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Resistance in the Genus Spodoptera: Key Insect Detoxification Genes

Université Côte D’Azur, INRAE, CNRS, ISA, F-06903 Sophia Antipolis, France
Institut D’Ecologie et des Sciences de L’Environnement de Paris, Sorbonne Université, CNRS, INRAE, IRD, iEES-Paris, F-75005 Paris, France
Author to whom correspondence should be addressed.
Insects 2021, 12(6), 544;
Original submission received: 18 May 2021 / Revised: 4 June 2021 / Accepted: 8 June 2021 / Published: 11 June 2021



Simple Summary

The moth larvae are among the most damaging pest species on crops worldwide. In this review, we focus on the genus Spodoptera, which can feed on many crops such as rice, cotton or corn. The massive use of insecticides to control these insects has led to the development of resistance. Here, we aim to compare the resistance mechanisms of four species (Spodoptera exigua, Spodoptera frugiperda, Spodoptera littoralis and Spodoptera litura) and highlight the role of enzymes and transporters in resistance to help us understand the molecular basis of their origin.


The genus Spodoptera (Lepidoptera: Noctuidae) includes species that are among the most important crop pests in the world. These polyphagous species are able to feed on many plants, including corn, rice and cotton. In addition to their ability to adapt to toxic compounds produced by plants, they have developed resistance to the chemical insecticides used for their control. One of the main mechanisms developed by insects to become resistant involves detoxification enzymes. In this review, we illustrate some examples of the role of major families of detoxification enzymes such as cytochromes P450, carboxyl/cholinesterases, glutathione S-transferases (GST) and transporters such as ATP-binding cassette (ABC) transporters in insecticide resistance. We compare available data for four species, Spodoptera exigua, S. frugiperda, S. littoralis and S. litura. Molecular mechanisms underlying the involvement of these genes in resistance will be described, including the duplication of the CYP9A cluster, over-expression of GST epsilon or point mutations in acetylcholinesterase and ABCC2. This review is not intended to be exhaustive but to highlight the key roles of certain genes.

Graphical Abstract

1. Introduction

The genus Spodoptera (Lepidotera: Noctuidae) contains some of the most important insect crop pests, many of which are highly polyphagous species, able to feed on more than 100 host plants including maize, rice, cotton (e.g., Spodoptera litura (Fabricius) [1]). They are present on all continents and their potential invasiveness has been highlighted in recent years, notably with the species S. frugiperda (J.E. Smith). Native to the American continent, it was detected in Africa in 2016 [2] and has since invaded Asia and Australia (CABI, Wallingford, UK, 2021). With its flight capabilities [3,4] and under favorable climatic conditions, its invasion of Europe in the near future seems inevitable.
The control of these pests requires the massive use of insecticides. They have developed resistance to all chemical families and three of the four Spodoptera species present in the Arthropod Pesticide Resistance Database are in the top 15 most resistant arthropods on the planet: S. litura, S. frugiperda and S. exigua (Hübner) [5].
Table 1 shows the available data on the molecules for which these three species, as well as S. littoralis (Boisduval) have developed resistance. These insects have developed resistance to all the chemical families: organophosphates, carbamates, pyrethroids but also for a more recent family such as diamides. There are usually two main mechanisms in insecticide resistance, either target modification or mechanisms that reduce the amount of insecticide reaching the target (reduced penetration, sequestration or intervention of detoxification enzymes). Detoxification involves enzymes that catalyze successive reactions to make the insecticidal molecule less toxic and more easily excreted from the body. Cytochromes P450 (P450) and carboxyl/cholinesterases (CCE) are phase I enzymes that catalyze oxidation, hydrolysis and reduction. Glutathione S-transferases (GST) are phase II enzymes and catalyze the addition of a group such as glutathione. This step is called conjugation and is followed by phase III excretion involving ATP-binding cassette (ABC) transporters. These proteins use the energy of ATP hydrolysis to transport substrates across lipid membranes. The resistance caused by the enzymes involved in these three phases implies either a modification of their level of expression or their catalytic activity. The molecular process behind this can be of several kinds. A point mutation in the sequence of a gene can modify the catalytic activity of the enzyme. For example, in the mosquito Anopheles funestus, the increased activity of GSTe2 is due to a point mutation (L119F) that leads to an increase in the accessibility of the active site, allowing high resistance to DDT [6]. Overexpression is one of the other commonly demonstrated mechanisms, involving duplications, amplifications or cis-or trans-regulations [7]. A well-known example of gene duplication is the case of CYP6G1 in Drosophila melanogaster. The overexpression of this gene in multi-insecticide resistant populations is due to duplication as well as insertion of transposable elements in its promoter [8]. In the aphid Myzus persicae, it is a gene amplification of one or two F4 and FE4 esterases that is at the origin of the resistance to organosphosphates, carbamates and pyrethroids [9]. Up to 80 copies of the same gene are found in some resistant aphids. The esterase in this case represented up to 3% of the total proteins [10]. A recent example illustrates cis and trans regulation in a chlorpyrifos-resistant strain of S. exigua [11]. The authors showed that the resistance was mainly due to the overexpression of a P450, CYP321A8, a P450 capable of metabolizing chlorpyrifos, cypermethrin and deltamethrin. This overexpression is due to both constitutive overexpression of the transcription factors Cap’n’collar isoform C (CncC) and Maf (Muscle aponeurosis fibromatosis, trans-regulation), major regulators of detoxification in insects [12], and to a mutation in the promoter of this P450. The mutation creates a cis-regulatory element that promotes binding of a Knirps nuclear receptor.
This review highlights some examples that demonstrate the involvement of each of the four families of detoxification genes (P450, CCE, GST and ABC) in insecticide resistance and the molecular mechanisms identified for pests of the genus Spodoptera. We have chosen to focus on specific genes that we consider emblematic rather than an exhaustive review of their roles.

2. Phase I: Functionalization

Once a xenobiotic enters the cell, it is processed by enzymes that make it functional. The phase I enzymes carry out oxidation reactions, hydrolysis, etc., making the metabolite more polar and more easily excreted.

2.1. Cytochrome P450s

Cytochrome P450 genes (CYP) constitute one of the largest gene families with representatives in nearly all living organisms [13]. CYPs catalyze a large number of reactions including hydroxylation, epoxidation, oxidation [14]. In insects, CYPs are involved in endogenous metabolism such as the biosynthesis of ecdysteroid [15] or the production of hydrocarbon [16], as well as in detoxification mechanisms with the metabolization of xenobiotics (e.g., plant secondary metabolites and insecticides). Their number varies from 36 in the louse Pediculus humanus [17] to 196 in the mosquito, Culex quinquefasciatus [18]. Nomenclature has been proposed based on sequence homology [19,20]. P450s belong to the same family designated by an Arabic numeral when they share 40% identity and to the same subfamily designated by capital letters when this percentage is higher than 55%. The nomenclature was revised and the notion of clan was proposed as a higher level of classification to take into account the increasing number of available sequences [21]. Initially, the arthropod P450s were divided into four clans, but a recent study brings this number to 6 with the historical clans, clan 2, clan 3, clan 4, mitochondrial clan to which clans 16 and 20 were added [22]. Clans 3 and 4 are the two main clans known to have P450s involved in resistance mechanisms, but not exclusively, for example, CYP12D1 from D. melanogaster which belongs to the mitochondrial clan is over-expressed in resistant strains [23] and has been shown to confer resistance to DDT and dicyclanil [24]. We will highlight the role of some of P450s in insecticide resistance and evolution of the CYP9A subfamily within the four Spodoptera species.

2.1.1. Phylogeny of CYP9A

CYP9A belongs to clan 3. In Lepidoptera, CYP9As were identified as a cluster in a comparative study of BAC sequences from three species Bombyx mori, Helicoverpa armigera and S. frugiperda [25] with the presence of four, five and nine genes respectively, showing the rapid evolution of the cluster. In the two more closely related species (H. armigera and S. frugiperda which diverged about ±20 MYA), only three orthologous pairs were identified [26]. The availability of the genome of the most closely related species in the genus Spodoptera enabled the evolution of this group to be assessed. In S. frugiperda, obtaining the chromosomally resolved genome of the maize strain helped identify 14 CYP9A genes (genome Version 6 [27]), 12 of which were in a cluster (Figure 1), i.e., three more than the nine genes previously identified in the BAC study, whereas 15 genes were identified in the rice strain (genome version 3 [28]). These numbers are very similar to those found in other species, 15 genes in S. litura (Figure 1) [1], 12 genes in S. exigua and 11 genes in S. littoralis (genome in progress, transcriptome available). In the latter, the number is potentially underestimated and improved genome resolution could lead to increases.
Among these genes, eight were true orthologs, CYP9A25, CYP9A26, CYP9A27, CYP9A28, CYP9A30, CYP9A31, CYP9A58 and CYP9A59 in S. frugiperda, S. litura and S. littoralis, probably already clustered in their common ancestor (Figure 2). CYP9A24 is only found in S. frugiperda. Some genes of the cluster are S. frugiperda corn strain specific (CYP9A32, CYP9A60, and CYP9A76), S. frugiperda rice strain specific (CYP9A91) or S. litura specific (CYP9A39-157-158-161-162). Orthologs of CYP9A26, CYP9A59 and CYP9A27 were also found in the genome of S. exigua.
The synteny of the CYP9 cluster is conserved between S. frugiperda (corn and rice variants) and S. litura (Figure 1) where the same genes, alcohol dehydrogenase (ADH), fucosyl transferase (FT) and E26 transformation specific (ETS, a transcription factor) were found on each side of the cluster. In S. litura, two gustatory receptors from the GR29 family are part of the CYP9 cluster. They form a head to tail tandem between CYP9A58 and CYP9A25 and they replace CYP9A24. The GR29 family is specific to Spodoptera and based on the evolutionary history of S. litura and S. littoralis [29], we expect GR genes to be present in the CYP9 cluster of S. littoralis. The mechanisms resulting in blooming in the CYP9A family in the Spodoptera complex still remain to be deciphered. Several origins are possible, one of which corresponds to transposable elements as they have been shown to be prevalent in the surrounding of CYP genes in Lepidoptera [25,30,31], Drosophila [32] or mosquito [33]. Other mechanisms include duplications (tandem, chromosome or genome duplications) and retropositions [26]. This CYP9A bloom likely provides Spodoptera species with a selective advantage and potentially diverse catalytic capabilities for each of these enzymes. Indeed, induction experiments show that within the CYP9A cluster, genes have their own induction profile depending on the xenobiotics used. For example, CYP9A31 is the most induced gene in response to xanthotoxin treatment in both S. frugiperda and S. litura whereas CYP9A28 is not induced by this molecule but by indole [34,35]. This suggests that within the cluster, each gene has its own catalytic competence unrelated to its position in the cluster or phylogeny as it has been suggested for the CYP6AE cluster in H. armigera [36]. Dermauw et al., (2020) suggested that there would be a selective advantage to keeping the cluster as a heritable unit, which would enable adaptation to new environments [22].

2.1.2. Resistance through Over-Expression of CYP9A

We next examine CYP9As in the light of insecticide resistance: the first CYP9A was found in Heliothis virescens and named CYP9A1 [37]. It was in a thiodicarb-resistant population of H. virescens. An elevated level of CYP9A1 mRNA was detected in the resistant strain compared to the susceptible strain. Several subsequent studies have associated over-expression of certain CYP9As with insecticide resistance, but very few studies have gone so far as to demonstrate their involvement in insecticide metabolism. An example came from H. armigera, where pyrethroid resistance was associated with constitutive over-expression of P450s in the laboratory-selected YGF strain [38]. In this strain, CYP9A12 and CYP9A14 were over-expressed 433-and 59-fold, respectively, in the fat body. The functional expression of these two P450s in Saccharomyces cerevisiae demonstrated their capacity to metabolize a pyrethroid insecticide, esfenvalerate [39]. Elevated levels of CYP9As associated with resistance have also been shown in Spodoptera spp. In a recent study, a Brazilian population of S. frugiperda resistant to Bacillus thuringiensis (Bt) Cry1 toxins was tested against 15 insecticide molecules with different modes of action. In addition to resistance to Bt toxins, the Sf-Des strain had showed 14- and eight-fold resistance to deltamethrin and chlorpyrifos, respectively [40]. P450 activity was increased in the Sf-Des strain compared to the susceptible Bt strain. RNAseq experiments confirmed the over-expression of several P450s, including CYP9As and RT-qPCR analysis validated that a CYP9A-like was expressed more than 200-fold in the resistant strain. In another study on a laboratory-selected population of S. frugiperda with lufenuron, a CYP9A-like gene was over-expressed 45-fold [41]. In both of these studies, the cause of the over-expression of the CYP9As was not identified, whereas Gimenez et al., (2020) reported the existence of two copies of the full CYP9A cluster of in a Puerto Rican resistant strain (PR) of S. frugiperda. PR was resistant to deltamethrin and the use of a P450 synergist, piperonil butoxide (PBO) abolished the resistance, confirming the primary role of P450. The CYP9A cluster locus is under positive selection in this strain [27]. Several field-collected strains of S. exigua resistant to organophosphates, pyrethroids and diamides also exhibited an over-expression of CYP9A. For a chlorpyrifos-resistant strain, the resistance was mainly associated with CYP321A8, which is over-expressed and capable of metabolising insecticides, however monitoring the expression of 68 P450s by RT-qPCR showed significant over-expression of several CYP9As including CYP9A11, CYP9A27, CYP9A97 and CYP9A98 [11]. Are these genes also involved in resistance? This remains to be demonstrated. However, there is some evidence of their involvement in resistance, such as the RNAi knockout (KO) of CYP9A98, which makes S. exigua larvae more sensitive to deltamethrin [42], the knockout of CYP9A10, which makes them more sensitive to alpha-cypermethrin [43], and the knockout of CYP9A105, which increases mortality to deltamethrin, alpha-cypermethrin, and fenvalerate [44]. Similar results were also obtained in the species S. litura. Indeed, strains resistant to fenvalerate, beta-cypermethrin and cyhalothrin had their resistance reduced by the use of PBO [45]. RNAseq experiments for these field-collected Chinese strains (LF and NJ) showed over-expression of several P450s including CYP9As. CYP9A40 was over-expressed 663.4- and 76.13-fold in LF and NJ, respectively, compared with the susceptible strain. Previous experiments showed that dsRNA silencing of CYP9A40 increased the sensitivity of S. litura to deltamethrin [46]. To our knowledge, no data suggesting a link between insecticide resistance and CYP9A of S. littoralis are available at this time. There is sufficient evidence across closely related species to suggest a major role for CYP9As at least in pyrethroid resistance; however, the ability of these enzymes to metabolise insecticides remains to be demonstrated. The cause of the over-expression of these genes also remains to be identified. In one case, it is due to the duplication of the CYP9A cluster [27] while in the study of Hu et al., (2021) it can be assumed that the over-expression of the transcription factor CncC, already shown in several insects to be the major regulator of detoxification genes, leads to the over-expression of CYP9A.

