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Agricultural and Biological Sciences » "Insecticides Resistance", book edited by Stanislav Trdan, ISBN 978-953-51-2258-6, Published: March 2, 2016 under CC BY 3.0 license. © The Author(s).

Chapter 18

Biochemical Insecticide Resistance in Tea Pests

By Dhiraj Saha
DOI: 10.5772/61949

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Insecticide detoxification pathways.
Figure 1. Insecticide detoxification pathways.
Metabolism of diazinon by cytochrome P450. Following an insertion of oxygen into the substrate, a reactive intermediate collapses (1) by desulphuration or (2) by cleavage of the ester linkage. DEP, diethylphosphate; DEPT, diethylphosphorothioate; P-ol, 2-isopropoxy-4-methyl-6-hydroxypyrimidine; [S], reactive form of sulphur released during the reaction. Adopted from Feyereisen, 2012.
Figure 2. Metabolism of diazinon by cytochrome P450. Following an insertion of oxygen into the substrate, a reactive intermediate collapses (1) by desulphuration or (2) by cleavage of the ester linkage. DEP, diethylphosphate; DEPT, diethylphosphorothioate; P-ol, 2-isopropoxy-4-methyl-6-hydroxypyrimidine; [S], reactive form of sulphur released during the reaction. Adopted from Feyereisen, 2012.
Metabolism of deltamethrin by insect P450 enzymes: (1) deltamethrin; (2) 4′ hydroxydeltamethrin; (3) trans-hydroxymethyl-deltamethrin; (4) cyano (3-hydroxyphenyl) methyl deltamethrate; (5) deltamethric acid. Adopted from Feyereisen, 2012.
Figure 3. Metabolism of deltamethrin by insect P450 enzymes: (1) deltamethrin; (2) 4′ hydroxydeltamethrin; (3) trans-hydroxymethyl-deltamethrin; (4) cyano (3-hydroxyphenyl) methyl deltamethrate; (5) deltamethric acid. Adopted from Feyereisen, 2012.
Chlorfenapyr and cypermethrin metabolism. The same P450 in Heliothis virescens probably activates the pyrrole and inactivates the pyrethroid, resulting in negative cross-resistance. Adopted from Feyereisen, 2012.
Figure 4. Chlorfenapyr and cypermethrin metabolism. The same P450 in Heliothis virescens probably activates the pyrrole and inactivates the pyrethroid, resulting in negative cross-resistance. Adopted from Feyereisen, 2012.
Metabolism of imidacloprid by insect P450 enzymes: (1) imidacloprid; (2) 5-hydroxyimidacloprid; (3) 4-hydroxyimidacloprid; (4) dihydroxyimidacloprid; (5) Non-enzymatically derived dehydroimidacloprid. Adopted from Feyereisen, 2005.
Figure 5. Metabolism of imidacloprid by insect P450 enzymes: (1) imidacloprid; (2) 5-hydroxyimidacloprid; (3) 4-hydroxyimidacloprid; (4) dihydroxyimidacloprid; (5) Non-enzymatically derived dehydroimidacloprid. Adopted from Feyereisen, 2005.

Biochemical Insecticide Resistance in Tea Pests

Dhiraj Saha1

1. Introduction

1.1. Tea, the plantation crop with economic value: Driving the shrub to cup

Tea is produced from young leaves and buds of tea plant, Camellia sinensis (L.) O. Kuntze is a native of China, and the Chinese are said to have discovered its use nearly 4,700 years ago. It is believed that Shen Nung, a Chinese emperor who lived some 4,700 years ago, discovered that tea leaves falling into boiling water make a refreshing drink. Tea then became a popular drink in China for both its flavour and medicinal qualities. Eventually, the habit of drinking tea spread throughout Asia and then throughout the world. Currently, tea is the second most preferred and popular drink after water. The word tea had its origin from t’e (pronounced as ‘tay’) in ‘Amoy’ dialect while in Cantonese it was called ch’a (‘chah’). This is the name by which this wonderful beverage is known in Japan, Iran, Russia, Indonesia, Malaysia, Vietnam and India. Tea was introduced in Japan about AD 800 and was regarded as medicine for about 500 years. Tea was introduced in Europe in the early 17th century with the beginning of trade between Europe and the Southeast Asia.

Now tea is produced in almost every region of the globe. The tea plant is predominantly grown in Asia followed by in Africa and to a very small extent in Europe, South America, Australia and New Zealand. Now, tea is grown in 36 tropical and subtropical countries. Major tea producing countries are India, China, Sri Lanka, Kenya, Japan, Indonesia, Thailand, Bangladesh, Nepal, Vietnam, Turkey and Argentina.

Tea plants are native to East and South Asia and probably originated around the point of confluence of the lands of northeast India, north Burma, southwest China and Tibet. The commercially cultivated tea plants are derived from small-leaved China plants, C. sinensis, the Assam plants, C. assamica (Masters), and the Cambod plants, C. assamica ssp. lasiocalyx (Planchon ex Watt) Wight and numerous hybrids among them. Tea plants require warm humid climate, well-distributed rainfall and long sunshine hours. Shoots, comprising two or three tender leaves and a bud, are harvested and processed in factories to manufacture different types of tea.

2. Insect pest occurrences in tea

Being a widespread perennial monoculture crop, tea plantations provide the most congenial microclimate as well as continuous food supply to a number of arthropods. Every part of the tea plant, i.e. leaf, stem, root, flower and seed is subjected to attack by at least one arthropod pest species. According to recent estimates, globally, more than a thousand (1,034 species) arthropods and 82 species of nematodes are associated with tea plantations [1]. Among the insect pests, 32% are from order Lepidoptera, followed by 27% pest species from Hemiptera [2]. In India, only 300 species of arthropods are recorded as pest and about 167 species are from tea-growing Northeast India (Table 1) [3]. The dynamic adaptations of insects have facilitated them to exploit every part of the tea plant and the maximum numbers of pests occur on the foliage.

Among the arthropods that attack tea plant, insect and mite pests are the most damaging [2], causing on average a 5%–55% yield loss [46]. Insect pests alone can cause on an average 11%–55% yield loss, if left unrestrained [1].

Tea mosquito bug, thrips, jassids, tea caterpillars (looper, redslug and bunch), aphids, termites, cockchafers and red spider mite are among the various insects and mite pests that cause severe loss in tea production. The damage caused by the tea pests frequently leads to a significant impact on quality and yield, while the degree of pest infestation differs in different tea-growing areas depending on altitude, climate, forest cover and local cultural practices. Tea pests can be broadly classified into the different categories based on their feeding nature, time of occurrence and severity. The major pests are mite pests: red spider mite (Oligonychus coffeae), scarlet mite (Brevipalpus phoenicis), pink mite (Acaphylla theae), purple mite (Calacarus carinatus); sucking pests: tea mosquito bug (Helopeltis theivora), tea thrips (Scirtothrips dorsalis and Myctrothrips setiventris), tea jassids (Empoasca flavescens), aphids, scale insects; leaf eaters: looper caterpillars (Hyposidra talaca, H. infixaria, Buzura suppressaria), red slug caterpillar (Eterusia magnifica), bunch caterpillar (Andraca bipunctata); and soil-borne pests: termites (Microcerotermes sp. Odontotermes sp.), cockchafers, nematodes, weevils, cricket [8].

