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BIO Magazine - Mechanisms of insecticide resistance Δεκέμβριος 2015
Δεκέμβριος 2015 No38

BIO Environment

Mechanisms of insecticide resistance
Mechanisms of insecticide resistance


Since the beginning of the miraculous insecticide era, around 1930, efforts focused on the production of species specific compounds with the least possible environmental impact. However, the widespread and persistent use of any insecticide will inevitably lead to the development of resistance. The vast majority of modern insecticides target the insect’s nervous system, either at the level of the cell’s axon or at the synapse. Most resistance mechanisms concern either the alteration of insecticide’s target or the increased metabolic detoxification and sequestration of the toxicant. At the molecular level this is achieved by: point mutations in the GABA receptor (cyclodiene insecticides); point mutations in the acetylcholinesterase active site (organophosphates and carbamates); amplification or up-regulation of esterase genes (organophosphates and carbamates); mutations in the sodium channel gene (DDT and pyrethroids); mutations leading to the up-regulation of detoxification enzymes such as cytochromoe P450 and glutathione S-transferase (many classes of insecticides). Nonetheless, molecular analysis of highly resistant insects still surprise us disclosing new mutations that indicate novel insecticide resistance mechanisms, such as the example of the olive fly. 


Pest Control

Humans have been battling insect and other arthropod pests since the beginning of agriculture, about 6000 B.C. The first recorded instance of pest control takes us back to 2500 B.C. when Sumerians used sulfur to control insects, a method still used to our days. There are also reports from around 500 B.C. of Chinese using mercury and arsenic compounds as a means to control body lice. Pest control methods improved throughout the years, including both physical or chemical agents (such as soap, tobacco or other herb infusions, coal tar, petroleum, as well as arsenic) and biological ways (e.g., ants to suppress caterpillars). 

However, it was not until the 1930s that we entered the miraculous era of chemical insecticides, which gave the hope that several serious insect pests, particularly those of public health concern (such as the devastating malaria mosquitoes), could be completely obliterated from the face of the earth. The discovery of the insecticidal properties of DDT (dichlorodiphenyltrichloroethane) marked the beginning of this era. DDT’s toxicity (particularly against mosquitoes, fleas and lice), its low production cost and its characteristic stability and long persistence gave the illusion that it could solve many of humanity’s major public health problems, such as malaria and typhus, by completely exterminating their insect vectors. In addition to its importance in medicine, DDT was used widely in agriculture. However, its characteristic stability and fat solubility resulted in the increased accumulation of active, non-metabolized pesticide in the higher levels of the trophic pyramid. At the same time, the resulted overuse, particularly in agriculture, lead to the development of insecticide resistance: more and higher doses of the pesticide were necessary for effective insect control, resulting in higher accumulation in the environment. DDT’s use was finally banned in the US in 1973, although it is still used in some parts of the world. Its place took several other “safer” insecticides that were already developed by one of the world’s largest chemical industries (Figure 1). Nonetheless, as long as they were also non-prudently overused, they also lead to the development of resistance (IRAC Organization: www.irac-online.org; Pedigo, 2002). 


Insecticide resistance

Resistance is the ability of certain individuals to tolerate a dose of toxicant that would be lethal to the majority of individuals in a normal (susceptible) population. Resistance generally evolves through the selection of resistance traits that pre-exist naturally in pest populations. It is a pre-adaptive phenomenon and not a mutational effect. 

When a pest population is exposed to an insecticide, some individuals manage to survive due to resistance trait(s) that they genetically predispose. A proportion of these rare individuals will survive after higher doses of insecticide applications. The offspring of the survivors will carry the genetic makeup of their parents and many of these will inherit the resistance trait(s). The part of the resistant individuals will become greater with each succeeding generation of the population and, as a result, in a short period of time the entire pest population will be able to survive the exposure to the insecticide (Figure 2). The development rate of insecticide resistance is a consequence of a number of factors, including reproduction rate (number of generations per year), genetic variability and gene flow in the wild population (existence and spreading out of the resistant trait), migration (introduction of susceptible individuals) and pest management strategies (rate, timing and number of insecticide applications). Moreover, phenomena of multiple and cross-resistance are potentially responsible for the quick resistance development. Multiple-resistance is the resistance to more than one chemical class of pesticides, while at cross-resistance the trait that confers resistance to one pesticide also confers resistance to other pesticides (IRAC).