2.2. Carboxyl/Cholinesterases (CCEs)

Insect carboxylesterases or carboxyl/cholinesterases (CCEs) have a wide range of physiological functions across living organisms, from metabolism of endogenous compounds (hormones, pheromones, neurotransmitters) to detoxification of various xenobiotics. They play a critical role in defense against various allelochemicals associated with plants and insecticides [47]. In particular, CCEs have been implicated in resistance to pyrethroids (PYRs), organophosphates and carbamates (CBs) in numerous pest species [48,49]. The mechanisms of esterase-mediated resistance involve either metabolic resistance or target site mutation [49]. Metabolic resistance can be based either on insecticide sequestration or insecticide hydrolysis: overproduction of CCEs through gene amplification or transcriptional up-regulation can lead to insecticide molecule sequestration without any (or very slow) hydrolysis of the insecticide, whereas point mutations can alter the catalytic properties of some CCEs, leading to increased hydrolysis towards insecticides. Target site resistance is due to mutation of acetylcholinesterases (AChEs) that renders them less sensitive to inhibition by insecticides.
Insect CCEs fall into three main functional groups, representing dietary/detoxification, hormone/semiochemical processing, and neuro/developmental functions [50]. A comprehensive phylogenetic analysis of CCE sequences isolated from H. armigera, H. zea, B. mori and Manduca sexta genomes has recently refined the CCE nomenclature in Lepidoptera [51]. The neuro/developmental class comprises seven clades (027 to 033) with generally catalytically incompetent proteins, excepted for AChEs (clade 027). The hormone/semiochemical processing class (clades 020 to 026) includes among others, juvenile hormone esterases, several pheromone-processing esterases, but also some CCEs (clade 026) previously associated with organophosphate (OP) resistance in Hemiptera (namely E4, FE4, [10]). Finally, the dietary/detoxification group (clades 001 to 019 and 034) is the most diversified: some enzymes have been implicated in semiochemical processing but most of them have been associated with dietary and detoxification functions, including insecticide resistance for some dipteran and hymenopteran CCEs (the α-esterases, reviewed in [47]). CCEs from this third group are the most abundant in Lepidoptera and in particular the three most numerous clades (001, 006 and 016) with species-specific radiations. Many clades in this third group are also Lepidoptera specific [51]. The total number of CCE genes identified so far in S. litura and S. frugiperda genomes were 110 and 96, respectively [1,28]. For S. exigua, we were able to retrieve 73 sequences from public database (GenBank:GCA 011316535.1). For S. littoralis, we have previously identified 30 CCE transcripts expressed in antennae [52,53,54]. This repertoire was supplemented with 26 new sequences by Walker et al., (2019) [55]. Searching in our RNAseq databases allowed us to identify an additional 19 CCE transcripts (Table S1), leading to a total of 65 SlitCCE sequences, which is likely underestimated compared to its sister species S. litura.
Although comprehensive complete repertoires of CCEs are now available in many lepidopteran species, including spodopterans, still very few CCE genes have been directly linked with insecticide resistance. Here, we will focus on some spodopteran CCEs for which a direct role in resistance is supported either by in vitro and/or in vivo approaches, i.e., enzymes both identified at the molecular level and directly involved in insecticide hydrolysis or insecticide susceptibility.

2.2.1. Resistance through Over-Expression of CCE

As reviewed previously in Farnsworth et al., (2010), there is a strong correlation between insecticide resistance and higher esterase activities in the four Spodoptera species studied here [56]. Esterase activities toward artificial substrates in vitro (generally α-or β-naphthyl acetate) were compared between homogenates of susceptible and resistant strains, which allowed for the detection of both higher overall CCE activities or higher staining intensities of certain isozymes after analysis by native PAGE electrophoresis. For example, higher staining intensities of several esterases isozymes have been associated with OP and PYR resistance in S. littoralis [57], S. litura [58], S. exigua [59] and S. frugiperda [60]. The underlying biochemical mechanisms are not yet known, but it is hypothesized that increased sequestration of insecticides by CCEs and catabolism apply [56].
In lepidopterans, over-expression of CCE associated with insecticide resistance has been intensively studied in H. armigera [61,62,63], especially through proteomic and real-time polymerase chain reaction (RT-PCR) approaches. Most of the over-expressed CCEs identified at the molecular level belonged to the 001 clade, which is very diverse in this species, with 21 CCE001 genes. This clade also shows a large expansion in spodopterans, with 23 and 19 CCE001 genes annotated in S. litura [1] and S. frugiperda [28], respectively. We counted 14 CCE001 sequences in S. littoralis ([52,53,54,55] Table S1), a number which is probably again underestimated in the absence of genome annotation. In S. litura, some of these CCE001 genes were inducible by imidacloprid [1]. Most interestingly, knock-down of two SlituCEE001s (SlituCOE57 and SlituCOE58) by siRNA injection increased the sensitivity of S. litura larvae to imidacloprid when fed with an insecticide-containing diet [1]. This is the first demonstration of a direct link between CCE induction and insecticide sensitivity at the molecular level in a lepidopteran species. After BLAST (Basic Local Alignment Search Tool) searches, we found orthologous sequences of SlituCOE57 and SlituCOE58 in S. littoralis (SlitCXE50 and SlitCXE48, respectively, Table S1) and S. frugiperda (SfruCCE001n and SfruCCE001f, respectively, [55]), suggesting that they might play a similar role in insecticide resistance. In S. litura, all of these CCE001s were grouped into a large cluster on chromosome 2 [1], as previously observed for the CCE001 of H. armigera [51].

2.2.2. Metabolic Resistance through Point Mutations of CCEs

Structural mutations in the active site of a carboxylesterase could result in a reduction in the ability of this enzyme to hydrolyze common carboxylesterase substrates (such as naphthyl acetate esters) but convert it to an OP hydrolase. This corresponds to the mutant ali-esterase mechanism [64]. Metabolic resistance by point mutation in CCE sequences has been observed for quite some time in several non-lepidopteran species, mostly in Diptera [49,65,66]. Two point mutations, G137D and W251L (or G151D and W271L as reported in Cui et al., (2011) following the sequence of D. melanogaster AChE [67]), were indeed found in field-resistant populations of several species, including Lucilia cuprina and Musca domestica, and involved in altering the substrate specificity of CCE, leading to increased activity towards OPs [65,66,67]. In addition to dipterans, another amino acid change at position 251 (W251G) was also found in an OP-resistant strain of the parasitoid wasp Anisopteromalus calandrae [68].
To the best of our knowledge, no point mutation-based resistance in CCE has been resolved at the molecular level in resistant Lepidopteran strains. However, an in vitro study conducted on CCEs from four insect orders including Lepidoptera, tested whether the change of substrate specificity associated with these two mutations could be a more general feature in OP-resistant insects [67]. The catalytic properties of seven CCEs mostly belonging to the dietary/detoxification group were analyzed, including one CCE from S. litura (GenBankEU783914). Recombinant mutant proteins were tested in vitro towards two OPs (paraoxon and chlorfenvinphos) and β-naphthyl acetate. For the seven enzymes tested, the G151D and W271L mutations conferred OP activity in 62.5% and 87.5% cases, respectively [67]. The same in vitro approach was developed more recently on eight H. armigera CCEs but with more contrasting results [62], with increased insecticide hydrolysis being observed only for some enzymes. However, for one esterase (HarmCCE001c), PYRs (fenvalerate and cypermethrine), hydrolysis was enhanced 14-fold after the leucine mutation. The S. litura CCE tested by [67] matched the SlituCOE082 genomic sequence identified later [1]. We found sequences orthologous to SlituCOE082 in S. frugiperda and S. littoralis, but not in S. exigua. SlituCOE82 presented 95.7% of amino acid identity with SlitCXE64 (Table S1) and 91.4% with SfruCCE16j [55], with residues conserved at the putative mutation positions (Figure 3).
SlituCOE082 belongs to the clade 016 which is also expanded in Lepidoptera [51]. It includes 16 genes in S. litura, 14 in S. frugiperda, eight in S. exigua and 16 in S. littoralis (including nine additional sequences from RNAseq data). Phylogenetic analysis of these 54 sequences (Figure 4) illustrates their diversification within the four Spodoptera species, and in particular the 1:1 orthologous relationship is clear for seven sequences from each species. In S. litura, the CCEs from in clade 016 showed a major expansion on chromosome 25, with a cluster of 12 adjacent sequences [1]. Six genes in this cluster showed induction by neonicotinoid insecticide, i.e., imidacloprid in S. litura larvae. In particular, SlituCOE82 is moderately induced by imidacloprid in Malpighian tubules [1].
To date, the presence of the G151D and/or W271L mutations in field-resistant spodopteran strains has not been found. However, although metabolic resistance by overproduction of CCE could induce broader resistance than qualitative mutation, the two studies discussed below suggest that point mutations could be a more common mechanism for insecticide resistance than expected. A systematic comparison of the corresponding CCE gene sequences in susceptible and resistant lepidopteran pest strains would be necessary to assess their precise role in resistance.

2.2.3. Target Site Resistance through Point Mutations; the Case of Acetylcholinesterases

Acetylcholinesterase (AChE or ace) terminates nerve impulses by catalyzing the hydrolysis of acetylcholine in cholinergic synapses. Irreversible inhibition of AChE by CBs and OPs thus causes acetylcholine to accumulate in synapses and acetylcholine receptors to open permanently, resulting in insect death. Most insects have two AChE genes but only the gene expressed in the central nervous system (namely ace-1 or AChE1) is essential for synapse functioning [69]. All four Spodoptera species possess two AChE genes as expected (Figure 4) with a clear orthologous relationship and highly conserved sequences (more than 98% of amino acid identity). Target site resistance mediated by AChE1 insensitivity to insecticides has been identified by biochemical approaches in several insect species, and subsequently elucidated at the molecular level for some species [49,69]. Sequencing of AChE genes from field-resistant strains and comparison with susceptible strains described several point mutations conserved across species, most of which modify the active site of the enzyme. Three substitutions (A201S, G227A and F290V, numbering corresponding to the mature enzyme of Torpedo californica) were first reported in CB- and/or OP-resistant strains of aphids (A. gossypii [70]), dipterans (Bactrocera dorsalis [71]) and lepidopterans (Chilo suppressalis [72] and in P. xylostella [73]).
The involvement of acetylcholinesterases in OP/CB resistance has been demonstrated in biochemical approaches on S. litura resistant strains from Korea [74] or India [75,76]. Similarly, in S. exigua, the AChE enzyme of a carbamate resistant strain from California was found to be approximately 30-fold more insensitive to methomyl compared to the enzyme from the susceptible laboratory strain [77]. However, within Spodoptera species, molecular data on the AChE point mutation are mostly available for S. frugiperda. In this species, Yu et al., (2003) showed that acetylcholinesterase of a field strain collected from corn fields in Florida was clearly far less sensitive (up to 85-fold) than that of a susceptible strain to inhibition by CBs and OPs [60]. Kinetic data also showed that the apparent Km value of acetylcholinesterase from the field strain was 56% of that from the susceptible strain. By comparing the predicted amino acid sequences of a S. frugiperda ace-1 fragment from a chlorpyrifos susceptible strain with the corresponding sequence isolated from an 18.1-fold resistant strain from Brazil, Carvalho et al., (2013) were then able to identify the three previously described point mutations (Figure 5) [78].
Genotyping revealed that of the three, the A201S allele was present at relatively low frequency (17.5%) while G227A and F290V were present at higher frequency (67.5% and 32.5%, respectively) [78]. More recently, Boaventura et al., (2020) searched for the presence of these mutations in 34 different resistant populations of S. frugiperda collected from four different continents [79] and showed that F290V was the most frequent substitution in all populations tested. Similar results were obtained by Guan et al., (2020) on S. frugiperda individuals collected from China and Africa, except that only two positions (A201S, F290V) were found, with the F290V allele at higher frequencies [80].

3. Phase II: Conjugation

Following oxido-reduction and hydrolysis reactions performed by phase I enzymes, xenobiotic metabolites (including insecticides) are then conjugated to small hydrophilic molecules by phase II enzymes, a group of transferases that metabolize hydrophobic compounds containing nucleophilic or electrophilic groups [81].

3.1. Glutathione-S-Transferases (GSTs)

Glutathione-S-transferases (GSTs, EC are one of the most important classes of this group whose biotransformation reaction leads to the generation of hydrophilic metabolites, which are readily excreted by efflux transporters such as ABC transporters. In arthropods, GSTs are highly diverse and are involved in a variety of cellular functions, from the detoxification of a wide range of both endogenous and xenobiotic compounds, to intracellular transport, hormone biosynthesis and reduction of oxidative stress [82]. GSTs primarily catalyze the conjugation of electrophilic lipophilic compounds with the thiol group of reduced glutathione (GSH) but are also capable of catalyzing a dehydrochlorination reaction using reduced glutathione as a cofactor. The enzymatic structure of cytosolic GSTs classically consists of hetero- or homo-dimeric proteins, with each monomer consisting of a highly conserved amino-terminal domain providing the GSH-binding site (G-site) while the carboxyl terminal domain interacts with the hydrophobic substrate (H-site) [82].
In insects, cytosolic GSTs belong to a diverse gene family divided into six classes (Delta, Epsilon, Omega, Sigma, Theta, and Zeta) based on their substrate specificities and phylogenetic relationships [83]. The growing number of available genomes reveals a large disparity in the total number of GSTs among insect species, with diversity ranging from low in hymenopterans (eight GSTs in Apis mellifera) to high in dipterans (39 in Culex quinquefasciatus) [84]. This variability is mainly due to genetic expansions observed in the Insecta-specific Delta and Epsilon classes due to multiple duplication events [85].
Insect GSTs have been intensively studied for their role in insecticide resistance: reports correlating high levels of GST activity with high resistance to various pesticides, including organophosphates, organochlorines, cyclodienes, and pyrethroids, exist for many species and among them S. littoralis and S. frugiperda [86]. This phenomenon relies on the precise regulation or induction of GST expression by xenobiotic compounds, and in particular the Epsilon class (GSTe) [87,88]. Here, we will focus on spodopteran GSTes that have been shown to contribute to insecticide response and resistance.

