-CAGCTATCCTGAGGGAAAC-3′ [26]. All PCRs were performed in a Qiagen™ Rotor-Gene® thermal cycler (Hesse, Germany). Subsequently, the following amplification conditions were used for 28S PCR: pre-denaturation at 94 °C for 5 min, 35 cycles of denaturation at 94 °C, annealing at 50 °C, extension at 72 °C for 1 min at each temperature, and final extension at 72 °C for 10 min. The PCR products were verified by electrophoresis on a 1% agarose gel containing 1x TAE buffer at 90 V for 45 min on a BioRad Sub-Cell®GT (California, USA) Agarose Gel Electrophoresis System using a 1 KB molecular weight Promega® DNA Marker as a reference. The PCR products were visualised on a BioDoc-It® Imager (California, USA) and commercially sequenced by Macrogen Inc. (Geumcheon-gu, Seoul, Republic of Korea). The 28S consensus sequences were then aligned to the sequences obtained for each primer using Geneious Pro-4.8.4® (Biomatters Ltd., Auckland, New Zealand).

2.4. In Vitro Mass Production of EPNs

Isolation and Cultivation of Symbiotic Photorhabdus spp. for EPN Production

Based on virulence and pathogenicity assessments, one potential EPN symbiotic bacteria Photorhabus species was extracted, cultured on NBTA plates, and incubated for 48 h at 30 °C. A single isolated colony of phase I bacteria was then transferred to 50 mL of STB medium [27] and incubated for 36 h at 30 °C with agitation at 150 rpm. Subsequently, 150 mL of production medium (PM) was inoculated with 5% (v/v) of the Photorhabus culture broth prepared in STB and incubated at 30 °C for 60 h with shaking at 150 rpm. The STB medium was prepared as follows: 3% (w/v) trypticase soy broth and 0.5% (w/v) yeast extract, adjusted to pH 7. This medium facilitated the growth of Photorhabus spp. for inoculating the nematode production medium [28]. The PM composition consisted of 2.3% (w/v) yeast extract, 1.25% (w/v) dried egg yolk, 0.5% (w/v) sodium chloride, and 4% (v/v) canola oil. Fermentation was carried out in specially designed shake flasks with modified shapes and sizes, as previously described by [29]. To initiate production, IJs were disinfected using 0.125% hyamine and rinsed thoroughly with sterilised tap water. The symbiotic bacterium (Photorhabdus spp.) was cultured for 36 h in 100 mL of STB medium, after which it was replaced with 200 mL of fresh production medium and incubated for an additional 48 h. Once the bacterial culture reached the stationary phase, containing approximately 3,000 IJs/mL, it was transferred to specially designed shake flask bioreactors for nematode production (10 bioreactors, each with 10 mL of disinfected IJ production medium). The efficiency of in vitro liquid culture for mass EPN production is highly dependent on physical and mechanical factors. Key parameters include the flask size, optimal temperature, agitation speed, and liquid medium volume, with slight modifications made during the process [30]. Each experiment was conducted in duplicate, with bioassays repeated four times. The results from duplicate assays were combined for a final statistical analysis.

2.5. Statistical Analysis

The values of survivability and pathogenicity assay were expressed as mean ± SD for three replicates. The difference in IJs survivability and pathogenicity was assessed by multifactorial ANOVA using STATGRAPHICS software 29 April 2025, is Version 19.7.01. ANOVA single analysis with Tukey’s post hoc test, (a) assuming there was no statistical significance, were performed. The number of specimens—first-generation male 10 and second-generation female 10—was measured.