At present, insecticide resistance is extremely widespread. Around 500 species of arthropods have been recorded resistant to at least one class of pesticides. About 40% of these species are pests of medical or veterinary importance, seriously complicating public health measures. The remaining 60% affects agricultural pests, causing extensive economical damage (Georgiou 1990).


Mechanisms of resistance

In general, there are three potential types of resistance that insects have evolved in order to decrease their susceptibility to the main classes of insecticides (see Box 1) (IRAC): 

  • Behavioral resistance involves a change in the behavior of the insect such that contact with the insect control product is avoided. The resistance to DDT of an Anopheles gambiae sub-species is an example of behavioral resistance. The malaria control was based on the indoor wall spraying with DDT. As a result, the endophilic (indoor) subpopulation that was susceptible to DDT was substituted by the exophilic (outdoor) subpopulation, which became dominant.
  • Biochemical – metabolic resistance is a result of enhanced metabolism. This is due to an increase in either the quantity or efficiency of an enzyme that breaks down the insect control product. Insects with metabolic resistance will be cross-resistant to other insect control products metabolized by the same enzyme.    
  • In physiological resistance the insect alters its vital physiological functions in order to survive the exposure to a particular insecticide via different ways such as:
    • Altered target-site resistance. The structure of the insecticide target is modified due to alterations of amino acids in a way that affects insecticide sensitivity. This type of mechanism usually confers cross-resistance to all insecticides of the same chemical class as well as other insecticides acting at the same site. Examples include knockdown resistance, which confers resistance to both DDT and pyrethroids, and dieldrin resistance, which confers cross-resistance to lindane, fipronil and all cylclodienes (Bloomquist, 1996). 
    • Decreased penetration. In resistant insects, the cuticle is altered structurally or alters its composition, thus reducing the absorption of the insecticide.  


Insecticide targets

Most modern, highly effective insect control products act on the insect nervous system (see Box 2). In theory, insecticides could act at many points of the nervous system. In practice, however, only a handful of sites have shown sufficient vulnerability. Insecticides interfere with signal transmission either along the axon or at synapses (Box 2, Figure 3). Signal transmission in the axon is electrical. Signal transmission at the synaptic cleft is relayed by neurotransmitters. Central role in the electrical transmission play the voltage-dependent sodium channels. Central role in the synapse transmission play two proteins, the nicotinic acetylcholine receptor (nAChR) and the acetylcholinesterase (AChE). Such molecules are the major insecticide targets (ffrench-Constant, 1999). 


a. Axonic insecticides

Voltage-gated sodium channels are transmembrane proteins responsible for the rapid rising phase of action potentials and are critical for electrical signaling in most excitable cells (Figure 3B). Voltage-gated channels are triggered by changes in membrane voltage. Inactive at the resting potential, these proteins sense membrane potential changes and open when the potential reaches a threshold value. Sodium ions flow into the cell and their positive charge drives the internal potential positive. At the end of this influx, the sodium channels become temporarily inactivated (halting any further influx) and voltage-gated potassium channels open, allowing K+ ions to flow out of the cell and restore the negativity inside the cell. DDT and pyrethroid insecticides bind to the sodium channel and slow the rate at which it can close or inactivate. These insecticides are very potent because even if only a small fraction of sodium channels (+ influx can generate a second action potential, and eventually one action potential becomes multiplied into many. Responsible for the “sodium channel”-resistance appears to be a single amino acid substitution in a region that forms the channel pore (Dong, 2007). 


b. Synaptic insecticides

An action potential in the pre-synaptic terminal results in the release of the neurotransmitter acetylcholine (ACh) into the synaptic cleft (Figure 3C). ACh molecules diffuse across to the post-synaptic membrane, where they bind to nicotinic ACh receptors (nAChR). When two molecules of ACh bind to a nAChR molecule (which itself is a sodium channel), they open the channel through which sodium ions can enter the post-synaptic cell, leading to depolarization and excitation. The action of ACh in the cleft is terminated by the enzyme acetylcholinesterase (AChE). AChE cleaves ACh into acetic acid and choline, which are transported back into the pre-synaptic cell and recycled. 