Sl. no. Name of the pests Nature of damage Family: Order
1. Andraca bipunctata Walk.Bunch caterpillarFolivoresBombycidae: Lep.
2. Eterusia magnifica Butl.Red slug caterpillarFolivoresZygaenidae: Lep.
3. Buzura suppressaria Guen.Looper caterpillarFolivoresGeometridae: Lep.
4. Hyposidra talaca, H. infixaria Walk.Black inch wormFolivoresGeometridae: Lep.
6. Lymantria albulunata Mre.Sungma caterpillarFolivoresLymantridae: Lep.
7. Gracillaria theivora Walsh.Tea leaf rollerFolivoresGracilariidae: Lep.
8 Homona coffearia NietnerTea tortrixFolivoresTorticidae:Lep.
9. Laspeyresia leucostoma Mayer.FlushwormFolivoresEucosmidae: Lep.
10. Holotrichia impressa Burm.Cockchafer grubsRoot of young teaScarabaeidae: Col.
11. Serica assamensis BrenskeLeaf-eating cockchaferLeaf eaterScarabaeidae: Col.
12. Asticus chrysochlorus Wied.Large green weevilLeaf eaterCurculionidae: Col.
13. Agromyzidae (Bigot) MeijTea leaf minerLeaf eaterAgromyzidae: Dip.
14. Microtermes spp.Live wood-eating termiteStem and root eaterTermitidae: Iso.
15. Odontotermes sp.Scavenging termiteStem and root eaterTermitidae: Iso.
16. Helopeltis theivora WaterhouseTea mosquito bugLeaf suckerMiridae: Hem.
17. Empoasca flavescens Fabr.Tea greenfly/tea jassidLeaf suckerJassidae: Hem.
18. Toxoptera aurantii Boyer de Fons.Tea aphidLeaf suckerAphididae: Hem.
19. Scirtothrips dorsalis HoodYellow tea thripsLeaf and bud suckerThripidae: Thy.
20. Mycterothrips (Teaniothrips) setiventris BagnallCommon thripsLeaf and bud suckerThripidae: Thy.
21 Oligonychus coffeae NietnerRed spider miteLeaf suckerTetranychidae: Aca.
22. Brevipalpus phoenicis GeijskesScarlet miteLeaf suckerTenuipalpidae: Aca.

Table 1.

Major insect and mite pests that occur on tea in India.

3. Insect pests of tea and management problems

Tea is produced from the young foliage, i.e. young leaves and a bud, and foliage production is increased by seasonal pruning which enhances the leaf cover. The major pests of the crop are those associated with the young foliage. The most important insect pest groups are the folivores (chewing) and sap suckers of the young tender leaves, buds and stems (sucking pest), which damage the most economic part of tea plant that is processed in tea industry for making the tea. These pests cause substantial loss in yield to the tea industry.

In India, different management practices are followed to protect the tea crop against different insect pest groups. Most of the plantations are managed conventionally i.e. using different organosynthetic insecticides, whereas some organic plantations use plant- and animal-based herbal and microbial insecticides. In conventional tea plantations, organo-synthetic insecticides of different functional groups such as organochlorines, organophosphates, synthetic pyrethroids (SPs) and neonicotinoids (NNs) are regularly used throughout the year to control the invasion of different insect pest groups (sucking, folivores and others) [8]. The use of insecticide is cost-effective to planters and a major concern for the environmental degradation due to contamination as well as in resurgence of primary pests [6], outbreak of secondary pests [9], development of insecticide resistance [10, 11], including undesirable residues in made tea [12]. Regular spraying of insecticides leads to the development of higher level of tolerance or resistance to insecticides in many insects [11, 13].

From the early forties onwards, dichlorodiphenyltrichloroethane (DDT) (organochlorine) was regularly used to manage the infestation of H. theivora, the major sucking pest in Northeast India [14]. In 1968, endosulfan (cyclodiene: organochlorine) was introduced in the tea plantations of the Dooars region of West Bengal, India, in the form of thiodan 35 EC [15]. Currently, in different conventional tea plantations of tea-growing regions of India, cypermethrin, deltamethrin, quinalphos, monocrotophos, chlorpyriphos, imidacloprid, etc. are extensively used during cropping season to control insect pests [1618].

Recently, a number of insecticides have been found to be ineffective in controlling the insect and mite pests in different tea-growing regions of India [8]. The development of resistance to different classes of insecticides is one of the causes for persistence and resurgence of insect pests on tea crop [8, 1921]. A major concern in managing the major insect pests of tea is its high potential to develop resistance rapidly to regularly used insecticides [11]. Continuous and repeated exposure to different classes of insecticides for many years, in addition to their high reproductive potential, short life cycle and numerous annual generations, has limited the management of major pests of tea [11]. Recently, there are reports on the development of resistance to many commonly used synthetic insecticides and consequent failure in controlling many tea pests [10, 2226]. Such failures are already known in case of organochlorines (OCs), organophosphorus (OP) and synthetic pyrethroid (SP) insecticides and more recently for the newer compound such as neonicotinoids [1921, 26]. The development of resistance in H. theivora populations to different classes of insecticides has been in the range of 1.47–62.99-fold for males and 1.25–62.82-fold for females in Northeast India [19]. Relative toxicity to commonly used insecticides has been observed to vary in H. theivora populations from Jorhat, Assam [20], Darjeeling [27], and from sub-Himalayan Dooars region of Northeast India [28]. LC50 values of insecticides, when compared with the field dose against H. theivora recommended by TRA (Tea Research Association, Tocklai, Assam, India), revealed a pronounced shift in the level of susceptibility of H. theivora to all insecticides except acephate [2021].

For the management of other sucking insects such as yellow tea thrips, S. dorsalis, insecticides are also used in conventional tea plantations. In tea ecosystem, control failure and the development of biochemical resistance have been reported in S. dorsalis [11, 24]. In India, S. dorsalis populations have developed a high degree of resistance to a range of organochlorine (DDT, BHC and endosulfan), organophosphate (acephate, dimethoate, phosalone, methyl-O-demeton and triazophos) and carbamate insecticide (carbaryl) in chili ecosystem [31]. S. dorsalis has also developed a high degree of resistance to various insecticides, viz. monocrotophos, acephate, dimethoate, phosalone, carbaryl and triazophos [32]. Recently, several insecticides have been tested on S. dorsalis in chili ecosystem in USA and found limited success with chlorfenpyr, spinosad and imidacloprid [33, 34]. The performance of novaluron, abamectin, spiromesifen, cyfluthrin, methiocarb and azadirachtin failed to provide effective control of this pest [35].

Similarly, in another emerging sucking insect pest of tea, tea greenfly, E. flavescens, repeated management failure and biochemical insecticide resistance in tea ecosystem from Northeast India have been reported [11, 24]. In China, chemical insecticides including fenvalerate, cyfluthin, cypermethrin and imidacloprid are sprayed to control the leafhoppers as frequently as seven times annually or even more frequently [3637]. A high level of resistance against many insecticides has been reported in related species, E. vitis [38]. The resistance to thiamethoxam was highest and to cyperemethrin was lowest in E. vitis. Recently, in Fujian province of China, a regional diversity of resistance to eight insecticides in E. vitis has been reported in tea ecosystem with higher resistance level to bifenthrin, acetamiprid, imidacloprid, cartap and chlorfenapyr [39].

