We challenged Locusta migratoria (Meyen) grasshoppers with simultaneous doses of both the insecticide chlorantraniliprole and the fungal pathogen, Metarhizium anisopliae. Our results showed synergistic and antagonistic effects on host mortality and enzyme activities. To elucidate the biochemical mechanisms that underlie detoxification and pathogen-immune responses in insects, we monitored the activities of 10 enzymes. After administration of insecticide and fungus, activities of glutathione-S-transferase (GST), general esterases (ESTs) and phenol oxidase (PO) decreased in the insect during the initial time period, whereas those of aryl acylamidase (AA) and chitinase (CHI) increased during the initial period and that of acetylcholinesterase (AChE) increased during a later time period. Activities of superoxide dismutase (SOD), catalase (CAT) and peroxidase (POD) decreased at a later time period post treatment. Interestingly, treatment with chlorantraniliprole and M. anisopliae relieved the convulsions that normally accompany M. anisopliae infection. We speculate that locust mortality increased as a result of synergism via a mechanism related to Ca2+ disruption in the host. Our study illuminates the biochemical mechanisms involved in insect immunity to xenobiotics and pathogens as well as the mechanisms by which these factors disrupt host homeostasis and induce death. We expect this knowledge to lead to more effective pest control.
Controlling insect pests, which continue to be agricultural, medical, and economic threats worldwide, requires constant human innovation1. One promising pest control method is to simultaneously attack insects with both an insecticide and an insect pathogen. This “dual-attack” approach can result in much higher insect mortality than using either of these methods independently2,3. Leveraging this combined approach to improve pest control requires a better understanding of the molecular and physiological mechanisms underlying the synergism between pesticides and pathogens. In this study, we attempted to understand the enzymatic consequences for the insect host of synergism between the insect-pathogenic fungus Metarhizium anisopliae and the insecticide chlorantraniliprole.
Metarhizium anisopliae (Deuteromycota: Hyphomycetes) is a well-known entomopathogen that has been successfully employed to control grasshoppers and other insect pests4,5,6. For example, applications of approximately 2.0 × 1012 to 1.0 × 1013 M. anisopliae conidia per hectare in nature can produce 80–98% mortality in locusts and grasshoppers in 7 to 21 d4,5,7,8,9. However, entomopathogenic fungi require long time periods to induce sufficient insect mortality. The dual-attack technique can overcome this problem, because the pathogen and insecticide often interact synergistically to both increase mortality and shorten the time until death in insects2,3,10,11,12. Chlorantraniliprole is a novel anthranilic diamide insecticide that causes feeding cessation, lethargy, muscle paralysis, and ultimately death13,14. Interestingly, both M. anisopliae and chlorantraniliprole harm insects, in part, by disrupting Ca2+ balance. Chlorantraniliprole activates the ryanodine receptor and affects calcium homeostasis by triggering Ca2+ release from the sarcoplasmic reticulum in host cells15. M. anisopliae produces numerous fungal secondary metabolites, including cyclic hexadepsipeptides, also known as destruxins, which induce membrane depolarization by opening Ca2+ channels, leading to tetanic paralysis and death of the host insect16,17. We hypothesized that a synergistic effect may exist between M. anisopliae and chlorantraniliprole and hence chose this specific pathogen and insecticide to study.
Insects defend against both insecticide and pathogen assault via multiple enzyme systems18,19,20,21. Multi-function oxidases (MFO), glutathione-S-transferase (GST) and general esterases (ESTs) are most commonly involved in defence processes in insects22. Because of their abundance, genetic diversity, broad substrate specificity and catalytic versatility, MFO, such as those involved in the P450 pathways in insects, are responsible for detoxification of natural and synthetic xenobiotics23. GST plays a pivotal role in detoxification and cellular antioxidant defences against oxidative stress by conjugating reduced glutathione to the electrophilic centres of natural and synthetic exogenous xenobiotics, including insecticides, allelochemicals and endogenously activated compounds24,25,26. ESTs perform important functions in insects through their involvement in the catabolism of the esters of higher fatty acids that influence flight as well in the degradation of inert metabolic esters19,20 and various xenobiotics, including insecticides. Changes in the activities of these enzymes are reflected in insect resistance to insecticides as well as in degradation of toxic molecules produced during M. anisopliae infection and, therefore, play a key role in the protection of insects against pathogens18,21.
Numerous other insect enzymes, including antioxidant enzymes such as superoxide dismutase (SOD), catalase (CAT) and peroxidase (POD)27 provide defences against pathogens and insecticides. Previous studies have demonstrated that these enzymes can be quickly up-regulated in response to xenobiotic threats and that increases in the activities of these enzymes are related to pesticide resistance and melanisation in insects28,29,30,31. The phenol oxidase (PO) cascade protects insects from microbial infections through its involvement in the melanisation of haemocytes attached to parasite surfaces32,33. Increased PO activity can strengthen an insect’s immunity to xenobiotics and promote wound healing34,35,36. Acetylcholinesterase (AChE) is a key enzyme catalysing the hydrolysis of the neurotransmitter acetylcholine in the nervous system of various organisms37. AChE is affected by organophosphate and carbamate insecticides, botanical insecticides and secondary fungal metabolites38,39,40,41,42. Conversely, aryl acylamidase (AA) has been found to confer resistance to chlorantraniliprole and fufenozide, whereas chitinase (CHI) confers resistance to fufenozide43,44. Although it is well known that both insecticide poisoning and fungal infections alter enzyme activities in insects18,21,42,43,45, few studies have explored the complex enzymatic consequences of simultaneous insecticide and pathogenic assaults in insects46, and no detailed investigations have been reported concerning the synergistic effect of M. anisopliae and insecticides on enzyme activities in insects.
The present study sought to understand the effects of simultaneous M. anisopliae infection and chlorantraniliprole poisoning on multiple insect enzyme systems. To this end, we evaluated the co-toxicity of chlorantraniliprole and M. anisopliae infection. Because many previous studies using the dual-attack strategy (simultaneous treatment with insecticide and pathogen) have used only one dose-level of each component2,3,10,11,12, we tested multiple dose-levels and measured the effects of different concentrations of chlorantraniliprole, M. anisopliae and their combination on MFO, GST, ESTs, PO, AChE, AA, CHI, CAT, POD and SOD activities in the locust Locusta migratoria. Our results establish a theoretical basis for the interaction between M. anisopliae and insecticides and should inform future studies investigating the mechanism by which M. anisopliae overcomes host immunity. We hope to use the information from this study to design better control methods for harmful insects.