The action of ACh on nAChR and its destruction by AChE are targets of insecticides. Imidacloprid and the natural insecticide nicotine mimic the action of ACh on the receptor. Resistance is the result of specific amino acid substitutions at conserved positions of the nAChR. On the other hand, the most important insecticide target of the last 40 years has been the enzyme AChE. Organophosphate and carbamate insecticides bind to the catalytic site of this enzyme and inhibit cleavage of ACh, prolonging the action of ACh on the postsynaptic cell. In this case, resistance arises from a variety of single amino acid replacements, alone or in combination. The presence of these mutations results in decrease affinity of the insecticide to the binding site of AChE (Fournier, 2005).


c. Other neurotransmitters

There are many other neurotransmitters in addition to ACh, each having its own specific synthesis, reception and inactivation mechanisms. Of importance to insect control are the neurotransmitters γ-aminobutyric acid (GABA), glutamate and octopamine. In contrast to ACh receptors, which are sodium channels, inhibitory GABA receptors and glutamate receptors are channels that allow negatively charged chloride ions to flow into the cell. The influx of negative charge inhibits the activity of neurons and counteracts the action of excitatory neurotransmitters such as ACh. Many insecticides (e.g., dieldrin, endosulfan, lindane and fipronil) block the GABA-activated chloride channels, resulting in widespread over-excitation of the nervous system. On the other hand, activation of inhibitory glutamate receptor-linked chloride channels (by avermectins and milbemycins), leads to over-inhibition and paralysis. At least 300 species have been documented to be resistant at these pesticides. The resistance is conferred by a replacement of alanine at position 302 with either a serine or a glycine. This mutation is located in the second membrane-spanning region of the channel subunit, lining the chloride ion pore. The presence of alanine at position 302 is crucial for insecticide binding and the replacement of only this amino acid can cause resistance due to a unique dual effect on insecticide binding (Buckingham et al., 2005).  


d. Detoxification mechanisms

Before an insecticide (or any other foreign substance) reaches its target inside the insect, it first has to overcome the effort of the insect’s detoxifying systems to break it down. This process is termed xenobiotic metabolism and involves three major groups of genes encoding metabolic enzymes, i.e., esterases, glutathione-S-transferases (GSTs) and cytochrome P450 monooxygenases (P450s). Xenobiotic resistance in insects has evolved predominantly by increasing the metabolic capability of detoxifying systems and/or reducing xenobiotic target site sensitivity. In contrast to the limited range of nucleotide changes that lead to target site insensitivity (that was described above), many molecular mechanisms lead to enhancements in xenobiotic metabolism. 


Esterases. They catalyze the addition of a water molecule to an ester, resulting in the formation of the corresponding acid and alcohol. Esterases play an important role in maintaining normal physiology and metabolism, detoxifying a wide range of insecticides. Resistance based on esterase metabolism is due to over-expression of esterase genes either as a result of changes in gene regulation or after gene amplification. Gene regulation may be affected by mutations in upstream regulatory sequences that lead to increased gene transcription or mutations in the processing rate of mRNA, which leads to increased translation. On the other hand, DNA amplification creates additional copies of functional esterase genes. In either way, resistance is the result of a very high titer of esterases and the noncatalytic reaction with, or the sequestering of, insecticides (Hemingway, 2000). Finally, qualitative changes in the structural gene have also been reported (e.g., altered catalytic center activity). A single mutation in the structural gene has been identified that converts the specificity of an esterase to an insecticide hydrolase: the resistant mutants presented a reduced ability to hydrolyse carboxylesterase substrates but an increased ability to hydrolyse OP substrates (Newcomb, 1997). 


GST. Glutathione S-transferases (GSTs) catalyze the conjugation of glutathione with compounds having a reactive electrophilic centre, leading to the formation of water-soluble, less reactive products that are more readily excreted from the insect’s body. GSTs belong to a superfamily of more than one hundred GST-like proteins. Elevated GST activity has been associated with resistance to all the major classes of insecticides. Resistance has been attributed to increases in the amount of one or more GST enzymes through increases in transcriptional rate, rather than qualitative changes in individual enzymes (Enayati et al., 2005).


P450s. The monooxygenase, or mixed function oxidase (MFO), complex involves a reductase and one or more cytochrome P450s and requires NADPH as cofactor. An increase in MFO activity is one of the most versatile mechanisms of resistance in insects, and commonly involves the overtranscription (rather than amplification) of cytochrome P450. Cytochrome P450s belong to a vast superfamily (Scott et al., 1998). It has been shown that overexpression of P450 is conferred by an insertion of a transposable element into cyp6g1, one of the 90 genes in the D. melanogaster genome that encode different cytochrome P450 enzymes (Daborn et al., 2002).