A high level of insecticide resistance in folivores, such as black hairy caterpillar, bunch caterpillar, looper pest complex (Hyposidra talaca, H. infixaria, Buzura suppressaria, Eturesia magnifica) and in termite of tea ecosystem, has been reported with reduced susceptibility against different insecticides.

Detoxification of insecticides is an important toxicokinetic mechanism for insect pests to tolerate regularly applied insecticides [8, 4042]. Susceptibility levels against insecticides change mainly due to metabolic detoxification of the insecticides through the induction of some detoxifying enzymes under the stress of different management practices [4345].

Generally, three principal enzymes, general esterases (GEs), glutathione S-transferases (GSTs) and cytochrome P450-mediated monooxygenases (CYP450s), are involved in the process of metabolic detoxification of insecticides [41]. Estimation of the activities of these metabolic defence-related detoxifying enzymes gives information on the level of tolerance/resistance of the insect pest population to insecticides and is a useful tool in monitoring the tolerance/resistance to insecticides at population level of the pest. The early detection of metabolic threats related to tolerance/resistance to insecticides in pest specimens is of crucial importance for devising pest control techniques that would minimize the development of tolerant/resistant forms and prevent any undesirable wastage of insecticide, money and manpower.

4. Insecticide resistance mechanisms in insect pests of tea

Insects come across with numerous toxins as they go through their life cycle. Some of these toxins are naturally produced by plants (plant allelochemical) and others by humans (synthetic insecticides). To protect themselves against the natural toxins, insects have evolved various detoxification mechanisms [41] These mechanisms also cross-protect insect pests when they are exposed to synthetic insecticides [25]. Herbivorous insect groups (agricultural pests) are significantly more diverse than their non-herbivorous sister groups [46]. The role of plant in promoting diversification in insects has occurred through co-evolutionary ‘defence strategies’ among them [47]. This diversification could also have been a result of insects ‘tracking’ plant phylogenies, with minor chemical changes in plants allowing the evolving populations of insects to change and speciate accordingly, which probably has occurred long after chemical changes in plants [48]. Evolution to herbivory preceded via mixed feeding on reproductive parts or spores, dead tissues of plants and animals and fungi. This progression implies that omnivory preceded generalized herbivory and the evolution of specialization on specific plant taxa was a later accomplishment [49].

Among sucking insect herbivores, the actual food used, i.e. digested whole tissue particularly parenchyma (as in Helopeltis sp.), cell content (thrips) and phloem flow (Empoasca sp.), influences both the feeding mechanism and feeding behaviour [48]. While the chewing insects (looper caterpillar complex) cause extensive damage, the sucking insects cause modest to barely perceptible damage. However, sucking insects, particularly phloem and digested tissue feeder,impose an additional challenge to the plants as they deplete photosynthates, act as vector of viruses and introduce chemical and protein effectors that alter plant defence mechanisms (signalling) and development [50]. When these attributes are combined with a broad host range, breeding strategies that promote invasiveness, highly evolved feeding strategies, the ability to adapt to a wide range plant habitats and the emergence of insecticide resistance, it is not surprising that sucking insects cause heavy losses in agriculture and horticulture [51].

Insecticide resistance is a genetic change in response to selection pressure of toxicants that impair pest control in the field [52]. Insecticide resistance does not occur unless a structural genetic change occurs that is heritable. Therefore, insecticide resistance is an evolutionary phenomenon that results under the selection pressure of a new toxicant in the environment [8].Thus insecticide resistance is different from insecticide tolerance. Insecticide tolerance is the natural ability of a population to withstand the toxic effect of a particular insecticide. It can develop within one generation as a result of physiological adaptation, i.e. induction of xenobiotic detoxifying enzymes. Hence, variation within a population may include individuals with genetic traits that make them better adapted to survive in exposure to an insecticide. If these individuals survive the insecticide exposure, then the tolerance traits can be passed on to the next generation, thereby enriching the gene pool with those genes. The mechanisms of development of insecticide tolerance can be divided into four levels:

Altered behaviour – avoidance of contact with the insecticide

Development of barrier tissues – reduced penetration of the insecticide though the integument

Enhanced detoxification – higher metabolism of the insecticide

Alteration of receptor – at the target site for the insecticide.

The first level, at which insecticide tolerance can develop, is when the insect encounters an insecticide. An altered behaviour helps the insect to avoid coming into contact with the insecticide. Once the insect comes in contact with the insecticide, a reduced and delayed penetration through the cuticle will reduce the effect of the insecticide at the target site; this is yet another level of resistance. Within the insect’s body, the insecticide may be enzymatically metabolized and thereby inactivated. At the third level of resistance mechanism, three systems of xenobiotic detoxification enzymes operate: esterases, glutathione S-transferases and cytochrome P450-dependent monooxygenases. The increased activity of one of these enzyme systems in metabolizing insecticides will result in insecticide tolerance. Alterations at the target site for the insecticide are the last level of insecticide resistance mechanisms. Different classes of insecticides bind to specific target sites and reduced binding at the target site, or increased number of target site molecules may confer insecticide resistance.

4.1. Behavioural resistance

Behavioural resistance mechanisms are the least studied resistance mechanisms in insects, but this is not to say that behavioural resistance is the least significant. Behavioural resistance can be defined as ‘evolved behaviours that will reduce an insect’s exposure to toxic compounds or that allows an insect to survive in what would otherwise be a toxic and fatal environment’ [53]. Behavioural resistance has been observed in more than 30 species of insects [53]. Avoidance is the first step in the evolution of behavioural resistance [54]. In H. theivora, this kind of resistance has been seen [55]. H. theivora shows a different egg-laying strategy to avoid insecticide exposure. Even E. flavescens avoids exposure to direct sunlight and therefore prefers to stay on the underside of the tea leaves. This behaviour cross-protects it from the direct insecticide exposure in conventional tea plantations during spraying [8]. The same has been found in S. dorsalis which resides inside the leaf bud during development and underside of the leaf during adult stage, thereby avoiding direct exposure to insecticides [8].