3.1.1. Phylogeny of GST Epsilon

GST-mediated resistance could be triggered by different mechanisms, including gene amplification by multiple duplication events could lead to enhanced detoxification of insecticides underlying of the resistance process. Based on the recent availability of chromosome-level assembly genomes from spodopterans, we re-annotated GST epsilon class. For convenience, we reconciled the previous nomenclature with our new annotation to generate a unified dataset (see Supplementary Materials Table S2) [1,28]. Overall, GSTe accounts for nearly half of the total number of GSTs identified in the Spodopteran genomes (20 over 40 on average). Our phylogenetic analysis (Figure 6) revealed clear orthologous relationships among the four species studied with 1:1 orthologs in almost all cases, and with epsilon GSTs of H. armigera, suggesting that this great diversification is occurring in the Noctuoidea clade, and especially in highly polyphagous species.
Such diversity could be explained by tandem and segmental gene duplications as demonstrated by the phylogenetic branching that corresponds to the genomic clustering of GSTe (Figure 6). Furthermore, synteny studies support this diversification, with half of the GSTe derived from only two duplication events, arguing that these clades share a common GST ancestor (in an ancient Lepidoptera ancestor) even though their genetic expansions occurred independently (Figure 7, [1]).
This extensive genetic amplification does not directly support a demonstration of insecticide resistance per se; however, this highly diverse class is expected to have ancient roles unique to polyphagous lepidopterans, such as the removal of harmful chemical compounds from natural sources. According to the pre-adaptation hypothesis, phytophagous insect species with more diverse diets are likely to acquire resistance to more diverse insecticides [89], and thus genetic diversity may underlie the origin and evolution of insecticide resistance.

3.1.2. GST Activity against Insecticides

The biochemical mechanisms of GST-based insecticide resistance are classically associated with the conjugation of GSH to the pre-inactivated phase I compound. Such conjugation resistance has been demonstrated for different classes of organophosphate or pyrethroid insecticides [48,82]. Moreover, dehydrochlorination reactions using GSH as a cofactor have been implicated in resistance to the organochlorine DDT [6] making GSTs potential phase I enzymes for hydrochloric compounds. In addition to these direct modes of action, GST has also been shown to reduce lipid hydroperoxides produced as a result of insecticide-induced damages [90]. This peroxidase activity of GSTs protects tissues from an excess of reactive oxygen species (ROS), which can be more harmful than the pesticide itself [91]. Finally, GSTs are also capable of simple binding and sequestration activity towards various compounds, such as pyrethroids [86].
In spodopterans, most in vitro insecticide studies rely on the use of competitive 1-chloro-2,4-dinitrobenzene (CDNB) assays, a method that compares the amount of CDNB, the universal substrate of GST, in the presence or absence of a competitor, i.e., an insecticide. While this technique clearly indicates an interaction between a GST and a pesticide, it does not decipher the mechanism by which this interaction occurs. S. litura SlGSTe2 and SlGSTe3 were the first spodotperan GSTes to be characterized in this manner, showing differential activity toward DDT and deltamethrin between the two enzymes, with SlGSTe2 having the greater activity toward the insecticide [92]. The SeGSTe2 ortholog was further characterized showing activity toward metaflumizone, indoxacarb, monosultap, chlorpyrifos, malathion, cyhalothrin, and imidacloprid, suggesting a conserved insecticide metabolizing function [93]. Similarly, GSTe1 was carefully analyzed in S. litura and S. exigua species [94,95]. CDNB competition assays showed activity towards chlopyrifos, malathion, phoxim, deltamethrin and cypermethrin and the specific activities of GSTE1 were further analyzed using high-performance liquid chromatography (HPLC), confirming their active binding to chlorpyrifos and cypermethrin. Interestingly, GSTe1 is also active towards secondary plant metabolites [96] and has strong peroxidase activity. Therefore, this enzyme has multi-detoxification properties with a wide range of substrates, potentially conferring insecticide resistance.
In vivo studies have also confirmed the direct involvement of GSTs in insecticide tolerance. In Cheng et al., (2019), injection of siRNAi against SlGSTe20 and SlGSTe3 increased the sensitivity of S. litura larvae to imidacloprid, and the recombinant proteins subsequently showed activity towards diazinon, permethrin, chlorfenapyr, and bendiocarb [97].

3.1.3. Resistance through Over-Expression of GST Epsilon

GST overexpression has often been a prerequisite for the identification of enzymes potentially involved in insecticide resistance [48]. Thus, many studies have identified candidate GSTes whose expression could be altered upon exposure to insecticides. In S. litura, SlGSTe1, e2, e3, e4, e10, e11, e12, e13, e15 and e20 showed overexpression following exposure to various pesticides such as tebufenozide, carbaryl, DDT, malathion, deltamethrin, chlorpyrifos and imidacloprid [1,92,98,99]. It is noteworthy that among these 10 over-expressed genes, no specific phylogeny-related pattern could be assigned (Figure 6) indicating that, rather than defining an insecticide-specific clade, the induced GSTe genes are diverse with potentially redundant and/or complementary activities. Similar patterns are observed in S. littoralis and S. exigua when exposed to deltamethrin or lambda-cyhalothrin, chlorpyrifos and chlorantraniliprole, respectively [94,100]. Nevertheless, induction after exposure may be only part of the animal’s response to a given toxicant and does not necessarily imply involvement in resistance [90]. Therefore, in an effort to find genes directly associated with insecticide catabolism, studies focusing on the expression profiles of genes highly expressed in resistant strains compared to the susceptible population have indicated potential candidates in GSTe as directly responsible for insecticide resistance. In S. frugiperda, SfGSTe5 was associated with an OP-resistant strain using a transcriptomic approach [78]. SfGSTe5 is the ortholog of Plutella xylostella PxylGSTe3, an epsilon GST induced and capable of metabolizing the organophosphate insecticides parathion and methylparathion [101,102]. In an RNAseq experiment comparing indoxacarb-resistant and susceptible S. litura populations [103], SlGSTe4 and SlGSTe20 were the only GSTs overexpressed. Interestingly, both of these GSTes are active against imidacloprid, diazinon, permethrin, chlorfenapyr, or bendiocarb [97,101], indicating potential cross-activity of SlGSTe for various insecticides. This complex pattern of gene expression regulation must be orchestrated by specific regulatory operators that can effectively govern down-or overexpression upon exposure but could also be affected by mutations in resistant strains to account for specific activations. Promoter sequence analysis revealed that some S. litura GSTs harbor the same cap ‘n’ collar ‘C/muscle aponeurosis fibromatosis (CncC/Maf) binding site and the aryl hydrocarbon receptor/nuclear aryl hydrocarbon receptor translocator (AhR/ARNT) binding site [84,94]. These transcription factors coordinately regulate GST expression through intracellular production of ROS induced by insecticide exposure. In a CncC RNAi experiment, Shi et al., (2020) demonstrate that this transcription factor not only enhances indoxacarb sensitivity in susceptible and resistant strains of S. litura, but is also involved in the down-regulation of detoxification genes related to indoxacarb resistance, confirming its central role in insecticide-resistance mechanisms [104].

3.1.4. Resistance through Point Mutations in GST Epsilon

Insecticide resistance by GSTe has been clearly established in Anopheles where a key residue, Phe120, has been implicated in the increased binding of DDT to GSTe2 [6]. Such a substitution is also observed in 5 GSTes from spodopteran species (GSTe2, e12, e15, e17, e18, Figure 8) indicating potential conservation of resistance processes between these species. Thus, further investigations are needed to fully understand the detailed mechanisms of pesticide-GSTes interaction and their precise contribution to insecticide resistance.

4. Phase III: Elimination/Export

The last phase concerns the elimination or excretion of metabolites outside the cell. Metabolites have been rendered less toxic by the reactions they underwent in phase I and/or phase II but they can also be directly excreted without prior chemical modification. This phase mainly involves ATP-binding cassette transporters.

4.1. ATP-Binding Cassette Transporters (ABCs)

ATP-binding cassette (ABC) transporters are proteins that use the energy of ATP hydrolysis to transport substrates such as amino acids, lipids, peptides, sugars and drugs across cell membranes. ABC transporters are classified into eight subfamilies from letter A to H based on similarities in their ATP binding domain. Full transporters (FT) consist of two cytosolic nucleotide-binding domains (NBDs) and two transmembrane domains (TMDs) whereas half transporters (HT) have only one of each domain and must dimerize to form a functional transporter. The NBD is involved in ATP binding and hydrolysis. The TMD consists of 5-6 transmembrane segments and is responsible for substrate specificity. In insects, their number varies from 32 in Nilaparvata lugens [105] to 82 in Plutella xylostella [106] while in the available genomes of Noctuidae about 50 ABCs have been found so far. Some subfamilies are conserved well from humans to arthropods. For example, ABCD, ABCE, ABCF have clear orthology and similar suspected physiological roles such as the transport of acyl-CoA molecules into the peroxisome, ribosome biogenesis and translation. Moreover, some ABC transporters subfamilies are involved in chemotherapy resistance in humans and pesticides resistance in insects. Their names reflect these roles, the B subfamily is known as multidrug-resistance protein (MDR) or P-glycoprotein (P-gp) while the C subfamily is known as multidrug-resistance associated protein (MRP). For example, the human MDR1 (ABCB1) excludes chemotherapeutic agents and confers resistance in several types of cancer when over-expressed [107]. MDR1 orthologs in Drosophila Mdr50 and Mdr65 are over-expressed in DDT-resistant strain 91-R. Knockdown of each gene in this strain by RNAi increases the susceptibility of flies to DDT [108]. These examples, in humans and Drosophila, illustrate one of the mechanisms of resistance involving ABC transporters: over-expression of the transporter which allows exclusion of a greater amount of xenobiotics. An additional mechanism in the case of insecticide resistance can be mentioned and corresponds to the mutation of the target. Indeed, ABCs are one of the receptor proteins of Bacillus thuringiensis (Bt) toxins as are cadherin-like, aminopeptidase-N or alkaline phosphatase, and a modification in the sequence of these receptors can confer resistance (for a recent review see [109]).