Insecticide resistance in the olive fly

The olive tree is an integral part of the lives of Mediterranean people. Its role is multiple: nutritional, social, cultural and political. Olive oil and table olives are basic agricultural products for most Mediterranean countries and in many mountainous regions they constitute the only source of income. Ninety-eight percent of olive trees grow in the Mediterranean region, 70% of which grow in Spain, Italy, Greece and Turkey. Greece is the third biggest olive oil producer (after Spain and Italy), possessing the 17% of world olive oil and table olive production (FAOSTAT Agricultural data). 

The olive fly, Bactrocera oleae, is the most important pest of the olive fruit. More than 30% annual loss of olive oil production is attributed to the olive fly with corresponding damages in table olives (Mazomenos, 1989). Female insects pierce the skin of the fruit and deposit their eggs in the pulp. The hatched larvae feed on the fruit sap and after a few days the larvae abandon the fruit to pupate in the soil. In addition, egg laying creates a source for bacterial or fungal infection of the fruit, thus exacerbating the damage of the fruit. In either case the fruit is spoiled or it is rendered non-commercial (Figure 4A). 

Control of the olive fly has taken place during the last four decades mostly by organophosphates, either in bait or in cover sprays. In general, fenthion is used early in the season and dimethoate later, closer to the harvest, due to its low residual persistence in olive oil. This heavy OP use against the olive fly inevitably lead to the development of resistance. Early studies in the 1970 indicated that the ability of olive fruit fly to overcome the exposure to OPs was evolved through the biochemical-metabolic mechanism. The liable enzyme was the acetylcholinesterase. It was hypothesized that there was a duplication of the AChE gene and consequently there were more enzyme molecules to break down the chemical compound (Zouros and Krimbas, 1969). Three decades later, it has been shown that the underlying resistance mechanism is the qualitative modification (altered target-site resistance) of the enzyme rather that its quantitative alteration. 

These alterations involve two point mutations in the active site of AChE and a small deletion in the C-terminal domain of the enzyme. 

Two point mutations in the catalytic domain. The isoleucine at position 214 and the glycine at position 488 are replaced by valine and serine, respectively. The two new amino acid residues point to a mechanism that involves the steric alteration of the active site. The neck of the gorge is constricted in such a way that affects the entrance and the binding of the insecticides to the base of the active site (Figure 4B). As a result, the molecules of acetylcholinesterase remain available to degrade ACh normally (Vontas et al., 2002).  

A small deletion in the C-terminal domain. The search for new mutations in highly resistant olive flies in our laboratory lead to the discovery of a short deletion at the carboxyl-terminal domain of AChE (Figure 4C). The C-terminal domain of AChE is responsible for the precise location of the enzyme. The correct localization of AChE regards the enzymatic cleavage of the enzyme’s hydrophobic carboxyl-terminal domain and its substitution for a glycosylphosphatidylinositol (GPI) anchor. The deletion of three glutamates at this domain is speculated to confer resistance through the improvement of GPI anchoring. The resistant insect undergoes a more efficient GPI modification and consequently there are more precisely located AChE molecules that degrade the pesticide than in the susceptible fly (Kakani et al., 2008). It is the first time such a mechanism is described. 



Pest management strategies have changed throughout the years. Until 1950s, the discovery of the “miraculous” insecticides had provided the illusion that a noxious insect could be completely obliterated and therefore the “complete kill” approach was adopted by the use of excessive amounts of insecticides. Entomologists soon realized that such an approach was on one hand extremely damaging to the environment and on the other was leading to problems of insecticide resistance and practically to the abandonment of several control products. After 1960s, the “complete kill” approach gave its position to the “reduction of pest impact” and the development of Integrated Pest Management (IPM). Despite this change in mentality, the two major problems, environmental impact and insecticide resistance, still remain to our days. However, the understanding and appreciation of resistance mechanisms at the molecular level can contribute to the means towards the reduction of the adverse effects of insecticides. On one hand, classical entomological practices such as less frequent applications, lower doses, use of insecticides with short residual activity, monitoring of pest populations, rotated application of different insecticides etc still remain extremely useful. On the other, the ability of modern molecular techniques to trace rare, resistance-associated alleles in a population could help model more efficient IPM practices. 