4.2. Reduced penetration

Reduced penetration of insecticides through barrier tissues of insects is another way in which an insect can modify the effective dose of insecticide at the target site. The mechanism may not prevent the insecticide from eventually entering the insect, but it can reduce the rate at which the insecticide reaches the target site. Reduced penetration has been shown to function as a resistance mechanism to many different insecticides, and, by the nature of this mechanism, cross-resistance is often found [56]. The rate of penetration of insecticides through the insect cuticle or other barriers (peritrophic membrane) depends on the physicochemical properties of the insecticide and the barrier. A reduced penetration contributes to DDT resistance in the tobacco budworm, Heliothis virescens F. (Lepidoptera: Noctuidae). DDT-resistant larvae had an altered composition of the cuticle. The protein and lipid contents are greater in the cuticle of resistant larvae and, furthermore, the cuticle of the resistant larvae probably had a higher degree of sclerotization [57]. In M. domestica, two resistant strains, with reduced penetration as one of the resistance mechanisms, also showed increased cuticular lipid content; more total lipids, mono-glycerides, fatty acids, sterols and phospholipids were present in the resistant strains compared to a susceptible strain [58]. Reduced penetration has been documented as a resistance mechanism only at the level of the insect cuticle, but any biological membrane may serve as a barrier and thereby give resistance [59]. As a single resistance mechanism it usually only confers low levels (less than threefold) of resistance [59]. Reduced penetration has been shown to function as a resistance mechanism to many different insecticides, including insecticides of the three major classes, OPs, carbamates and pyrethroids. Reduced penetration of OPs through the cuticular barrier has been reported, for example, diazinon in M. domestica [60], azinphos-methyl in the pear psylla, Psylla pyricola Foester (Hemiptera: Psyllidae) [61] and profenofos in H. virescens [62]. However, by slowing the penetration rate of insecticides, this mechanism reduces the risk that the insects’ detoxification systems become overloaded, and the dose of insecticide reaches to a lethal level at the target site. In female H. theivora, a higher level of body lipid has been found which effectively reduces the penetration of insecticide to the target site [63]. No studies on resistance due to reduced penetration in E. flavescens and S. dorsalis or any other tea pests have been reported to date. The studies had shown that when different resistance mechanisms are combined in the same individuals, a synergistic effect, resulting in a high level of resistance, may arise [64, 65]. Therefore, even a small degree of reduced penetration can contribute significantly to the overall insecticide resistance of the insect pests.

4.3. Metabolic detoxification

Metabolic detoxification of insecticides is an important toxicokinetic mechanism for insects to tolerate the toxic effects of insecticides. Generally, lipophilic (hydrophobic) insecticides are rapidly detoxified. Organophosphates, organochlorines, carbamates and pyrethroids are lipophilic compounds, and detoxification enzymes transform these insecticides to more hydrophilic and less biologically active compounds so that can be eliminated more easily by excretion. Increased detoxification of insecticides has often been reported in many resistant populations [40]. Three enzyme systems are generally recognized as the major detoxification systems involved in insecticide resistance in insects. These are carboxylesterases, cytochrome P450-dependent monooxygenases and glutathione S-transferases [40, 41].

4.4. Alteration at the target site for insecticide (target site insensitivity)

The biochemical sites for insecticide action differ for different insecticides and are a potential field of research for developing insecticides, which can act specifically or more efficiently on insect biochemical sites compared to mammals. The target site receptor for action of organophosphates, carbamates, organochlorines and pyrethroids is in the nervous system. The enzyme acetylcholinesterases (AChEs) (EC are the target sites for organophosphates and carbamates, and voltage-gated sodium channel of the nerve membrane is the target of pyrethroids and DDT. Neurotoxic insecticides such as cyclodienes (e.g. dieldrin and endosulfan), y-HCH (lindane) and fipronil target gamma-aminobutyric acid (GABA)-receptor [68, 69] and nicotinyl insecticides (imidacloprid and nicotine) target the nicotinic acetylcholine receptor (nACHR) [70]. Alteration at the target site, to less a sensitive target for neurotoxic insecticides, is an important toxicodynamic resistance mechanism in insects [71].

In insects, the potent inhibitors of AChE are organophosphates and carbamates. These compounds inhibit the activity of AChE by forming a stable covalent intermediate, preventing the enzyme to hydrolyse acetylcholine. An accumulation of acetylcholine keeps the ion channel of the receptor permanently open, which eventually kills insect. OPs and carbamates are quasi-irreversible inhibitors of AChE. The organophosphates and carbamates phosphorylate and carbamylate the active site serine of AChE, respectively [72]. Generally, the reactivation time of phosphorylated or carbamylated AChE is long. However, the half-lives of reactivation vary considerably, from minutes to several days, depending on the compound interacting with AChE [73]. Carbamylated AChE generally reactivates faster than phosphorylated AChE. Reduced sensitivity of AChE to inhibition by OPs and carbamates is an important resistance mechanism in insects and is often referred to as altered or insensitive AChE [74]. The presence of insensitive AChE conferring resistance was first noticed in OP-resistant mites, Tetranychus urticae Koch (Acari: Tetranycidae) [74] and also found in several insect populations resistant to these compounds [7577]. Insecticide susceptible and resistant insect pest populations differ in the level of AChE activity [7880]. A higher level of AChE activity has been reported in H. theivora sampled from conventional tea plantations than from organic garden indicating the presence of resistance to insecticides in conventional tea ecosystems [81, 82]. There is no such report on S. dorsalis and E. flavescens in conventional tea ecosystems to date.

5. Major metabolic detoxifying enzymes in insects

Carboxylesterases, glutathione S-transferases and cytochrome P450-mediated monooxygenases are the three principal enzymes that facilitate the insects to metabolize different kind of toxins. These large enzyme families contain multiple forms with overlapping substrate specificities. Knowledge of insecticide detoxification helps in understanding the mechanism of insecticide resistance, hence the development of a sound resistance management strategy. Detoxification can be divided into phase I (primary) and phase II (secondary) processes (Figure 1).


Figure 1.

Insecticide detoxification pathways.

Phase I reactions consist of oxidation, hydrolysis and reduction. The phase I metabolites are sometimes polar enough to be excreted but are usually further converted by phase II reactions. In phase II reactions, the polar products are conjugated with a variety of endogenous compounds such as sugars, sulphate, phosphate, amino acids or glutathione and subsequently excreted. Phase I reactions are usually responsible for decreasing the biological activity of toxins, and therefore the enzymes involved are rate limiting with respect to toxicity. The most important function of biotransformation is to decrease the lipophilicity of insecticides, so that they can be excreted quickly [83].

5.1. Phase I reactions

5.1.1. Cytochrome P450 monooxygenases (E.C. 1.14.-.-)

Oxidation is considered the most important among phase I reactions. The oxidative reactions are carried out mainly by a group of enzymes called cytochrome P450 monooxygenases [also known as mixed function oxidases (MFO) or polysubstrate monooxygenases (PSMO), microsomal oxidase, P450 enzymes]. Cytochrome P450, or CYP genes, constitutes one of the largest family of genes, with representatives in virtually all living organisms, from bacteria to protists, plants, fungi and animals [84].

In insects, P450 monooxygenases are involved in many processes including roles in the metabolism of plant allelochemicals by herbivores and in detoxification of insecticides. The human genome carries about 57 CYP genes, and insect genomes can carry from 36 CYP genes in the body louse Pediculus humanus [85] to 170 in a mosquito [86]. Each P450 protein is the product of a distinct CYP gene, and P450 diversity is the result of successive gene (or genome) duplications followed by sequence divergence [84].

The typically 45–55-kDa P450 proteins are heme-thiolate enzymes. Their essential common feature is the absorbance peak near 450 nm of their FeII–CO complex for which they are named [87]. P450 enzymes are best known for their monooxygenase role, catalysing the transfer of one atom of molecular oxygen to a substrate and reducing the other to water. The simple stoichiometry commonly describes the monooxygenase or mixed function oxidase reaction of P450:


However, oxygen atom transfer is not the only catalytic function of P450 enzymes. They also show activities such as oxidases, reductases, desaturases, isomerases, etc. and collectively are known to catalyse at least 60 chemically distinct reactions (Table 2).

