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Article

Evaluation of Entomopathogenic Nematodes against Common Wireworm Species in Potato Cultivation

by
Arife Gümüş Askar
1,*,
Ebubekir Yüksel
2,*,
Refik Bozbuğa
3,
Atilla Öcal
4,
Halil Kütük
5,
Dilek Dinçer
6,
Ramazan Canhilal
2,
Abdelfattah A. Dababat
7 and
Mustafa İmren
5
1
Istanbul Directorate of Agricultural Quarantine, Bakırköy, 34149 Istanbul, Türkiye
2
Department of Plant Protection, Faculty of Agriculture, Kayseri Erciyes University, Melikgazi, 38030 Kayseri, Türkiye
3
Department of Plant Protection, Faculty of Agriculture, Eskişehir Osmangazi University, Odunpazarı, 26160 Eskişehir, Türkiye
4
Atatürk Horticultural Central Research Institute, Merkez, 77100 Yalova, Türkiye
5
Department of Plant Protection, Faculty of Agriculture, Bolu Abant Izzet Baysal University, Gölköy, 14030 Bolu, Türkiye
6
Biological Control Research Institute, Yüreğir, 01321 Adana, Türkiye
7
International Maize and Wheat Improvement Centre (CIMMYT) 39, Emek, 06511 Ankara, Türkiye
*
Authors to whom correspondence should be addressed.
Pathogens 2023, 12(2), 288; https://doi.org/10.3390/pathogens12020288
Submission received: 4 January 2023 / Revised: 13 January 2023 / Accepted: 18 January 2023 / Published: 9 February 2023
(This article belongs to the Special Issue Entomopathogenic Fungi and Nematodes in Modern Agriculture)
Browse Figure
Review Reports Versions Notes

Abstract

:
Wireworms (Coleoptera: Elateridae) are common insect pests that attack a wide range of economically important crops including potatoes. The control of wireworms is of prime importance in potato production due to the potential damage of the larvae to tuber quantity and quality. Chemical insecticides, the main control strategy against wireworms, generally fail to provide satisfactory control due to the lack of available chemicals and the soil-dwelling habits of the larvae. In the last decades, new eco-friendly concepts have emerged in the sustainable control of wireworms, one of which is entomopathogenic nematodes (EPNs). EPNs are soil-inhabitant organisms and represent an ecological approach to controlling a great variety of soil-dwelling insect pests. In this study, the susceptibility of Agriotes sputator Linnaeus and A. rufipalpis Brullé larvae, the most common wireworm species in potato cultivation in Türkiye, to native EPN strains [Steinernema carpocapsae (Sc_BL22), S. feltiae (Sf_BL24 and Sf_KAY4), and Heterorhabditis bacteriophora (Hb_KAY10 and Hb_AF12)] were evaluated at two temperatures (25 and 30 °C) in pot experiments. Heterorhabditis bacteriophora Hb_AF12 was the most effective strain at 30 °C six days post-inoculation and caused 37.5% mortality to A. rufipalpis larvae. Agriotes sputator larvae were more susceptible to tested EPNs at the same exposure time, and 50% mortality was achieved by two EPNs species, Hb_AF12 and Sc_BL22. All EPN species/strains induced mortality over 70% to both wireworm species at both temperatures at 100 IJs/cm2, 18 days post-treatment. The results suggest that tested EPN species/strains have great potential in the control of A. sputator and A. rufipalpis larvae.

1. Introduction

Wireworms, Agriotes spp. (Coleoptera: Elateridae), are one of the most destructive soil-dwelling insect pests that can cause severe economic losses in many crops including potato [1,2]. Wireworms generally live under soil during their larval development and feed on underground parts of plants such as seeds, roots, and tubers, leading to severe yield and tuber quality losses [3]. In addition to direct damage to tubers, the feeding holes of wireworm larvae on tubers predispose plants to subsequent secondary infections by other invertebrates and microbial pathogens [4]. Wireworm damage leads frequently to a drop in market value due to the visual quality of potatoes. In the absence of appropriate control measures, nearly half of the production can be downgraded in terms of quality [5]. The average price reductions due only to wireworm damage were estimated at 12% [4]. Therefore, potato crops are at greater risk for wireworm damage and require particular attention because of the damage potential of wireworms that render the tubers unmarketable [6]. Up to now, more than 39 wireworm species have been reported to attack potatoes around the world [7]. However, 9 species, Agriotes brevis Candèze, Agriotes lineatus Linnaeus, Agriotes litigiosus Rossi, Agriotes obscurus Linnaeus, Agriotes proximus Linnaeus, Agriotes rufipalpis Brullé, Agriotes sordidus Illiger, Agriotes sputator Linnaeus, and Agriotes ustulatus Schäller, are considered the most devastating in Europe [8,9].
The control of wireworms is quite challenging due to their extensive and hidden life cycle in the soil, and non-uniform distribution of the larvae in the fields [10]. For many years, chemical insecticides with broad-spectrum active ingredients such as organochlorine, organophosphates, and carbamates have been the main strategy to control wireworms by most producers. However, restrictions and bans in recent decades on the use of many of these synthetic chemicals, due to environmental and health concerns, have led many researchers to search for eco-friendly alternatives to synthetic insecticides for the management of wireworm populations [11,12,13,14]. Among new ecologically based approaches, entomopathogenic nematodes (EPNs) (Steinernematidae and Heterorhabditidae) have recently attracted a great deal of attention due to their biocontrol potential against many economically important insect pests [15,16,17]. Being soil-originated organisms, EPNs are natural suppressors of insect pest populations in the soil environment [18,19]. The infective juveniles (IJs) of Steinernema (Steinernematidae) and Heterorhabditis (Heterorhabditidae) species, stress-resistant stage surrounded by a protective sheath, seek out a potential host in soil without feeding and initiate the infection process by penetrating their hosts via body openings (mouth, anus, or spiracles) or cuticle [20,21]. Following penetration, the IJs move toward the host hemocoel and release their endosymbiotic bacteria (Xenorhabdus spp. and Photorhabdus spp., respectively) into the host hemolymph which induces toxemia and kills the host within 24–48 h [22,23]. The IJs feed on the bacterial cells and degraded host tissues through several generations until the depletion of food sources and then leave the host dead body to search for a new potential host [20].
The soil-dwelling characteristics and an efficient host-searching mechanism of EPNs make them a perfect candidate for biological control of soil-borne insect pests such as wireworms [24]. However, the success of EPNs against any kind of insect pest is heavily dependent on the adaptation capability of EPNs in the application area and matching the most appropriate EPN species/strains with the target pests [25]. In general, local EPNs are considered well-adapted to climatic and environmental conditions where they are isolated and can effectively suppress the pest populations without adverse effects on non-target species [26,27,28,29]. Therefore, in a previous study, a wide-ranging field survey was conducted in major potato cultivation areas of Türkiye to identify EPN species for the control of major potato pests [30]. In the present study, the effectiveness of isolated EPN species/strains was evaluated against the larvae of A. sputator and A. rufipalpis which are the two predominant wireworm species in potato growing areas of Türkiye [31].