Bloomquist JR. (1996). Ion channels as targets for insecticides. Ann Rev Entomol 41: 163-190. 

Buckingham SD, Biggin PC, Sattelle BM, Brown LA, Sattelle DB. (2005). Insect GABA Receptors: Splicing, Editing, and Targeting by Antiparasitics and Insecticides. Mol Pharmacol 68: 942-951.

Daborn PJ, Yen JL, Bogwitz MR, Le Goff G, Feil E, Jeffers S, Tijet N, Perry T, Heckel D, Batterham P, Feyereisen R, Wilson TG, ffrench-Constant RH. (2002). A single P450 allele associated with insecticide resistance in global population of Drosophila. Science 297: 2252-2256. 

Dong K. (2007). Insect sodium channels and insecticide resistance. Invert Neurosci 7: 17-30.

Enayati AA, Ranson H, Hemingway J. (2005). Insect glutathione transferases and insecticide resistance. Insect Mol Biol 14: 3-8.

FAOSTAT Agricultural Data, http://apps.fao.org/default/hpm

ffrench-Constant RH. (1999). Target site mediated insecticide resistance: what questions remain? Insect Biochem Mol Biol 29: 397-403. 

Fournier D. (2005). Mutations of acetylcholinesterase which confer insecticide resistance in insect populations. Chem-Biol Interact 15: 257-261. 

Hemingway J. (2000). The molecular basis of two contrasting metabolic mechanisms of insecticide resistance. Insect Biochem Mol Biol 30: 1009-1015.  

Georgiou GP. (1990). Overview of insecticide resistance. ACS Symp Ser 421: 18-41.

IRAC Organization. Resistance : The Facts-History & overview of resistance. www.irac-online.org

Kakani EG, Ioannides IM, Margaritopoulos JT, Seraphides NA, Skouras PJ, Tsitsipis JA, Mathiopoulos KD. (2008). A small deletion in the olive fruit fly acetylcholinesterase gene associated with high levels of organophosphate resistance. Insect Biochem Mol Biol 38: 782-788. 

Mazomenos BE. (1989). Dacus oleae. In: World crop pests. Vol.3B. Edited by A.S. Robinson and G. Hooper. Elsevier Science Publishers B.V., Amsterdam. pp: 169-177.

Newcomb RD, Campbell PM, Ollis DL, Cheah E, Russell RJ, Oakeshott JG. (1997).  A single amino acid substitution converts a carboxylesterase to an organophosphorus hydrolase and confers insecticide resistance on a blowfly. Proc. Natl. Acad Sci 94: 7464-7468.

Pedigo LP (2002). Entomology and Pest Management, 4th Edition. Prentice Hall, Pearson Education Inc, New Jersey. 

Scott JG, Liu N, Wen Z. (1998). Insect cytochromes P450: diversity, insecticide resistance and tolerance to plant toxins. Comp Biochem Physiol C 121: 147-155,

Stasinakis P, Katsares V, Mavragani-Tsipidou P. (2001). Organophosphate resistance and allelic frequencies of esterases in the olive fruit fly Bactrocera oleae (Diptera: Tephritidae). J Agric Urban Entomol 18: 157-168. 

Vontas JG, Hejazi MJ, Hawkes NJ, Cosmidis N, Loukas M, Hemingway J. (2002). Resistance-associated point mutations of organophosphate insensitive acetylcholinesterase, in the olive fruit fly Bactrocera oleae. Insect Mol Biol 11: 329-336.

Zouros E, Krimbas CB. (1969). The genetics of Dacus oleae. III. Amounts of variation at two esterase loci in a Greek population. Genet Res 14: 249-258.

BOX 1: Major classes of synthetic insecticides


Insecticides have been grouped or classified in several ways, according to application or their nature and source. If application is the main focus, they can be grouped as stomach poisons, contact poisons and fumigants. If nature and source is the main focus, insecticides can be grouped as inorganic and organic. An example of inorganic insecticide is compounds of arsenic that have long been used against insect pests but they are highly toxic to non-target organisms and persist in the environment. Most modern insecticides are organic and are subdivided into natural (botanical insecticides, mineral oils) and synthetic. Synthetic insecticides constitute the major group broadly used today. The six major classes are chlorinated hydrocarbons, cyclodienes, organophoshpates, carbamates, pyrethroids and neonicotinoids.  