3. Results

3.1. Distribution and Prevalence of EPNs

Out of 160 soil samples, 12 (7.5%) were identified as EPN-positive and were white-trapped using the Galleria baiting technique (Figure 1 and Figure 2). In the initial screening, based on EPN infectivity, the cadaver exhibited different dark black coloration for the 12 isolates (Figure 3): AMZX02, AMZX05, AMZX08, AMZX10, AMZX13, AMZX18, AMZX29, AMZX32, AMZX63, AMZX78, AMZ382, and AMZ91. Isolates obtained from infected cadavers that were dark red in colour resembled the typical morphological characteristics of the genus Heterorhabditis (Table 1). In the soil sample analysis, positive isolates were found in pH ranges of 6.74–8.32 with EC of 0.72–1.40% and organic matter contents of 3.5–4.6%. The overall survey data showed that the 12 isolates of Heterorhabditis spp. were common and distributed in all the sampling zones.

3.2. Survivability and Pathogenicity Percentage of Recovered Isolates at Different Storage Time Intervals Against G. mellonella and T. molitor

Significant differences were observed in the pathogenicity of H. bacteriophora and H. atacamensis across the two insect hosts. H. bacteriophora (AMZX 13) showed higher virulence in G. mellonella, which is consistent with previous studies highlighting this nematode’s effectiveness against soft-bodied insect pests (Figure 4). However, the mortality rate of H. bacteriophora (AMZX 13) proved more effective against G. mellonella than T. molitor, indicating that this species may have specialised adaptations for infecting coleopteran hosts (Figure 5). These findings have important implications for biological pest control. Depending on the target insect pest, one Heterorhabditis species may be preferable over another. For instance, H. bacteriophora could be deployed for pests similar to G. mellonella, while H. atacamensis could be better suited for coleopteran pests, such as T. molitor.

3.3. Morphometrics and Developmental Stages of H. atacamensis and H. bacteriophora

The morphometric features of isolates AMZX05 and AMZX10 identified them as H. atacamensis. The measurements for IJs are as follows in Table 2. Similarly, isolate AMZX13 was identified as H. bacteriophora (Table 3). Compared to H. atacamensis, H. bacteriophora males exhibit shorter spicules and a smaller gubernaculum (Figure 6). Supplementary Tables S1 and S2: General morphometrics and developmental stages of H. atacamensis and H. bacteriophora. This Supplementary Tables summarises the key morphometric parameters (e.g., body length, width, tail length, and oesophageal length) recorded across different developmental stages of H. atacamensis and H. bacteriophora. Measurements are presented as mean ± standard deviation, highlighting interspecific variation and developmental progression.

3.4. Molecular Characterisation

The molecular characterisation revealed that all the isolates exhibited the typical developmental patterns consistent with the genus Heterorhabditis, a well-known group of EPNs. Among the 12 isolates, a subset was further characterised based on their high survivability and pathogenicity, identifying them as potentially significant strains. Of these, two isolates, AMZX05 (accession number: PQ189283) and AMZX10 (accession number: PQ189703), were molecularly characterised as belonging to H. atacamensis (strains GChX5 and GChX10, respectively). Another isolation, AMZX13 (accession number: PQ189461), was identified as H. bacteriophora GChX13 (Figure 7). These findings were supported by BLAST (NCBI BLAST+ version 2.16.0) analysis, which revealed that the PCR products from the recovered samples shared more than 100% sequence similarity, indicating an exact match between the sequences. A phylogenetic tree was constructed using the ClustalW (Clustal W version 1.83) alignment algorithm (Slow/Accurate, IUB) to determine the relationship between these isolates.

3.5. Mass EPN Production

During the production process, IJs rapidly transitioned through developmental stages. On day 1, second-stage juveniles (IJ2) were predominant; by day 3, fourth-stage juveniles (J4) emerged, followed by young or hermaphroditic females on day 5. By day 6, adult nematodes were observed, and by day 8, egg-producing adults became dominant. The highest growth dynamics in the nematode population occurred around day 5, correlating with the emergence of reproductive females. At the end of the production period, the initial nep density was 3 × 103 IJs mL, and the final nep concentrations were as follows: A: 110 × 103 IJs/mL, B: 108 × 103 IJs/mL, and C: 109 × 103 IJs/mL, with 95% of them in the juvenile infective stage (IJ) (Figure 8).

