**2. Insecticide resistance mechanisms**

**1. Introduction**

244 Insecticides Resistance

**1.1. Insecticides and mode of action**

Insecticides are traditionally employed in several human activities with the purpose of eliminating or controlling the density of undesired insect populations. At present, albeit the obvious environmental impact, the control of agricultural pests and disease vectors is still largely based on the use of those substances. Moreover, in several cases, chemical compounds represent the principal approach to interrupt the transmission of pathogens. Before the Second World War, most insecticides were constituted of inorganic compounds, and a few organic substances, such as nicotine, pyrethrin, and rotenone [1]. The modern era of organic insecti‐ cides began in the 1940s, a period known as the age of the "pesticide revolution", when DDT

Currently, there are 25 groups of insecticides and acaricides based on available evidence about their target sites and mode of action, according to the Insecticide Resistance Action Committee (IRAC) [3]. The World Health Organization Pesticide Scheme (WHOPES) promotes and coordinates the testing and evaluation of pesticides for public health purposes, since 1960. Its recommendations are generally adopted for national campaigns in several countries. The main insecticide classes used for vector control are: organochlorine (OC), organophosphates (OP), carbamates (CA), pyrethroids (PY), insect growth regulators (IGR), spinosyns (SP), and toxins derived from bacteria (*Bacillus thuringiensis* var. *israelensis* and *Bacillus sphaericus*) [4, 5]. The classes OC, OP, CA, PY, and SP include a broad range of compounds that act on the insect

The target site of OP and CA is the acetylcholinesterase (AChE), a conserved enzyme present in a wide variety of animals, including mammals, birds, reptiles, fish, and insects. This enzyme is responsible for the rapid hydrolytic degradation of the acetylcholine neurotransmitter at synapses, causing momentary interruption of the nerve impulse. The OP and CA insecticides bind in the AChE active site, compromising the acetylcholine hydrolysis and then accumulat‐

The PY and OC (DDT and analogues) maintain the sodium channels in their opened confor‐ mation, generating a continuous influx of ions throughout the axons. Cyclodienes, another group of OC insecticides, act directly on the gamma-aminobutyric acid receptor (GABA), preventing the normal input of chlorine ions in the neurons, just after the nervous impulse. In all cases, regardless of the target site, OP, OC, and PY promote a continuous nerve impulse

Unlike neurotoxic insecticides, IGRs do not induce an immediate death of the insects. How‐ ever, they are toxic mainly against immature stages, affecting the moulting, metamorphosis processes, besides commitments in viability and reproduction of adults [7]. Based on the mode of action, the IGRs are classified into three major categories: (i) juvenile hormone mimics; (ii)

Concerning the bacterial toxins, *Bacillus thuringiensis* (*Bs*) var. *israelensis* (*Bti*) and *B. sphaeri‐ cus* are the most employed as insecticides. When ingested by larvae, the *Bt* toxins are activated

ing the neurotransmitter at the synapses, causing repetitive nerve impulses.

transmission that culminates in paralysis, convulsions, and death [6].

ecdysone agonists; and (iii) chitin synthesis inhibitors [8].

(dichlorodiphenyltrichloroethane) was used for the first time as an insecticide [2].

central nervous system and, thus, have an immediate effect.

Insecticide resistance is considered the major challenge for control programs involving the use of chemicals. Up to 2014, populations of at least 590 species of insects were diagnosed as resistant to insecticides. Resistance to around 300 compounds, including the neurotoxics (OC, OP, CA, PY, and SP), the IGRs, and *Bt* toxins, has already been registered to one or more insect species [3].

Insecticide resistance has a genetic basis. Randomly arisen mutations can prompt several alterations in aspects of behavior, metabolism, and physiology of the insects, which may gain adaptive advantages in an insecticide-treated environment. Such alterations can be classified as: (i) behavioral changes; (ii) altered penetration (increased production of cuticular compo‐ nents that reduces intake of insecticide); (iii) target site modification; and (iv) metabolic resistance (detoxification enzymes and ABC transporters) [10]. Although evidenced, the two first aspects are less reported, whilst several studies have described and evaluated the target site and metabolic resistance mechanisms. These two, alone or combined, potentially induce a wide range of resistance levels to virtually all available insecticides [11].

Most insecticides target a single protein in the insect organism. The interaction between these molecules disrupts a normal biological process, leading to the toxicant effects. However, some mutations that induce structural alterations in the target protein can change the insecticide levels of toxicity. Moreover, most of these alterations are conserved among distinct insect orders. For instance, cyclodienes inhibit chloride ion transport by keeping the gama-amino‐ butyric acid (GABA) receptor in a close conformation [12]. The replacement of an alanine to a serine or glycine at the aminoacid position 302 (A302S/G) in the GABA, generally referred to as *rdl* mutations (resistance to dieldrin), confer resistance in several species, such as *Drosophila melanogaster*, *Musca domestica*, *Hametobia irritans, Lucilia cuprina, Tribolium castaneum, Peripla‐ neta americana*, and *Anopheles mosquitos* [13].

The glycine-to-serine substitution (G119S)2 in the AChE (AChE-1, encoded by the *ace-1* gene) confers resistance to OP and CA in *Anopheles* and *Culex* mosquitoes. Interestingly, this mutation was never found in *Aedes* mosquitoes, regardless of the intense use of OP against their populations. The most accepted hypothesis for this relies on the fact that the AChE-1 119 glycine is encoded by a GGA, differently from the GGC in other species. It means that in other mosquitoes a serine substitution (AGC) requires only one nucleotide change. By contrast, two

<sup>1</sup> A complete review about insecticides and their mode of action can be found at Sparks and Nauen (2015).

