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Rants’o, T.A.;  Koekemoer, L.L.;  Panayides, J.;  Zyl, R.L.V. Essential Oil-Based Anticholinesterase Insecticides against Anopheles Vectors. Encyclopedia. Available online: (accessed on 24 June 2024).
Rants’o TA,  Koekemoer LL,  Panayides J,  Zyl RLV. Essential Oil-Based Anticholinesterase Insecticides against Anopheles Vectors. Encyclopedia. Available at: Accessed June 24, 2024.
Rants’o, Thankhoe A., Lizette L. Koekemoer, Jenny-Lee Panayides, Robyn L. Van Zyl. "Essential Oil-Based Anticholinesterase Insecticides against Anopheles Vectors" Encyclopedia, (accessed June 24, 2024).
Rants’o, T.A.,  Koekemoer, L.L.,  Panayides, J., & Zyl, R.L.V. (2022, October 28). Essential Oil-Based Anticholinesterase Insecticides against Anopheles Vectors. In Encyclopedia.
Rants’o, Thankhoe A., et al. "Essential Oil-Based Anticholinesterase Insecticides against Anopheles Vectors." Encyclopedia. Web. 28 October, 2022.
Essential Oil-Based Anticholinesterase Insecticides against Anopheles Vectors

The insect's nervous system is critical for its functionality. The cholinergic system, of which acetylcholinesterase (AChE) is a key enzyme, is essential to the Anopheles (consisting of major malaria vector species) nervous system. Furthermore, the nervous system is also the primary target site for insecticides used in malaria vector control programs. Insecticides, incorporated in insecticide-treated nets and used for indoor residual spraying, are a core intervention employed in malaria vector control. 

malaria insecticides terpenoids acetylcholinesterase

1. Introduction

Malaria is a devastating disease caused by a protozoan parasite, namely Plasmodium falciparum which is the major causative agent in the pathogenesis of this infectious disease[1][2]. Anopheles vectors are infected with malaria after they ingest blood from an infected human host. The female Anopheles vectors effectively bite the human hosts between dusk and dawn[3] and it is during this time that she ingests gametocytes. The Plasmodium gametocytes develop into an oocyst in the mosquito midgut, which then matures into sporozoites. The sporozoites are released into the hemolymph and migrate to the salivary glands[1][4]. This parasite developmental process within the vector takes approximately 11–16 days before the female mosquito is able to transmit the parasite to the next human host during a blood feeding. This means that a long lifespan of the Anopheles vector is required for the successful completion of the parasite development and reinfection of the human host. Vertebrate blood is needed every 2–3 days by the female mosquito for nutrition, as well as egg development. The eggs are oviposited into water and fertile eggs hatch into larvae a few days later. Larvae will develop into pupae and finally adults will emerge after a few days[3]. There are more than 400 Anopheles species of which about 30 are major malaria vectors. The African Anopheles vectors have both long lifespans and a higher preference for human feeding and, collectively, these account for the high malaria cases and mortality that is recorded in Africa[3][5]. Other factors, such as climate conditions and political and economic stability, also affect the intensity of transmission and enhance the problem[3].
The malaria vectors have long been controlled by using insecticides. Insecticide classes include organophosphates, carbamates, pyrethroids, organochlorines and neonicotinoids[6]. Larvicides including insect growth inhibitors as well as bacterial larvicides, such as Bacillus thuringiensis subspecies israelensis, Bacillus sphaericus and spinosyns from Saccharopolyspora species, have also gained popularity in mosquito control activities[6][7][8]. The implementation of large-scale larviciding is however challenging in Sub-Saharan Africa and these may be used as a complementary intervention[6][9][10][11]. The Anopheles vectors have developed substantial resistance against almost all current insecticides[12][13][14][15]. To compound the issue, the commercial development of insecticides through various and often complicated synthetic mechanisms is expensive and time-consuming[16][17]. The researchers propose that the identification of potential insecticides from natural product resources, such as essential oils (EOs), is a relatively cost-effective and faster alternative. Target identification and the corresponding mechanism of action are critical components of the drug discovery process[18]. Acetylcholinesterase (AChE) is a validated target in the insect nervous system and inhibitors of this critical enzyme have been useful in the control of malaria vectors for over eighty years[6][19][20].