4. Discussions

This study provides novel insights into the occurrence, diversity, and pathogenic potential of native entomopathogenic nematodes (EPNs) in the agricultural region of Amazcala, Querétaro. The findings support the potential integration of locally adapted EPNs into sustainable pest management strategies, offering a valuable alternative to synthetic pesticides. In Mexico, a country with high biodiversity and diverse agroclimatic regions, annual crops such as maize (Zea mays), avena (Avena sativa), sorghum (Sorghum bicolor), and alfalfa (Medicago sativa) are central to the agricultural economy. The ability of EPNs to persist in the soil and infect multiple larval instars makes them an attractive alternative to conventional insecticides. This work determined the molecular identification and phylogenetic diversity of native EPNs isolated from maize, avena, sorghum, and alfalfa crops, providing a detailed exploration of their ecological roles and potential in biocontrol. The isolation of twelve EPN strains from different agricultural fields confirms the natural presence and adaptability of these organisms to local edaphoclimatic conditions. Notably, species from the genus Heterorhabditis were more prevalent than Steinernema, which contrasts with findings in other regions of Mexico and Latin America where Steinernema often predominates. This may reflect the specific soil characteristics, vegetation type, and agricultural practices in Amazcala, suggesting that Heterorhabditis species may be better adapted to these local environments. Pathogenicity assays revealed differential virulence among the recovered strains. Heterorhabditis bacteriophora (AMZX10) and Heterorhabditis atacamensis (AMZX05 and AMZX13) caused significantly higher mortality in Galleria mellonella than in Tenebrio molitor due to its lack of strong immune defences and soft body tissue. The strong virulence shown by these strains, particularly H. atacamensis, is noteworthy, as this species has been underexplored and may offer new avenues for biocontrol. The mechanisms of the pathogenicity of EPNs from natural habitats typically involve baiting techniques, where susceptible insect hosts such as G. mellonella larvae are exposed to soil samples to attract nematodes. These nematodes enter the insect hosts, releasing symbiotic bacteria that kill the host, allowing the nematodes to reproduce and emerge. Both H. bacteriophora and H. atacamensis have been successfully isolated using such methods from diverse soil types, particularly from regions with variable climatic conditions [31,32].
Characterising Heterorhabditis species involves both morphological and molecular techniques. Morphologically, the IJs of H. bacteriophora and H. atacamensis are similar. However, in this study, the isolated strains of AMZX05, AMZX10, and AMZX13 from H. bacteriophora were smaller in body length compared to H. Atacamensis, as reported by [29]. As for the male specimens, distinct traits, such as the shape and structure of the spicules in adult males, caudal bursa, and the pattern of papillae, can differentiate species since they are key taxonomic characteristics [33]. Molecular characterisation typically involves the analysis of ribosomal RNA genes, particularly the 18S, 28S, and ITS (Internal Transcribed Spacer) regions, which provide clear genetic distinctions between species [34,35]. These characterisation techniques provide critical insights into the life cycle, infective potential, and ecological adaptability of the nematodes, making them valuable for biological control programmes. The genetic variability observed within these EPN populations highlights their ability to adapt to specific environmental factors, such as soil composition, climatic conditions, and the availability of insect hosts. Investigating the phylogenetic relationships of EPNs is crucial for deciphering their evolutionary pathways and ecological functions. In Mexico, EPN diversity is likely influenced by the nation’s diverse agricultural landscapes and the unique insect species associated with different annual crops. Phylogenetic studies frequently reveal species clustering based on geographic regions or host crops, suggesting processes of localised evolution and ecological adaptation.
Assessing the pathogenicity of H. bacteriophora and H. atacamensis involves testing their ability to infect and kill insect hosts under laboratory and field conditions. G. mellonella larvae are commonly used as model organisms due to their high susceptibility to EPN infection. H. bacteriophora, for instance, has demonstrated high pathogenicity against G. mellonella, killing the larvae within 48–72 h under optimal laboratory conditions. The nematodes invade the insect’s hemocoel, releasing symbiotic Photorhabdus bacteria, which leads to septicaemia and rapid host death. In contrast, T. molitor exhibits slightly greater resistance to EPN infection. Although H. bacteriophora is still effective, it often takes longer for infection to result in mortality, likely due to the mealworm’s thicker cuticle and different immune responses [36]. Studies have shown that H. atacamensis, which has been isolated from more arid environments, may have evolved specific adaptations that allow it to infect insect hosts in harsher conditions, which could extend its utility as a biocontrol agent in areas with extreme temperatures or a lower moisture content [37]. The rapid host mortality and significant nematode reproduction within the host cadavers emphasise the capacity of these species to establish and persist in the field, thereby reducing the need for repeated applications—a key advantage in IPM programmes.