Virulence of chlorantraniliprole in combination with M. anisopliae against L. migratoria
The efficacy of chlorantraniliprole mixed with M. anisopliae against L. migratoria is presented in Table 1. The LC50 value of M. anisopliae alone against this insect was 0.15 mg/L (at ~7.5 × 106 spores/mL). Mixing chlorantraniliprole with M. anisopliae in Treatments 1 and 2 resulted in higher mortality rates with LC50 values of 0.01 and 0.02 mg/L and co-toxicity coefficients of 1646 and 1619, respectively. In contrast, when formulated in Treatment 3, an antagonistic interaction was observed between chlorantraniliprole and M. anisopliae, resulting in a co-toxicity coefficient of 34.
In a separate experiment, L. migratoria was treated with M. anisopliae alone for 3 d before introducing wheat sprouts treated with chlorantraniliprole in Treatment 4, the resulting co-toxicity coefficient was 127 (Table 2).
Effect of chlorantraniliprole and M. anisopliae on enzyme activities in L. migratoria
The activities of ESTs in L. migratoria are shown in Fig. 1. Chlorantraniliprole increased the activities of ESTs in L. migratoria during the initial post-treatment period; likewise, the activities of ESTs significantly increased during the initial period following M. anisopliae infection. However, the activities of ESTs markedly decreased after treatment with a mixture of M. anisopliae and chlorantraniliprole during the initial days of the experiment.
The activities of GSTs with different substrates were assessed, as shown in Figs 2 and 3. When treated with chlorantraniliprole, L. migratoria locust nymphs exhibited high GST activity (with CDNB or DCNB) during the early period, but this GST activity decreased during the later period. The activities of GSTs also increased during the initial period following M. anisopliae infection. However, GST activities decreased significantly during the initial period of the experiment when L. migratoria was treated with a combination of M. anisopliae and chlorantraniliprole simultaneously.
The activities of PO in L. migratoria after single and dual treatments were assessed, as shown in Fig. 4. When locust nymphs were treated with chlorantraniliprole alone, PO activities increased during the initial days of the experiment. When locust nymphs were treated with M. anisopliae, PO activities were high during the early period but decreased during the later period. In contrast, PO activities decreased during the initial days but increased during the later period when locust nymphs were treated with a combination of M. anisopliae and chlorantraniliprole.
The activities of MFO under single and dual treatment conditions were assessed, as shown in Fig. 5. MFO activity increased during the later period for locusts treated with M. anisopliae alone. MFO activity also increased during the later period when M. anisopliae and chlorantraniliprole were administered simultaneously; however, this increase in MFO activity was much lower than that obtained for treatment with M. anisopliae alone.
The activities of AChE in L. migratoria under different treatment conditions are shown in Fig. 6. AChE activity increased in locusts during the early period following treatment with chlorantraniliprole alone. When treated with M. anisopliae alone, locust AChE activities increased during the early period but decreased during the later period. When locusts were treated with a combination of M. anisopliae and chlorantraniliprole, AChE activities increased throughout the entire period, except the third day.
The activities of CHI (Fig. 7) in L. migratoria increased during the initial days of the experiment when locusts were treated with chlorantraniliprole alone. When locusts were treated with M. anisopliae alone, CHI activities increased throughout the entire analysis period. When locusts were treated with a combination of M. anisopliae and chlorantraniliprole, CHI activities increased during the early period but decreased during the later period.
The activities of AA (Fig. 8) increased during the initial days after independent treatment with chlorantraniliprole or M. anisopliae. AA activities also increased during the initial days after locusts were treated with a combination of M. anisopliae and chlorantraniliprole.
SOD, POD and CAT activities (Figs 9, 10, 11) increased during the initial days after independent chlorantraniliprole or M. anisopliae treatment. SOD and POD activities increased to high levels when locusts were treated with a combination of M. anisopliae and chlorantraniliprole; however, under these conditions, CAT activities decreased to low levels during the initial period, and, near the end of the experiment, CAT, SOD and POD activities decreased to low levels.
Previous studies have indicated that combinations of entomopathogenic fungi and insecticides can have synergistic, antagonistic or additive physiological and mortality effects on insects10,12,47. However, these investigations of fungal-insecticide interactions, mortality, and/or LC50 values were obtained using only one dose of pathogen or insecticide, such that the most efficacious synergistic formulations could not be determined10,11,12. The present study utilised co-toxicity coefficients to calculate insecticide incorporation rates and estimate the efficacies of different chlorantraniliprole+ M. anisopliae formulations. We found both synergistic and antagonistic interactions when chlorantraniliprole and M. anisopliae were administered together. Treatments 1, 2, and 4 revealed synergistic interactions with co-toxicity coefficients of 1646, 1619, and 127, respectively. These high co-toxicity coefficients, which were accompanied by insect mortalities >97% for some treatments, illustrate the effectiveness of this dual-attack method of insect pest control. In contrast, an antagonistic interaction (co-toxicity coefficient = 33) was observed for Treatment 3, possibly because of the high proportion of insecticide in this treatment. Our results demonstrate that chlorantraniliprole influences the virulence of M. anisopliae against L. migratoria relatively early in the initial infection period, producing a strong synergistic interaction with very high co-toxicity coefficients that vary depending on the proportions of the two agents and the time of application. As such, co-toxicity coefficients can be used to evaluate the effect of interactions between entomopathogenic fungi and insecticides, such that the best dose-combinations and time of application can be determined.
Our study also illuminated some of the biochemical consequences of a paired insecticide-pathogen challenge in insects. The accompanying enzymatic responses are numerous and complex, in part because they include attempts to detoxify the insecticide as well as the xenobiotic (the insect’s immune response to fungal attack) and because they involve biochemical/physiological disruptions due to the insecticide-and pathogen-induced pathology. Overall, most of the enzymes studied showed moderate to strong responses.
MFO, GST and ESTs are the major enzymes involved in detoxifying penetrating xenobiotics in insects18,42. Chlorantraniliprole has been found to increase the activities of ESTs and GST in Cry1Ac-susceptible and -resistant strains of Helicoverpa armigera caterpillars and to inhibit GST and MFO activities in Plutella xylostella larvae43,48. Sublethal doses of chlorantraniliprole have been observed to increase MFO and EST activities but to decrease GST activity in the Lepidoptera H. armigera, Spodoptera exigua and Choristoneura rosaceana45,49,50. In the present study, increased activities of GST and ESTs following chlorantraniliprole treatment suggest that these enzymes may act to detoxify this insecticide. Reported exceptions to these phenomena may be due to differences in the insect species as well as the concentrations of chlorantraniliprole used45,48,50.