Reaction catalysedP450
Oxidase activity
O2 to H2O, H2O2, O2
CYP6A1 (and probably most P450 enzymes)
Aliphatic hydroxylation
C–H hydroxylation
CYP4C7, CYP6A1, CYP6A2, CYP6A8, CYP6G1, CYP6M2, CYP6CM1vQ, CYP9T2, CYP12A1, CYP18A1, CYP302A1 CYP306A1, CYP312A1, CYP314A1, CYP315A1
O-dealkylationCYP6A1, CYP6D1, CYP6A5, CYP6B4, CYP6B17, CYP6B21, CYP6G1, CYP6Z2, CYP6CM1vQ, CYP9A12, CYP9A14, CYP12A1, CYP321A1,
EpoxidationCYP6A1, CYP6A2, CYP6B8, CYP6B27, CYP6AB3, CYP6 CYP6AB11, CYP9E1, CYP12A1, CYP15A1, CYP321A1
Aromatic hydroxylationCYP6D1, CYP6G1, CYP6M2
Heteroatom oxidation and dealkylation
Phosphorothioate ester oxidation
N-dealkylationCYP6A5, CYP12A1
Aldehyde oxidationCYP18A1
Complex and atypical reactions
Cyanogenic glucoside biosynthesis:
Val/Ile to oximes
Oximes to cyanohydrins
Aryl ether cleavage
Carbon–carbon cleavage
Decarbonylation with C–C cleavage
Endoperoxide isomerisation
+?(Sterols, ecdysteroid)
+(Defensive steroids)

Table 2.

Enzymatic reactions catalysed by insect P450 enzymes (adapted from Feyereisen, 2005).

The first insect P450 cloned and sequenced was CYP6A1 from Musca domestica in 1989 [88]. The P450 gene complement (CYPome) size of an insect genome is not a definite number [89]. Insects can survive with small CYPomes even in toxic environments. The human body louse Pediculus humanus, with 36 CYP genes, is known to become highly resistant to many classes of insecticides [85], and the honeybee, with just 46 CYP genes [90], is not more sensitive than other species in a comparison to the toxicity of 62 insecticides [91]. The main driver of CYPome evolution is of course gene duplication, followed by divergence (by neofunctionalization or subfunctionalization) or death (pseudogenization or deletion) [84].

5.1.2. Carboxylesterases (EC

Carboxylesterase or esterase is a collective term for the enzymes that hydrolyse carboxylic esters [92]. Classification of these enzymes is difficult because of their overlapping substrate specificity [93]. However, the esterase classification of Aldridge [94] is generally recognized. According to that classification, esterases inhibited by paraoxon in a progressive and temperature-dependent manner are called B-esterases and those which are not inhibited are A-esterases [94]. Some A-esterases can hydrolyse OPs, through an acylated cysteine in their active site, and are termed phosphoric triester hydrolases (EC 3.1.8.) [95, 96]. The term carboxylesterase is now mainly attributed to B-esterases [95, 96]. These enzymes have an active site serine residue, hence the terms B-esterase and serine hydrolase are synonymous. Insecticides such as organophosphates, carbamates, pyrethroids and some juvenoids, which contain ester linkages, are susceptible to hydrolysis. Esterases are hydrolases that split ester compounds by the addition of water to yield an acid and alcohol.

Esterases that metabolize organophosphates can be divided into three groups: A-esterases which are not inhibited by organophosphates but hydrolyse them; B-esterases, which are susceptible to organophosphate inhibition; and C-esterases which are uninhibited by organophosphates and do not degrade them [41].

There are two types of esterases that are important in metabolizing insecticides, namely, carboxylesterases and phosphatases (also called phosphorotriester hydrolases or phosphor-triesterases). Carboxyl esterases, which are B-esterases, play a significant role in degrading organophosphates, carbamates, pyrethroids, and some juvenoids in insects. The best example is malathion hydrolysis, which yields both α- and β-monoacids and ethanol [41]. Phosphatases are A-esterases that detoxify many organophosphorous insecticides especially phosphates in insects. In houseflies, paraoxon can be hydrolysed to diethyl phosphoric acid and p-nitrophenol. Phosphatases also hydrolyse the alkyl groups of organophosphates. Paraoxon is hydrolysed by the enzyme in houseflies. Several amides containing organophosphorous insecticides such as dimethoate and acephate have been shown to be hydrolysed by carboxylamidases to their corresponding carboxylic acid derivatives [41].

5.2. Phase II reactions

Phase I reactions with xenobiotics result in the addition of functional groups such as hydroxyl, carboxyl and epoxide. These phase I products can further undergo conjugation reactions with endogenous molecules. These conjugations are called phase II reactions. The endogenous molecules include sugars, amino acids, glutathione, phosphate and sulphate. Conjugation products are usually more polar, less toxic and more readily excreted than their parent compounds. Thus, the process with only a few exceptions results in detoxifications.

Three types of conjugation reactions occur in insects. Type I requires an activated conjugating agent that then combines with the substrate to form the conjugated product. Type II involves the activation of the substrate to form an activated donor that then combines with an endogenous molecule to yield a conjugated product. In Type III, conjugation can proceed directly between the substrate and the conjugating agent without involving activation. Thus, Type I and II require information of high-energy intermediates before the conjugation reactions proceed. The chemical groups required for Type I are –OH, NH2, COOH and SH (glucose conjugation, sulphate conjugation and phosphate conjugation); for Type II COOH (amino acid conjugation); and for Type III, halogens, alkenes, NO2, epoxides, ethers and esters (glutathione conjugation).

5.2.1. Glutathione S-transferases (EC

Glutathione conjugations are performed by a group of multifunctional enzymes known as glutathione S-transferases and are involved in detoxification mechanisms of many molecules. GSTs are involved in the transport of physiologically important lipophilic compounds. These enzymes catalyse reactions in which the sulphur atom of glutathione provides electron for nucleophilic attack on a second electrophilic substrate; the latter can be endogenous natural substrates such as epoxides, organic hydroperoxides, or activated alkenals resulting from oxidative metabolism. These enzymes catalyse the conjugation of reduced glutathione (GSH) with electrophilic substrates. Glutathione S-transferases perform a variety of reactions including:

The S-alkylation of GSH by alkyl halides and related compounds.

The replacement of labile aryl halogen or nitro groups by GSH.

The replacement of labile arakyl halogen and ester groups by GSH.

The addition of GSH to various epoxides.

The addition of GSH to α-, β-unsaturated compounds including aldehydes, ketones, lactones, nitriles and nitro compounds.

The O-alkyl and O-aryl conjugation of phosphorothiotes and phosphates with GSH.