The efficacy of EPNs as biocontrol agents depends on several factors, such as environmental conditions, host susceptibility, and nematode virulence. For example, temperature and soil moisture have a significant impact on the survival and infectivity of EPNs. Studies have shown that H. bacteriophora thrives in moderately moist soils, with optimal temperatures ranging from 25 to 30 °C [38]. H. atacamensis, isolated from drier environments, demonstrates a greater tolerance to lower moisture levels and higher temperatures, making it a more versatile agent in regions with extreme conditions. However, the susceptibility of the target insect host also plays a role. For instance, G. mellonella is widely recognised as a highly susceptible host, making it an excellent model for testing the virulence of EPN species. T. molitor, although still susceptible, often requires more nematodes for effective control due to its tougher exoskeleton. The virulence of different Heterorhabditis species and strains can vary. H. bacteriophora is considered a highly virulent species, while H. atacamensis, though less studied, shows promise for its ability to infect hosts under challenging environmental conditions. Their virulence is closely tied to the potency of their symbiotic bacteria, Photorhabdus, which produces a variety of toxins and enzymes that aid in host degradation and the suppression of the insect immune system [39]. Consequently, both H. bacteriophora and H. atacamensis have proven effective in laboratory settings, but they exhibit differences in environmental adaptability and virulence across host species. Although H. bacteriophora has a broad geographical distribution and is widely studied for use in biocontrol, H. atacamensis, which was recently discovered in more arid regions, has the potential to expand the range of environments where EPNs can be used effectively. Their ability to target different insect pests, including G. mellonella and T. molitor, highlights their versatility as biocontrol agents. The integration of EPNs in these annual crops aligns with sustainable agricultural practices, reducing reliance on synthetic pesticides while maintaining crop productivity. Further research on formulation technologies, application timing, and synergistic interactions with other biocontrol agents will enhance the adoption of EPNs in annual cropping systems.
Globally, our results contribute to the growing body of evidence supporting the use of EPNs as effective, environmentally safe biological control agents. The demonstrated pathogenicity, infectivity, and reproductive potential of H. bacteriophora and H. atacamensis reaffirm the role of EPNs as viable alternatives to chemical pesticides, aligning with international efforts to reduce pesticide usage, mitigate resistance development, and preserve non-target organisms and soil health. For IPM, our study offers valuable insight into the potential inclusion of native and climate-adapted EPNs within diversified pest control programmes [40]. Their compatibility with other biological and cultural practices enhances the robustness and sustainability of IPM frameworks across various cropping systems. From a sustainable agriculture perspective, our findings support a shift toward ecologically grounded pest management strategies that maintain productivity while preserving ecosystem services. By highlighting the potential of a lesser-known native species like H. atacamensis, we also underscore the importance of exploring and utilising local biodiversity to develop context-specific, scalable solutions for pest control.
Our findings provide valuable insight into the pathogenicity, virulence, and reproductive potential of these entomopathogenic nematodes’ integration into sustainable pest management strategies. The comparative pathogenicity trials revealed that both H. bacteriophora and H. atacamensis exhibit high infectivity and virulence against G. mellonella, resulting in their potential as reliable bio-insecticides. Interestingly, H. atacamensis, a species adapted to arid environments, also demonstrated strong pathogenicity against T. molitor, suggesting it could be particularly valuable for pest control in dryland and semi-arid agricultural ecosystems. In the future, research will evaluate the field performance and persistence of these nematodes under various climatic and soil conditions.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/agriculture15151603/s1, Supplementary Table S1. Morphometric measurements of IJs and adults of H. atacamensis isolates AMZX05 and AMZX10 (all values in µm). Supplementary Table S2. Morphometric measurements of IJs and adult males of H. bacteriophora isolate AMZX13 (all values in µm).