Increased detoxifying enzyme activities against mycoses and other infections represent the insect’s response to bodily intoxication by metabolites or the host-tissue-degrading products of pathogens51. In the present study, we found that the activities of ESTs and GST were significantly increased during the initial period following M. anisopliae infection. MFO activity significantly increased during the later period following M. anisopliae infection. In Lepidopteran Galleria mellonella caterpillars, the activities of GST and ESTs in the haemolymph significantly increase during the first three days after M. anisopliae infection18. In the Hemipteran Eurygaster integriceps, the activities of GST and ESTs increase four to five days after infection with B. bassiana spores and secondary metabolites42. Alterations in the activities of GST and ESTs in G. mellonella and L. migratoria infected with entomopathogenic fungi are considered to constitute nonspecific body responses to integument damage, induction of additional isoenzymes or fungal toxins21,51,52. Based on these observations, GST and ESTs appear to participate in the defensive reaction of L. migratoria to M. anisopliae during the initial period of infection.
When L. migratoria were treated with a combination of M. anisopliae and chlorantraniliprole simultaneously, the activities of ESTs and GST decreased significantly during the initial period. However, MFO activity increased during the later period. Our results are consistent with those from a previously published investigation of the system components involved in insect detoxification. For example, when the Colorado Potato Beetle Leptinotarsa decemlineata is treated with a mixture of M. anisopliae and an organophosphate insecticide, EST and GST activities significantly decrease 2 d after inoculation46. Hall has reported that the pathogen sickens the pest, thereby lowering its chemical resistance, and the chemical, in turn, sufficiently weakens the pest and increases its susceptibility to pathogen infection53. Furlong and Groden reported that the interactions between fungi and insecticides are probably mediated during certain periods of the infection process, such as conidial attachment, conidial germination, cuticular penetration, or the initial proliferation of hyphal bodies in the haemocoel54. We believe that chlorantraniliprole can be used as a stressor in L. migratoria to increase its susceptibility to M. anisopliae and that toxic metabolites produced during the initial period of M. anisopliae infection may block the activation of detoxification components in the host.
PO is a key enzyme in insect immunity against pathogens. PO converts phenols to quinones, which subsequently polymerise to form melanin, which functions against both parasites and pathogens32,34. For example, in Spodoptera exempta, melanic larvae have higher PO activities and are more resistant to the entomopathogenic fungus, Beauveria bassiana, than non-melanic larvae55,56. Interestingly, high PO levels in insects might also increase resistance to insecticides. For example, insecticide-resistant diamondback moths display higher PO activities than susceptible moths36. In the present study, PO activities increased following treatment with either chlorantraniliprole or M. anisopliae, alone. In stark contrast, PO activities decreased significantly during the initial period after combined treatment with chlorantraniliprole and M. anisopliae, suggesting an antagonistic interaction of the insecticide and the pathogen in the host. Hiromori and Nishigaki have reported that mixed application of M. anisopliae and the insecticides teflubenzuron or fenitrothion inhibit PO activity in larvae of the beetle Anomala cuprea, and that granular cells are supressed3. These authors speculated that the observed interaction might be due to inhibition of the larval humoral defence and cellular immune systems. Moreover, joint treatment with M. anisopliae and an insecticide caused significant decrease in encapsulation intensity in the beetle Leptinotarsa decemlineata46. The interaction observed in our L. migratoria experiments might also be caused by PO inhibition and weakened humoral defence or cellular immune systems.
AChE is a key enzyme that terminates nerve impulses by catalysing the hydrolysis of the neurotransmitter acetylcholine in the nervous system of various organisms57. As such, AChE is the primary target site in the central nervous system for organophosphate and carbamate insecticides41. In our study, AChE activities decreased during the later period after M. anisopliae infection. A previous study has demonstrated the inhibitory effect of secondary metabolites on AChE in synapses of B. bassiana42. Inhibition of AChE causes acetylcholine to accumulate at synapses, such that post-synaptic membranes remain in a state of permanent stimulation; the result is paralysis, ataxia, a general lack of coordination in the neuromuscular system and eventual death58. In our experiment, however, AChE activities significantly increased throughout the entire experimental period, except for the third day, when locusts were treated with a combination of M. anisopliae and chlorantraniliprole. We speculated that the synergy between chlorantraniliprole and M. anisopliae affected AChE activity, thus mitigating symptoms of paralysis and ataxia.
CHIs are present in both fungi and insects. In fungi, they are involved in cell growth and division. In insects, they degrade chitin in the peritrophic membrane and exoskeletal cuticle and play an important role in moulting. Accumulating evidence suggests that CHI may also have roles in organismal defence against pathogenic fungi and parasites59,60,61. AA catalyses the hydrolysis of various anilide derivatives and esters and transfers an acetyl group to aniline, which acts as an acetyl acceptor62. In larvae of the diamondback moth, the specific activity of AA is significantly inhibited by chlorantraniliprole43. In our study, CHI and AA activities increased during the initial period after combined chlorantraniliprole and M. anisopliae treatment, thus helping the fungi to colonise the host. However, CHI activities decreased later in the experiment. We speculated that the synergy between chlorantraniliprole and M. anisopliae affected CHI and AA activities and disrupted the insect’s moulting process.
The major components of an insect’s antioxidant defence system include several antioxidant enzymes, such as superoxide dismutase (SOD), catalase (CAT) and peroxidase (POD)27. These enzymes remove damaging reactive oxygen species (ROS), such as superoxide anions (O2•−), hydrogen peroxide (H2O2), singlet oxygen molecules (O2•) and hydroxyl radicals (OH•−)30. ROS are regularly synthesised in the guts of insects following natural bacterial infections. The dynamic cycle of ROS generation and elimination appears to be a continuous and essential process in insects63,64. Studies in the grasshopper Oxya chinensis have shown increased SOD and CAT activities after treatment with phoxim, malathion and chlorpyrifos insecticides65,66. Likewise, SOD, CAT and POD activities have been shown to increase after selection by chlorpyrifos for eight generations in Nilaparvata lugens planthoppers67. SOD activity increases in the midgut of Galleria mellonella L. moth larvae on days 1–3 after infection by Bacillus thuringiensis, whereas CAT activity decreases during this time68. In our study, when L. migratoria were treated with M. anisopliae alone, CAT, SOD and POD activities increased during the initial period of infection. It should be noted that SOD and CAT, together, take part in stepwise oxygen reduction. Enhanced SOD activity in the infected nymph should elevate H2O2 concentrations and increase CAT activity. When chlorantraniliprole was applied in combination with M. anisopliae, SOD and POD activities increased but CAT activities decreased during the initial period post infection. We assumed that CAT was inhibited by the accumulation of superoxide radicals generated during the destruction processes. Research has shown that SOD1 protects calcineurin (CaN) from inactivation by ROS69. The enhanced activities of SOD and CAT then lead to the elimination of ROS68. In our study, the decreased activities of SOD, POD and CAT suggested a sharp decline in the ability to eliminate ROS during the later phase of infection. Large quantities of generated ROS can rapidly denature a wide range of biomolecules, including lipids, proteins and nucleic acids, thereby threatening virtually all cellular processes and leading to insect death27. Our results indicate a clear synergistic interaction between chlorantraniliprole and M. anisopliae infection when applied in combination.