The glutathione conjugate is subsequently transformed to mercapturic acid through the stepwise loss of glutamic acid and glycine to a cysteine conjugate, which is finally acetylated before excretion. Because of their broad substrate specificities, glutathione S-transferases are responsible for the detoxification of numerous xenobiotics [97]. More than 40 GST genes have been identified in insects [98, 99]. Mammalian GSTs have been classified into eight cytosolic classes (alpha, mu, pi, theta, sigma, zeta, kappa and omega) and a microsomal class on the basis of their amino acid sequence, immunological properties and substrate specificity. Each class shares 40% or higher amino acid in common. The classification of insect GST is not clear. The majority of insect GSTs do not belong to mammalian classes. Insect glutathione S-transferases consist of two subunits (homodimers and heterodimers) of molecular weight between 19 and 35 kD. Two classes of insect GSTs (Class I and Class II) were reported [100], which have been referred to as the Delta and Sigma class, respectively. Recently, a new class of insect GSTs, referred to as Epsilon has been described in several species of insects including Anopheles gambiae [101]. Purified cytosolic and microsomal glutathione S-transferase isozymes from fall armyworm larvae possessed cumenehydrperoxide peroxidase [102]. A Delta class GST purified from German cockroaches also showed high peroxidase activity [103]. The name of each GST is composed of the initials of the species scientific name, followed by the acronym GST, a capital letter to designate the class name and an Arabic number for the individual protein, such as AgGSTD2.

Glutathione S-transferases are important in the metabolism of organophosphorous insecticides resulting in detoxification [99, 104]. For example, methyl parathion is dealkylated by glutathione S-tranferases to form desmethyl parathion and methyl glutathione [41]. On the contrary, parathion can be de-arylated by glutathione S-transferases to produce diethyl phosphorothioic acid and S-(p-nitrophenyl) glutathione [41]. Interestingly, a glutathione S-transferase isozyme from the housefly exhibits DDT-dehydrochlorinase activity, showing that DDT-dehydrochlorinase (DDTase) is one of the glutathione S-tranferases [104]. DDT-dehydrochlorinase converts DDT to DDE, resulting in detoxification [41].

6. Detoxifying enzymes and insecticide metabolism

The metabolism of insecticides by P450 enzymes is very often a key factor in determining toxicity to insects and to non-target species. The importance of monooxygenases in insecticide resistance became evident in the early 1960s, when it was shown that resistance to carbaryl could be abolished by the P450 inhibitor sesame [106]. Additional evidence of monooxygenase-based resistance quickly amassed [107, 108]. Monooxygenase-mediated detoxification is frequently found as a major mechanism of resistance, and unlike target site resistance, detoxification has the potential to confer cross-resistance to toxins independent of their target sites [109, 110]. Most cases of monooxygenase-mediated resistance result from an increase in detoxification (Table 3). However, in cases where the parent insecticide must undergo monooxygenase-mediated bioactivation, as is the case for many organophosphates, it is also possible that resistance could be achieved through decreased activation [111]. Although this has been reported once, it does not appear to be a common mechanism of resistance. This may explain why esterases are relatively more common than monooxygenases in resistance to some organophosphates [110, 112]. The classical example is probably the metabolism of phosphorothioate insecticides. In many cases, the active ingredients of organophosphorus insecticides are phosphorothioate (P=S) compounds (also known as phosphorothionates), whereas the molecule active at the acetylcholinesterase target site is the corresponding phosphate (P=O) (Figure 2).


Figure 2.

Metabolism of diazinon by cytochrome P450. Following an insertion of oxygen into the substrate, a reactive intermediate collapses (1) by desulphuration or (2) by cleavage of the ester linkage. DEP, diethylphosphate; DEPT, diethylphosphorothioate; P-ol, 2-isopropoxy-4-methyl-6-hydroxypyrimidine; [S], reactive form of sulphur released during the reaction. Adopted from Feyereisen, 2012.

P450 enzymes that metabolize OPs can metabolize other insecticides as well, and this sometimes leads to potentially useful interactions. Thus, enhanced detoxification of dicofol in spider mites can lead to enhanced chlorpyriphos activation, and hence negative cross-resistance [113]. Similarly, permethrin resistance in horn flies is suppressible by piperonyl butoxide, and negatively related to diazinon toxicity [114]. In H. armigera populations from West Africa, triazophos shows negative cross-resistance with pyrethroids, and in this case, the synergism shown by the OP towards the pyrethroid appears due to an enhanced activation to the oxon form [115]. Organophosphorus compounds (disulfoton and fenthion) are also activated by thioether oxidation (formation of sulphoxide and sulfone). The metabolism of pyrethroid by P450 enzymes is well studied in insects. Hydroxylations and further metabolism make pyrethroid metabolism and has been noticed for the single enantiomer of deltamethrin (Figure 3).


Figure 3.

Metabolism of deltamethrin by insect P450 enzymes: (1) deltamethrin; (2) 4′ hydroxydeltamethrin; (3) trans-hydroxymethyl-deltamethrin; (4) cyano (3-hydroxyphenyl) methyl deltamethrate; (5) deltamethric acid. Adopted from Feyereisen, 2012.

The currently banned cyclodiene insecticides aldrin, heptachlor and isodrin are epoxidized by P450 enzymes to the environmentally stable toxic epoxides dieldrin, heptachlor epoxide and endrin, respectively [116]. Recombinant CYP6A1, -A2, -A8, -B8 and -B27; CYP12A1; and CYP321A1 can catalyse these epoxidations. Examples of pro-insecticide metabolism include the activation of chlorfenapyr by N-dealkylation [117] and diafenthiuron by S-oxidation [118]. In each case, the insect P450-dependent activation is a key in the selective toxicity of these pro-insecticides that target mitochondrial respiration. Recombinant housefly CYP6A1 catalyses the activation of chlorfenapyr (Figure 4).


Figure 4.

Chlorfenapyr and cypermethrin metabolism. The same P450 in Heliothis virescens probably activates the pyrrole and inactivates the pyrethroid, resulting in negative cross-resistance. Adopted from Feyereisen, 2012.

In H. virescens, the toxicity of chlorfenapyr is negatively correlated with cypermethrin toxicity [119]. The metabolism of imidacloprid is also of interest, particularly in relation to resistance. Piperonylbutoxide can synergize the toxicity of imidacloprid, and two P450 enzymes, CYP6G1 of D. melanogaster and CYP-6CM1vQ of Bemisia tabaci, have been shown to metabolize this neonicotinoid [120, 121] (Figure 5). Hydroxylations at the 4 and 5 positions can lead to the olefinic metabolite or to the dihydroxylated metabolite. In the whitefly, the 5-hydroxy metabolite is not toxic, but the 4-hydroxy metabolite is as toxic as the parent compound, so region selectivity may be of importance.

Despite the continuous use of insecticides, there are repeated failures in controlling the sucking insect pest species in recent years [11, 21] in different conventional tea plantations of Terai, the Dooars and Darjeeling foothill regions. Such a failure occurs due to changes in the susceptibility level of the pest species to the applied insecticides. Susceptibility level changes mainly due to metabolic detoxification of the insecticides through higher level of activity of some insecticide detoxifying enzymes under the stress of different management practices [8, 10, 11, 22]. In another mirid pest, Lygus lineolaris, metabolic resistance to insecticides due to enhanced level of activity has been reported by many authors [177, 178].


Figure 5.

Metabolism of imidacloprid by insect P450 enzymes: (1) imidacloprid; (2) 5-hydroxyimidacloprid; (3) 4-hydroxyimidacloprid; (4) dihydroxyimidacloprid; (5) Non-enzymatically derived dehydroimidacloprid. Adopted from Feyereisen, 2005.