2. Materials and Methods

2.1. Source of Nematodes

Three EPN species that were previously recovered from potato fields were employed in in vitro bioassays [30] (Table 1). In order to obtain a fresh batch of IJs, EPN species were multiplied in vitro on the last instar larvae of Galleria mellonella (L.) (Lepidoptera: Pyralidae) [25]. The IJs were suspended in 1 mL of sterile water at a concentration 200 IJs/Petri dish and inoculated to Petri plates (Ø9 cm) with autoclaved sandy loamy soil (20 g). The Petri plates were covered with parafilm and maintained at 25 °C and 65% relative humidity (RH). Dead Galleria larvae were collected daily with soft forceps and placed into modified white traps [32]. The harvested IJs were washed several times with sterile water and stored at 15 °C horizontally in cell culture flasks (250 mL) until bioassays were performed. The initial culture of G. mellonella larvae was obtained from the Entomology Laboratory of Erciyes University and reared on an artificial diet as described by Metwally et al. [33]. The larvae were reared in glass wide-neck jars (1 L) which were sterilized by autoclaving. The diet consisted of the following ingredients: wheat flour, wheat bran, milk powder, maize flour, dried yeast powder, honey, and glycerin. Approximately 100 1st instar larvae were put into each jar and the jars were maintained under laboratory conditions (30 °C and 65% relative humidity). The diet was refreshed every 20 days until the last instar larvae were obtained [33].

2.2. Source of Wireworms

Agriotes rufipalpis and A. sputator larvae were assembled from potato fields in different parts of Türkiye and identified based on the mitochondrial cytochrome c oxidase subunit I (COI) sequences in a previous study [31]. The collected larvae were brought individually to the laboratory in plastic containers (50 mL) containing autoclaved sandy loamy soil (20 g) with a slice of potato (Ø2 cm) and kept at 25 °C. The 4th and 5th larvae were separated according to their head width and observed for one week for any sign of infection [9,34,35]. Only healthy larvae were included in the pathogenicity bioassays. The pathogenicity of EPNs was tested against the mixed groups of 4th and 5th larval instars.

2.3. Evaluation of the Pathogenicity of EPNs in Pot Experiment

The effectiveness of EPNs was evaluated at two temperatures (25 and 30 °C) in pots (1.5 L) (Surface area 240 cm2) including 1 kg autoclaved (121 °C for 60 min) sandy loamy soil and a slice of fresh potato (Ø2 cm). The soil used in our experiment was obtained from Bolu Abant İzzet Baysal University (Department of Seed Science and Technology) and consisted of 81% sand, 14% silt, and 5% clay. The organic matter content of the soil was 2.1% with a pH of 6.5. Prior to the inoculation of IJs, each pot was irrigated with 100 mL of distilled water to provide an adequate amount of moisture for IJs. One larva of 4th or 5th instar was placed into pots and allowed to move deeper in the soil profile. The IJs suspended in 5 mL distilled water were applied uniformly to the soil surface at the concentration of 25, 50, 100, and 150 IJs/cm2 with the help of an automatic pipette (corresponding to 6000, 12,000, 24,000, and 36,000 IJs per pot, respectively). Then, the pots were maintained at 25 °C and 30 °C, 65% RH. Since moisture is one of the most important factors affecting the movement of IJs, the pots were irrigated with 50 mL of distilled water daily during the experiment. The larval mortality was checked at 6 and 18 days after treatment. Dead larvae that were transferred to white traps were observed under a stereomicroscope to confirm the nematode infection. There were four replicates of each treatment with ten larvae per replicate. All experiments were repeated twice on different dates.

2.4. Statistical Analysis

No mortality occurred in the control groups. Data pooled from two experiments were analyzed using IBM SPSS Statistics, Version 20.0 for Windows (SPSS Inc., Chicago, IL, USA). Prior to analyses, a normality test was performed and data were subjected to arcsine transformation. To determine significant differences among treatments, full-factorial model repeated-measures ANOVA was applied. The effects of the main factors (Nematode, Temperature, and IJs concentrations) and their interactions were considered significant at α = 0.05. Post-hoc comparisons were performed using Tukey’s multiple comparison test (p ≤ 0.05).

3. Results

The results revealed that the larvae of A. rufipalpis and A. sputator showed varying degrees of susceptibility to all tested EPN species and strains (Figure 1). All main factors had a significant effect on the mortality rates of both wireworm species (Table 2). Increasing concentrations of IJs and exposure time generally led to higher mortalities in the A. rufipalpis larvae, and mortality rates ranged between 2.5 and 37.5% at both temperatures tested. The susceptibility of A. rufipalpis larvae to EPNs tended to increase at 30 °C, and three EPN species/strains induced mortality over 30% at 6 days after treatment (DAT). Heterorhabditis bacterophora AF12 was the most efficient strain at 6 DAT and yielded 37.5% mortality at the highest concentration (150 IJs/cm2) (Table 3).
At the lowest concentration, only two H. bacteriophora isolates (Hb_KAY10 and Hb_AF12) were able to cause mortalities of over 40% at 18 DAT. A remarkable increase occurred in the mortality rates of A. rufipalpis larvae at 100 and 150 IJs/cm2 concentrations at 30 °C, and all EPN species/strains induced mortality of over 80% (Table 4). All EPN species/strains were able to kill the larvae of A. sputator at tested IJ concentrations and temperatures except for S. feltiae BL24 and H. bacteriophora KAY10 which induced no mortality at 25 IJs/cm2 at 25 °C. However, with increasing IJ concentrations, H. bacteriophora KAY10 yielded the highest mortality (47.5%) at 100 and 150 Ijs/cm2 at 25 °C and 6 DAT followed by H. bacteriophora AF12 (45.0%) and S. feltiae KAY4 (42.5%). Although S. feltiae BL22 was the least efficient isolate at 25 °C, the efficiency of BL22 significantly increased at 30 °C and achieved the highest mortality (50.0%) at 6 DAT along with H. bacteriophora KAY10 (Table 5).
After 18 days of exposure to the Ijs, the larval mortality substantially increased and all H. bacteriophora strains caused mortality over 50% at the lowest concentration (25 Ijs/cm2) at 25 °C. At 100 Ijs/cm2 concentrations, KAY10 and N3 strains of H. bacteriophora performed better than other EPN species/strains and induced 80 and 82.5% mortality, respectively. All EPN species/strains did not differ significantly at 100 and 150 Ijs/cm2 concentrations at 30 °C, and mortality ranged between 82.5 and 87.5% at 18 DAT. KAY10 and AF12 strains of H. bacteriophora were the only EPN species/strains that cause mortality over 65% at 25 Ijs/cm2 concentrations at 30 °C at 18 DAT. The highest efficacy (87.5%) was obtained from H. bacteriophora AF12 strain at 150 Ijs/cm2 at 18 DAT (Table 6).