Summarising locust enzyme responses to combined insecticide and pathogen treatment, it appears that chlorantraniliprole increased the probability of M. anisopliae infection and that M. anisopliae weakened L. migratoria defence against chlorantraniliprole. Our results suggest that combined treatment with chlorantraniliprole and M. anisopliae decreased GST, EST and PO activities during the initial post-infection period. Likewise, our results suggest that L. migratoria immunity to chlorantraniliprole and M. anisopliae was weakened by increased activities of AA and CHI during the initial post-infection period, whereas the activities of SOD, POD, CAT decreased during the later period. At the same time, combined treatment with chlorantraniliprole and M. anisopliae relieved the convulsions that normally accompany M. anisopliae infection by increasing the activity of AChE.
Based on previous and current results, we suggest that L. migratoria normally activates enzymatic defences against either pathogen or insecticide assault. Although these biochemical defences are sometimes successful, the combination of these two agents is able to overcome this defence. What is the chemical basis of this synergistic effect? One possibility is the disruption of Ca2+ concentrations in locust cells, given that both chlorantraniliprole and M. anisopliae are known to strongly disrupt Ca2+ balances in insects. For example, PO functions in an insect’s immune response to pathogens, but PO activity is regulated by Ca2+ concentration70,71. Hence, disrupting the Ca2+ balance could disrupt PO activity, thereby weakening the host’s immunity to M. anisopliae. Our results also reveal some of the complex biochemical processes that underlie the synergistic action of the two agents tested herein and illuminate the enzymatic mechanisms involved in insect immunity to pathogens and detoxification of insecticides. Understanding the response and activity of each enzyme alone and in combination will allow the design of novel and more effective means for pest control. Our results should provide guidance for future studies of the biochemical mechanisms underlying M. anisopliae’s disruption of host immunity.
Healthy 3rd-instar Oriental Migratory Locusts, Locusta migratoria (Meyen), were used in this study. Locusts were originally collected from Cangzhou, Hebei Province, China, then reared for 13 generations in the laboratory without exposure to insecticides. The eggs were hatched in a growth chamber maintained at 30 ± 2 °C and 60 ± 5% relative humidity (RH) for 2 wks. After hatching, the nymphs were fed fresh wheat sprouts and were maintained in a cage (60 × 50 × 70 cm) in the laboratory at 30 ± 2 °C and 60 ± 5% RH under a 14:10-h light:dark photoperiod.
Fungus and synthetic insecticide
The entomopathogenic fungus Metarhizium anisopliae (Metschnikoff) Sorokin IMI330189 was cultured on potato dextrose agar yeast extract (PDAY) at 27 ± 1 °C and 75 ± 10% RH for 7–10 d under constant light. Conidia were harvested from culture plates by scraping the surface of the PDAY with a sterile mounted needle and were then placed into plastic centrifuge tubes containing 0.1% Tween in sterile water. An oscillator was used to break up any aggregates. The spore concentration was determined using an improved Neubauer haemocytometer and then adjusted to a concentration of 2.5 × 108 spores/mL. For each experiment, this procedure was repeated to obtain a fresh suspension of spores. Chlorantraniliprole powder (96%) was provided by the Pesticide Science Group of the Institute of Plant Protection, Chinese Academy of Agricultural Sciences, Beijing, China. Preliminary trials showed that, at the doses tested (below) chlorantraniliprole did not significantly positively or negatively influence fungal growth on agar plates, nor did it influence fungal enzymes.
Efficacy of chlorantraniliprole against L. migratoria under laboratory conditions. The leaf-dip bioassay method described by Shelton et al. and Liang et al.72,73 was adopted for the toxicity bioassay of chlorantraniliprole. Wheat sprout bundles (3-cm diameter) were cut near the roots and dipped in various concentrations of chlorantraniliprole (shown in Table 3) prepared with distilled water. A bundle dipped in distilled water was used as a control. Each bundle was dipped for 10 s and allowed to air dry at room temperature, and three replicates were performed for each concentration analysed. Each bundle was placed inside a separate plastic container (30 × 22 × 9 cm) in which 30 third-instar nymphs were confined. Nymph mortality was recorded at 24-h intervals for 6 d. Nymphs were recorded as dead if they did not move when probed with a camel-hair brush. L. migratoria were fed only wheat sprouts treated with chlorantraniliprole for 1 d; on the other 5 d they were fed fresh, non-contaminated wheat sprouts. We used a similar protocol to obtain insecticide-challenged locusts (chlorantraniliprole, 5 mg/L, 0.5 mg/L and 0.05 mg/L) for our enzyme studies. Any surviving nymphs were used for the enzyme activity analyses (see below).
Efficacy of M. anisopliae against L. migratoria under laboratory conditions. Five conidial suspensions were prepared at various concentrations in sterile water containing 0.1% Tween 80 (shown in Table 3). Sterile water containing 0.1% Tween 80 was used as a control. Third-instar nymphs were used for all experiments. A total of 30 third-instar nymphs were treated per conidial concentration, with 2 mL of each conidial suspension applied as a spray under a Potter Precision Spray Tower (Burkard Manufacturing, Rickmansworth, UK). Three replicates each of five concentrations were tested. After being subjected to the conidial suspension spray treatment, nymphs were confined in each container and fed fresh wheat sprouts. Mortality was recorded at 24-h intervals for 6 d; nymphs were recorded as dead if they did not move when probed with a camel-hair brush. We used a similar protocol to obtain M. anisopliae-challenged locusts (M. anisopliae, 2.5 × 108, 2.5 × 107 and 2.5 × 106 spores/mL) for our enzyme studies. Any surviving nymphs were used for the enzyme activity analyses (see below).
Efficacy of chlorantraniliprole combined with M. anisopliae against L. migratoria under laboratory conditions. Chlorantraniliprole was combined with M. anisopliae in four dose-combinations for each of three proportions (Treatment groups 1, 2, & 3) (Table 3). A total of 30 third-instar nymphs were treated per solution by spraying under a Potter Precision Spray Tower with 2 mL of conidial suspension. Treatments were performed in triplicate. After being sprayed with a conidial suspension, nymphs were confined in containers and fed wheat sprouts treated with various concentrations of chlorantraniliprole for 1 d; on the other 5 d they were fed fresh, non-contaminated wheat sprouts. Mortality was recorded at 24-h intervals for 6 d; nymphs were recorded as dead if they did not move when probed with a camel-hair brush. We used a similar protocol to obtain chlorantraniliprole+ M. anisopliae-challenged locusts (1 mg/L + 2.0 × 108 spores/mL, 0.1 mg/L + 2.0 × 107 spores/mL and 0.01 mg/L + 2.0 × 106 spores/mL) for our enzyme studies. Surviving nymphs were used for the enzyme activity analyses (see below).