Author Contributions

G.C.: Writing—review and editing, writing—original draft. M.B.Á.L. and M.G.: review and editing, visualisation, validation, methodology, data curation. G.M.S.Z., A.E.S.G., and B.M.C.: supervision, resources, conceptualization. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding.

Institutional Review Board Statement

Not applicable.

Data Availability Statement

The original contributions presented in this study are included in the article. Further inquiries can be directed to the corresponding author.

Acknowledgments

The authors thank the technical assistance of Manuel Aguilar Franco from the LaNCaM, at the centre for Applied Physics and Advanced Technology (CFATA), UNAM campus Juriquilla, for the availability to carry out the microscopy characterisation analyses.

Conflicts of Interest

The authors declare no conflicts of interest.

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Agriculture 15 01603 g001
Figure 1. Annual crops, such as alfalfa, maize, sorghum, and avena, at the location of the Natural Sciences Campus of the Autonomous University of Queretaro (Amazcala campus), Queretaro state, Mexico.
Figure 1. Annual crops, such as alfalfa, maize, sorghum, and avena, at the location of the Natural Sciences Campus of the Autonomous University of Queretaro (Amazcala campus), Queretaro state, Mexico.
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Figure 2. Soil sample processing and entomopathogenic nematode (EPN) extraction using G. mellonella and T. molitor as bait insects. (A) Healthy G. mellonella larvae used as bait; (B) G. mellonella eggs placed on artificial diet for rearing; (C) G. mellonella larvae actively feeding in the diet; (D) G. mellonella pupae stage; (E) adult G. mellonella moths emerging from pupae; (F) T. molitor larvae growing in artificial diet; (G) healthy T. molitor larvae used for baiting; (H) soil sample contained in a plastic container with perforated lid to allow aeration; (I) setup of Galleria baiting method, showing larvae placed in soil samples to detect EPN presence.
Figure 2. Soil sample processing and entomopathogenic nematode (EPN) extraction using G. mellonella and T. molitor as bait insects. (A) Healthy G. mellonella larvae used as bait; (B) G. mellonella eggs placed on artificial diet for rearing; (C) G. mellonella larvae actively feeding in the diet; (D) G. mellonella pupae stage; (E) adult G. mellonella moths emerging from pupae; (F) T. molitor larvae growing in artificial diet; (G) healthy T. molitor larvae used for baiting; (H) soil sample contained in a plastic container with perforated lid to allow aeration; (I) setup of Galleria baiting method, showing larvae placed in soil samples to detect EPN presence.
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Figure 3. White-trap method. EPN-infected larvae cadaver in white trap. (A) H. bacteriophora and (B) H. atacamensis; (C) EPN outcome in the larvae; (D) EPN microscopic view.
Figure 3. White-trap method. EPN-infected larvae cadaver in white trap. (A) H. bacteriophora and (B) H. atacamensis; (C) EPN outcome in the larvae; (D) EPN microscopic view.
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Figure 4. Percentage of survivability at different storage time intervals (days) against G. mellonella (IJs per mL). Twelve recovered EPNs were tested—AMZX02, AMZX05, AMZX08, AMZX10, AMZX13, AMZX18, AMZX29, AMZX32, AMZX63, AMZX78, AMZX82, and AMZX91. Among the 12, the following 3 EPNs have potential survivable rates from 10 to 90 days—AMZX05—Heterorhabditis atacamensis; AMZX10—Heterorhabditis bacteriophora; AMZX13—Heterorhabditis atacamensis.
Figure 4. Percentage of survivability at different storage time intervals (days) against G. mellonella (IJs per mL). Twelve recovered EPNs were tested—AMZX02, AMZX05, AMZX08, AMZX10, AMZX13, AMZX18, AMZX29, AMZX32, AMZX63, AMZX78, AMZX82, and AMZX91. Among the 12, the following 3 EPNs have potential survivable rates from 10 to 90 days—AMZX05—Heterorhabditis atacamensis; AMZX10—Heterorhabditis bacteriophora; AMZX13—Heterorhabditis atacamensis.