Chlorinated hydrocarbons

Are the oldest major insecticide class, having been the first widely used synthetic organic insecticides. All insecticides of this group (such as DDT and lindane) contain chlorine, hydrogen and carbon. Occasionally, these insecticides also contain oxygen and sulfur. Although very effective, the use of chlorinated hydrocarbons in Europe and the US is negligible. Their characteristic stability and fat solubility generated many environmental and human safety concerns. Chlorinated hydrocarbons interfere with sodium channel function, rapidly opening them, causing hyperactivity and nerve block due to continual influx of sodium ions. 



They were developed after DDT, beginning about 1945. The major compounds in this group are aldrin, dieldrin, chlordane and heptachlor. They are persistent chemicals, stable in soil and relatively so in sunlight. Therefore, many were used in great quantities against soil insects. Most cyclodienes have higher levels of mammalian toxicity than DDT and are more dangerous to apply. Growing ineffectiveness from insecticide resistance and problems with residue uptake in harvested produce have caused the elimination of most cyclodienes in agriculture. They bind to the GABA-gated chloride channel receptor complex and inhibit the binding of GABA. Consequently, the GABA channel cannot act and thus the formation of new action potentials is inhibited, causing neuronal hyperactivity.



Organophosphates were developed in Germany during World War II and their name is linked to the “nerve gases” used in concentration camps. They are derived from phosphoric acid and are some of the most toxic insecticides. Due to their instability and low persistence in the environment they replaced the chlorinated hydrocarbons in many programs. In fact, they are the most widely used group of insecticides today. Malathion, dimethoate, fenthion and diazinon are some of the most common representatives. They act as inhibitors of AChE at nerve synapses. These insecticides bind to acetylcholinesterase and inhibit the binding and degradation of acetylcholine. As a result, acetylcholine remains bound to receptors, causing hyperactivity in the nervous system. Due to the extensive and widespread use, many insect species have developed resistance to them. 



Are broad-spectrum insecticides that have had wide application in agriculture. They are produced from carbamic acid and are similar in environmental persistence to the OPs. A distinct limitation of carbamates in pest management is their toxicity to Hymenoptera, including both pollinators and parasitoids. Carbaryl and carbofuran are two of the oldest carbamates. Similarly to OPs, they act as inhibitors of AChE at nerve synapses. 



Are the fastest developing group of modern insecticides. They are replacing many older insecticides because of their great effectiveness and safety of application. Pyrethroids are not new. Allethrin was developed in 1949; it was synthesized to duplicate the insecticidal activity of the botanical insecticide pyrethrum. Their high toxicity to insects and fast breakdown by UV in the environment make them very useful. Newer pyrethroids (permethrin, cypermethrin, deltamethrin) are even more potent insect poisons and therefore very tiny doses are needed for equal effectiveness. Pyrethroids are sodium channel modulators, interfering with the influx of sodium ions in nerve cells. 



They constitute a new class of synthetic insecticides. They resemble the natural product nicotine. The class is most prominently represented by imidacloprid. It is a systemic and contact insecticide with primary activity on piercing-sucking insects such as aphids, leafhoppers, thrips and whiteflies. Additionally, it has relatively low mammalian toxicity and generally good environmental characteristics. Neonicotinoids act as agonists of acetylcholine at the postsynaptic nicotinic acetylcholine receptor (nAChR). They are bound to receptors and maintain them opened, leading to overstimulation and hyperactivity. Due to this distinct mode of action, they have the potential of managing insects that have become insecticide resistant to the other major classes of insecticides. 



BOX 2: The insect nervous system


The nervous system of insects functions to generate and transport impulses, to integrate information received and to stimulate muscles for movement. The basic unit of the nervous system that functions in nerve-impulse transmission is the nerve cell, or neuron (Figure 3A). A neuron is composed of a cell body, one or more receptor fibrils, and an axon that branches at the tip. 