Efficacy against L. migratoria after 3-d exposure to M. anisopliae and chlorantraniliprole under laboratory conditions. Treatment 4: The concentrations of M. anisopliae solutions applied were similar to those used in Treatment 1 (shown in Table 3). After being sprayed with a conidial suspension, nymphs were confined in each container and fed fresh wheat sprouts for 3 d. On the 4th day, the nymphs were fed wheat sprouts treated with various concentrations of chlorantraniliprole (shown in Table 3, Treatment 4), and for the following 2 d the nymphs were fed fresh wheat. Mortality, which was assessed as described above for earlier experiments, was recorded at 24-h intervals for 6 d.
For enzyme assays, we took three 3rd-instar nymphs per insecticide x pathogen dose-combination with three replicates. Nymphs were homogenised in various solutions of 0.1 M ice-cold phosphate buffer (PBS) at different pH levels as follows: pH 7 (ESTs: 1.5 mL PBS with 0.1% TritonX-100; AChE and PO: 1.5 mL PBS), pH 7.3 (MFO: 1 mL PBS with 1 mM EDTA and 1 mM DTT; SOD, POD and CAT: 1.5 mL PBS), and pH 7.5 (AA, CHI and GST: 1.5 mL PBS). Extracted samples were centrifuged at 10,000 × g for 10 min at 4 °C. The resulting supernatant was transferred to a new Eppendorf tube and centrifuged at 15,000 × g for 20 min at 4 °C. The supernatant from this final centrifugation was used to determine enzyme activities and protein concentrations with five replicates were performed for each sample.
Enzyme activity assays
The activities of ESTs were assayed using a modification of the method described by Han et al.74. Enzyme activity was determined by kinetic analysis using a microplate reader (Molecular Devices, LA, CA, USA), with 100 μL of 1-naphthyl acetate solution (10 mM), 100 μL Fast Blue RR salt (1 mM) and 90 μL PBS placed in each microplate well. The reaction was initiated by the addition of 10 μL of enzyme solution. Optical densities (ODs) at 450 nm were recorded at 25-s intervals for 10 min. All reactions were carried out at 27 °C. Enzyme activities were calculated as the rate of absorbance change per mg protein (mOD/min/mg).
GST activity was measured using a modification of the method described by Oppenoorth and Welling75. After pipetting 100 μL of 1-chloro-2,4-dinitrobenzene (CDNB) (20 mM) or 3,4-dichloronitrobenzene (DCNB) (40 mM) and 100 μL of GSH (40 mM) into microplate wells, we added 50 μL of enzyme solution (for DCNB) or 10 μL of enzyme solution and 90 μL of PBS (for CDNB). The OD values at 340 nm were recorded at 25-s intervals for 10 min. Enzyme activities were calculated as the rate of absorbance change per mg protein (mOD/min/mg).
MFO activity was assayed using a method modified from Hansen and Hodgson76. After pipetting 100 μL of p-nitroanisole (2 mM) and 50 μL of nicotinamide adenine dinucleotide 2′-phosphate reduced tetrasodium salt (9.6 mM) into microplate wells, we added 50 μL of enzyme solution. The OD values at 405 nm were recorded at 25-s intervals for 10 min. Enzyme activities were calculated as the rate of absorbance change per mg protein (mOD/min/mg).
AChE activity was assayed using a modification of the method described by Han et al.74. After pipetting 100 μL of 5,5′-dithiobis-(2-nitrobenzoic acid) (45 μM), 100 μL of acetylthiocholine iodide and 90 μL of PBS into microplate wells, we added 50 μL of enzyme solution. The OD values at 405 nm were recorded at 30-s intervals for 40 min. Enzyme activities were calculated as the rate of absorbance change per mg protein (mOD/min/mg).
PO activity was measured using a method modified from that described by Luo and Xue77. We pipetted 180 μL of catechol (10 mM) and 20 μL of enzyme solution into microplate wells and recorded the OD values at 420 nm at 1-min intervals for 1 h. Enzyme activities were calculated as the rate of absorbance change per mg protein (mOD/min/mg).
SOD, POD and CAT activities were determined using commercial assay kits (Nanjing Jiancheng, Nanjing, China) according to the manufacturer’s instructions. SOD, POD and CAT enzyme activities were measured in units of U/mg, U/mg and U/g, respectively.
CHI activity was measured by using a reducing-sugar assay. A 400-μL reaction mixture containing 300 μL of 1% (w/v) colloidal chitin (prepared based on the methods of Hsu and Lockwood78) and 100 μL of enzyme solution was incubated at 37 °C in an Eppendorf tube. After 4 h of incubation, the mixture was centrifuged at 8,000 × g for 5 min to precipitate the remaining chitin. We mixed 200 μL of supernatant with 80 μL of potassium tetraborate (0.8 M, pH 9.1). The reaction was terminated by boiling at 100 °C for 5 min, after which it was cooled to room temperature under running water. The mixture was mixed with 1.2 mL of dimethylamine borane (DMAB) (10 g DMAB diluted to 1,000 mL with distilled water and then diluted 10-fold with glacial acetic acid). This mixture was then incubated at 37 °C for 20 min and subsequently cooled to room temperature under running water. The absorbance was detected at 585 nm, with CHI activity measured as the change of absorbance per mg protein (ΔOD/min/mg).
A method modified from Tang et al.44 was used to assay AA activity. A reaction mixture comprising 50 μL enzyme solution, 50 μL p-nitroacetanilide (1.2 mM in absolute ethanol) and 150 μL PBS was incubated in water at 35 °C for 30 min. The reaction was terminated by boiling in water for 10 min, and the mixture was then centrifuged at 10,000 × g for 15 min. The p-nitroaniline released into the supernatant was measured at 405 nm, and boiled enzyme was used as a control. AA activity was expressed as the change of absorbance per mg protein (ΔOD/min/mg).
Sample protein concentrations were estimated by using the method described by Bradford79. Bovine serum albumin was used for the calibration curve. Measurements were performed at 595 nm using a microplate reader with SoftMax Pro 6.1 software.
Co-toxicity coefficient values were calculated as described by Sun and Johnson80. Co-toxicity coefficients for the mixed formulation were calculated after calculating the LC50 of each incorporated component. Calculations were performed using the following equations: (1) toxicity index of agent = (LC50 of standard agent/LC50 of supplied agent) ×100; (2) theoretical toxicity index of the mixed formulation = (toxicity index of agent 1 × percentage of agent 1 in the mixed formulation) + (toxicity index of agent 2 × percentage of agent 2 in the mixed formulation); (3) co-toxicity coefficient = (actual toxicity index of the mixed formulation/theoretical toxicity index of the mixed formulation) ×100.