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Figure 5. Three-day mortality rates (%) of entomopathogenic nematode (EPN) isolates against (A) G. mellonella larvae and (B) T. molitor larvae at three concentrations: 25, 50, and 100 (IJs)/mL. AMZX02, AMZX05, AMZX08, AMZX10, AMZX13, AMZX18, AMZX29, AMZX32, AMZX63, AMZX78, AMZX82, and AMZX91. Among 12 species, the following EPNs showed virulent activity: AMZX05—Heterorhabditis atacamensis; AMZX10—Heterorhabditis bacteriophora; AMZX13—Heterorhabditis atacamensis.
Figure 5. Three-day mortality rates (%) of entomopathogenic nematode (EPN) isolates against (A) G. mellonella larvae and (B) T. molitor larvae at three concentrations: 25, 50, and 100 (IJs)/mL. AMZX02, AMZX05, AMZX08, AMZX10, AMZX13, AMZX18, AMZX29, AMZX32, AMZX63, AMZX78, AMZX82, and AMZX91. Among 12 species, the following EPNs showed virulent activity: AMZX05—Heterorhabditis atacamensis; AMZX10—Heterorhabditis bacteriophora; AMZX13—Heterorhabditis atacamensis.
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Figure 6. Scanning electron microscopy images of Heterorhabditis bacteriophora and Heterorhabditis atacamensis. Note: (A–C) IJs of Heterorhabditis bacteriophora. (D,E) Heterorhabditis atacamensis—tail of hermaphroditic female and amphimictic female, respectively. (F) Heterorhabditis bacteriophora—tail of hermaphroditic female and amphimictic female, respectively. (G) Heterorhabditis atacamensis—excretory pore. (H,I) Heterorhabditis atacamensis—lip region of hermaphroditic female and amphimictic male, respectively.
Figure 6. Scanning electron microscopy images of Heterorhabditis bacteriophora and Heterorhabditis atacamensis. Note: (A–C) IJs of Heterorhabditis bacteriophora. (D,E) Heterorhabditis atacamensis—tail of hermaphroditic female and amphimictic female, respectively. (F) Heterorhabditis bacteriophora—tail of hermaphroditic female and amphimictic female, respectively. (G) Heterorhabditis atacamensis—excretory pore. (H,I) Heterorhabditis atacamensis—lip region of hermaphroditic female and amphimictic male, respectively.
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Figure 7. (A) Heterorhabditis atacamensis GChX5 (AMZX05) ♦ and (B) Heterorhabditis atacamensis GChX10 (AMZX10) ♦ and (C) Heterorhabditis bacteriophora GChX13 (AMZX13) ♦.
Figure 7. (A) Heterorhabditis atacamensis GChX5 (AMZX05) ♦ and (B) Heterorhabditis atacamensis GChX10 (AMZX10) ♦ and (C) Heterorhabditis bacteriophora GChX13 (AMZX13) ♦.
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Figure 8. In vitro mass production of Photorhabdus spp.: Bacterial growth curve showing the change in Photorhabdus spp. cell density (measured as optical density at 600 nm or CFU/mL) over time during different phases of the production process. The timeline includes key stages such as initial culture in STB medium, transfer to production medium (PM), and incubation phases. The data illustrate bacterial growth kinetics over a 60 h fermentation period under controlled shaking and temperature conditions.
Figure 8. In vitro mass production of Photorhabdus spp.: Bacterial growth curve showing the change in Photorhabdus spp. cell density (measured as optical density at 600 nm or CFU/mL) over time during different phases of the production process. The timeline includes key stages such as initial culture in STB medium, transfer to production medium (PM), and incubation phases. The data illustrate bacterial growth kinetics over a 60 h fermentation period under controlled shaking and temperature conditions.
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Table 1. Occurrence and distributions of EPNs and their soil characteristics of different sampling zones in Amazcala, Queretaro, Mexico.
Table 1. Occurrence and distributions of EPNs and their soil characteristics of different sampling zones in Amazcala, Queretaro, Mexico.
Soil SampleSoil TypeRecovered EPNs (Heterorhabditis sp.) Soil Temperature Organic Contact (%) pHElectrical Conductivity (mS/cm)Total (%) (Heterorhabditis sp.)
Avenaclay++313.67.130.837.5%
loam+283.38.131.19
sandy loam++293.96.900.82