<sup>2</sup> This denomination refers to the aminoacid in the position 119 of the AChE protein (AChE1), based on the Torpedo nomenclature (Toutant, 1989).

concomitantly selected mutations would be necessary in *Aedes* mosquitoes, an unlikely situation referred to as codon constraint [14].

Similarly, several mutations associated with PY and DDT resistance are present in distinct insect orders: the *kdr* mutations, that impair the *knockdown effect* provoked by those insecticides. The most common *kdr* (*knockdown resistance*) mutation is a leucine-to-phenylalanine substitu‐ tion in the 1014 codon3 , although serine, histidine, cysteine, and tryptophan replacements are also found (reviews presented in Rinkevich et al., 2013 [15]). Several PY-resistant populations of major arthropod pests and disease vectors were found harboring *kdr* mutations. In this sense, for diagnostic purposes, different well-established tools for *kdr* genotyping have been imple‐ mented, specific for an increasing number of insect species. This allows a rapid and accurate access of the genetic background for PY resistance in natural populations [16].

The recent commercially introduced SP insecticides, which target the nicotinic acetylcholine receptors (nAChRs) [17], have been used for crop protection, animal health, and against human disease vectors. Three formulations of SP were approved by WHOPES for use in drinking water, increasing the chemical arsenal against mosquitoes [4]. However, resistance to this class of insecticides was already detected in a variety of insect species. A target-site point mutation (glycine-to-glutamate substitution G275E), for example, was identified in the nAChR of a Western flower thrips (*Frankliniella occidentalis*) in association with SP resistance [18]. Besides this single amino acid substitution, alternative splicing in the nAChRα 6 subunit seemed to be the mechanism selected in an SP-resistant population of the diamondback moth *Plutella xylostella* [19].

As exemplified above, mutations selected for resistance in the molecular targets of insecticides generally share homologous sites among different insects. These molecules are components of the nervous system, which are highly conserved among animals. Therefore, it is expected that few mutations can be maintained without impairing the essential physiological role of that molecule [20]. Target-site-resistant alleles are increasing in frequency and rapidly spreading, as well-recorded for malaria and dengue vectors. An interactive compilation of these data, organized in time and space scales, can be currently accessed on two distinct online platforms: IR Mapper (http://www.irmapper.com) and Popbio (https://www.vectorbase.org/ popbio/).

Detoxifying enzymes are naturally present in living organisms with a protective function against potential damages caused by xenobiotics and endogenous metabolites. In many cases, insecticide resistance occurs due to an increased activity of such enzymes, a mechanism known as metabolic resistance. In general, this mechanism is related with the intense use of insecti‐ cides. However, other toxic compounds, such as chemical pollutants and plant toxins can also select for metabolic resistance mechanisms in insect populations. In this sense, different xenobiotics present in the environment are probably related, at least in part, with a preadap‐ tation for insecticide resistance in disease vector and agricultural pests [21, 22]. Basically, xenobiotics pass through a series of enzymatic steps that transform them in polar substances,

<sup>3</sup> In the case of the voltage gated sodium channel (NaV), the *M domestica* aminoacid sequence is most commonly taken as reference.

soluble in water for an easier excretion [23]. The biotransformation is divided into three phases, with the participation of three main groups of enzymes. Phase I includes multiple function oxidases enzymes (MFO or P450) that carry out chemical modifications of a broad variety of xenobiotics. In phase II, glutathione S-transferases (GST) usually conduct conjugation reac‐ tions in the products resulting from the previous phase. The esterases (EST) can participate in both phase I and II, hydrolyzing ester bonds present in the xenobiotics. Finally, during phase III, the metabolites produced in the two first phases are actively exported out of the cells via ATP-binding cassette (ABC) transporters [24-27].

The metabolic resistance mechanisms are characterized by a gain in the ability for detoxifying molecules of insecticides, preventing them from reaching their targets. This acquisition can be selected by either an increase in the enzymatic activity over the insecticide (mutations that improve the detoxifying power) or an augment in the amount of copies of a specific enzyme (due to an increase in the transcription rate, for instance). Glutathione S-transferases, EST and MFO P450 enzymes are each comprised of tens of genes, composing supergene families, possibly resulting from duplication events along the evolutionary process, as well as inde‐ pendent gene duplications inside distinct species [28, 29]. Differently from target site mutations that can arise in homologous sites among different insect groups, several detoxifying genes are unique for some species and may be selected for insecticide resistance in a particular way.

The main questions that lie upon the molecular basis of insecticide resistance mechanisms are how many (and which) genes control the phenotype of resistance, how many mutations were selected within that gene(s), and if they are just spreading from one origin or appearing multiple times [30]. The advent of high-throughput screening molecular tools expands the searches for selected resistance mechanisms and their overall effects, toward beyond the target site mechanisms. Recent advances have revealed the complexity of metabolic systems enrolled in insecticide resistance at transcriptomic and genomic levels. Comparisons of the whole transcriptional profile between susceptible and resistant individuals generally indicate the participation of several genes in the physiological process of resistance [31-33]. In addition, genetic loci influencing the resistance can be physically mapped in the chromosomes through quantitative trait loci (QTL) approaches [34-36]. Likewise, a recent study identified several single nucleotide polymorphisms (SNPs), as well as an important and previously neglected copy number variation (CNV) related to insecticide resistance in *Aedes aegypti*, by combining genomic target enrichment with next-generation sequencing technologies [37].