A co-toxicity coefficient of 100 indicates that the effect of the mixture is identical to that predicted from the proportions of the two components. A co-toxicity coefficient significantly greater than 100 corresponds to a synergistic effect. In contrast, when a mixture is characterised by a co-toxicity coefficient less than 100, the effect of the mixture is antagonistic.
LC50 values and concentration-mortality slopes for each bioassay were estimated via probit analysis81 using POLO-PC software82. Significant differences among LC50 values were determined on the basis of non-overlapping 95% confidence limits. In the figures, each symbol point represents average fold change values of five sub-replicates of enzyme activities compare with control. One-way ANOVAs were also used to analyse the activities of detoxification enzymes (ESTs, GST and MFO), protective enzymes (SOD, CAT and POD) and PO, AChE, CHI and AA in L. migratoria nymphs. Differences among means were compared by using the LSD test at P < 0.05.
How to cite this article: Jia, M. et al. Biochemical basis of synergism between pathogenic fungus Metarhizium anisopliae and insecticide chlorantraniliprole in Locusta migratoria (Meyen). Sci. Rep. 6, 28424; doi: 10.1038/srep28424 (2016).
Hill, D. S. Agricultural insect pests of the tropics and their control 2nd ed. 1–118 (Cambridge, 1983).
Bitsadze, N. et al. Joint action of Beauveria bassiana and the insect growth regulators diflubenzuron and novaluron, on the migratory locust, Locusta migratoria . J. Pest Sci. 86, 293–300 (2013).
Hiromori, H. & Nishigaki, J. Factor analysis of synergistic effect between the entomopathogenic fungus Metarhizium anisopliae and synthetic insecticides. Appl. Entomol. Zoo . 36, 231–236 (2001).
Lomer, C. J. et al. Field Infection of Zonocerus variegatus following application of an oil-based formulation of Metarhizium flavoviride conidia. Biocontrol Sci. Techn. 3, 337–346 (1993).
Peng, G. X., Wang, Z. K., Yin, Y. P., Zeng, D. Y. & Xia, Y. X. Field trials of Metarhizium anisopliae var. acridum (Ascomycota: Hypocreales) against oriental migratory locusts, Locusta migratoria manilensis (Meyen) in Northern China. Crop Prot. 27, 1244–1250 (2008).
Mnyone, L. L., Ng’habi, K. R., Mazigo, H. D., Katakweba, A. A. & Lyimo, I. N. Entomopathogenic fungi, Metarhizium anisopliae and Beauveria bassiana reduce the survival of Xenopsylla brasiliensis larvae (Siphonaptera: Pulicidae). Parasite. Vector . 5, 204 (2012).
Price, R. E., Bateman, R. P., Brown, H. D., Butler, E. T. & Müller, E. J. Aerial spray trials against brown locust (Locustana pardalina, Walker) nymphs in South Africa using oil-based formulations of Metarhizium flavoviride . Crop Prot. 16, 345–351 (1997).
Zhang, Z. H. et al. Using Metarhizium flavoviride Oil Spray to Control Grasshoppers in Inner Mongolia Grassland. Chinese J. of Biol. Control . 16, 49–52 (2000).
Lomer, C. J., Bateman, R. P., Johnson, D. L., Langewald, J. & Thomas, M. Biological control of locusts and grasshoppers. Annu. Rev. Entomol. 46, 667–702 (2001).
Jaramillo, J. et al. Effect of combined applications of Metarhizium anisopliae (Metsch.) Sorokin (Deuteromycotina: Hyphomycetes) strain CIAT 224 and different dosages of imidacloprid on the subterranean burrower bug Cyrtomenus bergi Froeschner (Hemiptera: Cydnidae). Biol. Control. 34, 12–20 (2005).
Purwar, J. P. & Sachan, G. C. Synergistic effect of entomogenous fungi on some insecticides against Bihar hairy caterpillar Spilarctia obliqua (Lepidoptera: Arctiidae). Microbiol. Res. 161, 38–42 (2006).
Sharififard, M., Mossadegh, M. S., Vazirianzadeh, B. & Zarei-Mahmoudabadi, A. Interactions between Entomopathogenic fungus, Metarhizium anisopliae and sublethal doses of spinosad for control of house fly, Musca domestica . Iran. J. Arthropod-Bor . 5, 28–36 (2011).
Bandyopadhyay, A., Shin, D., Ahn, J. O. & Kim, D. H. Calcineurin regulates ryanodine receptor/Ca2+-release channels in rat heart. Biochem. J. 352, 61–70 (2000).
Lahm, G. P. et al. Rynaxypyr: A new insecticidal anthranilic diamide that acts as a potent and selective ryanodine receptor activator. Bioorg. Med. Chem. Lett. 17, 6274–6279 (2007).
Cordova, D. et al. Anthranilic diamides: a new class of insecticides with a novel mode of action, ryanodine receptor activation. Pestic. Biochem. Phys . 84, 196–214 (2006).
Pedras, M. S. C., Zaharia, L. I. & Ward, D. E. The destruxins: synthesis, biosynthesis, biotransformation, and biological activity. Phytochemistry 59, 579–596 (2002).
Ruiz-Sanchez, E., Lange, A. B. & Orchard, I. Effects of the mycotoxin destruxin A on Locusta migratoria visceral muscles. Toxicon 56, 1043–1051 (2010).
Serebrov, V. V. et al. Effect of entomopathogenic fungi on detoxification enzyme activity in greater wax moth Galleria mellonella L. (Lepidoptera, Pyralidae) and role of detoxification enzymes in development of insect resistance to entomopathogenic fungi. Biol. Bull. 33, 581–586 (2006).
Roslavtseva, S. A., Bakanova, E. I. & Eremina, O. Y. Esterases in Arthropods and their role in the mechanisms of Insect acaricide detoxication. Izv. Akad. Nauk Biol. 3, 368–375 (1993).
Terriere, L. C. Induction of Detoxication Enzymes in Insects. Annu. Rev. Entomol. 29, 71–88 (1984).
Dubovskiy, I. M. et al. The activity of nonspecific esterases and glutathione- S-transferase in Locusta migratoria larvae infected with the fungus Metarhizium anisopliae (Ascomycota, Hypocreales). Entomol. Rev . 92, 27–31 (2012).
Li, X., Schuler, M. A. & Berenbaum, M. R. Molecular mechanisms of metabolic resistance to synthetic and natural xenobiotics. Annu. Rev. Entomol. 52, 231–253 (2007).
Feyereisen, R. Insect cytochrome P450 in Comprehensive Molecular Insect Science, Vol. 4 (eds Gilbert, L. I. et al. ) 1–77 (Elsevier, 2005).
Enayati, A. A., Ranson, H. & Hemingway, J. Insect glutathione transferases and insecticide resistance. Insect Mol. Biol. 14, 3–8 (2005).