4. Discussion

In the present study, native EPN species were tested for biocontrol potential against the most abundantly collected wireworm species from potato fields which is a major concern for potato growers in Europe including Türkiye [31,32,33,34,35,36]. As compared to other host species of EPNs, wireworms generally exhibited lower susceptibility to nematode infection, and mortality generally occurred over a longer period of time [37]. For instance, Williams et al. [38] tested the efficacy of S. feltiae, S. carpocapsae, H. bacteriophora, and H. indica on the larvae of Melanotus communis (G.) at 100 Ijs/cm2 concentration and the highest larval mortality did not exceed 15%. In another study, Forgia et al. [39] conducted a laboratory bioassay against Agriotes sordidus (Illiger) larvae in well plates at 2 Ijs/cm2 concentration and reported 8.3 and 16.7% of mortality for S. carpocapsae and H. bacteriophora strains, respectively. Campos-Herrera and Gutiérrez [40] evaluated the pathogenicity of different strains of S. feltiae and S. carpocapsae in well plates at 250 Ijs/cm2 and reported 9% mortality in larvae of A. sordidus 12 DAT which was induced by only one S. feltiae strain. In contrast to aforementioned studies, Toba et al. [41] reported 58% mortality in the 7–10th instars larvae of Limonius californicus (M.) when S. feltiae was applied at a concentration of 393 Ijs/cm2. In another study, Morton and Garcia-del-Pino [42] conducted a laboratory bioassay with different S. feltiae, S. carpocapsae, and H. bacteriophora strains against the 5th/6th larvae of A. obscurus, and mortality rates ranged between 17% and 35% for H. bacteriophora strains while the highest mortality (75%) was obtained with S. carpocapsae B14 strain at a concentration of 100 IJs/cm2. In the present study, all tested EPN species/strains were highly pathogenic to A. rufipalpis and A. sputator larvae at 18 DAT, and mortality over 70% was achieved by all EPN species/strains. The high virulence of tested EPNs in this study is in line with Ansari et al. [34], Morton and Garcia-del-Pino [42], and Sandhi et al. [43] who reported mortality between 50 and 75% in wireworm species. It is a well-known fact that the pathogenicity of EPNs on the same host species differs greatly among species and even strains [44,45]. Therefore, preliminary pathogenicity screening tests provide valuable information before evaluation of performance of EPNs [46]. In the present study, the most pathogenic EPN species that proved to be highly virulent on the larvae of G. mellonella in an earlier study were used [30]. In the aforementioned studies, different wireworm and EPN species were utilized in the bioassays which might be one possible reason leading to variations in the mortality rates. Earlier studies illustrated species-dependent nematode infection among a range of hosts [47,48]. Unsuitable EPN species-host combinations may be partially responsible for the differences in the mortality rates of wireworm larvae. In the current study, all EPN species and strains that were collected from potato fields infested with A. sputator and A. rufipalpis were used in the bioassays. It is reasonable to assume that tested EPN species/strains were predisposed to successfully infect and kill wireworm larvae. The co-existence of the tested EPNs within the host habitat may have helped to give additional positive responses to wireworm-derived cues and eventually to explain the high infection rates [47,49]. On the other hand, infection of EPNs requires successful penetration of IJs into the host body. Wireworms are considered to have strong morphological structures that help them to avoid or limit the penetration of IJs [50]. The differences in the morphological structures of wireworm species that function as physical barriers to EPNs might be another reason behind differences in the mortality rates. In addition, the immune systems of host species that detect the presence of microbial infection play a key role in the pathogenicity of IJs. Rahatkhah et al. [51] indicated that there could be a great variation in the recognition of IJs by the immune system of different wireworm species. Conversely, EPNs and their bacterial associates produce a large number of metabolites in the host hemocoel that exhibit immunosuppressant activity with varying levels of efficiency as well as toxicity to the host intestine [52,53,54]. A great variation in the chemical composition of secondary metabolites produced by different species and strains of Xenorhabdus and Photohabdus bacteria was also reported in earlier studies which may have contributed to the differences in the mortality of wireworm larvae [55,56]. Furthermore, the developmental stage of the host insect is one of the key factors affecting the pathogenicity of EPNs. Williams et al. [38] reported that smaller M. communis larvae showed higher susceptibility to EPNs compared to larger wireworm larvae. Morton and Garcia-del-Pino [42] reported 75% mortality against the 5th/6th instars larvae of A. obscurus which is in line with our study. The immune responses of insects show variation among developmental stages and late instars larvae may have more immune responses against pathogens [57,58]. Ebssa and Koppenhöfer [59] reported that the 4th and 5th instars of Agrotis ipsilon (H.) (Lepidoptera: Noctuidae) were the most susceptible stages to EPN species. In another study, Abdolmaleki et al. [58] stated that the 4th instar larvae of Pieris brassicae (L.) (Lepidoptera: Pieridae) demonstrated higher immune responses to bacterial associates of EPNs Photorhabdus temperata subsp. temperate than the 3rd instars. In the aforementioned studies, the efficacy of EPNs was tested against mixed instars of wireworm species and this might have also affected the effectiveness of EPNs. In addition, considering the number of larval instars of wireworm species (up to 13 instars), the 4th/5th instars larvae of A. sputator and A. rufipalpis may be the most susceptible stages against tested EPNs [35].
Environmental variables also have an influence on the effectiveness of EPNs, and the adaptation capability of EPNs to environmental factors varies greatly among species and strains [60,61]. Temperature and humidity are among the major environmental factors that can either enhance or diminish the survival, mobility, and virulence of EPNs [62,63,64]. In the present study, a significant increase was observed in the efficacy of H. bacteriophora strains, particularly at low concentrations at 18 DAT. In previous studies, the optimum temperatures were reported to range between 22 and 24 °C for S. feltiae and 14 and 35 °C for S. carpocapsae, while H. bacteriophora reported to perform better between 25 and 30 °C [65,66]. The higher performance of H. bacteriophora strains at low concentrations may be explained by the optimal temperatures needed by the strains to successfully infect their host.

5. Conclusions

In this study, all tested EPN species/strains were highly pathogenic to A. rufipalpis and A. sputator larvae, and mortality over 70% was achieved at 18 DAT by all EPN species/strains. The results obtained indicated that EPN species that were recovered from the potato fields where EPNs and wireworms co-exist have the potential to provide better control against wireworms. However, field evaluation of these EPN species and strains will provide better insights into the performance of EPNs. In field conditions, several wireworm species in different development stages may be present with varying susceptibility to EPNs. Therefore, combining several EPN species may help suppress the wireworm populations.