4.1.1. Resistance through Point Mutations in ABCs, the Case of ABCC2

ABCC2 was first identified as a receptor for one of the Bt crystal toxins (Cry), Cry1Ac in the lepidopteran Heliothis virescens [110]. In this study, a strain of H. virescens was selected in the laboratory for several years with Cry1Ab and Cry1Ac toxins. Very high levels of resistance were obtained as well as a loss of membrane binding for the toxins. A deletion in exon 2 of ABCC2 resulting in a 99 amino acid protein truncation was identified as the cause of resistance to Cry1Ac [110]. This study paved the way to investigating the role of ABC transporters in resistance to Bt-toxin. Special attention is given to the four species of interest in this review. Annotations of ABC transporters in the genome of S. litura, S. frugiperda, S. exigua and in the transcriptomic data of S. littoralis were mainly performed automatically and only a few subfamilies were annotated manually by experts (G. Le Goff, personal communication). In a recent publication, Liu et al., (2018) showed that the sensitivity towards Cry1Ac toxin differed by a factor of 65 between the more tolerant S. litura, and S. frugiperda [111]. SlABCC2 and SfABCC2 share 97% identity. Fragment substitutions and point mutations in the transporter expressed in insect cells allowed the authors to identify the amino acid at position 125 as the key to this difference in sensitivity. A glutamine is present at this position in S. frugiperda while it is a glutamic acid in S. litura. By aligning the sequences of ABCC2 between the four Spodoptera species (S. exigua, S. frugiperda, S. littoralis and S. litura), it is possible to predict their susceptibility to Cry1Ac toxin. S. exigua should be susceptible like S. frugiperda while S. littoralis should be more tolerant like S. litura (Figure 9).
Three-dimensional structure predictions suggest that the G or E at position 125 is localized in the extracellular loop 1 (ECL1). Liu et al., hypothesize that it may have a role in toxin binding, which would explain these differences in sensitivity [111]. The importance of ECL1 in Bt toxin selectivity was previously demonstrated in B. mori, where replacement of certain amino acids in the ECL1 loop between BmABCC2 and BmABCC3 resulted in increased binding affinity of Cry1A toxins [112]. These variations in toxin receptor sequence may also explain the spectrum of specificity of Bt toxins. Indeed, some toxins primarily target lepidopterans, for example Cry1 toxins, while others are specific to Coleoptera, such as Cry3 [113].
Larger changes in the sequence of ABCC2 (Figure 10A) have been associated with resistance in some of these Spodoptera spp., especially in field-resistant S. frugiperda populations. Indeed, resistance to transgenic maize expressing the Cry1Fa toxin has been reported since 2007 in Puerto Rico for S. frugiperda. Banerjee and colleagues have demonstrated that the resistance was due to mutations in the SfABCC2 toxin receptor [114]. These mutations result in a truncated protein that loses the second transmembrane domain and at the same time toxin binding. A nine-base deletion (position 39–47) and a two-base insertion (GC at position 2218) lead to a frameshift and the occurrence of a premature stop codon. The truncated protein is 746 amino acids (Figure 10B) whereas the ABCC2 of the susceptible strain encodes a protein of 1349 amino acids (Figure 10A). The frequency of mutated SfABCC2 in Puerto Rico increased between 2007 and 2009 from 1% to 42% and stabilized at 55% in 2017. This truncated receptor was not found by the authors in the populations from Florida or the Dominican Republic [114].
Another group confirmed these results. Flagel et al., isolated a population of S. frugiperda from a site in Puerto Rico in 2010 and this population was selected in the laboratory with Cry1Fa for 50 generations [115]. The resulting strain had a resistance factor of 500 for Cry1Fa and 87-fold cross-resistance to Cry1A.105 compared to the susceptible reference strain. They found the same GC insertion in the ABCC2 sequence that creates a stop codon at position 747 and results in a truncated protein. By conducting a sampling campaign in various localities of Puerto Rico and Brazil, the mutation was to be found in Puerto Rico only. In addition to this allele, the authors mention the existence of a second resistance allele and the insertion of a sequence in the fourth exon of the gene. However, sequencing difficulties due to repeated sequences did not allow them to identify precisely the effect of this insertion on the ABCC2 sequence (Figure 10C) although an aberrant splicing of this allele 2 was suspected.
Furthermore, a recent study reported resistance to Cry1F toxin in isolated Brazilian populations of S. frugiperda [116]. The authors identified two mutations in the extracellular loop 4 (ECL4) of ABCC2: a deletion of two amino acids (GY) at positions 788 and 789 and the change of a proline to either lysine or arginine at position 799 (Figure 10D). Unlike the resistance reported in Puerto Rico, these mutations do not cause premature termination of the sequence. Expression of mutated ABCC2 in insect cells confirmed the role of these mutations in toxin binding. Analysis of populations collected in different regions of Brazil showed a high frequency of the GY deletion as well as many rare alleles that result in sequence changes between amino acids 783–799. According to the authors, these results reinforce the primary role of the ECL4 extracellular loop in Cry1F toxicity. Further analysis is needed to demonstrate the role that these mutations may have in resistance to other toxins. Indeed, in addition to resistance to Cry1F other studies on Puerto Rican populations with truncated SfABCC2 have shown cross-resistance for Cry1A.105, Cry1Ac, and Cry1Ab toxins [114,115]. Do mutations in the ECL4 loop also confer these cross-resistances? Another mutation in ABCC2 has been identified in Brazilian populations corresponding to the insertion of 12 bases at the intracellular loop between transmembrane domains 8 and 9 resulting in a premature stop codon [80]. The truncated protein lost the last four transmembrane segments and the second intracellular ATP-binding domain (Figure 10E). The study was performed on insects collected three years after those of Boaventura et al., (2020) and changes in the use of transgenic crops may have occurred. The authors did not investigate the effect of this mutation on resistance and of Bt toxin binding. Instead, they tested whether invasive populations of S. frugiperda (also sampled in Africa and Asia) carried mutations known to cause insecticide resistance.
None of the mutations identified so far in ABCC2 were found in these invasive populations. This truncated protein retains an intact extracellular loop 4 and thus potentially to bind to the Cry1F toxin. Further experiments are needed to determine if it can confer resistance to some of the Bt toxins.
Looking at other Spodoptera species, resistance to some Bt toxins was reported as early as 1994–1995 for S. littoralis and S. exigua respectively [118,119]. However, in both cases, these were laboratory selected strains. No resistance has been reported to date for S. litura in the Arthropod Pesticide Resistance Database although one study mentioned the development of resistance to Cry1C and Vip3A toxins in a laboratory selected strain [120]. But these studies did not involve ABCC2. In S. exigua, a strain was selected with a commercial B. thuringiensis product, Xentari. The main toxins contained in this product are Cry1a and Cry1Ca. The selected strain (Xen-R) showed a resistance factor of more than 1000 [121]. Bulk segregant analysis based on high-throughput sequencing identified the region of the genome carrying the major resistance loci, which contained three genes encoding ABCC1, ABCC2 and ABCC3 [117]. ABCC2 and ABCC3 played a major role in resistance. ABCC2 contained a mutation in the resistant strain, which resulted in the loss of the ATP binding domain II caused by the truncation of 82 terminal amino acids (Figure 10F). However, the binding of Cry1Ca to brush border membrane vesicles was not significantly different between resistant and susceptible strains. RNAi experiments confirmed the role of ABCC2 and ABCC3 in the resistance to Cry1A and Cry1Ca in S. exigua [117]. In addition, Pinos and co-workers showed that the loss of the second nucleotide binding domain did not affect the binding of Cry1A toxins [122]. Indeed, expression of the truncated transporter in Sf21 insect cells conferred sensitivity to Cry1A, as specific binding of the toxin was still effective. These results suggest that this domain is not required for a functional toxin receptor [122]. Nevertheless, the use of techniques such as CRISPR/Cas9 validated the main role of ABCC2 as a receptor for Cry1Ac and Cry1Fa toxin in S. exigua [123]. The knock-out (KO) strain for SeABCC2 was more resistant than the parental strain by a factor of 470 and 240 for Cry1Ac and Cry1Fa, respectively. Using the same technique in S. frugiperda to produce a KO for ABCC2 conferred resistance to Cry1F toxin [124].

4.1.2. Resistance through Over- or Reduced Expression of ABCs

Data on the involvement of ABC transporters in resistance to chemical insecticides in Spodoptera spp. are scarce, as it is in Lepidoptera in a broader sense. A few examples exist, such as the involvement of H. armigera Pgp1 insecticide transport. HaPgp1 expression was induced in the larval gut after exposure to abamectin [125]. When the expression of this transporter is suppressed by RNAi, larvae became more sensitive to the insecticide and mortality increased from 26% to 84% for a fixed abamectin dose of 0.4 µg/g diet [125]. Another study confirmed the role of HaPgp in abamectin transport, as the use of a specific Pgp inhibitor, verapamil, increased the sensitivity of larvae towards abamectin [126].
The development of the CRISPR/Cas9 technique has paved the way to study the role of ABCs in the transport of chemical insecticides and potentially in the development of resistance. Here, we report the few examples in which an increase in LD50 was observed for insecticides after KO of certain ABC transporters in Spodoptera spp. In S. exigua, the ability of ABCB1 to transport 12 insecticidal molecules from 10 different chemical families was investigated by knocking out this protein using CRISPR/Cas 9 [127]. The ABCB1 KO strain showed a 2.73-fold and 3.01-fold increase in sensitivity to abamectin and emamectin benzoate, respectively, while no significant difference was observed for the other insecticides tested. In S. frugiperda, the KO of ABCC2 rendered the insects 7.8 and 3.1 times more sensitive to abamectin and spinosad, respectively. Similarly, a reduction in tolerance to these two insecticides was observed for the ABCC3 KO by factors of 4.5 and 2 [128]. Another study reported that SfABCC2 KO does not induce resistance to insecticide molecules such as bifenthrin, chlorantraniliprole, spinetoram and acephate [124]. To our knowledge, there are no CRISPR/Cas9-mediated ABC KO data for S. litura and S. littoralis to date.

4.1.3. Resistance and Regulation of ABCs Expression

While the mechanisms leading to the acquisition of chemotherapy resistance through changes in ABC transporters expression of are well known in humans, those leading to pesticide resistance are still largely unexplored in insects. For example, over-expression of MDR1 (ABCB1) in human tumors has been associated with increased gene copy number through chromosomal region amplification, epigenetic modifications and single nucleotide polymorphisms (SNPs) [129]. In insects, the acquisition of resistance may involve either under- or over-expression of a given ABC transporter. Indeed, in the case of Bt toxins, decreased expression of the ABC receptor can confer resistance, while conversely, an over-expression of ABCs that can transport insecticides would allow the development of resistance. MicroRNAs (miRNA), which are small non-coding RNA sequences (between 19 and 24 nucleotides in length) are known to regulate gene expression. The miRNA miR-998-3p has target sites in the ABCC2 coding sequence (CDS) in several Lepidoptera species [130]. The involvement of these sites in the regulation of ABCC2 has been demonstrated in P. xylostella. MiR-998-3p was over-expressed by a factor of two in a Cry1Ac-resistant strain (GX-R) compared to a susceptible strain and ABCC2 expression in the larval midgut was reduced by approximately 50%, demonstrating the involvement of this miRNA in Bt resistance. A conserved target site for miR998-3p had been found in the four Spodoptera spp. as well as one and two non-conserved sites for S. exigua and the other three species respectively ([130] and the present study). MiR-998-3p could play a role in the acquisition of resistance in these species but to the best of our knowledge there has been no study has so far reported such cases and more generally the role of miRNAs in insecticide resistance remains largely unexplored.
Another regulator of ABCC2 expression is the transcription factor Forkhead box protein A (FoxA). Its transfection into Sf9 cells (S. frugiperda cell model) induces the expression of ABCC2 and ABCC3 and results in increased susceptibility of the cells to Cry1Ac toxin [131]. Potential resistance could arise through under-expression of this transcription factor but has not been described at this time.
Cap ‘n’ collar isoform C (CncC) is a major transcription factor for controlling the expression of detoxification genes in insects [12]. CncC is constitutively overexpressed in a Chinese field population of S. exigua resistant to chlorpyrifos and cypermethrin [94]. Several studies on this resistant strain showed that CncC is involved in controlling the expression of detoxification genes including P450s (CYP321A8, CYP321A16, and CYP332A1) [11,132] and GSTs (GSTo2, GSTe6, GSTd3) [94]. However, none of these studies mention the role of CncC in the expression of ABC transporters. Nevertheless, this role is not excluded and an RNAseq analysis would allow a global approach to identify CncC-regulated genes in this resistant population. In Drosophila, a DDT-resistant strain (91R) constitutively overexpressing CncC exhibits multifactorial resistance involving some P450 and GST in addition to ABC transporters [108]. None of these studies investigated the underlying cause of CncC overexpression. In other Spodoptera species, CncC regulates the expression of several detoxification genes involved in indoxacarb resistance, including the ABC transporter SlituABCH-1 in S. litura [104]. Although the CncC gene from S. frugiperda has been cloned [133], it has not been shown to control of the expression of resistance-related ABC transporters and, to date, this information is also lacking in S. littoralis.
A number of questions remain unanswered. What is the mechanism behind the over-expression of CncC in some resistant populations? What other regulatory mechanisms could control the expression of ABC transporters involved in resistance? Do epigenetic regulations play a major role in the expression of certain ABC transporters and the acquisition of resistance, as has been observed in humans?

Supplementary Materials

The following are available online at, Table S1: Protein sequences of S. littoralis carboxyl/cholinesterases, Table S2: Spodopteran GSTes reconciled nomenclature, with new annotations combined with previous studies.

Author Contributions

All authors contribute equally to the writing. All authors have read and agreed to the published version of the manuscript.


This research received no external funding.


The authors would like to thank Abby Cuttriss (Université Côte d’Azur, Office of International Visibility). Le Goff would like to thank Dries Amezian for his very helpful comments.

Conflicts of Interest

The authors declare no conflict of interest.