Lumjuan, N., McCarroll, L., Prapanthadara, L., Hemingway, J. & Ranson, H. Elevated activity of an Epsilon class glutathione transferase confers DDT resistance in the dengue vector, Aedes aegypti . Insect Biochem. Molec . 35, 861–871 (2005).
Ortelli, F., Rossiter, L. C., Vontas, J., Ranson, H. & Hemingway, J. Heterologous expression of four glutathione transferase genes genetically linked to a major insecticide-resistance locus from the malaria vector Anopheles gambiae. Biochem. J. 373, 957–963 (2003).
Felton, G. W. & Summers, C. B. Antioxidant Systems in insects. Arch. Insect Biochem . 29, 187–197 (1995).
Müller, P., Donnelly, M. J. & Ranson, H. Transcription profiling of a recently colonised pyrethroid resistant Anopheles gambiae strain from Ghana. BMC Genomics 8, 36 (2007).
Wu, Q. J., Zhang, Y. J., Xu, B. Y. & Zhang, W. J. The defending enzymes in abamectin resistant Plutella xylostella . Chinese J. Appl. Entomol. 48, 291–295 (2011).
Campa-Córdova, A. I., Hernández-Saavedra, N. Y. & Ascencio, F. Superoxide dismutase as modulator of immune function in American white shrimp (Litopenaeus vannamei). Comp. Biochem. Physiol. C Pharmacol. Toxicol. Endocrinol . 133, 557–565 (2002).
Gopalakrishnan, S. et al. Modulation and interaction of immune-associated parameters with antioxidant in the immunocytes of Crab Scylla paramamosain Challenged with lipopolysaccharides. Evid.-Based Compl. Alt . 2011, 1–8 (2011).
Söderhäll, K. & Cerenius, L. Role of the prophenoloxidase-activating system in invertebrate immunity. Curr. Opin. Immunol. 10, 23–28 (1998).
Carton, Y., Poirié, M. & Nappi, A. J. Insect immune resistance to parasitoids. Insect Sci. 15, 67–87 (2008).
Cerenius, L. & Söderhäll, K. The prophenoloxidase-activating system in invertebrates. Immunol. Rev. 198, 116–126 (2004).
Chang, C. C., Rahmawaty, A. & Chang, Z. W. Molecular and immunological responses of the giant freshwater prawn, Macrobrachium rosenbergii, to the organophosphorus insecticide, trichlorfon. Aquat. Toxicol. 130, 18–26 (2013).
Liu, S. et al. Does phenoloxidase contributed to the resistance? Selection with butane-fipronil enhanced its activities from diamondback moths. Open Biochem. J. 3, 9–13 (2009).
Oehmichen, M. & Besserer, K. Forensic significance of acetylcholine esterase histochemistry in organophosphate intoxication. Int. J. Legal Med. 89, 149–165 (1982).
Fournier, D. & Mutero, A. Modification of acetylcholinesterase as a mechanism of resistance to insecticides. Comp. Biochem. Physiol. C Pharmacol. Toxicol. Endocrinol . 108, 19–31 (1994).
Bretaud, S., Toutant, J. P. & Saglio, P. Effects of Carbofuran, Diuron, and Nicosulfuron on Acetylcholinesterase Activity in Goldfish (Carassius auratus). Ecotox. Environ. Safe . 47, 117–124 (2000).
Yoo, J. K., Lee, S. W., Ahn, Y. J. & Shono, T. Altered acetylcholinesterase as a resistance mechanism in the brown planthopper (Homoptera: Delphacidae), Nilaparvata lugens Stal . Appl. Entomol. Zool. 37, 37–42 (2002).
Senthil, N. S. et al. Effect of azadirachtin on acetylcholinesterase (AChE) activity and histology of the brown planthopper Nilaparvata lugens (Stål). Ecotox. Environ. Safe . 70, 244–250 (2008).
Zibaee, A., Bandani, A. R. & Tork, M. Effect of the entomopathogenic fungus, Beauveria bassiana, and its secondary metabolite on detoxifying enzyme activities and acetylcholinesterase (AChE) of the Sunn pest, Eurygaster integriceps (Heteroptera: Scutellaridae). Biocontrol Sci. Techn . 19, 485–498 (2009).
Xing, J., Liang, P. & Gao, X. W. Effects of sublethal concentrations of chlorantraniliprole on insecticide susceptibility and detoxifying enzyme activity in Plutella xylostella . Chinese J. Pestic. Sci. 13, 464–470 (2011).
Tang, B. Z., Sun, J. Y., Zhou, X. G., Gao, X. W. & Liang, P. The stability and biochemical basis of fufenozide resistance in a laboratory-selected strain of Plutella xylostella . Pestic. Biochem. Phys. 101, 80–85 (2011).
Ou, S. S., Liang, P., Song, D. L., Shi, X. Y. & Gao, X. W. Effects of sublethal dosage of chlorantraniliprole on development and detoxifying enzymes activity of Helicoverpa armigera . Plant Prot . 38, 1–8 (2012).
Dubovskiy, I. M. et al. Activity of the detoxificative enzyme system and encapsulation rate in the Colorado potato beetle Leptinotarsa decemlineata (Say) larvae under organophosphorus insecticide treatment and entomopathogenic fungus Metharizium anisopliae (Metsch.) infection. Euroasian Entomol. J . 9, 577–582 (2010).
Anderson, T. E., Hajek, A. E., Roberts, D. W., Preisler, H. K. & Robertson, J. L. Colorado potato beetle (Coleoptera: Chrysomelidae): effects of combinations of Beauveria bassiana with insecticides. J. Econ. Entomol. 82, 83–89 (1989).
Cao, G. C. et al. Toxicity of chlorantraniliprole to Cry1Ac-susceptible and resistant strains of Helicoverpa armigera . Pestic. Biochem. Physiol. 98, 99–103 (2010).
Lai, T. C., Li, J. & Su, J. Y. Monitoring of beet armyworm Spodoptera exigua (Lepidoptera: Noctuidae) resistance to chlorantraniliprole in China. Pestic. Biochem. Physiol. 101, 198–205 (2011).
Sial, A. A., Brunner, J. F. & Garczynski, S. F. Biochemical characterization of chlorantraniliprole and spinetoram resistance in laboratory-selected obliquebanded leafroller, Choristoneura rosaceana (Harris) (Lepidoptera: Tortricidae). Pestic. Biochem. Physiol. 99, 274–279 (2011).
Serebrov, V. V., Alekseev, A. A. & Glupov, V. V. Changes in Activity and Pattern of Hemolymph Esterases in Larvae of Wax Moth Galleria mellonella L. (Lepidoptera, Pyralidae) during Mycosis. Izv. Akad. Nauk Ser. Biol. 28, 588–592 (2001).
Androsov, G. K. & Alieva, M. I. Protective Responses of Insect Hemolymph to Mycotoxicosis. Zh. Obshch. Biol. 41, 726–733 (1980).