Cornsilt soil+284.26.900.86
loam+313.87.011.23
loamy clay+293.77.130.83
Sorghumloam+293.46.780.78
loam+303.08.310.91
Alfalfasilt soil+274.126.820.92
clay+304.07.130.71
++ shows two positive isolates of Heterorhabditis sp. in the different soil samples; + shows one positive isolate of Heterorhabditis sp. in the different soil samples.
Table 2. Morphometric measurements of H. bacteriophora IJ and H. atacamensis IJs (µm). Measurements are in µm and in the form of mean ± standard deviation (range).
Table 2. Morphometric measurements of H. bacteriophora IJ and H. atacamensis IJs (µm). Measurements are in µm and in the form of mean ± standard deviation (range).
Recovered EPNsMorphometric Characteristics
TBLMBWEPESTLD%E%
AMZX05 1535 ± 10 a20 ± 1 a120 ± 2 a118 ± 12 a90 ± 9 a84 ± 4 a 101 ± 7 a
AMZX10 1539 ± 16 a20 ± 1 a120 ± 1 a118 ± 12 a89 ± 11 a84 ± 5 a104 ± 9 a
AMZX13 2542 ± 8 a20 ± 1 a121 ± 2 a119 ± 13 a93 ± 8 a86 ± 4 a103 ± 7 a
H. bacteriophora
[28] 1		527 ± 620 ± 1119 ± 2110 ± 1084 ± 1086 ± 5101 ± 6
H. atacamensis
[29] 2		529 ± 1221 ± 1118 ± 2115 ± 1179 ± 1285 ± 698 ± 8
TBL = total body length; MBW = maximum body width; EP = distance from the head to the excretory pore; ES = distance from the head to the base of the oesophagus; TL = total body length/distance from the head to the oesophagus; D% = distance from the head to the excretory pore/distance from the head to the base of the oesophagus × 100; E% = distance from the head to the excretory pore/length of tail × 100. Ten IJ specimens measured. H. atacamensis (AMZX05 1 and AMZX10 1) is the same species with different strains, H. bacteriophora strain AMZX13 2. Values are presented as mean ± SE. Superscript letters (a) within the same row indicate statistical comparison using [Tukey’s HSD]. Means sharing the same letter are not significantly different at p > 0.05. In this case, all values share the same letter, indicating no significant differences. Superscripts 1 and 2 indicate measurements corresponding to two different entomopathogenic nematode species: 1 Heterorhabditis bacteriophora, 2 Heterorhabditis atacamensis.
Table 3. Morphometric measurements of H. bacteriophora males and H. atacamensis females (µm). Measurements are in µm and in the form of mean ± standard deviation (range).
Table 3. Morphometric measurements of H. bacteriophora males and H. atacamensis females (µm). Measurements are in µm and in the form of mean ± standard deviation (range).
Recovered EPNsMorphometric Characteristics
ABWSPLGuLTBLMBWEPTLES
AMZX05 121 ± 3 a44 ± 4 a20 ± 1 a906 ± 36 a104 ± 7 a118 ± 5 a30 ± 1 a109 ± 3 a
AMZX10 121 ± 3 a44 ± 5 a20 ± 2 a916 ± 34 a103 ± 6 a119 ± 6 a29 ± 2 a106 ± 6 a
AMZX13 221 ± 2 a45 ± 6 a20 ± 1 a914 ± 37 a102 ± 6 a121 ± 8 a29 ± 2 a106 ± 6 a
H. bacteriophora
[28] 1		19 ± 3 44 ± 421 ± 1898 ± 32105 ± 5122 ± 432 ± 199 ± 7
H. atacamensis
[29] 2		22 ± 245 ± 420 ± 2918 ± 30106 ± 6124 ± 729 ± 2108 ± 4
ABW = anal body width; SPL = spicule length; GuL = gubernaculum length; TBL = total body length; MBW = maximum body width; EP = excretory pore distance from anterior end; TL = tail length; ES = distance from the head to the base of the oesophagus. H. atacamensis (AMZX05 1 and AMZX10 1) is the same species with different strains, H. bacteriophora strain AMZX13 2. Values are presented as mean ± SE. Superscript letters (a) within the same row indicate statistical comparison using [Tukey’s HSD]. Means sharing the same letter are not significantly different at p > 0.05. In this case, all values share the same letter, indicating no significant differences. Superscripts 1 and 2 indicate measurements corresponding to two different entomopathogenic nematode species: 1 Heterorhabditis bacteriophora, 2 Heterorhabditis atacamensis.
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Chandrakasan, G.; Ávila López, M.B.; Gastauer, M.; Soto Zarazua, G.M.; Sánchez Gutiérrez, A.E.; Martinez Cano, B. Distribution, Characterization, and Pathogenicity of Entomopathogenic Nematodes in Agricultural Crops in Amazcala, Querétaro. Agriculture 2025, 15, 1603. https://doi.org/10.3390/agriculture15151603