  1. Cheng, T.; Wu, J.; Wu, Y.; Chilukuri, R.V.; Huang, L.; Yamamoto, K.; Feng, L.; Li, W.; Chen, Z.; Guo, H.; et al. Genomic adaptation to polyphagy and insecticides in a major East Asian noctuid pest. Nat. Ecol. Evol. 2017. [Google Scholar] [CrossRef]
  2. Stokstad, E. New crop pest takes Africa at lightning speed. Science 2017, 356, 473–474. [Google Scholar] [CrossRef] [PubMed]
  3. Johnson, S.J. Migration and life history strategy of the fall armyworm Spodoptera frugiperda in the Western hemisphere. Insect Sci. Appl. 1987, 8, 543–549. [Google Scholar] [CrossRef]
  4. Westbrook, J.K. Noctuid migration in Texas within the nocturnal aeroecological boundary layer. Integr. Comp. Biol. 2008, 48, 99–106. [Google Scholar] [CrossRef] [PubMed][Green Version]
  5. Sparks, T.C.; Crossthwaite, A.J.; Nauen, R.; Banba, S.; Cordova, D.; Earley, F.; Ebbinghaus-Kintscher, U.; Fujioka, S.; Hirao, A.; Karmon, D.; et al. Insecticides, biologics and nematicides: Updates to IRAC’s mode of action classification-a tool for resistance management. Pestic. Biochem. Physiol. 2020, 167, 104587. [Google Scholar] [CrossRef]
  6. Riveron, J.M.; Yunta, C.; Ibrahim, S.S.; Djouaka, R.; Irving, H.; Menze, B.D.; Ismail, H.M.; Hemingway, J.; Ranson, H.; Albert, A.; et al. A single mutation in the GSTe2 gene allows tracking of metabolically based insecticide resistance in a major malaria vector. Genome Biol. 2014, 15, R27. [Google Scholar] [CrossRef][Green Version]
  7. Feyereisen, R.; Dermauw, W.; van Leeuwen, T. Genotype to phenotype, the molecular and physiological dimensions of resistance in arthropods. Pestic. Biochem. Physiol. 2015, 121, 61–77. [Google Scholar] [CrossRef]
  8. Schmidt, J.M.; Good, R.T.; Appleton, B.; Sherrard, J.; Raymant, G.C.; Bogwitz, M.R.; Martin, J.; Daborn, P.J.; Goddard, M.E.; Batterham, P.; et al. Copy number variation and transposable elements feature in recent, ongoing adaptation at the Cyp6g1 locus. PLoS Genet. 2010, 6, e1000998. [Google Scholar] [CrossRef] [PubMed][Green Version]
  9. Devonshire, A.L.; Field, L.M. Gene amplification and insecticide resistance. Annu. Rev. Entomol. 1991, 36, 1–23. [Google Scholar] [CrossRef] [PubMed]
  10. Field, L.M.; Blackman, R.L.; Tyler-Smith, C.; Devonshire, A.L. Relationship between amount of esterase and gene copy number in insecticide-resistant Myzus persicae (Sulzer). Biochem. J. 1999, 339 Pt 3, 737–742. [Google Scholar] [CrossRef]
  11. Hu, B.; Huang, H.; Hu, S.; Ren, M.; Wei, Q.; Tian, X.; Elzaki, M.E.A.; Bass, C.; Su, J.; Palli, S.R. Changes in both trans- and cis-regulatory elements mediate insecticide resistance in a lepidopteron pest, Spodoptera exigua. PLoS Genet. 2021, 17, e1009403. [Google Scholar] [CrossRef] [PubMed]
  12. Amezian, D.; Nauen, R.; le Goff, G. Transcriptional regulation of xenobiotic detoxification genes in insects-An overview. Pestic. Biochem. Physiol. 2021, 174, 104822. [Google Scholar] [CrossRef] [PubMed]
  13. Werck-Reichhart, D.; Feyereisen, R. Cytochromes P450: A success story. Genome Biol. 2000, 1, REVIEWS3003. [Google Scholar] [CrossRef] [PubMed][Green Version]
  14. Mansuy, D. The great diversity of reactions catalyzed by cytochromes P450. Comp. Biochem. Physiol. C Pharmacol. Toxicol. Endocrinol. 1998, 121, 5–14. [Google Scholar] [CrossRef]
  15. Rewitz, K.F.; Rybczynski, R.; Warren, J.T.; Gilbert, L.I. The Halloween genes code for cytochrome P450 enzymes mediating synthesis of the insect moulting hormone. Biochem. Soc. Trans. 2006, 34 Pt 6, 1256–1260. [Google Scholar] [CrossRef]
  16. Qiu, Y.; Tittiger, C.; Wicker-Thomas, C.; le Goff, G.; Young, S.; Wajnberg, E.; Fricaux, T.; Taquet, N.; Blomquist, G.J.; Feyereisen, R. An insect-specific P450 oxidative decarbonylase for cuticular hydrocarbon biosynthesis. Proc. Natl. Acad. Sci. USA 2012, 109, 14858–14863. [Google Scholar] [CrossRef][Green Version]
  17. Kirkness, E.F.; Haas, B.J.; Sun, W.; Braig, H.R.; Perotti, M.A.; Clark, J.M.; Lee, S.H.; Robertson, H.M.; Kennedy, R.C.; Elhaik, E.; et al. Genome sequences of the human body louse and its primary endosymbiont provide insights into the permanent parasitic lifestyle. Proc. Natl. Acad. Sci. USA 2010, 107, 12168–12173. [Google Scholar] [CrossRef] [PubMed][Green Version]
  18. Yang, T.; Liu, N. Genome analysis of cytochrome P450s and their expression profiles in insecticide resistant mosquitoes, Culex quinquefasciatus. PLoS ONE 2011, 6, e29418. [Google Scholar] [CrossRef][Green Version]
  19. Nebert, D.W.; Adesnik, M.; Coon, M.J.; Estabrook, R.W.; Gonzalez, F.J.; Guengerich, F.P.; Gunsalus, I.C.; Johnson, E.F.; Kemper, B.; Levin, W.; et al. The P450 gene superfamily: Recommended nomenclature. DNA 1987, 6, 1–11. [Google Scholar] [CrossRef] [PubMed]
  20. Nebert, D.W.; Nelson, D.R.; Coon, M.J.; Estabrook, R.W.; Feyereisen, R.; Fujii-Kuriyama, Y.; Gonzalez, F.J.; Guengerich, F.P.; Gunsalus, I.C.; Johnson, E.F.; et al. The P450 superfamily: Update on new sequences, gene mapping, and recommended nomenclature. DNA Cell Biol. 1991, 10, 1–14. [Google Scholar] [CrossRef]
  21. Nelson, D.R. Metazoan cytochrome P450 evolution. Comp. Biochem Physiol C Pharmacol Toxicol Endocrinol 1998, 121, 15–22. [Google Scholar] [CrossRef]
  22. Dermauw, W.; van Leeuwen, T.; Feyereisen, R. Diversity and evolution of the P450 family in arthropods. Insect Biochem. Mol. Biol. 2020, 127, 103490. [Google Scholar] [CrossRef]
  23. Le Goff, G.; Boundy, S.; Daborn, P.J.; Yen, J.L.; Sofer, L.; Lind, R.; Sabourault, C.; Madi-Ravazzi, L.; ffrench-Constant, R.H. Microarray analysis of cytochrome P450 mediated insecticide resistance in Drosophila. Insect Biochem. Mol. Biol. 2003, 33, 701–708. [Google Scholar] [CrossRef]
  24. Daborn, P.J.; Lumb, C.; Boey, A.; Wong, W.; Ffrench-Constant, R.H.; Batterham, P. Evaluating the insecticide resistance potential of eight Drosophila melanogaster cytochrome P450 genes by transgenic over-expression. Insect Biochem. Mol. Biol. 2007, 37, 512–519. [Google Scholar] [CrossRef] [PubMed]
  25. D’Alencon, E.; Sezutsu, H.; Legeai, F.; Permal, E.; Bernard-Samain, S.; Gimenez, S.; Gagneur, C.; Cousserans, F.; Shimomura, M.; Brun-Barale, A.; et al. Extensive synteny conservation of holocentric chromosomes in Lepidoptera despite high rates of local genome rearrangements. Proc. Natl. Acad. Sci. USA 2010, 107, 7680–7685. [Google Scholar] [CrossRef][Green Version]
  26. Feyereisen, R. Arthropod CYPomes illustrate the tempo and mode in P450 evolution. Biochim. Biophys. Acta 2011, 1814, 19–28. [Google Scholar] [CrossRef]
  27. Gimenez, S.; Abdelgaffar, H.; le Goff, G.; Hilliou, F.; Blanco, C.A.; Hanniger, S.; Bretaudeau, A.; Legeai, F.; Negre, N.; Jurat-Fuentes, J.L.; et al. Adaptation by copy number variation increases insecticide resistance in the fall armyworm. Commun. Biol. 2020, 3, 664. [Google Scholar] [CrossRef] [PubMed]
  28. Gouin, A.; Bretaudeau, A.; Nam, K.; Gimenez, S.; Aury, J.M.; Duvic, B.; Hilliou, F.; Durand, N.; Montagne, N.; Darboux, I.; et al. Two genomes of highly polyphagous lepidopteran pests (Spodoptera frugiperda, Noctuidae) with different host-plant ranges. Sci. Rep. 2017, 7, 11816. [Google Scholar] [CrossRef] [PubMed]
  29. Kergoat, G.J.; Goldstein, P.Z.; le Ru, B.; Meagher, R.L., Jr.; Zilli, A.; Mitchell, A.; Clamens, A.L.; Gimenez, S.; Barbut, J.; Negre, N.; et al. A novel reference dated phylogeny for the genus Spodoptera Guenee (Lepidoptera: Noctuidae: Noctuinae): New insights into the evolution of a pest-rich genus. Mol. Phylogenet. Evol. 2021, 107161. [Google Scholar] [CrossRef]
  30. Klai, K.; Chenais, B.; Zidi, M.; Djebbi, S.; Caruso, A.; Denis, F.; Confais, J.; Badawi, M.; Casse, N.; Khemakhem, M.M. Screening of Helicoverpa armigera mobilome revealed transposable element insertions in insecticide resistance genes. Insects 2020, 11, 879. [Google Scholar] [CrossRef] [PubMed]
  31. Yu, L.; Tang, W.; He, W.; Ma, X.; Vasseur, L.; Baxter, S.W.; Yang, G.; Huang, S.; Song, F.; You, M. Characterization and expression of the cytochrome P450 gene family in diamondback moth, Plutella xylostella (L.). Sci. Rep. 2015, 5, 8952. [Google Scholar] [CrossRef] [PubMed]
  32. Carareto, C.M.; Hernandez, E.H.; Vieira, C. Genomic regions harboring insecticide resistance-associated Cyp genes are enriched by transposable element fragments carrying putative transcription factor binding sites in two sibling Drosophila species. Gene 2014, 537, 93–99. [Google Scholar] [CrossRef] [PubMed][Green Version]
  33. Wilding, C.S.; Smith, I.; Lynd, A.; Yawson, A.E.; Weetman, D.; Paine, M.J.; Donnelly, M.J. A cis-regulatory sequence driving metabolic insecticide resistance in mosquitoes: Functional characterisation and signatures of selection. Insect Biochem. Mol. Biol. 2012, 42, 699–707. [Google Scholar] [CrossRef] [PubMed]
  34. Cheng, D.; Guo, Z.; Riegler, M.; Xi, Z.; Liang, G.; Xu, Y. Gut symbiont enhances insecticide resistance in a significant pest, the oriental fruit fly Bactrocera dorsalis (Hendel). Microbiome 2017, 5, 13. [Google Scholar] [CrossRef][Green Version]
  35. Giraudo, M.; Hilliou, F.; Fricaux, T.; Audant, P.; Feyereisen, R.; le Goff, G. Cytochrome P450s from the fall armyworm (Spodoptera frugiperda): Responses to plant allelochemicals and pesticides. Insect Mol. Biol. 2015, 24, 115–128. [Google Scholar] [CrossRef] [PubMed]
  36. Shi, Y.; Wang, H.; Liu, Z.; Wu, S.; Yang, Y.; Feyereisen, R.; Heckel, D.G.; Wu, Y. Phylogenetic and functional characterization of ten P450 genes from the CYP6AE subfamily of Helicoverpa armigera involved in xenobiotic metabolism. Insect Biochem. Mol. Biol. 2018, 93, 79–91. [Google Scholar] [CrossRef] [PubMed]
  37. Rose, R.L.; Goh, D.; Thompson, D.M.; Verma, K.D.; Heckel, D.G.; Gahan, L.J.; Roe, R.M.; Hodgson, E. Cytochrome P450 (CYP)9A1 in Heliothis virescens: The first member of a new CYP family. Insect Biochem. Mol. Biol. 1997, 27, 605–615. [Google Scholar] [CrossRef]
  38. Yang, Y.; Chen, S.; Wu, S.; Yue, L.; Wu, Y. Constitutive overexpression of multiple cytochrome P450 genes associated with pyrethroid resistance in Helicoverpa armigera. J. Econ. Entomol. 2006, 99, 1784–1789. [Google Scholar] [CrossRef]
  39. Yang, Y.; Yue, L.; Chen, S.; Wu, Y. Functional expression of Helicoverpa armigera CYP9A12 and CYP9A14 in Saccharomyces cerevisiae. Pestic. Biochem. Physiol. 2008, 92, 101–105. [Google Scholar] [CrossRef]
  40. Boaventura, D.; Buer, B.; Hamaekers, N.; Maiwald, F.; Nauen, R. Toxicological and molecular profiling of insecticide resistance in a Brazilian strain of fall armyworm resistant to Bt Cry1 proteins. Pest. Manag. Sci. 2020. [Google Scholar] [CrossRef]
  41. Do Nascimento, A.R.; Fresia, P.; Consoli, F.L.; Omoto, C. Comparative transcriptome analysis of lufenuron-resistant and susceptible strains of Spodoptera frugiperda (Lepidoptera: Noctuidae). BMC Genom. 2015, 16, 985. [Google Scholar] [CrossRef] [PubMed][Green Version]
  42. Hafeez, M.; Liu, S.; Yousaf, H.K.; Jan, S.; Wang, R.L.; Fernandez-Grandon, G.M.; Li, X.; Gulzar, A.; Ali, B.; Rehman, M.; et al. RNA interference-mediated knockdown of a cytochrome P450 gene enhanced the toxicity of alpha-cypermethrin in xanthotoxin-fed larvae of Spodoptera exigua (Hubner). Pestic. Biochem. Physiol. 2020, 162, 6–14. [Google Scholar] [CrossRef]
  43. Hafeez, M.; Liu, S.; Jan, S.; Shi, L.; Fernandez-Grandon, G.M.; Gulzar, A.; Ali, B.; Rehman, M.; Wang, M. Knock-down of gossypol-inducing cytochrome P450 genes reduced deltamethrin sensitivity in Spodoptera exigua (Hubner). Int. J. Mol. Sci. 2019, 20, 2248. [Google Scholar] [CrossRef][Green Version]
  44. Wang, R.L.; Liu, S.W.; Baerson, S.R.; Qin, Z.; Ma, Z.H.; Su, Y.J.; Zhang, J.E. Identification and functional analysis of a novel cytochrome P450 gene CYP9A105 associated with pyrethroid detoxification in Spodoptera exigua Hubner. Int. J. Mol. Sci. 2018, 19, 737. [Google Scholar] [CrossRef][Green Version]
  45. Xu, L.; Mei, Y.; Liu, R.; Chen, X.; Li, D.; Wang, C. Transcriptome analysis of Spodoptera litura reveals the molecular mechanism to pyrethroids resistance. Pestic. Biochem. Physiol. 2020, 169, 104649. [Google Scholar] [CrossRef] [PubMed]
  46. Wang, R.L.; Staehelin, C.; Xia, Q.Q.; Su, Y.J.; Zeng, R.S. Identification and characterization of CYP9A40 from the tobacco cutworm moth (Spodoptera litura), a cytochrome P450 gene induced by plant allelochemicals and insecticides. Int. J. Mol. Sci. 2015, 16, 22606–22620. [Google Scholar] [CrossRef] [PubMed][Green Version]