2. Malaria Vector Control

The early vector control strategies adopted vast activities to reduce larval populations, which included, amongst others, the drainage of breeding sites such as swamps or the application of copper (II) acetoarsenite (Paris green), a highly toxic inorganic compound to the breeding sites[21][22]. In addition, the screening of windows and doors to prevent vectors entering houses and the use of mosquito nets have been at the forefront in protecting people against mosquito bites[6]. The plant-based insecticides were the first preparations used historically. Pyrethrins extracted from the flowers of Chrysanthemum cinerariifolium and Chrysanthemum roseum were used against indoor Anopheles mosquitoes in the 19th century[23][24]. However, the structural modifications of the natural pyrethrins and the generation of first synthetic pyrethroids were first reported in the period 1924 to 1970[25]. The discovery of an organochloride, namely, dichlorodiphenyltrichloroethane (DDT), was reported in 1939[25][26]. DDT has been highly effective against malaria vectors. However, in recent times increasing safety concerns have seen it being replaced in many countries by newer insecticides with reduced toxicity profiles[27][28][29].
Currently, malaria vector control adopts an integrated vector management program through the use of insecticides targeting both the larval and adult stages[6][27]. This is achieved through two main interventions, namely, insecticide-treated mosquito nets and indoor residual spraying, with additional interventions including larviciding[6]. The insecticide-treated nets provide both a physical barrier and insecticidal activity against Anopheles vectors. Indoor residual spraying (IRS) on the other hand, provides host protection through the Anopheles insecticidal effect[3]. Pyrethroids, pyrethroid-PBO combinations and pyrroles are the only insecticide classes used for the insecticide-treated nets, as the latter insecticides pose a low toxicity risk to humans[30]. The pyrethroids used in IRS include deltamethrin, alpha-cypermethrin, etofenprox, lambda-cyhalothrin, bifenthrin and cyfluthrin, while the organochlorines include DDT. On the other hand, the organophosphates approved by WHO for IRS include malathion, fenitrothion, pirimiphos-methyl and the carbamates such as propoxur and bendiocarb[6][31][32][33].

2.1. Insecticide Resistance in Main African Malaria Vectors

Insecticide resistance has been reported in all of the main African malaria vectors and this resistance against WHO approved insecticidal agents is rapidly increasing in intensity and geographical distribution[5][34]. An overview of mosquito resistance has been highlighted below. To keep this brief, only a few examples will be provided to explain the extent of the problem mainly on the African continent. Insecticide resistance in the main vector species has been reported for pyrethroids[34][35][36][37], organochlorides[36][38][39][40], organophosphates[12][41] and carbamates[20][36][40][42].
Common insecticide resistance markers associated with pyrethroid and organochloride resistance include the L1014F and L1014S mutation of the voltage-gated sodium channel gene, known as knockdown resistance (kdr). These mutations shift activation voltage dependence of sodium channels stabilizing them in the closed state. This antagonizes the action of pyrethroids and organochlorines since these compounds bind to open sodium channels[25][38][43]. Apart from the kdr mutations, elevated metabolic enzymes, including P450 monooxygenases, glutathione-S-transferases and non-specific esterases, also convey high resistance to pyrethroids and organochlorides[13][37][44][41]. On the other hand, the resistance mechanism commonly conferring organophosphate and carbamate resistance is a single point polymorphism resulting from glycine conversion to a serine residue at position 119 (G119S; Torpedo californica AChE numbering) or more precisely, position 280 (G280S; Anopheles gambiae AChE numbering) in the AChE target[38][43][45][46]. Resistance mechanisms often prevent the intended biological activity of a specific insecticide.

2.2. Modes of Action of Main Insecticides Used in Malaria Vector Control

Regardless of their small size, insects have a high surface area for the penetration and subsequent systemic distribution of an insecticide from contact exposure. Furthermore, the small size generates short pathways to the insect’s nervous system and as a result, most insecticides act on the insect’s nervous system[47]. Organochlorines act specifically on the peripheral nervous system, where they bind and stabilize the open voltage-gated sodium channels[25]. The stabilized open state of the sodium channels allows for continuous sodium influx and prolonged action potentials leading to spontaneous neuronal firings succeeded by muscle twitches and sustained body tremors[48]. In contrast to the organochlorines, pyrethroids act on both the peripheral and central nervous systems; however, they act in a similar manner to prevent the closing of the voltage-gated sodium channels, resulting in continuous neuronal discharges followed by paralysis[48]. Similarly, the organophosphates and carbamates also exert their effects on the central nervous system. However, these insecticide classes inhibit acetylcholinesterase, a principal enzyme in the insect nervous system, which leads to an increase in the neurotransmitter ACh levels in the synapse. This leads to enhanced ACh effects on the cholinergic receptors resulting in constant neurotransmission and neuronal hyperexcitation[49]. On the other hand, the neonicotinoids enhance cholinergic activity by acting as agonists on nicotinic acetylcholine receptors (nAChRs). Similarly, this disrupts neuronal transmission in the insect nervous system, causing paralysis and subsequent insect death[50].