Author Contributions

Conceptualization, A.G.A., R.B., M.İ. and E.Y.; methodology, A.G.A., R.B., H.K., R.C., E.Y., A.Ö., M.İ. and A.A.D.; validation, A.G.A., H.K., R.C., E.Y., M.İ., A.Ö. and A.A.D.; formal analysis, A.G.A., R.C., E.Y., D.D. and A.A.D.; writing—review and editing, A.G.A., R.B., E.Y., M.İ. and A.A.D. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by The Scientific and Technological Council of Turkey (TUBITAK/TOVAG), grant number 115R025.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Data generated in this study are available upon reasonable request to the corresponding author.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Furlan, L.; Contiero, B.; Chiarini, F.; Colauzzi, M.; Sartori, E.; Benvegnù, I.; Fracasso, F.; Giandon, P. Risk assessment of maize damage by wireworms (Coleoptera: Elateridae) as the first step in implementing IPM and in reducing the environmental impact of soil insecticides. Environ. Sci. Pollut. Res. 2017, 24, 236–251. [Google Scholar] [CrossRef] [PubMed]
  2. Vernon, R.S.; van Herk, W.G. Wireworms as Pests of Potato. In Insect Pests of Potato: Global Perspectives on Biology and Management; Alyokhin, A., Vincent, C., Giordanengo, P., Eds.; Academic Press: Cambridge, MA, USA; Elsevier: Amsterdam, The Netherlands, 2013; pp. 103–164. [Google Scholar]
  3. Chalfant, R.B.; Seal, D.R. Biology and management of wireworms on sweet potato. In Sweet Potato Pest Management; Jansson, R.K., Raman, K.V., Eds.; CRC Press: Boca Raton, FA, USA, 2019; pp. 303–326. [Google Scholar] [CrossRef]
  4. Keiser, A.; Häberli, M.; Stamp, P. Quality deficiencies on potato (Solanum tuberosum L.) tubers caused by Rhizoctonia solani, wireworms (Agriotes ssp.) and slugs (Deroceras reticulatum, Arion hortensis) in different farming systems. Field Crop. Res. 2012, 128, 147–155. [Google Scholar] [CrossRef]
  5. Jansson, R.K.; Lecrone, S.H. Effects of summer cover crop management on wireworm (Coleoptera: Elateridae) abundance and damage to potato. J. Econ. Entomol. 1991, 84, 581–586. [Google Scholar] [CrossRef]
  6. Nikoukar, A.; Rashed, A. Integrated Pest Management of Wireworms (Coleoptera: Elateridae) and the Rhizosphere in Agroecosystems. Insects 2022, 13, 769. [Google Scholar] [CrossRef]
  7. Kroschel, J.; Mujica, N.; Okonya, J.; Alyokhin, A. Insect pests affecting potatoes in tropical, subtropical, and temperate regions. In The Potato Crop: Its Agricultural, Nutritional and Social Contribution to Humankind; Campos, H., Ortiz, O., Eds.; Springer: New York, NY, USA, 2020; pp. 80–123. [Google Scholar] [CrossRef]
  8. Furlan, L.; Tóth, M. Occurrence of click beetle pest spp. (Coleoptera, Elateridae) in Europe as detected by pheromone traps: Survey results of 1998–2006. IOBC/WPRS Bull. 2007, 30, 19. [Google Scholar]
  9. Furlan, L.; Benvegnù, I.; Bilò, M.F.; Lehmhus, J.; Ruzzier, E. Species Identification of Wireworms (Agriotes spp.; Coleoptera: Elateridae) of Agricultural Importance in Europe: A New “Horizontal Identification Table”. Insects 2021, 12, 534. [Google Scholar] [CrossRef]
  10. Poggi, S.; Le Cointe, R.; Lehmhus, J.; Plantegenest, M.; Furlan, L. Alternative strategies for controlling wireworms in field crops: A review. Agriculture 2021, 11, 436. [Google Scholar] [CrossRef]
  11. Reddy, G.V.; Tangtrakulwanich, K.; Wu, S.; Miller, J.H.; Ophus, V.L.; Prewett, J.; Jaronski, S.T. Evaluation of the effectiveness of entomopathogens for the management of wireworms (Coleoptera: Elateridae) on spring wheat. J. Invertebr. Pathol. 2014, 120, 43–49. [Google Scholar] [CrossRef]
  12. Eckard, S.; Ansari, M.A.; Bacher, S.; Butt, T.M.; Enkerli, J.; Grabenweger, G. Virulence of in vivo and in vitro produced conidia of Metarhizium brunneum strains for control of wireworms. J. Crop. Prot. 2014, 64, 137–142. [Google Scholar] [CrossRef]
  13. Furlan, L.; Contiero, B.; Chiarini, F.; Benvegnù, I.; Tóth, M. The use of click beetle pheromone traps to optimize the risk assessment of wireworm (Coleoptera: Elateridae) maize damage. Sci. Rep. 2020, 10, 8780. [Google Scholar] [CrossRef]
  14. La Forgia, D.; Bruno, P.; Campos-Herrera, R.; Turlings, T.; Verheggen, F. The lure of hidden death: Development of an attract-and-kill strategy against Agriotes obscurus (Coleoptera: Elateridae) combining semiochemicals and entomopathogenic nematodes. Turk. J. Zool. 2021, 45, 347–355. [Google Scholar] [CrossRef]
  15. Gulzar, S.; Wakil, W.; Shapiro-Ilan, D.I. Potential use of entomopathogenic nematodes against the soil dwelling stages of onion thrips, Thrips tabaci Lindeman: Laboratory, greenhouse and field trials. Biol. Control 2021, 161, 104677. [Google Scholar] [CrossRef]
  16. Vicente-Díez, I.; Blanco-Pérez, R.; Chelkha, M.; Puelles, M.; Pou, A.; Campos-Herrera, R. Exploring the use of entomopathogenic nematodes and the natural products derived from their symbiotic bacteria to control the grapevine moth, Lobesia botrana (Lepidoptera: Tortricidae). Insects 2021, 12, 1033. [Google Scholar] [CrossRef] [PubMed]
  17. Peçen, A.; Kepenekci, İ. Efficacy of entomopathogenic nematode isolates from Turkey against wheat stink bug, Aelia rostrata Boheman (Hemiptera: Pentatomidae) adults under laboratory conditions. Egypt. J. Biol. Pest Control 2022, 32, 1–6. [Google Scholar] [CrossRef]