  47. Oakeshott, J.; Claudianos, C.; Campbell, P.M.; Newcomb, R.; Russell, R.J. Biochemical genetics and genomics of insect esterases. In Comprehensive Molecular Insect Science-Pharmacology; Gilbert, L.I., Iatrou, K., Gill, S.S., Eds.; Elsevier: Amsterdam, The Netherlands, 2005; pp. 309–381. [Google Scholar]
  48. Li, X.; Schuler, M.A.; Berenbaum, M.R. Molecular mechanisms of metabolic resistance to synthetic and natural xenobiotics. Annu. Rev. Entomol. 2007, 52, 231–253. [Google Scholar] [CrossRef]
  49. Oakeshott, J.G.; Devonshire, A.L.; Claudianos, C.; Sutherland, T.D.; Horne, I.; Campbell, P.M.; Ollis, D.L.; Russell, R.J. Comparing the organophosphorus and carbamate insecticide resistance mutations in cholin- and carboxyl-esterases. Chem. Biol. Interact. 2005, 157–158, 269–275. [Google Scholar] [CrossRef]
  50. Oakeshott, J.G.; Johnson, R.M.; Berenbaum, M.R.; Ranson, H.; Cristino, A.S.; Claudianos, C. Metabolic enzymes associated with xenobiotic and chemosensory responses in Nasonia vitripennis. Insect Mol. Biol. 2010, 19 (Suppl. 1), 147–163. [Google Scholar] [CrossRef][Green Version]
  51. Pearce, S.L.; Clarke, D.F.; East, P.D.; Elfekih, S.; Gordon, K.H.J.; Jermiin, L.S.; McGaughran, A.; Oakeshott, J.G.; Papanikolaou, A.; Perera, O.P.; et al. Genomic innovations, transcriptional plasticity and gene loss underlying the evolution and divergence of two highly polyphagous and invasive Helicoverpa pest species. BMC Biol. 2017, 15, 63. [Google Scholar] [CrossRef][Green Version]
  52. Durand, N.; Carot-Sans, G.; Chertemps, T.; Montagne, N.; Jacquin-Joly, E.; Debernard, S.; Maibeche-Coisne, M. A diversity of putative carboxylesterases are expressed in the antennae of the noctuid moth Spodoptera littoralis. Insect Mol. Biol. 2010, 19, 87–97. [Google Scholar] [CrossRef] [PubMed]
  53. Durand, N.; Chertemps, T.; Maibeche-Coisne, M. Antennal carboxylesterases in a moth, structural and functional diversity. Commun. Integr. Biol. 2012, 5, 284–286. [Google Scholar] [CrossRef] [PubMed][Green Version]
  54. Merlin, C.; Rosell, G.; Carot-Sans, G.; Francois, M.C.; Bozzolan, F.; Pelletier, J.; Jacquin-Joly, E.; Guerrero, A.; Maibeche-Coisne, M. Antennal esterase cDNAs from two pest moths, Spodoptera littoralis and Sesamia nonagrioides, potentially involved in odourant degradation. Insect Mol. Biol. 2007, 16, 73–81. [Google Scholar] [CrossRef][Green Version]
  55. Walker, W.B., 3rd; Roy, A.; Anderson, P.; Schlyter, F.; Hansson, B.S.; Larsson, M.C. Transcriptome Analysis of gene families involved in chemosensory function in Spodoptera littoralis (Lepidoptera: Noctuidae). BMC Genom. 2019, 20, 428. [Google Scholar] [CrossRef] [PubMed]
  56. Farnsworth, C.A.; Teese, M.G.; Yuan, G.; Li, Y.; Scott, C.; Zhang, X.; Wu, Y.; Russell, R.J.; Oakeshott, J.G. Esterase-based metabolic resistance to insecticides in heliothine and spodopteran pests. J. Pestic. Sci. 2010, 35, 275–289. [Google Scholar] [CrossRef][Green Version]
  57. El-Guindy, M.A.; Saleh, W.S.; El-Refai, A.R.A.; Abou-Donia, S.A. The role of heamolymph esterases as protectants against intoxication by fenitrothion in the cotton leafworm Spodoptera littoralis (Boisd.). Bull. Entomol. Soc. Egypt. Econ. Ser. 1984, 14, 177–197. [Google Scholar]
  58. Cho, J.R.; Kim, Y.J.; Kim, J.J.; Kim, H.S.; Yoo, J.K.; Lee, J.O. Electrophoretic Pattern of Larval Esterases in field and laboratory-selected strains of the tobacco cutworm, Spodoptera litura (Fabricius). J. Asia-Pac. Entomol. 1999, 2, 39–44. [Google Scholar] [CrossRef]
  59. Kim, Y.G.; Lee, J.I.; Kang, S.Y.; Han, S.C. Variation in insecticide susceptibilities of the beet armyworm, Spodoptera exigua (Hubner): Esterase and acetylcholinesterase activities. Korean J. Appl. Entomol. 1997, 36, 172–178. [Google Scholar]
  60. Yu, S.J.; Nguyen, S.N.; Abo-Elghar, G.E. Biochemical characteristics of insecticide resistance in the fall armyworm, Spodoptera frugiperda (J.E. Smith). Pestic. Biochem. Physiol. 2003, 77, 1–11. [Google Scholar] [CrossRef]
  61. Han, Y.; Wu, S.; Li, Y.; Liu, J.; Campbell, P.M.; Farnsworth, C.A.; Scott, C.; Russell, R.J.; Oakeshott, J.G.; Wu, Y. Proteomic and molecular analyses of esterases associated with monocrotophos resistance in Helicoverpa armigera. Pestic. Biochem. Physiol. 2012, 104, 243–251. [Google Scholar] [CrossRef]
  62. Li, Y.; Farnsworth, C.A.; Coppin, C.W.; Teese, M.G.; Liu, J.W.; Scott, C.; Zhang, X.; Russell, R.J.; Oakeshott, J.G. Organophosphate and pyrethroid hydrolase activities of mutant Esterases from the cotton bollworm Helicoverpa armigera. PLoS ONE 2013, 8, e77685. [Google Scholar] [CrossRef][Green Version]
  63. Teese, M.G.; Farnsworth, C.A.; Li, Y.; Coppin, C.W.; Devonshire, A.L.; Scott, C.; East, P.; Russell, R.J.; Oakeshott, J.G. Heterologous expression and biochemical characterisation of fourteen esterases from Helicoverpa armigera. PLoS ONE 2013, 8, e65951. [Google Scholar] [CrossRef]
  64. Campbell, P.M.; Trott, J.F.; Claudianos, C.; Smyth, K.A.; Russell, R.J.; Oakeshott, J.G. Biochemistry of esterases associated with organophosphate resistance in Lucilia cuprina with comparisons to putative orthologues in other Diptera. Biochem. Genet. 1997, 35, 17–40. [Google Scholar] [CrossRef]
  65. Claudianos, C.; Russell, R.J.; Oakeshott, J.G. The same amino acid substitution in orthologous esterases confers organophosphate resistance on the house fly and a blowfly. Insect Biochem. Mol. Biol. 1999, 29, 675–686. [Google Scholar] [CrossRef]
  66. Newcomb, R.D.; Campbell, P.M.; Ollis, D.L.; Cheah, E.; Russell, R.J.; Oakeshott, J.G. A single amino acid substitution converts a carboxylesterase to an organophosphorus hydrolase and confers insecticide resistance on a blowfly. Proc. Natl. Acad. Sci. USA 1997, 94, 7464–7468. [Google Scholar] [CrossRef][Green Version]
  67. Cui, F.; Lin, Z.; Wang, H.; Liu, S.; Chang, H.; Reeck, G.; Qiao, C.; Raymond, M.; Kang, L. Two single mutations commonly cause qualitative change of nonspecific carboxylesterases in insects. Insect Biochem. Mol. Biol. 2011, 41, 1–8. [Google Scholar] [CrossRef]
  68. Zhu, Y.; Dowdy, A.K.; Baker, J.E. Detection of single-base substitution in an esterase gene and its linkage to malathion resistance in the parasitoid Anisopteromalus calandrae (Hymenoptera: Pteromalidae). Pestic. Sci. 1999, 55, 398–404. [Google Scholar] [CrossRef]
  69. Fournier, D. Mutations of acetylcholinesterase which confer insecticide resistance in insect populations. Chem. Biol. Interact. 2005, 157–158, 257–261. [Google Scholar] [CrossRef] [PubMed]
  70. Andrews, M.C.; Callaghan, A.; Field, L.M.; Williamson, M.S.; Moores, G.D. Identification of mutations conferring insecticide-insensitive AChE in the cotton-melon aphid, Aphis gossypii Glover. Insect Mol. Biol. 2004, 13, 555–561. [Google Scholar] [CrossRef]
  71. Hsu, J.C.; Haymer, D.S.; Wu, W.J.; Feng, H.T. Mutations in the acetylcholinesterase gene of Bactrocera dorsalis associated with resistance to organophosphorus insecticides. Insect Biochem. Mol. Biol. 2006, 36, 396–402. [Google Scholar] [CrossRef] [PubMed]
  72. Jiang, X.; Qu, M.; Denholm, I.; Fang, J.; Jiang, W.; Han, Z. Mutation in acetylcholinesterase1 associated with triazophos resistance in rice stem borer, Chilo suppressalis (Lepidoptera: Pyralidae). Biochem. Biophys. Res. Commun. 2009, 378, 269–272. [Google Scholar] [CrossRef] [PubMed]
  73. Lee, D.W.; Choi, J.Y.; Kim, W.T.; Je, Y.H.; Song, J.T.; Chung, B.K.; Boo, K.S.; Koh, Y.H. Mutations of acetylcholinesterase1 contribute to prothiofos-resistance in Plutella xylostella (L.). Biochem. Biophys. Res. Commun. 2007, 353, 591–597. [Google Scholar] [CrossRef] [PubMed]
  74. Kim, Y.; Cho, J.R.; Lee, J.I.; Kang, S.Y.; Han, S.C.; Hong, K.J.; Kim, H.S.; Yoo, J.K.; Lee, J.O. Insecticide resistance in the tobacco cutworm, Spodoptera litura (Fabricius) (Lepidoptera: Noctuidae). J. Asia-Pac. Entomol. 1998, 1, 115–122. [Google Scholar]
  75. Kranthi, K.R.; Jadhav, D.R.; Kranthi, S.; Wanjari, R.R.; Ali, S.S.; Russell, D.A. Insecticide resistance in five major insect pests of cotton in India. Crop. Prot. 2002, 21, 449–460. [Google Scholar] [CrossRef]
  76. Muthusamy, Shivakumar, Karthi, and Ramkumar, Pesticide detoxifying mechanism in field population of Spodoptera litura (Lepidoptera: Noctuidae) from South India Egypt. Acad. J. Biolog. Sci. 2011, 3, 51–57.
  77. Byrne, F.J.; Toscano, N.C. An insensitive acetylcholinesterase confers resistance to methomyl in the beet armyworm Spodoptera exigua (Lepidoptera: Noctuidae). J. Econ. Entomol. 2001, 94, 524–528. [Google Scholar] [CrossRef]
  78. Carvalho, R.A.; Omoto, C.; Field, L.M.; Williamson, M.S.; Bass, C. Investigating the molecular mechanisms of organophosphate and pyrethroid resistance in the fall armyworm Spodoptera frugiperda. PLoS ONE 2013, 8, e62268. [Google Scholar] [CrossRef][Green Version]
  79. Boaventura, D.; Martin, M.; Pozzebon, A.; Mota-Sanchez, D.; Nauen, R. Monitoring of target-site mutations conferring insecticide resistance in Spodoptera frugiperda. Insects 2020, 11, 545. [Google Scholar] [CrossRef]
  80. Guan, F.; Zhang, J.; Shen, H.; Wang, X.; Padovan, A.; Walsh, T.K.; Tay, W.T.; Gordon, K.H.J.; James, W.; Czepak, C.; et al. Whole-genome sequencing to detect mutations associated with resistance to insecticides and Bt proteins in Spodoptera frugiperda. Insect Sci. 2020. [Google Scholar] [CrossRef]
  81. Jakoby, W.B.; Ziegler, D.M. The enzymes of detoxication. J. Biol. Chem. 1990, 265, 20715–20718. [Google Scholar] [CrossRef]
  82. Enayati, A.A.; Ranson, H.; Hemingway, J. Insect glutathione transferases and insecticide resistance. Insect Mol. Biol. 2005, 14, 3–8. [Google Scholar] [CrossRef] [PubMed][Green Version]
  83. Sheehan, D.; Meade, G.; Foley, V.M.; Dowd, C.A. Structure, function and evolution of glutathione transferases: Implications for classification of non-mammalian members of an ancient enzyme superfamily. Biochem. J. 2001, 360 Pt 1, 1–16. [Google Scholar] [CrossRef]
  84. Hu, B.; Hu, S.; Huang, H.; Wei, Q.; Ren, M.; Huang, S.; Tian, X.; Su, J. Insecticides induce the co-expression of glutathione S-transferases through ROS/CncC pathway in Spodoptera exigua. Pestic. Biochem. Physiol. 2019, 155, 58–71. [Google Scholar] [CrossRef]
  85. Friedman, R. Genomic organization of the glutathione S-transferase family in insects. Mol. Phylogenet. Evol. 2011, 61, 924–932. [Google Scholar] [CrossRef] [PubMed]
  86. Kostaropoulos, I.; Papadopoulos, A.I.; Metaxakis, A.; Boukouvala, E.; Papadopoulou-Mourkidou, E. Glutathione S-transferase in the defence against pyrethroids in insects. Insect Biochem. Mol. Biol. 2001, 31, 313–319. [Google Scholar] [CrossRef]
  87. Alias, Z.; Clark, A.G. Studies on the glutathione S-transferase proteome of adult Drosophila melanogaster: Responsiveness to chemical challenge. Proteomics 2007, 7, 3618–3628. [Google Scholar] [CrossRef] [PubMed]
  88. Lumjuan, N.; Rajatileka, S.; Changsom, D.; Wicheer, J.; Leelapat, P.; Prapanthadara, L.A.; Somboon, P.; Lycett, G.; Ranson, H. The role of the Aedes aegypti Epsilon glutathione transferases in conferring resistance to DDT and pyrethroid insecticides. Insect Biochem. Mol. Biol. 2011, 41, 203–209. [Google Scholar] [CrossRef]
  89. Hardy, N.B.; Peterson, D.A.; Ross, L.; Rosenheim, J.A. Does a plant-eating insect’s diet govern the evolution of insecticide resistance? Comparative tests of the pre-adaptation hypothesis. Evol. Appl. 2018, 11, 739–747. [Google Scholar] [CrossRef][Green Version]
  90. Pavlidi, N.; Vontas, J.; van Leeuwen, T. The role of glutathione S-transferases (GSTs) in insecticide resistance in crop pests and disease vectors. Curr. Opin. Insect Sci. 2018, 27, 97–102. [Google Scholar] [CrossRef] [PubMed]
  91. Vontas, J.G.; Small, G.J.; Hemingway, J. Glutathione S-transferases as antioxidant defence agents confer pyrethroid resistance in Nilaparvata lugens. Biochem. J. 2001, 357 Pt 1, 65–72. [Google Scholar] [CrossRef]
  92. Deng, H.; Huang, Y.; Feng, Q.; Zheng, S. Two epsilon glutathione S-transferase cDNAs from the common cutworm, Spodoptera litura: Characterization and developmental and induced expression by insecticides. J. Insect Physiol. 2009, 55, 1174–1183. [Google Scholar] [CrossRef] [PubMed]