Hall, I. M. Microbial Control in Insect Pathology An Advanced Treatise (ed. Steinhaus, E. A. ) 477–511 (Academic, 1963).
Furlong, M. J. & Groden, E. Evaluation of synergistic interactions between the Colorado potato beetle (Coleoptera: Chrysomelidae) pathogen Beauveria bassiana and the insecticides, imidacloprid, and cyromazine. J. Econ. Entomol. 94, 344–356 (2001).
Reeson, A. F., Wilson, K., Gunn, A., Hails, R. S. & Goulson, D. Baculovirus resistance in the noctuid Spodoptera exempta is phenotypically plastic and responds to population density. Proceedings of the Royal Society of London Series B:Biological Sciences . 265, 1787–1791 (1998).
Whitman, D. W. & Ananthakrishnan, T. N. Phenotypic Plasticity of Insects: Mechanisms and Consequences (eds Whitman, D. W. et al. ) 202–243 (Science, 2009).
Wang, J. J., Cheng, W. X., Ding, W. & Zhao, Z. M. The Effect of the Insecticide Dichlorvos on Esterase Activity Extracted from the Psocids, Liposcelis bostrychophila and L. entomophila . J. Insect Sci. 4, 1–5 (2004).
Singh, K. & Singh, D. K. Toxicity to the Snail Limnaea acuminata of Plant-derived Molluscicides in Combination with Synergists. Pest Manag. Sci. 56, 889–898 (2000).
Wang, X. et al. Characterization of a 46 kDa insect chitinase from transgenic tobacco. Insect Biochem. Molec . 26, 1055–1064 (1996).
Kramer, K. J. & Muthukrishnan, S. Insect chitinases: molecular biology and potential use as biopesticides. Insect Biochem. Molec . 27, 887–900 (1997).
Ding, X. et al. Insect resistance of transgenic tobacco expressing an insect chitinase gene. Transgenic Res. 7, 77–84 (1998).
Yoshioka, H., Nagasawa, T. & Yamada, H. Purification and characterization of aryl acylamidase from Nocardia globerula. Eur. J. Biochem. 199, 17–24 (1991).
Ahmad, S. Biochemical defense of pro-oxidant plant allelochemicals by herbivorous insects. Biochem. Syst. Ecol. 20, 269–296 (1992).
Felton, G. Antioxidant defenses of invertebrates and vertebrates in Oxidative Stress and Antioxidant Defenses in Biology (ed. Ahmad, S. ) 356–434. (Springer, 1995).
Wu, H. et al. Biochemical effects of acute phoxim administration on antioxidant system and acetylcholinesterase in Oxya chinensis (Thunberg) (Orthoptera: Acrididae). Pestic. Biochem. 100, 23–26 (2011).
Wu, H. H., Zhang, R., Liu, J. Y., Guo, Y. P. & Ma, E. B. Effects of malathion and chlorpyrifos on acetylcholinesterase and antioxidant defense system in Oxya chinensis (Thunberg) (Orthoptera: Acrididae). Chemosphere 83, 599–604 (2011).
Ling, S. & Zhang, H. Influences of chlorpyrifos on antioxidant enzyme activities of Nilaparvata lugens . Ecotox. environ. Safe . 98, 187–190 (2013).
Dubovskiy, I. M. et al. Effect of bacterial infection on antioxidant activity and lipid peroxidation in the midgut of Galleria mellonella L. larvae (Lepidoptera, Pyralidae). Comp. Biochem. Physiol . C Pharmacol. Toxicol. Endocrinol . 148, 1–5 (2008).
Wang, X., Culotta, V. C. & Klee, C. B. Superoxide dismutase protects calcineurin from inactivation. Nature 383, 434–437 (1996).
Gollas-Galván, T., Hernández-López, J. & Vargas-Albores, F. Effect of calcium on the prophenoloxidase system activation of the brown shrimp (Penaeus californiensis, Holmes). Comp. Biochem. Physiol. A Physiol . 117, 419–425 (1997).
Perdomo-Morales, R., Montero-Alejo, V., Perera, E., Pardo-Ruiz, Z. & Alonso-Jiménez, E. Phenoloxidase activity in the hemolymph of the spiny lobster Panulirus argus . Fish shellfish immune . 23, 1187–1195 (2007).
Shelton, A. M. et al. Resistance of diamondback moth (Lepidoptera: Yponomeutidae) to Bacillus thuringiensis subspecies in the field. J. Econ. Entomol. 86, 697–705 (1993).
Liang, P., Gao, X. W. & Zheng, B. Z. Genetic basis of resistance and studies on cross-resistance in a population of diamondback moth, Plutella xylostella (Lepidoptera: Plutellidae). Pest Manag. Sci. 59, 1232–1236 (2003).
Han, Z. J., Moores, G., Devonshire, A. & Denholm, L. Association between biochemical marks and insecticide resistance in the cotton aphid, Aphis gossypii . Pesti. Biochem. Phys . 62, 164–171 (1998).
Oppenoorth, F. J. & Welling, W. Biochemistry and physiology of resistance. Insecticide Biochemistry and Physiology (ed. Wilkinson, C. F. ) Ch. 13, 507–551 (Plenum, 1976).
Hansen, L. G. & Hodgson, E. Biochemical characteristics of insect microsomes N- and O-demethylation. Biochem. Pharmacol. 20, 1569–1573 (1971).
Luo, W. C. & Xue, C. B. Insects phenol oxidase and inhibitor 30 (Science, 2010).
Hsu, S. C. & Lockwood, J. L. Powdered chitin agar as a selective medium for enumeration of actinomycetes in water and soil. Appl. Microbiol . 29, 422–426 (1975).
Bradford, M. M. A rapid and sensitive method for the quantification of microgram quantities of protein utilizing the principle of protein binding. Anal. Biochem. 72, 248–254 (1976).
Sun, Y. P. & Johnson, E. R. Analysis of joint action of insecticides against houseflies. J. Econ. Entomol. 53, 887–892 (1960).
Finney, D. J. Probit Analysis, third ed. (Cambridge, 1971).
LeOra Software. POLO-PC: A User’s Guide to Probit Analysis or Logit Analysis. LeOra Software, Berkeley, CA (1987).
This research was supported by National Natural Science Foundation of China (No. 31471824) and The Special Fund for Agro-scientific Research in the Public Interest of China (201003079) and the earmarked fund for China Agriculture Research System (CARS-35-07) and Innovation Project of Chinese Academy of Agricultural Sciences.
The authors declare no competing financial interests.
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Jia, M., Cao, G., Li, Y. et al. Biochemical basis of synergism between pathogenic fungus Metarhizium anisopliae and insecticide chlorantraniliprole in Locusta migratoria (Meyen). Sci Rep 6, 28424 (2016). https://doi.org/10.1038/srep28424
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