Role of nicotinic acetylcholine receptor subunits in the mode of action of neonicotinoid, sulfoximine and spinosyn insecticides in Drosophila melanogaster
Graphical abstract
Introduction
For more than a century insecticide resistance has reduced the capacity of successive generations of insecticides to control pest insects that damage crops and threaten human and animal health (Ffrench-Constant, 2013; McKenzie and Batterham, 1998; Nauen and Denholm, 2005; Whalon et al., 2008). Insecticides are still pivotal for the control of many pest insects, although a wide range of other practices are routinely implemented. Efforts to prevent or stall the evolution and spread of resistance to current and future insecticidal compounds are facilitated by a detailed understanding of their mode of action (Nauen et al., 2019; Sparks and Nauen, 2015). This knowledge is increasingly valuable in light of losses caused by ineffective pest control and the increasing technological and regulatory costs of bringing new insecticides to market (Sparks and Lorsbach, 2017).
The insect nervous system provides targets for a majority of the most commercially successful insecticides, including acetylcholinesterases (organophosphates and carbamates), voltage gated sodium channels (pyrethroids and DDT) and γ-aminobutyric acid (GABA) and Glutamate-gated chloride channels (cyclodienes, avermectins, and phenylpyrazoles) (Ffrench-Constant et al., 2016; Sparks and Nauen, 2015; Sparks et al., 2019b). The nicotinic acetylcholine receptor family (nAChRs) has, over time, been targeted by a number of insecticides of diverse chemical structures including nicotine, nereistoxin analogs, the neonicotinoids, spinosyns, sulfoximines, butenolids, and most recently, the mesoionics (Sparks and Nauen, 2015; Casida, 2018; Nauen et al., 2019; Matsuda et al., 2020). At present insecticides acting at the nAChR account for approximately 29% of the global insecticide market, rendering the nAChR the most used target site for insect pest control (Sparks et al., 2020).
Insect nAChRs are members of the cys-loop ligand-gated ion-channel superfamily. In the model insect Drosophila melanogaster there are 10 receptor subunit genes (seven α-subunits and three β-subunits) (Sattelle et al., 2005). The insect nAChR is a pentameric assembly of these subunits, expressed mainly in the central nervous systems of insects (Sattelle and Breer, 1990). Thus, many possible combinations of these subunits could exist producing an array of nAChR subtypes, each potentially having different physiological properties. The endogenous ligand, acetylcholine (ACh), binds at the interface between two receptor subunits. The principal face of the binding pocket for ACh is formed by loops A-C of an α-subunit, while the complementary face is formed from loops D-F of the adjacent α or β subunit (Corringer et al., 2000). This subunit interface is also believed to be the primary site of binding for the neonicotinoid and sulfoximine insecticide classes (Ihara et al., 2015; Matsuda et al., 2005; Wang et al., 2016). In contrast other nAChR acting insecticides, the spinosyns, appear to act at an allosteric site (Geng et al., 2013; Puinean et al., 2013; Silva et al., 2016; Somers et al., 2015; Watson et al., 2010).
The Insecticide Resistance Action Committee (IRAC) (Nauen et al., 2019; Sparks and Nauen, 2015) classifies insecticides according to their mode of action. Compounds are also allocated to distinct subgroups based on their chemical structure and information demonstrating differential metabolism and reduced resistance from testing of resistant insect strains. Whereas the insecticides imidacloprid (IMI), nitenpyram (NIT) and sulfoxaflor (SFX) are categorized as nAChR competitive modulators; the neonicotinoids (including IMI & NIT) are both in the same subgroup (4A) (Nauen et al., 2019; Sparks and Nauen, 2015), while the sulfoximine, SFX, is a different subgroup (4C) (Sparks and Nauen 2015) due to differences in chemistry and metabolic resistance (Sparks et al., 2013; Watson et al., 2011, 2017). The spinosyns, (including spinosad (SPIN)) are nAChR allosteric modulator – site I compounds and are thus in an entirely different group (Group 5) (Sparks and Nauen, 2015).
Given the contributions of two different subunits to a single ligand binding pocket and that five interfaces are present in a pentameric receptor, it is important to determine which particular nAChR subunits are present and their arrangement. Heterologous expression studies have provided some insights into the interaction of insecticides at specific residues/possible receptor interfaces when insect nAChR subunits are co-expressed with vertebrate β2 or β4 subunits (Lansdell et al., 2008; Lansdell and Millar, 2000a, b; Matsuda et al., 1998; Matsuda et al., 2005) or with an accessory protein from Caenorhabditis elegans, RIC3 (Lansdell et al., 2012; Watson et al., 2010). Until recently, difficulties expressing insect nAChRs limited investigation via these methods (Lansdell et al., 2012; Watson et al., 2010); however co-expression of additional cofactors has led to robust functional expression of several combinations of nAChR subunits from D. melanogaster, Apis mellifera and Bombus terrestris in Xenopus laevis oocytes (Ihara et al., 2020).