  93. Wan, H.; Zhan, S.; Xia, X.; Xu, P.; You, H.; Jin, B.R.; Li, J. Identification and functional characterization of an epsilon glutathione S-transferase from the beet armyworm (Spodoptera exigua). Pestic. Biochem. Physiol. 2016, 132, 81–88. [Google Scholar] [CrossRef] [PubMed]
  94. Hu, B.; Huang, H.; Wei, Q.; Ren, M.; Mburu, D.K.; Tian, X.; Su, J. Transcription factors CncC/Maf and AhR/ARNT coordinately regulate the expression of multiple GSTs conferring resistance to chlorpyrifos and cypermethrin in Spodoptera exigua. Pest. Manag. Sci. 2019, 75, 2009–2019. [Google Scholar] [CrossRef]
  95. Xu, Z.B.; Zou, X.P.; Zhang, N.; Feng, Q.L.; Zheng, S.C. Detoxification of insecticides, allechemicals and heavy metals by glutathione S-transferase SlGSTE1 in the gut of Spodoptera litura. Insect Sci. 2015, 22, 503–511. [Google Scholar] [CrossRef] [PubMed]
  96. Zou, X.; Xu, Z.; Zou, H.; Liu, J.; Chen, S.; Feng, Q.; Zheng, S. Glutathione S-transferase SlGSTE1 in Spodoptera litura may be associated with feeding adaptation of host plants. Insect Biochem Mol. Biol. 2016, 70, 32–43. [Google Scholar] [CrossRef]
  97. Hirowatari, A.; Chen, Z.; Mita, K.; Yamamoto, K. Enzymatic characterization of two epsilon-class glutathione S-transferases of Spodoptera litura. Arch. Insect Biochem. Physiol. 2018, 97. [Google Scholar] [CrossRef]
  98. Huang, Y.; Xu, Z.; Lin, X.; Feng, Q.; Zheng, S. Structure and expression of glutathione S-transferase genes from the midgut of the common cutworm, Spodoptera litura (Noctuidae) and their response to xenobiotic compounds and bacteria. J. Insect Physiol. 2011, 57, 1033–1044. [Google Scholar] [CrossRef]
  99. Zhang, N.; Liu, J.; Chen, S.N.; Huang, L.H.; Feng, Q.L.; Zheng, S.C. Expression profiles of glutathione S-transferase superfamily in Spodoptera litura tolerated to sublethal doses of chlorpyrifos. Insect Sci. 2016, 23, 675–687. [Google Scholar] [CrossRef]
  100. Lalouette, L.; Pottier, M.A.; Wycke, M.A.; Boitard, C.; Bozzolan, F.; Maria, A.; Demondion, E.; Chertemps, T.; Lucas, P.; Renault, D.; et al. Unexpected effects of sublethal doses of insecticide on the peripheral olfactory response and sexual behavioral in a pest insect. Environ. Sci. Pollut. Res. 2016, 23, 3073–3085. [Google Scholar] [CrossRef]
  101. Chen, X.; Zhang, Y.L. Identification and characterisation of multiple glutathione S-transferase genes from the diamondback moth, Plutella xylostella. Pest. Manag. Sci. 2015, 71, 592–600. [Google Scholar] [CrossRef]
  102. Huang, H.S.; Hu, N.T.; Yao, Y.E.; Wu, C.Y.; Chiang, S.W.; Sun, C.N. Molecular cloning and heterologous expression of a glutathione S-transferase involved in insecticide resistance from the diamondback moth, Plutella xylostella. Insect Biochem. Mol. Biol. 1998, 28, 651–658. [Google Scholar] [CrossRef]
  103. Shi, L.; Shi, Y.; Zhang, Y.; Liao, X. A systemic study of indoxacarb resistance in Spodoptera litura revealed complex expression profiles and regulatory mechanism. Sci. Rep. 2019, 9, 14997. [Google Scholar] [CrossRef][Green Version]
  104. Shi, L.; Shi, Y.; Liu, M.F.; Zhang, Y.; Liao, X.L. Transcription factor CncC potentially regulates the expression of multiple detoxification genes that mediate indoxacarb resistance in Spodoptera litura. Insect Sci. 2020. [Google Scholar] [CrossRef]
  105. Li, Z.; Cai, T.; Qin, Y.; Zhang, Y.; Jin, R.; Mao, K.; Liao, X.; Wan, H.; Li, J. Transcriptional Response of ATP-Binding Cassette (ABC) Transporters to Insecticide in the Brown Planthopper, Nilaparvata lugens (Stal). Insects 2020, 11, 280. [Google Scholar] [CrossRef]
  106. Qi, W.; Ma, X.; He, W.; Chen, W.; Zou, M.; Gurr, G.M.; Vasseur, L.; You, M. Characterization and expression profiling of ATP-binding cassette transporter genes in the diamondback moth, Plutella xylostella (L.). BMC Genom. 2016, 17, 760. [Google Scholar] [CrossRef][Green Version]
  107. Briz, O.; Perez-Silva, L.; Al-Abdulla, R.; Abete, L.; Reviejo, M.; Romero, M.R.; Marin, J.J.G. What “The Cancer Genome Atlas” database tells us about the role of ATP-binding cassette (ABC) proteins in chemoresistance to anticancer drugs. Expert Opin Drug Metab. Toxicol. 2019, 15, 577–593. [Google Scholar] [CrossRef] [PubMed]
  108. Gellatly, K.J.; Yoon, K.S.; Doherty, J.J.; Sun, W.; Pittendrigh, B.R.; Clark, J.M. RNAi validation of resistance genes and their interactions in the highly DDT-resistant 91-R strain of Drosophila melanogaster. Pestic. Biochem. Physiol. 2015, 121, 107–115. [Google Scholar] [CrossRef] [PubMed][Green Version]
  109. Jurat-Fuentes, J.L.; Heckel, D.G.; Ferre, J. Mechanisms of Resistance to Insecticidal Proteins from Bacillus thuringiensis. Annu. Rev. Entomol. 2021, 66, 121–140. [Google Scholar] [CrossRef] [PubMed]
  110. Gahan, L.J.; Pauchet, Y.; Vogel, H.; Heckel, D.G. An ABC transporter mutation is correlated with insect resistance to Bacillus thuringiensis Cry1Ac toxin. PLoS Genet. 2010, 6, e1001248. [Google Scholar] [CrossRef][Green Version]
  111. Liu, L.; Chen, Z.; Yang, Y.; Xiao, Y.; Liu, C.; Ma, Y.; Soberon, M.; Bravo, A.; Yang, Y.; Liu, K. A single amino acid polymorphism in ABCC2 loop 1 is responsible for differential toxicity of Bacillus thuringiensis Cry1Ac toxin in different Spodoptera (Noctuidae) species. Insect Biochem. Mol. Biol. 2018, 100, 59–65. [Google Scholar] [CrossRef] [PubMed]
  112. Endo, H.; Tanaka, S.; Adegawa, S.; Ichino, F.; Tabunoki, H.; Kikuta, S.; Sato, R. Extracellular loop structures in silkworm ABCC transporters determine their specificities for Bacillus thuringiensis Cry toxins. J. Biol. Chem. 2018, 293, 8569–8577. [Google Scholar] [CrossRef] [PubMed][Green Version]
  113. De Maagd, R.A.; Bravo, A.; Crickmore, N. How Bacillus thuringiensis has evolved specific toxins to colonize the insect world. Trends Genet. 2001, 17, 193–199. [Google Scholar] [CrossRef]
  114. Banerjee, R.; Hasler, J.; Meagher, R.; Nagoshi, R.; Hietala, L.; Huang, F.; Narva, K.; Jurat-Fuentes, J.L. Mechanism and DNA-based detection of field-evolved resistance to transgenic Bt corn in fall armyworm (Spodoptera frugiperda). Sci. Rep. 2017, 7, 10877. [Google Scholar] [CrossRef] [PubMed][Green Version]
  115. Flagel, L.; Lee, Y.W.; Wanjugi, H.; Swarup, S.; Brown, A.; Wang, J.; Kraft, E.; Greenplate, J.; Simmons, J.; Adams, N.; et al. Mutational disruption of the ABCC2 gene in fall armyworm, Spodoptera frugiperda, confers resistance to the Cry1Fa and Cry1A.105 insecticidal proteins. Sci. Rep. 2018, 8, 7255. [Google Scholar] [CrossRef][Green Version]
  116. Boaventura, D.; Ulrich, J.; Lueke, B.; Bolzan, A.; Okuma, D.; Gutbrod, O.; Geibel, S.; Zeng, Q.; Dourado, P.M.; Martinelli, S.; et al. Molecular characterization of Cry1F resistance in fall armyworm, Spodoptera frugiperda from Brazil. Insect Biochem. Mol. Biol. 2020, 116, 103280. [Google Scholar] [CrossRef]
  117. Park, Y.; Gonzalez-Martinez, R.M.; Navarro-Cerrillo, G.; Chakroun, M.; Kim, Y.; Ziarsolo, P.; Blanca, J.; Canizares, J.; Ferre, J.; Herrero, S. ABCC transporters mediate insect resistance to multiple Bt toxins revealed by bulk segregant analysis. BMC Biol. 2014, 12, 46. [Google Scholar] [CrossRef][Green Version]
  118. Moar, W.J.; Pusztai-Carey, M.; van Faassen, H.; Bosch, D.; Frutos, R.; Rang, C.; Luo, K.; Adang, M.J. Development of Bacillus thuringiensis CryIC resistance by Spodoptera exigua (Hubner) (Lepidoptera: Noctuidae). Appl. Environ. Microbiol. 1995, 61, 2086–2092. [Google Scholar] [CrossRef][Green Version]
  119. Müller-Cohn, J.; Chaufaux, J.; Buisson, C.; Gilois, N.; Sanchis, V.; Lereclus, D. Spodoptera littoralis (Lepidoptera: Noctuidae) resistance to CryIC and cross-resistance to other Bacillus thuringiensis crystal toxins. J. Econ. Entomol. 1996, 89, 791–797. [Google Scholar] [CrossRef]
  120. Barkhade, U.P.; Thakare, A.S. Protease mediated resistance mechanism to Cry1C and Vip3A in Spodoptera litura. Egypt. Acad. J. Biol. Sci. 2010, 6, 43–50. [Google Scholar] [CrossRef]
  121. Hernandez-Martinez, P.; Navarro-Cerrillo, G.; Caccia, S.; de Maagd, R.A.; Moar, W.J.; Ferre, J.; Escriche, B.; Herrero, S. Constitutive activation of the midgut response to Bacillus thuringiensis in Bt-resistant Spodoptera exigua. PLoS ONE 2010, 5. [Google Scholar] [CrossRef]
  122. Pinos, D.; Martinez-Solis, M.; Herrero, S.; Ferre, J.; Hernandez-Martinez, P. The Spodoptera exigua ABCC2 acts as a Cry1A receptor independently of its nucleotide binding domain II. Toxins 2019, 11, 172. [Google Scholar] [CrossRef] [PubMed][Green Version]
  123. Huang, J.; Xu, Y.; Zuo, Y.; Yang, Y.; Tabashnik, B.E.; Wu, Y. Evaluation of five candidate receptors for three Bt toxins in the beet armyworm using CRISPR-mediated gene knockouts. Insect Biochem. Mol. Biol. 2020, 121, 103361. [Google Scholar] [CrossRef]
  124. Abdelgaffar, H.; Perera, O.P.; Jurat-Fuentes, J.L. ABC transporter mutations in Cry1F-resistant fall armyworm (Spodoptera frugiperda) do not result in altered susceptibility to selected small molecule pesticides. Pest. Manag. Sci. 2021, 77, 949–955. [Google Scholar] [CrossRef]
  125. Xiang, M.; Zhang, L.; Lu, Y.; Tang, Q.; Liang, P.; Shi, X.; Song, D.; Gao, X. A P-glycoprotein gene serves as a component of the protective mechanisms against 2-tridecanone and abamectin in Helicoverpa armigera. Gene 2017, 627, 63–71. [Google Scholar] [CrossRef]
  126. Jin, M.; Liao, C.; Chakrabarty, S.; Zheng, W.; Wu, K.; Xiao, Y. Transcriptional response of ATP-binding cassette (ABC) transporters to insecticides in the cotton bollworm, Helicoverpa armigera. Pestic. Biochem. Physiol. 2019, 154, 46–59. [Google Scholar] [CrossRef]
  127. Zuo, Y.Y.; Huang, J.L.; Wang, J.; Feng, Y.; Han, T.T.; Wu, Y.D.; Yang, Y.H. Knockout of a P-glycoprotein gene increases susceptibility to abamectin and emamectin benzoate in Spodoptera exigua. Insect Mol. Biol. 2017. [Google Scholar] [CrossRef]
  128. Jin, M.; Yang, Y.; Shan, Y.; Chakrabarty, S.; Cheng, Y.; Soberon, M.; Bravo, A.; Liu, K.; Wu, K.; Xiao, Y. Two ABC transporters are differentially involved in the toxicity of two Bacillus thuringiensis Cry1 toxins to the invasive crop-pest Spodoptera frugiperda (J. E. Smith). Pest. Manag. Sci. 2021, 77, 1492–1501. [Google Scholar] [CrossRef]
  129. Genovese, I.; Ilari, A.; Assaraf, Y.G.; Fazi, F.; Colotti, G. Not only P-glycoprotein: Amplification of the ABCB1-containing chromosome region 7q21 confers multidrug resistance upon cancer cells by coordinated overexpression of an assortment of resistance-related proteins. Drug Resist. Updat. 2017, 32, 23–46. [Google Scholar] [CrossRef]
  130. Zhu, B.; Sun, X.; Nie, X.; Liang, P.; Gao, X. MicroRNA-998-3p contributes to Cry1Ac-resistance by targeting ABCC2 in lepidopteran insects. Insect Biochem. Mol. Biol. 2020, 117, 103283. [Google Scholar] [CrossRef] [PubMed]
  131. Li, J.; Ma, Y.; Yuan, W.; Xiao, Y.; Liu, C.; Wang, J.; Peng, J.; Peng, R.; Soberon, M.; Bravo, A.; et al. FOXA transcriptional factor modulates insect susceptibility to Bacillus thuringiensis Cry1Ac toxin by regulating the expression of toxin-receptor ABCC2 and ABCC3 genes. Insect Biochem. Mol. Biol. 2017, 88, 1–11. [Google Scholar] [CrossRef] [PubMed]
  132. Bo, H.; Miaomiao, R.; Jianfeng, F.; Sufang, H.; Xia, W.; Elzaki, M.E.A.; Chris, B.; Palli, S.R.; Jianya, S. Xenobiotic transcription factors CncC and maf regulate expression of CYP321A16 and CYP332A1 that mediate chlorpyrifos resistance in Spodoptera exigua. J. Hazard. Mater. 2020, 398, 122971. [Google Scholar] [CrossRef] [PubMed]
  133. Pan, Y.; Zeng, X.; Wen, S.; Liu, X.; Shang, Q. Characterization of the Cap ‘n’ Collar Isoform C gene in Spodoptera frugiperda and its Association with Superoxide Dismutase. Insects 2020, 11, 221. [Google Scholar] [CrossRef] [PubMed][Green Version]
Figure 1. CYP9A cluster in the genome of S. litura (A), S. frugiperda corn strain (B) and S. frugiperda rice strain (C). Numbers in blue correspond to CYP9A genes. Arrows correspond to gene orientation, ADH: alcohol dehydrogenase, FT: fucosyl transferase, EST TF: E26 transformation specific transcription factor, * pseudogene.
Figure 1. CYP9A cluster in the genome of S. litura (A), S. frugiperda corn strain (B) and S. frugiperda rice strain (C). Numbers in blue correspond to CYP9A genes. Arrows correspond to gene orientation, ADH: alcohol dehydrogenase, FT: fucosyl transferase, EST TF: E26 transformation specific transcription factor, * pseudogene.