2.3. Insect Nervous System

The insect nervous system is composed of central, visceral and peripheral nervous systems[51]. The insect central nervous system (CNS) is composed of the ventral nerve cord and brain connected to various ganglia including supra- and sub-esophageal ganglia, thoracic ganglia and abdominal ganglia (Figure 1). The sub-esophageal ganglion transmits impulses to the mouthparts and salivary glands. The insect brain is composed of three cephalic neuromeres, including the protocerebrum, deutocerebrum and tritocerebrum. The deutocerebrum carries out olfactory and sensory functions through the antennae; where the olfactory signal transduction is important in host identification and interaction by the insect. The tritocerebrum nerves innervate the ventral nerve cord and internal organs including the anterior digestive canal[51][52][53]. The insect’s peripheral nervous system, commonly referred to as a stomatogastric nervous system is composed of the peripheral ganglia complex and nerves that innervate visceral organs. This system mainly controls food intake and digestion. Generally, the insect CNS ganglia receive sensory impulses from the appendages and body cuticle after which the efferent signals are sent to the body muscles, internal organs and genitalia[54][55]. The protocerebrum controls the insect’s vision through compound eyes and ocelli. Most importantly, neurosecretory cells are located in the protocerebrum[55][56]. Most insecticides including the organophosphates and carbamates affect neurotransmitter secretion and action[56][57].
Figure 1. The nervous system ganglia of Anopheles[51].

2.4. Molecular Characterization of Acetylcholinesterase

There are apparent AChE structural differences between insects and mammals. These span from their distinct genomics, amino acid sequences to their active and peripheral anionic site conformations[58]. Recent biochemical studies have revealed critical differences between the Anopheles AChE and human AChE that could serve as potential drug targets for directed insecticide design.
The amino acid sequence of Anopheles AChE is reported to be 48–49% identical to that of the human AChE[59][60]. Unlike humans where there is a single ace gene coding for AChE, mosquitoes have two ace genes, ace-1 and ace-2, coding for AChE1 and AChE2 enzymes, respectively[61][62]. These genes are crucial in all life stages of the mosquito, ranging from egg through to adult stages[63]. AChE1 is the main catalytic enzyme, while AChE2 is involved in non-catalytic activities such as reproduction. As a result, target site insensitivity on insect AChEs, such as G280S genotype, is linked to mutations in ace-1 but not ace-2[46][61][57]. AChE is characterized by a deep and narrow active-site gorge (Figure 2). There are differences in these gorge structures between Anopheles and human AChEs and this may affect ligand binding and specificity[46][59][64]. Notably, a free cysteine residue (Cys447) is available at the entrance to the active site gorge of Anopheles AChE (Figure 2A,B), but not in human AChE. Instead, a human AChE has a bulky phenylalanine (Phe295) at the active site entrance (Figure 2C). Additionally, in Anopheles AChE, a smaller aspartic acid residue (Asp602; Figure 2A,B) replaces a larger tyrosine residue (Tyr449) at the base of the active site gorge[46]. Moreover, a conserved arginine residue (Arg339; not shown in order to maintain the catalytic side resolution) has also been identified in Anopheles AChE[65]. In addition, the displayed An. gambiae AChE catalytic site in Figure 2B shows a G280S mutated site (pointed).
Figure 2. Molecular comparison of An. gambiae wild-type (A) and resistant (B) AChE catalytic sites (PDB IDs: 5YDI and 6ARY, respectively) to the human AChE (PDB ID: 7E3H) (C). generated by Schrodinger’s Maestro 2018-2 software (New York, NY, USA). The G280S mutation is shown (red arrow) in the resistant Anopheles AChE phenotype (B)[46].

2.5 Acetylcholinesterase Inhibition in Anopheles

The catalytic site in Anopheles is characterized with a catalytic triad made of His-Ser-Glu (His600-Ser360-Glu359; Figure 3A) amino acid combination. The catalytic serine (Ser360;
Figure 2A) is the target for covalent insecticides, including organophosphates and carbamates[56][57]. These insecticides establish a covalent bond with AChE through phosphorylation and carbamoylation, respectively[66]. The Anopheles resistance to the anticholinesterase insecticide classes is usually caused by ace-1 G280S mutation (Figure 3B) and metabolic resistance resulting from the elevated levels of monooxygenases, glutathione S-transferases and general esterases[57][67][68][69]. Given the widespread resistance that has largely rendered organophosphates and carbamates noneffective, there is an urgent need to identify novel anticholinesterase insecticides. AChE has proven to be a valid target in Anopheles vectors[59] and EOs have also shown to be the promising sources of novel insecticides[70][71][72]


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