Genetic studies offer another line of investigation into the mode of action of insecticides to improve our understanding of the roles and functions of the different nAChR subunits. For example, in D. melanogaster, loss of function mutants were used to show that the Dα1 and Dβ2 subunits are involved in binding of neonicotinoid insecticides including IMI and NIT (Perry et al., 2008). The modest levels of resistance measured for these mutants, singly and in combination, suggested that receptors comprised of additional subunits are targeted by neonicotinoids (Perry et al., 2008). Because the Dα1 and Dβ2 mutants were not highly resistant to SFX or SPIN insecticides (Perry et al., 2012), this suggested that the neonicotinoids, sulfoximines and spinosyns target different nAChR subtypes. In contrast to the neonicotinoids, the Dα6 subunit has been established as the main target of the spinosyns in Drosophila (Perry et al., 2007; Watson et al., 2010). Mutations in Dα6 or its orthologues are associated with high levels of resistance in Drosophila (Crouse et al., 2018; Perry et al., 2007; Sparks et al., 2019a; Watson et al., 2010) and a wide range of pest insects (Bao and Xu, 2011; Baxter et al., 2010; Geng et al., 2013; Puinean et al., 2013; Silva et al., 2016). One caveat to the findings of previous studies is that only a limited set of receptor mutations known to confer resistance were identified and tested, making it unclear as to whether other subunits play a role, either major or minor. The advent of the capability to specifically manipulate genomes at will provides a new avenue to understand the target site of insecticides and their respective resistance mechanisms (Perry and Batterham, 2018; Somers et al., 2015; Zimmer et al., 2016), and hence options for insecticide resistance management (IRM) programs.
Herein we report our findings from the first systematic investigation of the potential involvement of Drosophila melanogaster nAChR subunits in the mode of action of these three insecticide classes (neonicotinoids, sulfoximines, spinosyns). We took advantage of gene editing techniques using Clustered regularly Interspaced Palindromic Repeats (CRISPR) and the CRISPR Associated Protein 9 (Cas9) nuclease to disrupt the function of nine of the ten individual nAChR subunit genes in D. melanogaster. Our findings help to refine the list of subunits that are involved in the insecticidal activity of the tested nAChR-acting compounds and the level of resistance conferred from loss of function. They also provide insights into the potential combinations of subunits that may co-assemble into native insect nAChR subtypes, an area highly relevant to the understanding of insecticide mode of action and resistance as well as insect neurophysiology. A detailed understanding of an insecticide's mode of action is an important component for the development of effective IRM strategies (Nauen et al., 2019; Sparks and Nauen, 2015), especially those involving the rotation of groups possessing a different mode of action, in an effort to reduce selection for resistance and cross-resistance (Roush, 1989).
Section snippets
Compounds
Spinosad (Success®; Dow AgroSciences), imidacloprid 99% (Pestanal®; Sigma-Aldrich), nitenpyram 99% (Pestanal®; Sigma-Aldrich) were purchased commercially. Sulfoxaflor (99%, racemic mixture) was synthesised and provided by Dow AgroSciences (now Corteva Agriscience).
Toxicology bioassays
D. melanogaster larval bioassays were performed as described previously (Perry et al., 2012). Briefly, 1st instar larvae were collected and reared on semolina-based fly media containing selected doses of the insecticides. Control
nAChR subunit deletions created using CRISPR/Cas9
Deletions of varying lengths were isolated for nine nAChR subunit genes using a variety of CRISPR/Cas9 methods (Fig. 1). Flies homozygous for a deletion of Dα2, Dα3 or Dβ3 were viable. Consistent with published data, the Dα1, Dβ2, Dα4, Dα6 and Dα7 mutants were also viable (Fayyazuddin et al., 2006; Perry et al., 2007, 2008; Shi et al., 2014). With the exception of Dβ1, the mean 1st instar to adult viability of these mutants on untreated media ranged from 84% to 112.8% when compared to the
Germline and somatic CRISPR/Cas9 gene editing can be used to examine insecticide targets
The capacity of CRISPR/Cas9 methods to specifically target and manipulate genes to validate their role in insecticide resistance generated in the lab (Somers et al., 2015), or identified in the field (Zimmer et al., 2016), is clear. Here, we used CRISPR/Cas9 to systematically create mutations in nAChR subunit genes to examine the contribution of individual gene family members to resistance phenotypes. Microinjection of the CRISPR/Cas9 components in combination with transgenic approaches (Gratz
Declarations of competing interest
T. Perry and P. Batterham received the sulfoxaflor compound and funding assistance towards sulfoxaflor bioassays from Dow AgroSciences. T. C. Sparks currently acts as a consultant for the agrochemical industry including Corteva Agriscience.
Acknowledgements
We thank Mr. Gerald Watson and Dr. Melissa Siebert (Corteva Agriscience) for useful discussions and feedback during the preparation of this article. Dow AgroSciences synthesised sulfoxaflor and provided funding assistance for sulfoxaflor studies. Fly strains were sourced from the Bloomington Drosophila Stock Centre and the Australian Drosophila Biomedical Research Support Facility provided quarantine facilities for imported fly strains. Funding for this research was provided through an
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