  18. Shapiro-Ilan, D.I.; Hazir, S.; Glazer, I. Entomopathogenic Nematodes as Models for Inundative Biological Control. In Nematodes as Model Organisms; Glazer, I., Shapiro-Ilan, D.I., Sternberg, P.W., Eds.; CABI International: Boston, MA, USA, 2022; pp. 293–308. [Google Scholar]
  19. Usman, M.; Wakil, W.; Shapiro-Ilan, D.I. Entomopathogenic nematodes as biological control agent against Bactrocera zonata and Bactrocera dorsalis (Diptera: Tephritidae). Biol. Control 2021, 163, 104706. [Google Scholar] [CrossRef]
  20. Kaya, H.K.; Gaugler, R. Entomopathogenic nematodes. Annu. Rev. Entomol. 1993, 38, 181–206. [Google Scholar] [CrossRef]
  21. Koppenhöfer, A.M.; Shapiro-Ilan, D.I.; Hiltpold, I. Entomopathogenic nematodes in sustainable food production. Front. Sustain. Food Syst. 2020, 4, 125. [Google Scholar] [CrossRef]
  22. Stock, S.P. Partners in crime: Symbiont-assisted resource acquisition in Steinernema entomopathogenic nematodes. Curr. Opin. Insect Sci. 2019, 32, 22–27. [Google Scholar] [CrossRef]
  23. Abd-Elgawad, M.M. Xenorhabdus spp.: An overview of the useful facets of mutualistic bacteria of entomopathogenic nematodes. Life 2022, 12, 1360. [Google Scholar] [CrossRef]
  24. Hazir, S.; Kaya, H.K.; Stock, S.P.; Keskin, N. Entomopathogenic nematodes (Steinernematidae and Heterorhabditidae) for biological control of soil pests. Turk. J. Biol. 2003, 27, 181–202. [Google Scholar]
  25. Shapiro-Ilan, D.; Dolinski, C. Entomopathogenic nematode application technology. In Nematode Pathogenesis of Insects and Other Pests; Campos-Herrera, R., Ed.; Springer: Berlin/Heidelberg, Germany, 2015; pp. 231–254. [Google Scholar] [CrossRef]
  26. Grewal, P.S.; Selvan, S.; Gaugler, R. Nematodes: Niche breadth for infection, establishment, and reproduction. J. Therm. Biol. 1994, 19, 245–253. [Google Scholar]
  27. Lewis, E.E.; Campbell, J.; Griffin, C.; Kaya, H.; Peters, A. Behavioral ecology of entomopathogenic nematodes. Biol. Control 2006, 38, 66–79. [Google Scholar] [CrossRef]
  28. Noujeim, E.; Rehayem, M.; Nemer, N. Comparison of indigenous and exotic entomopathogenic nematode strains for control of the cedar web-spinning sawfly, Cephalcia tannourinensis in vitro. Biocontrol Sci. Technol. 2015, 25, 843–851. [Google Scholar] [CrossRef]
  29. Sandhi, R.K.; Reddy, G.V. Effects of entomopathogenic nematodes and symbiotic bacteria on non-target arthropods. In Microbes for Sustainable Insect Pest Management; Khan, A.M., Ahmad, W., Eds.; Springer: Berlin/Heidelberg, Germany, 2019; pp. 247–273. [Google Scholar] [CrossRef]
  30. Gümüş Askar, A.; Yüksel, E.; Öcal, A.; Özer, G.; Kütük, H.; Dababat, A.; İmren, M. Identification and control potential of entomopathogenic nematodes against the black cutworm, Agrotis ipsilon (Fabricius) (Lepidoptera: Noctuidae), in potato-growing areas of Turkey. J. Plant Dis. Prot. 2022, 129, 911–922. [Google Scholar] [CrossRef]
  31. Gümüş Askar, A.; Yüksel, E.; Dinçer, D.; Bozbuğa, R.; Öcal, A.; Kütük, H.; Dababat, A.A.; Özer, G.; İmren, M. Molecular Identification, Occurrence and Biodiversity of Wireworm Species (Agriotes spp.) (Coleoptera: Elateridae) in Major Potato Cultivated Areas of Türkiye. J. Insect Biodivers. 2023; submitted. [Google Scholar]
  32. Kaya, H.K.; Stock, S.P. Techniques in insect nematology. In Manual of Techniques in Insect Pathology, 1st ed.; Lacey, L.A., Ed.; Academic Press: Cambridge, MA, USA, 1997; pp. 281–324. [Google Scholar] [CrossRef]
  33. Metwally, H.M.; Hafez, G.A.; Hussein, M.A.; Hussein, M.A.; Salem, H.A.; Saleh, M.M.E. Low cost artificial diet for rearing the greater wax moth, Galleria mellonella L. (Lepidoptera: Pyralidae) as a host for entomopathogenic nematodes. Egypt. J. Biol. Pest Control 2012, 22, 15. [Google Scholar]
  34. Ansari, M.A.; Evans, M.; Butt, T.M. Identification of pathogenic strains of entomopathogenic nematodes and fungi for wireworm control. Crop Prot. 2009, 28, 269–272. [Google Scholar] [CrossRef]
  35. Sufyan, M.; Neuhoff, D.; Furlan, L. Larval development of Agriotes obscurus under laboratory and semi-natural conditions. Bull. Insectol. 2014, 67, 227–235. [Google Scholar]
  36. Ritter, C.; Richter, E. Control methods and monitoring of Agriotes wireworms (Coleoptera: Elateridae). J. Plant Dis. Prot. 2013, 120, 4–15. [Google Scholar]
  37. Öğretmen, A.; Yüksel, E.; Canhilal, R. Susceptibility of larvae of wireworms (Agriotes spp.)(Coleoptera: Elateridae) to some Turkish isolates of entomopathogenic nematodes under laboratory and field conditions. Biol. Control 2020, 149, 104320. [Google Scholar] [CrossRef]
  38. Williams, L.; Cherry, R.; Shapiro-Ilan, D. Effect of Host Size on Susceptibility of Melanotus communis (Coleoptera: Elateridae) Wireworms to Entomopathogens. J. Nematol. 2022, 54, 10.2478/jofnem-2022-0033. [Google Scholar] [CrossRef]
  39. La Forgia, D.; Jaffuel, G.; Campos-Herrera, R.; Verheggen, F.; Turlings, T. Efficiency of an Attract-and-Kill System with Entomopathogenic Nematodes against Wireworms (Coleoptera: Elateridae). IOBC/WPRS Bull. 2020, 150, 91–95. [Google Scholar]
  40. Campos-Herrera, R.; Gutiérrez, C. Screening Spanish Isolates of Steinernematid Nematodes for Use as Biological Control Agents through Laboratory and Greenhouse Microcosm Studies. J. Invertebr. Pathol. 2009, 100, 100–105. [Google Scholar] [CrossRef] [PubMed]