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Figure 2. CYP9A subtree: ML phylogeny with bootstrap value. SfC: S. frugiperda corn strain, SfR: S. frugiperda rice strain, Sl: S. litura, Slit: S. littoralis, Bm: B. mori. Slit9A58, Slit9A81, and Slit9A40-like are missing from the tree. Black square: ancient speciation representing B. mori split from Spodoptera species. Black dot: duplication events.
Figure 2. CYP9A subtree: ML phylogeny with bootstrap value. SfC: S. frugiperda corn strain, SfR: S. frugiperda rice strain, Sl: S. litura, Slit: S. littoralis, Bm: B. mori. Slit9A58, Slit9A81, and Slit9A40-like are missing from the tree. Black square: ancient speciation representing B. mori split from Spodoptera species. Black dot: duplication events.
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Figure 3. Alignment of two 50 amino acid sequence portions of SlituCOE082, SlitCXE64, SfruCCE16j and MdaE7 from Musca domestica (Rutgers diazinon-R resistant strain). The putative mutation positions G151D and W271L are shaded in grey. According to [67], the amino acid corresponding to G137D and W251L were G107 and W219 in S. litura sequence; for S. litura carboxyl/cholinesterase (CCE), the mutation tested in vitro was G151D. Genbank accession number, S. frugiperda (XP_035450257.1), S. litura (XP_022828113.1), M. domestica (AAD29685.1).
Figure 3. Alignment of two 50 amino acid sequence portions of SlituCOE082, SlitCXE64, SfruCCE16j and MdaE7 from Musca domestica (Rutgers diazinon-R resistant strain). The putative mutation positions G151D and W271L are shaded in grey. According to [67], the amino acid corresponding to G137D and W251L were G107 and W219 in S. litura sequence; for S. litura carboxyl/cholinesterase (CCE), the mutation tested in vitro was G151D. Genbank accession number, S. frugiperda (XP_035450257.1), S. litura (XP_022828113.1), M. domestica (AAD29685.1).
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Figure 4. Maximum-likelihood phylogeny of spodopteran CCEs from Clade (016) and acetylcholinesterases (AChEs). The tree was built from amino-acid sequences of CCE repertoires of S. littoralis (branches colored in red), S. frugiperda Corn strains (green), S. litura (blue), S. exigua (purple). Sequences from [1,28] for S. litura and S. frugiperda, respectively. For S. littoralis, from [52,55] and new sequence from our RNAseq database (Table S1). For S. exigua, GenBank numbers were inserted the sequence name, SexiAChE1 (AZB49078.1) and SexiAChE2 (AZB49079.1). Likelihood-ratio test values are only indicated in the internal nodes defining orthogroups, when aLRT > 0.9. The scale bar represents 0.2 expected amino-acid substitutions per site.
Figure 4. Maximum-likelihood phylogeny of spodopteran CCEs from Clade (016) and acetylcholinesterases (AChEs). The tree was built from amino-acid sequences of CCE repertoires of S. littoralis (branches colored in red), S. frugiperda Corn strains (green), S. litura (blue), S. exigua (purple). Sequences from [1,28] for S. litura and S. frugiperda, respectively. For S. littoralis, from [52,55] and new sequence from our RNAseq database (Table S1). For S. exigua, GenBank numbers were inserted the sequence name, SexiAChE1 (AZB49078.1) and SexiAChE2 (AZB49079.1). Likelihood-ratio test values are only indicated in the internal nodes defining orthogroups, when aLRT > 0.9. The scale bar represents 0.2 expected amino-acid substitutions per site.
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Figure 5. Alignment of a 120 amino acid sequence portion of AChE1 S. frugiperda susceptible (Genbank accession number KC435023.1) and resistant strain (KC435024.1) with orthologous sequences from S. litura (XP_022819835.1 = SlituCOE002), S. littoralis (Table S1) and S. exigua (AZB49078.1). The mutation positions are shaded in grey.
Figure 5. Alignment of a 120 amino acid sequence portion of AChE1 S. frugiperda susceptible (Genbank accession number KC435023.1) and resistant strain (KC435024.1) with orthologous sequences from S. litura (XP_022819835.1 = SlituCOE002), S. littoralis (Table S1) and S. exigua (AZB49078.1). The mutation positions are shaded in grey.
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Figure 6. Maximum-likelihood phylogeny of lepidopteran epsilon glutathione-S-transferases (GSTes). The tree was built from amino-acid sequences of GST repertoires of S. littoralis (branches colored in red), S. frugiperda Corn and Rice strains (green), S. litura (blue), S. exigua (purple), H. armigera, P. xylostella and B. mori (black). Likelihood-ratio test values are only indicated in the internal nodes defining orthogroups, when aLRT > 0.9. Black stars indicate that GSTes are overexpressed in insecticide-resistant strains or when exposed to pesticides. Red dots indicate GSTes with demonstrated in vitro/in vivo insecticide resistance. The scale bar represents 0.2 expected amino-acid substitutions per site.
Figure 6. Maximum-likelihood phylogeny of lepidopteran epsilon glutathione-S-transferases (GSTes). The tree was built from amino-acid sequences of GST repertoires of S. littoralis (branches colored in red), S. frugiperda Corn and Rice strains (green), S. litura (blue), S. exigua (purple), H. armigera, P. xylostella and B. mori (black). Likelihood-ratio test values are only indicated in the internal nodes defining orthogroups, when aLRT > 0.9. Black stars indicate that GSTes are overexpressed in insecticide-resistant strains or when exposed to pesticides. Red dots indicate GSTes with demonstrated in vitro/in vivo insecticide resistance. The scale bar represents 0.2 expected amino-acid substitutions per site.
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Figure 7. GSTe clusters derived from B. mori GSTe2 and GSTe3 identified in the genomes of S. litura, S. frugiperda corn and rice strains and S. exigua (adapted from [1]).
Figure 7. GSTe clusters derived from B. mori GSTe2 and GSTe3 identified in the genomes of S. litura, S. frugiperda corn and rice strains and S. exigua (adapted from [1]).
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Figure 8. Alignment of a subset of GSTes from S. litura (Sl) S. frugiperda corn and rice strains (SfC and SfR), S. exigua (Se), S. littoralis (Slit) and AfunGSTe2 from Anopheles funestus (Afun, genbank accession number KC800363.1) harboring a L119F substitution. The L119F mutation (in red) is associated with DDT resistance in Anophelines [6].
Figure 8. Alignment of a subset of GSTes from S. litura (Sl) S. frugiperda corn and rice strains (SfC and SfR), S. exigua (Se), S. littoralis (Slit) and AfunGSTe2 from Anopheles funestus (Afun, genbank accession number KC800363.1) harboring a L119F substitution. The L119F mutation (in red) is associated with DDT resistance in Anophelines [6].
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Figure 9. Alignment of a 20 amino acid sequence of ABCC2 from four Spodoptera species, presence of a glutamine or glutamic acid at position 119. Genbank accession number, S. exigua AIB06822, S. frugiperda AUO38740, S. littoralis, S. litura XM_022967434.
Figure 9. Alignment of a 20 amino acid sequence of ABCC2 from four Spodoptera species, presence of a glutamine or glutamic acid at position 119. Genbank accession number, S. exigua AIB06822, S. frugiperda AUO38740, S. littoralis, S. litura XM_022967434.
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Figure 10. Protein structure of ABCC2 of Spodoptera spp from susceptible or resistant populations. (A) Protein structure of SfABCC2 and SeABCC2 wild-type. (B,C) Protein structures of SfABCC2 in resistant populations from Puerto Rico, * corresponds to an insertion of amino acids [114,115]. (D,E) Protein structure of SfABCC2 in resistant populations from Brazil [80,116]. (F) Protein structure of SeABCC2 from a laboratory selected strain [117].
Figure 10. Protein structure of ABCC2 of Spodoptera spp from susceptible or resistant populations. (A) Protein structure of SfABCC2 and SeABCC2 wild-type. (B,C) Protein structures of SfABCC2 in resistant populations from Puerto Rico, * corresponds to an insertion of amino acids [114,115]. (D,E) Protein structure of SfABCC2 in resistant populations from Brazil [80,116]. (F) Protein structure of SeABCC2 from a laboratory selected strain [117].
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Table 1. Assessment of insecticide resistance worldwide for the four species of Spodoptera, data extracted from the Arthropod Pesticide Resistance Database (, accessed on 18 May 2021).
Table 1. Assessment of insecticide resistance worldwide for the four species of Spodoptera, data extracted from the Arthropod Pesticide Resistance Database (, accessed on 18 May 2021).
Chemical Class
S. exiguaS. frugiperdaS. littoralisS. litura
AvermectinsAbamectin (#31), emamectin benzoate (#48) Abamectin (#43), emamectin benzoate (#27)
BenzoylureaChlorfluazuron (#9), diflubenzuron, lufenuron (#10), teflubenzuronLufenuron (#2), triflumuronDiflubenzuron, teflubenzuron (#3)Lufenuron (#8)
Bacillus thuringensisBt var. unspecified (#10), var aizawai, var. kurstaki Bt var unspecified (#3), var aizawai
Bt toxinsCry1CaCry1Aa, Cry1A.105, Cry1Ab (#2), Cry1Ac (#3), Cry1F (#54), Cry2Ab2 (#2), Vip3A (#3)
CarbamatesMethomyl (#18), thiodicarbCarbaryl (#7), methomyl (#6), thiodicarb (#2)Carbaryl (#2), methomyl (#2),Carbaryl (#2), methomyl (#38), thiodicarb (#33)
CyclodienesBHC/cyclodiene—unspecified in literature (#3), endosulfan (#19)Aldrin, dieldrin, lindane (#2)Endrin, toxaphene (#2)Endosulfan (#31), lindane
DiamidesChlorantraniliprole (#26), cyantraniliprole, flubendiamide (#5)Chlorantraniliprole (#2), flubendiamide (#2) Chlorantraniliprole (#11)
DiacylhydrazinesMethoxyfenozide (#25), tebufenozide (#19) TebufenozideMethoxyfenozide (#36), tebufenozide
Neonicotinoids Acetamiprid
OrganochlorineDDT (#4)DDT (#3)DDTDDT (#2)
OrganophosphatesChlorpyrifos (#48), parathion-methyl (#3), profenofos (#22), quinalphos (#9)Acephate, chlorpurifos (#7), diazinon (#2), dichlorvos, malathion (#2), parathion-methyl (#4), sulprofos, trichlorfon,Acephate, azinphos-methyl, chlorpyrifos (#4), fenitrothion, leptophos, methamidophos, methidathion, monocrotophos (#4), parathion (#2), parathion-methyl (#3), profenofos, sulprofos, triazophos, trichlorfonChlorfenvinphos, chlorpyrifos (#55), diazinon, dichlorvos, malathion, monocrotophos (#5), phoxim (#4), profenofos (#56), quinalphos (#9), triazophos (#2), trichlorfon
OxadiazinesIndoxacarb (#44), metaflumizone (#4) IndoxacarbIndoxacarb (#50), metaflumizone
Phenylpyrazoles Fipronil (#15)
PyrethroidsBifenthrine (#13), cyfluthrin (#2), cyhalothrin-lambda (#2), cypermethrin (#56), cypermethrin-beta, deltamethrin (#43), fenpropathrin (#14), fenvalerate (#5), permethrin (#2), pyrethroids-unspecified in literature (#3)Bifenthrine, cyfluthrin, cyhalothrin, cyhalothrin-lambda (#11), cypermethrin (#2), cypermethrin-zeta, deltamethrin (#2), fenvalerate, fluvalinate, permethrin (#5), tau-fluvalinate, tralomethrinCypermethrin (#4), deltamethrin (#2), fenvalerate, flucythrinate, permethrinBifenthrine (#33), cyfluthrin (#12), cyfluthrin-beta (#9), cyhalothrin (#3), cyhalothrin-lambda (#11), cypermethrin (#49), cypermethrin-beta (#15), deltamethrin (#50), esfenvalerate (#9), fenpropathrin (#10), fenvalerate (#7), pyrethrins
PyrrolesChlorfenapyr (#11) Chlorfenapyr (#5)
SpinosynsSpinetoram, spinosad (#56) Spinetoram (#2), spinosad (#2)Spinosad (#39)
# number of cases reported in Arthropod Pesticide Resistance database.
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Hilliou, F.; Chertemps, T.; Maïbèche, M.; Le Goff, G. Resistance in the Genus Spodoptera: Key Insect Detoxification Genes. Insects 2021, 12, 544.

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Hilliou F, Chertemps T, Maïbèche M, Le Goff G. Resistance in the Genus Spodoptera: Key Insect Detoxification Genes. Insects. 2021; 12(6):544.

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Hilliou, Frédérique, Thomas Chertemps, Martine Maïbèche, and Gaëlle Le Goff. 2021. "Resistance in the Genus Spodoptera: Key Insect Detoxification Genes" Insects 12, no. 6: 544.

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