Species P450 Over-expressed Resistance pattern
Musca domestica CYP6A1OP, carbametes [122]
IGR [123, 124]
CYP6A5v2, CYP6A36Pyrethroids [125127]
CYP6D1, CYP6D3Pyrethroids [128130]
CYP6D1Pyrethroids [131]
CYP6D1, CYP6D3v2Pyrethroids [132]
CYP6A24Pyrethroids [133]
CYP12A1Pyrethroids [134]
CYP6A2DDT, malathion [135137]
CYP6A8Malathion [137, 138]
Malathion [137]
DDT [139]
CYP6G1DDT [140, 141]
DDT [142]
Lufenuron, propoxur [141]
Imidacloprid [141]
Imidacloprid [142]
DDT [143]
Diazinon [144]
CYP12D1/2DDT [143]
DDT [139]
CYP12A4Lufenuron [145]
CYP6G1DDT, imidacloprid, Malathion [139]
CYP6Z1Pyrethroids [146]
CYP325A3Pyrethroids [147]
CYP6M2, CYP6P3Pyrethroids [148]
CYP6P3Permethrin [149]
CYP6M2, CYP6Z2Permethrin [150]
CYP4C27, CYP4H15DDT [151]
CYP6Z1,2, CYP12F1, CYP314A1DDT [147]
A. stephensi CYP325C1Pyrethroids [152]
A. funetus CYP6P4, CYP6P9Pyrethroids [153, 154]
Aedesaegypti CYP9J10,27,32Pyrethroids [155]
Culex pipien squinquefasciatus CYP9M10Permethrin [156]
CYP6F1Permethrin [157]
CYP4H34, CYP6Z10, CYP9M10Permethrin [158]
CYP4H21, H22, H23, CYP4J4, CYP4J6Deltamethrin [159]
Heliothis virescens CYP9A1Thiodicarb [160]
Helicoverpa zea CYP6B8,B9Cypermethrin [161]
H. armigera CYP4G8Pyrethroids [162]
CYP6B7Pyrethroids [163]
CYP6B7, CYP9A12, CYP9A14Pyrethroids [164]
CYP4S1, CYP337B1Fenvalerate [165]
CYP4L5,11, CYP4M6,7, CYP6AE11, CYP9A14, CYP332A1, CYP337B1Deltamethrin [166]
Plutella xylostella CYP6BG1 Cypermethrin [167]
CYP4M20Cypermethrin [168]
Lygus lineolaris CYP6X1Permethrin [169]
Bemisia tabaci CYP6CM1vQImidacloprid [170]
Nilaparvata lugens CYP6ER1Imidacloprid [171]
Myzus persicae CYP6CY3Neonicotinoids [172]
Diabrotica virgifera CYP4Me-parathion [173]
Tribolium castaneum CYP6BQ8,9,10, CYP436B1, B2Deltamethrin [174]
Blattella germanica P450MA, Chlorpyriphos [175]
CYP4G19Pyrethroids [176]

Table 3.

Over-expressed CYP genes in insecticide-resistant strains.

In Western Flower Thrips, Frankliniella occidentalis, metabolic detoxification of insecticides has been reported by many authors [179]. In Bemisia tabaci, metabolic resistance due to enhanced activity of insecticide resistance-related enzymes has also been reported [180].

7. Host allelochemicals, induction of detoxifying enzymes and insecticide resistance

Understanding the diversity of insect responses to chemical pressures (plant allelochemicals and insecticides) in their local ecological context represents a key challenge in developing sustainable pest control strategies. Plants and insects have had co-existing relationships for a long time. Insects were suppressed either by other insects or toxins or by plant defence mechanisms in order to create a balance between the insect pest population and host. Each plant species has a unique set of defence traits ranging from morphological to phytochemical parameters that have behavioural and physiological ramifications for a potential herbivore consumer [181, 182]. Therefore, the resistance mechanisms evolved by insects to deal with the chemical defences of plants are similar to those mechanisms that have evolved to resist synthetic insecticides. The chemical structure of some synthetic insecticides is comparable to that of some plant-produced compounds (e.g. pyrethroids and nicotinoids). Insect resistance to plant allelochemicals interferes with their resistance to synthetic insecticides [183]. From the evolutionary perspective, despite the key role of the chemical ‘arms race’ in driving the co-evolution of plants and insects, much research has focused so far on describing the diversity of plant chemicals and their effects on herbivores. Hence, the understanding of insecticide resistance mechanisms as well as taking into account other ecological parameters is important in predicting the spread of insecticide resistance in natural pest populations and in choosing the optimum strategy for managing insect pest populations. Less is known about the multiple mechanisms evolved by insects to overcome these chemical defences (Table 4). These mechanisms include contact and ingestion avoidance, excretion, sequestration, degradation of the toxin and target site mutation.

Plant allelochemicals Target (mechanism of effect) Resistance mechanisms Species
AlkaloidsNeuroreceptors (inhibition), ion channels (antagonists), nucleic acids (disruption of DNA synthesis), feeding (deterrent owing to bitterness), enzymes (inhibition)Modification of nicotine synthesis by salivary glucose oxidase Helicoverpa zea (Lep.) [184]
CardenolidesNervous system (depressing activity); Na+, K+-ATPase (specific inhibitor)Canal trenching behaviour,
target site mutation
Danaus plexippus (Lep.) [185]
Chrysochus sp. (Col.) [186]
Cyanogenic glycosidesElectron transport (inhibition of mitochondrial cytochrome oxidase)Ingestion avoidance,
sequestration and detoxification
Schistocerca americana (Ort.),
Hypera brunneipennis (Col), Zygaena sp. (Lep.), Clossiana euphrosyne (Lep.), Heliconius sara (Lep.) [187]
GlucosinolatesRespiration (inhibition)Detoxification by GSTs,
detoxification by glucosinolate sulphatase, formation of nitriles instead of isothiocyanate detoxification by P450s, detoxification by N-oxidation and sequestration
Myzus persicae (Hem.) [188] Plutella xylostella (Lep.) [189]
Pieris rapae (Lep.) [190]
Drosophila melanogaster [191]
Estigmene acrea (Lep.) [192]
Tyria jacobaeae (Lep.) [193]
Flavonoids and phenolic acidsRespiration (inhibition),
growth (inhibition)
Ingestion avoidance,
decrease of toxin levels in gall tissue, glycosylation by UDP-glycosyl transferase, sequestration and/or excretion
Manduca sexta (Lep.) [194]
Potania sp. (Hym) [195]
Bombyx mori (Lep.) [196]
Iridoid glycosidesFeeding (deterrent owing to bitterness), nucleic acids (inhibition of DNA polymerase), proteins (denaturant and cross-linking activities)Sequestration Longitarsus sp. (Col.) [197]
Coumarins and furanocoumarinsNucleic acids (photoactivie DNA bonding),
pro-oxidant activity
Detoxification by P450s,
detoxification by GSTs
Papilio polyxenes (Lep.) [198]
Depressaria pastinacella [199]
Spodoptera frugiperda (Lep.) [200]
Protease inhibitorsDigestive system (inhibition of protease)Over-expression of insensitive protease Callosobruchus maculatus (Col.) [201]
TerpinoidsNervous system (inhibition of acetylcholine esterases); feeding (deterrent owing to physical barrier and bitterness); growth and development inhibitor (pheromone analogue)Repression of genes involved in biosynthetic pathways Spodoptera exigua (Lep.) [202]
TanninsFeeding (complexation of salivary and gut proteins); pro-oxidant activitySynthesis of anti-oxidant compounds Orgyia leucostigma (Lep.) [203]

Table 4.