  41. Toba, H.H.; Lindegren, J.E.; Turner, J.E.; Vail, P.V. Susceptibility of the Colorado Potato Beetle and the Sugarbeet Wireworm to Steinernema feltiae and S. glaseri. J. Nematol. 1983, 15, 597–601. [Google Scholar] [PubMed]
  42. Morton, A.; Garcia-del-Pino, F. Laboratory and Field Evaluation of Entomopathogenic Nematodes for Control of Agriotes obscurus (L.) (Coleoptera: Elateridae). J. Appl. Entomol. 2017, 141, 241–246. [Google Scholar] [CrossRef]
  43. Sandhi, R.K.; Shapiro-Ilan, D.; Sharma, A.; Reddy, G.V. Efficacy of entomopathogenic nematodes against the sugarbeet wireworm, Limonius californicus (Mannerheim) (Coleoptera: Elateridae). Biol. Control 2020, 143, 104190. [Google Scholar] [CrossRef]
  44. Abbas, M.S.T. Pathogenicity of entomopathogenic nematodes to dipteran leaf miners, house flies and mushroom flies. Egypt. J. Biol. Pest Control 2022, 32, 76. [Google Scholar] [CrossRef]
  45. Yüksel, E. Biocontrol potential of endosymbiotic bacteria of entomopathogenic nematodes against the tomato leaf miner, Tuta absoluta (Meyrick)(Lepidoptera: Gelechiidae). Egypt. J. Biol. Pest Control 2022, 32, 135. [Google Scholar] [CrossRef]
  46. Köhl, J.; Postma, J.; Nicot, P.; Ruocco, M.; Blum, B. Stepwise screening of microorganisms for commercial use in biological control of plant-pathogenic fungi and bacteria. Biol. Control 2011, 57, 1–12. [Google Scholar] [CrossRef]
  47. Simoes, N.; Rosa, J.S. Pathogenicity and host specificity of entomopathogenic nematodes. Biocontrol Sci. Technol. 1996, 6, 403–412. [Google Scholar] [CrossRef]
  48. Alonso, V.; Nasrolahi, S.; Dillman, A.R. Host-specific activation of entomopathogenic nematode infective juveniles. Insects 2018, 9, 59. [Google Scholar] [CrossRef]
  49. Zhang, X.; Li, L.; Kesner, L.; Robert, C.A.M. Chemical host-seeking cues of entomopathogenic nematodes. Curr. Opin. Insect Sci. 2021, 44, 72–81. [Google Scholar] [CrossRef] [PubMed]
  50. Eidt, D.C.; Thurston, G.S. Physical deterrents to infection by entomopathogenic nematodes in wireworms (Coleoptera: Elateridae) and other soil insects. Can. Entomol. 1995, 127, 423–429. [Google Scholar] [CrossRef]
  51. Rahatkhah, Z.; Karimi, J.; Ghadamyari, M.; Brivio, M.F. Immune defenses of Agriotes lineatus larvae against entomopathogenic nematodes. BioControl 2015, 60, 641–653. [Google Scholar] [CrossRef]
  52. Bode, H.B.; Müller, R. The impact of bacterial genomics on natural product research. Angew. Chem. Int. Ed. 2005, 44, 6828–6846. [Google Scholar] [CrossRef] [PubMed]
  53. Bode, H.B. Entomopathogenic bacteria as a source of secondary metabolites. Curr. Opin. Chem. Biol. 2009, 13, 224–230. [Google Scholar] [CrossRef] [PubMed]
  54. Shawer, R.; Donati, I.; Cellini, A.; Spinelli, F.; Mori, N. Insecticidal Activity of Photorhabdus luminescens against Drosophila suzukii. Insects 2018, 9, 148. [Google Scholar] [CrossRef]
  55. Tobias, N.J.; Shi, Y.M.; Bode, H.B. Refining the natural product repertoire in entomopathogenic bacteria. Trends Microbiol. 2018, 26, 833–840. [Google Scholar] [CrossRef]
  56. Hasan, M.A.; Ahmed, S.; Mollah, M.M.I.; Lee, D.; Kim, Y. Variation in pathogenicity of different strains of Xenorhabdus nematophila; Differential immunosuppressive activities and secondary metabolite production. J. Invertebr. Pathol. 2019, 166, 107221. [Google Scholar] [CrossRef]
  57. Feldhaar, H.; Gross, R. Immune reactions of insects on bacterial pathogens and mutualists. Microbes Infect. 2008, 10, 1082–1088. [Google Scholar] [CrossRef]
  58. Abdolmaleki, A.; Rafiee Dastjerdi, H.; Tanha Maafi, Z.; Naseri, B. Virulence of two entomopathogenic nematodes through their interaction with Beauveria bassiana and Bacillus thuringiensis against Pieris brassicae (Lepidoptera: Pieridae). J. Crop Prot. 2017, 6, 287–299. [Google Scholar]
  59. Ebssa, L.; Koppenhöfer, A.M. Efficacy and persistence of entomopathogenic nematodes for black cutworm control in turfgrass. Biocontrol Sci. Technol. 2011, 21, 779–796. [Google Scholar] [CrossRef]
  60. Shapiro-Ilan, D.I.; Gouge, D.H.; Piggott, S.J.; Fife, J.P. Application technology and environmental considerations for use of entomopathogenic nematodes in biological control. Biol. Control 2006, 38, 124–133. [Google Scholar] [CrossRef]
  61. Gulzar, S.; Usman, M.; Wakil, W.; Gulcu, B.; Hazir, C.; Karagoz, M.; Shapiro-Ilan, D.I. Environmental tolerance of entomopathogenic nematodes differs among nematodes arising from host cadavers versus aqueous suspension. J. Invertebr. Pathol. 2020, 175, 107452. [Google Scholar] [CrossRef] [PubMed]
  62. Hsiao, W.F.; All, J.N. Effects of temperature and placement site on the dispersal of the entomopathogenic nematode, Steinernema carpocapsae in four soils. Chin. J. Entomol. 1996, 16, 95–106. [Google Scholar]
  63. Hummel, R.L.; Walgenbach, J.F.; Barbercheck, M.E.; Kennedy, G.G.; Hoyt, G.D.; Arellano, C. Effects of production practices on soil-borne entomopathogens in western North Carolina vegetable systems. Environ. Entomol. 2002, 31, 84–91. [Google Scholar] [CrossRef]
  64. Shaurub, E.H.; Soliman, N.A.; Hashem, A.G.; Abdel-Rahman, A.M. Infectivity of four entomopathogenic nematodes in relation to environmental factors and their effects on the biochemistry of the Medfly Ceratitis capitata (Wied.)(Diptera: Tephritidae). Neotrop. Entomol. 2015, 44, 610–618. [Google Scholar]