Nerve impulses traveling along axons are electrical (Figure 3B). These impulses arise from the flow of positive sodium ions through the sodium channels of the cell membrane, causing depolarization. Depolarization creates an action potential that passes, as a wave, quickly down the axon. The action potential lasts only a few thousandths of a second, at which time sodium channels of the membrane close. When the sodium channels close, positive potassium ions flow out through the potassium channels and restore the cell’s resting potential (slightly negative). This is followed by an exchange of sodium inside with potassium outside the cell and a return of the original (resting) ion distribution. This type of nerve transmission is termed axonic transmission; it conveys an impulse from an arrival point along the axon to another neuron, muscle, or gland, or from a receptor cell. 

Another type of impulse transmission is synaptic transmission, which is mainly chemical (Figure 3C). Among other places, a synapse occurs at the junction between a sensory neuron and a motor neuron. When an impulse moving along the axon reaches a synapse, a chemical (neurotransmitter) is released into the synapse. The interaction between the neurotransmitter and its receptor (at the post-synaptic cell) causes the opening of ion channels and the diffusion of sodium ions into the post-synaptic cell. This change of ion balance induces and stimulates an impulse in an adjacent neuron or stimulates a muscle or gland. The most well known of these chemical transmitters is acetylcholine, although there are many others. Following transmission, the synapse is returned to the resting state by enzymes such as acetylcholinesterase, which break down the chemical transmitter acetylcholine. 

Figure legends


Figure 1. Global insecticide sales. 





Figure 2. Evolution of insecticide resistance in insect populations. By spraying crops with poisons to kill insects, humans have unwittingly favored the reproductive success of insects with inherent resistance to the poisons. Resistant individuals survive and reproduce, passing the gene for insecticide resistance to offspring. Additional applications of the same insecticide will be less effective and the frequency of resistant insects in the population will grow. 



Figure 3. The insect’s nervous system

A. A diagrammatic representation of a simple reflex loop in the insect nervous system. A stimulus to a sensory dendrite sends a signal into a central nervous system ganglion via the axon of the sensory cell. This signal is transmitted to other neurons at synaptic junctions and can be carried back out of the central nervous system in motor axons. The signal can be transmitted to a muscle cell at a neuromuscular synapse to produce a movement (not drawn to scale).



B. The action potential is a wave of depolarization and repolarization that, once initiated, self-propagates along the axon membrane. The two diagrams can be thought of as snapshots in time, depicting the same region of the membrane in different stages of the sequence, as an action potential travels toward the right.

  1. Sodium channels locally open in response to stimulus, generating an action potential here. 
  2. Some depolarizing current passively flows down axon. 
  3. Local depolarization causes neighboring sodium channels to open and generates an action potential here. 
  4. Upstream sodium channels inactivate, while potassium channels open. Membrane potential repolarizes and axon is refractory here. 
  • The process is repeated, propagating the action potential along the axon. 


C. Neurons communicate chemically at specialized structures called synapses. At this acetylcholine synapse, the action potential triggers the fusion of synaptic vesicles with the presynaptic membrane, resulting in the release of ACh into the cleft. The ACh activates postsynaptic nicotinic ACh receptors and is then split by acetylcholinesterase into acetate and choline. The action of insect control products is indicated on sodium channels, AChE, and the nicotinic acetylcholine receptor.



Figure 4. Insecticide resistance in the olive fly. 

A. The life cycle of the olive fly. The olive fruit fly typically has two to five generations a year. In early August, the female pierces the skin of the olive and deposits a single egg in the pulp. A female may lay from 50 to 400 eggs in her lifetime. The larva feeds throughout the olive and pupates in a hollow area just beneath the epidermis or outer skin. Development from egg to adult takes 30 to 40 days. Other generations during the year follow this same pattern but the last generation larvae abandon the fruit to pupate outside the olive. The fruit fly spends the winter either as a pupa in the soil or as an adult in or outside the olive grove.

B. The G488 mutation of B. oleae. The mutation hinders the binding of OP insecticides in the active site gorge of acetylcholinesterase. In this stereo representation are shown the positions of residues G488 and G365 in relation to the active site of Drosophila AChE (from Vontas et al., 2002). 


C. New mutations in olive fly’s carboxyl terminal domain of acetylcholinesterase. The circled mutation leads to a glutamine to lysine change, whereas the short boxed deletion of three glutamines points to a novel OP resistance mechanism (see text). 


Evdoxia G. Kakani and Kostas D. Mathiopoulos

Department of Biochemistry and Biotechnology, University of Thessaly, Larissa

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