AMA Style

Chandrakasan G, Ávila López MB, Gastauer M, Soto Zarazua GM, Sánchez Gutiérrez AE, Martinez Cano B. Distribution, Characterization, and Pathogenicity of Entomopathogenic Nematodes in Agricultural Crops in Amazcala, Querétaro. Agriculture. 2025; 15(15):1603. https://doi.org/10.3390/agriculture15151603

Chicago/Turabian Style

Chandrakasan, Gobinath, Mariana Beatriz Ávila López, Markus Gastauer, Genaro Martin Soto Zarazua, Arantza Elena Sánchez Gutiérrez, and Betsie Martinez Cano. 2025. "Distribution, Characterization, and Pathogenicity of Entomopathogenic Nematodes in Agricultural Crops in Amazcala, Querétaro" Agriculture 15, no. 15: 1603. https://doi.org/10.3390/agriculture15151603

APA Style

Chandrakasan, G., Ávila López, M. B., Gastauer, M., Soto Zarazua, G. M., Sánchez Gutiérrez, A. E., & Martinez Cano, B. (2025). Distribution, Characterization, and Pathogenicity of Entomopathogenic Nematodes in Agricultural Crops in Amazcala, Querétaro. Agriculture, 15(15), 1603. https://doi.org/10.3390/agriculture15151603

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