  65. Hirao, A.; Ehlers, R.U. Effect of temperature on the development of Steinernema carpocapsae and Steinernema feltiae (Nematoda: Rhabditida) in liquid culture. Appl. Microbiol. Biotechnol. 2009, 84, 1061–1067. [Google Scholar]
  66. Chung, H.J.; Lee, D.W.; Yoon, H.S.; Lee, S.M.; Park, C.G.; Choo, H.Y. Temperature and dose effects on the pathogenicity and reproduction of two Korean isolates of Heterorhabditis bacteriophora (Nematoda: Heterorhabditidae). J. Asia Pac. Entomol. 2010, 13, 277–282. [Google Scholar] [CrossRef]
Pathogens 12 00288 g001 550
Figure 1. Emerging infective juveniles (black arrows) of Steinernema feltiae from the body of Agriotes rufipalpis larvae.
Figure 1. Emerging infective juveniles (black arrows) of Steinernema feltiae from the body of Agriotes rufipalpis larvae.
Pathogens 12 00288 g001
Table
Table 1. List of entomopathogenic nematode species/strains used in the experiments.
Table 1. List of entomopathogenic nematode species/strains used in the experiments.
Entomopathogenic Nematodes StrainHabitatCoordinatesGenBank Accession Number
Steinernema carpocapsaeSc_BL22Potato 40°47′11″ N 31°38′78″ EOK632299
Steinernema feltiaeSf_BL24Potato40°47′14″ N 31°39′10″ EOK632300
Steinernema feltiaeSf_KAY4Potato38°20′28″ N 35°27′49″ EOK632306
Heterorhabditis bacteriophoraHb_N3Potato38°02′19″ N 34°44′18″ EOK632328
Heterorhabditis bacteriophoraHb_KAY10Potato38°16′49″ N 35°25′15″ EOK632308
Heterorhabditis bacteriophoraHb_AF12Potato37°55′31″ N 29°52′19″ EOK632288
Table
Table 2. Repeated-measures analysis of variance parameters for the main factors and associated interactions (Tukey, p ≤ 0.05).
Table 2. Repeated-measures analysis of variance parameters for the main factors and associated interactions (Tukey, p ≤ 0.05).
Sources Agriotes rufipalpisAgriotes sputator
Degree of FreedomF Valuep ValueF Valuep Value
Nematode (N)59.268<0.0117.455<0.01
Concentration (C)5295.432<0.01825.869<0.01
Temperature (T)142.688<0.0157.436<0.01
C × N251.7350.0203.327<0.01
C × T515.012<0.015.465<0.01
N × T52.7460.0202.0420.074
C × N × T251.5540.0514.895<0.01
Error1216
Exposure time (t)1491.683<0.01337.559<0.01
t × C594.844<0.0128.474<0.01
t × N51.2250.2982.9610.013
t × T177.7780.5403.1940.075
t × C × N250.7990.7421.4570.081
t × C × T55.3210.3581.6360.152
t × N × T55.0540.0113.2060.008
t × C × N × T251.7170.0221.3400.137
Error2216
Table
Table 3. Mortality rates (%) of 4th/5th instars larvae of Agriotes rufipalpis 6 days after application of different entomopathogenic nematode species/strains in the pot experiments.
Table 3. Mortality rates (%) of 4th/5th instars larvae of Agriotes rufipalpis 6 days after application of different entomopathogenic nematode species/strains in the pot experiments.
TemperaturesNematodes *Mortality Rates (%) 6 Days after Treatment (DAT)
Control25 IJs/cm250 IJs/cm2100 IJs/cm2150 IJs/cm2
25 °CSc_BL220.0 ± 0.0A a a b7.5 ± 5.0Ba7.5 ± 5.0Ba10.0 ± 8.1Ba12.5 ± 5.0Ba
Sf_BL240.0 ± 0.0Aa2.5 ± 5.0Aa5.0 ± 5.7Aa15.0 ± 5.7Ba25.0 ± 5.7Cb
Sf_KAY40.0 ± 0.0Aa2.5 ± 5.0Aa5.0 ± 5.7Aa25.0 ± 5.7Bb25.0 ± 5.7Bb
Hb_N30.0 ± 0.0Aa2.5 ± 5.0Aa2.5 ± 5.0Aa15.0 ± 10.0Ba27.5 ± 9.5Cb
Hb_KAY100.0 ± 0.0Aa5.0 ± 5.7Aa5.0 ± 10.0Aa15.0 ± 10.0Ba20.0 ± 8.1Bab
Hb_AF120.0 ± 0.0Aa7.5 ± 5.0Ba7.5 ± 9.5Ba15.0 ± 5.7Ca15.0 ± 5.7Ca
30 °CSc_BL220.0 ± 0.0Aa12.5 ± 5.0Ba12.5 ± 9.5Ba15.0 ± 8.1Ba22.5 ± 5.0Ca
Sf_BL240.0 ± 0.0Aa7.5 ± 9.5Ba17.5 ± 5.0Cab30.0 ± 8.1Db32.5 ± 5.0Dab
Sf_KAY40.0 ± 0.0Aa15.0 ± 5.7Ba25.0 ± 5.7Cb27.5 ± 5.0Cb32.5 ± 5.0Dab
Hb_N30.0 ± 0.0Aa2.5 ± 5.0Aa10.0 ± 11.5Ba25.0 ± 12.9Cb27.5 ± 9.5Ca
Hb_KAY100.0 ± 0.0Aa10.0 ± 0.0Ba10.0 ± 0.0Ba12.5 ± 5.0Ba25.0 ± 10.0Ca
Hb_AF120.0 ± 0.0Aa15.0 ± 10.0Ba20.0 ± 0.0Bb25.0 ± 5.7Bb37.5 ± 5.0Cb
* Sc_BL22: Steinernema carpocapsae; Sf_BL24 and Sf_KAY4: Steinernema feltiae; Hb_N3, Hb_KAY10, and Hb_AF12: Heterorhabditis bacteriophora. a Different capital letters show statistically significant differences among the infective juvenile concentrations (IJs) for each entomopathogenic nematode species. b Different lowercase letters show statistically significant differences among entomopathogenic nematode species/strains for each infective juvenile concentration (p < 0.05, Tukey).
Table
Table 4. Mortality rates (%) of 4th/5th instars larvae of Agriotes rufipalpis 18 days after application of different entomopathogenic nematode species/strains in the pot experiments.
Table 4. Mortality rates (%) of 4th/5th instars larvae of Agriotes rufipalpis 18 days after application of different entomopathogenic nematode species/strains in the pot experiments.
TemperaturesNematodes *Mortality Rates (%) 18 Days after Treatment (DAT)
Control25 Ijs/cm250 Ijs/cm2100 Ijs/cm2150 Ijs/cm2
25 °CSc_BL220.0 ± 0.0A a a b30.0 ± 0.0Bb45.0 ± 10.0Bab82.5 ± 5.0Cba85.0 ± 11.5Cb
Sf_BL240.0 ± 0.0Aa25.0 ± 5.7Bab37.5 ± 5.0Ba77.5 ± 5.0Ca85.0 ± 12.9Cb
Sf_KAY40.0 ± 0.0Aa25.0 ± 5.7Bab37.5 ± 5.0Ba80.5 ± 15.0Ca85.0 ± 5.7Cb
Hb_N30.0 ± 0.0Aa25.0 ± 5.7Bab55.5 ± 5.0Cb77.5 ± 5.0Da80.0 ± 8.1Da
Hb_KAY100.0 ± 0.0Aa20.0 ± 8.1Ba60.0 ± 8.1Cb80.0 ± 5.7Ca80.5 ± 12.5Da
Hb_AF120.0 ± 0.0Aa25.0 ± 5.7Bab50.0 ± 0.0Cab72.5 ± 5.0Da85.0 ± 12.9Db
30 °CSc_BL220.0 ± 0.0Aa30.0 ± 0.0Ba55.0 ± 10.0Ca82.5 ± 5.0Bda85.0 ± 11.5Da
Sf_BL240.0 ± 0.0Aa35.0 ± 5.7Aa67.5 ± 5.0Bab87.5 ± 5.0Ca87.5 ± 10.5Ca
Sf_KAY40.0 ± 0.0Aa25.0 ± 5.7Aa67.5 ± 5.0Bab82.5 ± 15.0Ca85.0 ± 5.7Ca
Hb_N30.0 ± 0.0Aa35.0 ± 5.7Ba65.5 ± 5.0Bab80.5 ± 5.0Ca80.0 ± 8.1Ca
Hb_KAY100.0 ± 0.0Aa40.0 ± 8.1Cab75.0 ± 8.1Db80.0 ± 5.7Da82.5 ± 12.5Da
Hb_AF120.0 ± 0.0Aa45.0 ± 5.7Cb75.0 ± 0.0Db85.5 ± 5.0Da85.0 ± 12.9Da
* Sc_BL22: Steinernema carpocapsae; Sf_BL24 and Sf_KAY4: S. feltiae; Hb_N3, Hb_KAY10, and Hb_AF12: Heterorhabditis bacteriophora. a Different capital letters show statistically significant differences among the infective juvenile concentrations (Ijs) for each entomopathogenic nematode species. b Different lowercase letters show statistically significant differences among entomopathogenic nematode species/strains for each infective juvenile concentration (p < 0.05, Tukey).
Table
Table 5. Mortality rates (%) of 4th/5th instars larvae of Agriotes sputator 6 days after application of different entomopathogenic nematode species/strains in the pot experiments.
Table 5. Mortality rates (%) of 4th/5th instars larvae of Agriotes sputator 6 days after application of different entomopathogenic nematode species/strains in the pot experiments.
TemperaturesNematodes *Mortality Rates (%) 6 Days after Treatment (DAT)
Control25 Ijs/cm250 Ijs/cm2100 Ijs/cm2150 Ijs/cm2
25 °CSc_BL220.0 ± 0.0A a a b5.0 ± 5.7Aa10.0 ± 0.0Aba12.5 ± 9.5Aba17.5 ± 5.0Ba
Sf_BL240.0 ± 0.0Aa0.0 ± 0.0Aa7.5 ± 9.5Ba27.5 ± 5.0Cab27.5 ± 5.0Cab
Sf_KAY40.0 ± 0.0Aa2.5 ± 5.0Aa5.0 ± 5.7Aa25.0 ± 5.7Bab42.5 ± 5.0Cb
Hb_N30.0 ± 0.0Aa7.5 ± 9.5Ba17.5 ± 5.0Cab20.0 ± 0.0Cab37.5 ± 5.0Db
Hb_KAY100.0 ± 0.0Aa0.0 ± 0.0Aa12.5 ± 9.5Bab40.0 ± 8.1Cb47.5 ± 9.5Cb
Hb_AF120.0 ± 0.0Aa7.5 ± 9.5Ba15.0 ± 5.7BCb32.5 ± 5.0Cab45.0 ± 5.7Db
30 °CSc_BL220.0 ± 0.0Aa5.0 ± 5.7Aa15.0 ± 5.7Ba32.5 ± 5.0Cb50.0 ± 8.1Db
Sf_BL240.0 ± 0.0Aa10.0 ± 11.5Ba22.5 ± 9.5Ca25.0 ± 12.9Cab37.5 ± 5.0Da
Sf_KAY40.0 ± 0.0Aa10.0 ± 8.1Ba20.0 ± 0.0Ca32.5 ± 5.0Db40.0 ± 8.1Da
Hb_N30.0 ± 0.0Aa7.5 ± 9.5Ba17.5 ± 5.0Ca20.0 ± 0.0Ca37.5 ± 5.0Da
Hb_KAY100.0 ± 0.0Aa15.0 ± 10.0Ba20.0 ± 8.1Ba35.0 ± 5.7Cb50.0 ± 8.1Db
Hb_AF120.0 ± 0.0Aa15.0 ± 5.7Ba22.5 ± 5.0Ba27.5 ± 5.0BCb40.0 ± 8.1Ca
* Sc_BL22: Steinernema carpocapsae; Sf_BL24 and Sf_KAY4: S. feltiae; Hb_N3, Hb_KAY10, and Hb_AF12: Heterorhabditis bacteriophora. a Different capital letters show statistically significant differences among the infective juvenile concentrations (Ijs) for each entomopathogenic nematode species. b Different lowercase letters show statistically significant differences among entomopathogenic nematode species/strains for each infective juvenile concentration (p < 0.05, Tukey).
Table
Table 6. Mortality rates (%) of 4th/5th instars larvae of Agriotes sputator 18 days after application of different entomopathogenic nematode species/strains in the pot experiments.
Table 6. Mortality rates (%) of 4th/5th instars larvae of Agriotes sputator 18 days after application of different entomopathogenic nematode species/strains in the pot experiments.
TemperaturesNematodes *Mortality Rates (%) 18 Days after Treatment (DAT)
Control25 Ijs/cm250 Ijs/cm2100 Ijs/cm2150 Ijs/cm2
25 °CSc_BL220.0 ± 0.0A a a b45.0 ± 5.7Ba55.0 ± 5.7Ba75.0 ± 5.7Ca85.0 ± 5.7Ca
Sf_BL240.0 ± 0.0Aa35.0 ± 5.7Ba65.0 ± 5.7Cb75.0 ± 5.7Ca75.0 ± 5.7Ca
Sf_KAY40.0 ± 0.0Aa42.5 ± 5.0Ca52.5 ± 5.0Ca72.5 ± 5.0Da72.5 ± 5.0Da
Hb_N30.0 ± 0.0Aa52.5 ± 5.0Cab62.5 ± 5.0Ca82.5 ± 5.0Da82.5 ± 5.0Da
Hb_KAY100.0 ± 0.0Aa60.0 ± 11.5Cb70.0 ± 11.5CDb80.0 ± 11.5Da 85.0 ± 11.5Da
Hb_AF120.0 ± 0.0Aa57.5 ± 5.0Cab77.5 ± 5.0Dc77.5 ± 5.0Da 80.0 ± 5.0Da
30 °CSc_BL220.0 ± 0.0Aa55.0 ± 5.7Bca75.0 ± 5.7Ca85.0 ± 5.7Ca85.0 ± 5.7Ca
Sf_BL240.0 ± 0.0Aa55.0 ± 5.7Ca75.0 ± 5.7Da85.0 ± 5.7Da85.0 ± 5.7Da
Sf_KAY40.0 ± 0.0Aa52.5 ± 5.0Ca72.5 ± 5.0Da82.5 ± 5.0Da82.5 ± 5.0Da
Hb_N30.0 ± 0.0Aa55.0 ± 5.0Ca80.0 ± 5.0Da82.5 ± 5.0Da82.5 ± 5.0Da
Hb_KAY100.0 ± 0.0Aa65.0 ± 11.5Cb80.0 ± 11.5Da85.0 ± 11.5Da85.0 ± 11.5Da
Hb_AF120.0 ± 0.0Aa67.5 ± 5.0Bb80.0 ± 5.0Ca82.5 ± 5.0Ca87.5 ± 5.0Ca
* Sc_BL22: Steinernema carpocapsae; Sf_BL24 and Sf_KAY4: S. feltiae; Hb_N3, Hb_KAY10, and Hb_AF12: Heterorhabditis bacteriophora. a Different capital letters show statistically significant differences among the infective juvenile concentrations (Ijs) for each entomopathogenic nematode species. b Different lowercase letters show statistically significant differences among entomopathogenic nematode species/strains for each infective juvenile concentration (p < 0.05, Tukey).
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Askar, A.G.; Yüksel, E.; Bozbuğa, R.; Öcal, A.; Kütük, H.; Dinçer, D.; Canhilal, R.; Dababat, A.A.; İmren, M. Evaluation of Entomopathogenic Nematodes against Common Wireworm Species in Potato Cultivation. Pathogens 2023, 12, 288. https://doi.org/10.3390/pathogens12020288

AMA Style

Askar AG, Yüksel E, Bozbuğa R, Öcal A, Kütük H, Dinçer D, Canhilal R, Dababat AA, İmren M. Evaluation of Entomopathogenic Nematodes against Common Wireworm Species in Potato Cultivation. Pathogens. 2023; 12(2):288. https://doi.org/10.3390/pathogens12020288

Chicago/Turabian Style

Askar, Arife Gümüş, Ebubekir Yüksel, Refik Bozbuğa, Atilla Öcal, Halil Kütük, Dilek Dinçer, Ramazan Canhilal, Abdelfattah A. Dababat, and Mustafa İmren. 2023. "Evaluation of Entomopathogenic Nematodes against Common Wireworm Species in Potato Cultivation" Pathogens 12, no. 2: 288. https://doi.org/10.3390/pathogens12020288

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