Plant allelochemicals and associated resistance mechanisms in insects.

[i] - Lep., Lepidoptera; Col., Coleoptera; Ort., Orthoptera; Hym., Hymenoptera; Dip., Diptera; Hem., Hemiptera.

Biotransformation of plant toxins is one of the major weapons that insects have evolved in their co-evolutionary arms race with plants [204]. To date, metabolic resistance to plant chemicals has been identified not only in herbivorous insects [194] but also in detrivorous insects such as mosquito larvae feeding plant debris [205]. Metabolic resistance often results from the overproduction of ‘detoxification enzymes’ that can metabolize plant xenobiotics (allelochemicals). This mechanism is often associated with phenotypic plasticity, as the production of detoxification enzymes is usually induced by the presence of plant xenobiotics in the diet of the insect.

Induction of insect detoxifying enzyme activities by plant allelochemicals is a clear manifestation of biochemical phenotypic plasticity and has been documented in several instances [206, 207]. Many of the theories and some of the experiments implicitly or explicitly deal with the insect’s ability to metabolize plant secondary substances by P450 and other enzymes. In those studies, a ‘higher activity of midgut microsomal oxidase enzymes in polyphagous than in monophagous species indicates that the natural function of these enzymes is to detoxify natural insecticides present in the larval food plants’. The estimation of aldrin epoxidation in gut homogenates of last instar larvae from 35 species of Lepidoptera showed that polyphagous species had on average a 15 times higher activity than monophagous species. This trend was seen in sucking insects as well. A 20-fold lower aldrin epoxidase activity was found in the oleander aphid Aphis nerii (specialist feeder on two plant families, Asclepiadaceae and Apocynaceae) when compared to the potato aphid Myzus euphorbiae or to the green peach aphid Myzus persicae (both are generalists found on 30–72 plant families) [208]. A similar type of observation was made for other detoxification enzymes. In mites, predatory mite has a five times lower aldrin epoxidase activity than its herbivorous prey [209]. The toxicity of the natural phototoxin α-terthienyl is inversely proportional to the level of its metabolism in Lepidoptera and is related to diet breadth. Metabolism is highest in Ostrinia nubilalis, which feeds on numerous phototoxic Asteraceae; lower in Helicoverpa virescens, which has a broad diet, including some Asteraceae that are non-phototoxic; and lowest in Manduca sexta, a specialist of Solanaceae [210].

In addition to insecticides, insect carboxylesterases also metabolize many glycosides. Β-glucosidase enzyme is active towards a variety of glucosides in fall armyworms, corn earworms, cabbage loopers and valvet bean caterpillars. The p-nitrophenyl β-D-glucoside, 4-methyl umbelliferyl β-D-glucoside, D (+)-cellobiose, D-amygdalin and helicon were preferred substrates whereas sinigrin, phloridzin, α-solanine, tomatine and linamarin were poor substrates for these insects and many other insects reported to date [211]. β-Glycosidases have been shown to play important roles in the survival of certain phytophagous insects [211]. The ability of peach tree borer, Synanthedon exitiosa, larvae to survive well on prunasin-containing peach tree is because they can metabolize cyanogenic glycosides through β-glucosidase and detoxify the released cyanide by β-cyanoalanine synthase, thereby allowing them to utilize peach trees [212]. Another example is the larvae of the tiger swallowtail, Papilio glaucus, which feeds on quacking aspen, which contains various phenolics glycosides (e.g. salicortin). These larvae hydrolyse the glycosides by β-glucosidase and detoxify the released phenolics aglycone by a highly active esterase, thereby allowing them to survive on aspen [213, 214].

Glutathione S-transferases are also involved in the metabolism of many toxic plant allelochemicals. These plant allelochemicals may be of many diverse groups including α, β-unsaturated carbonyl compounds (e.g. trans-cinnamaldehyde, trans-2-hexanal), isothiocyanates (e.g. allylisothiocyanate, benzyl isothiocyanate) and organothiocyanates (e.g. benzyl thiocyanate) as have been documented in some instances [41]. The glutathione S-transferase activities are lower in the specialist insects than in the generalists. In the crucifer-adapted cabbage looper for the metabolism of isothiocyanates (plant allelochemical), the activity of this enzyme wasfound to be two- to sixfold higher than that in the fall armyworm [215, 216]. These findings strongly advocate that glutathione S-transferases play an important role in developing resistance towards plant allelochemicals in phytophagous lepidopteran insects [215217]. Many plant allelochemicals are potent inhibitors of glutathione S-transferases in many insects [218]. Many flavonoids, other phenols and α-, β-unsaturated carbonyl compounds are also found to be potent inhibitors of the enzymes.

H. theivora, E. flavescens and S. dorsalis all are polyphagous in nature. H. theivora known to feed on at least sixteen different plant families reported till date [11]. Similarly, E flavescens is also polyphagous [11]. S. dorsalis has been documented to attack more than 150 hosts from at least 40 different plant families [219]. Hence, these pests are exposed to a wide variety of plant allelochemicals of diverse groups having the potential to induce the activity of these resistance-related enzymes. A higher level of detoxifying enzyme activity in H. theivora has been reported when reared on two alternative hosts, i.e. Mikania micrantha (Asteraceae) and Psidium guajava (Myrtaceae), than on tea [220]. Over the four hundred million years of co-evolution with plants, phytophagous insects have developed diverse resistance mechanisms to cope with plant chemical defences. Because insects face a geographical mosaic of chemical environments, from non-toxic to highly toxic plants, the costs associated with resistance traits vary with the probability of encountering a toxin. Moreover, other selection pressures, such as the presence or absence of competitors and predators, can also influence the costs and selection of particular resistance traits. Thus, the complexity of the local community composition is a key factor in maintaining the diversity of adaptive mechanisms to plant xenobiotics. These mechanisms are more plastic and complex compared with those involved in resistance to insecticides, perhaps because environments in which insecticides are heavily used also tend to have communities of low diversity and complexity. However, because some detoxification enzymes are involved in plant toxins and insecticides metabolism, cross-resistance mechanisms can be predicted to be observed under specific environmental conditions. Deciphering the impact of allelochemicals in cross-resistance mechanisms with insecticides at a local scale, and comparing the molecular and evolutionary mechanisms of resistance to phytotoxins and synthetic insecticides, represent promising areas of research for developing long-term sustainable insect control strategies for the effective management of pest concern [220].

8. Genetics and insecticide resistance in tea pests

Earlier, common visible markers including mophometrices, eye colour, body spots or bands and hairs or spines, wing venation were used as phenotypic markers in studying the pattern of dispersal, mating behaviour, population variability and inheritance of genetic traits in insects [221, 222]. Although the phenotypic markers are found at all times of life span of the organism and can be readily used for studies in field conditions, they suffer from many practical limitations. The major drawback is that these visible phenotypes are relatively infrequent and often hard to score. Because the phenotype markers are rare, use of these markers in mapping a trait is difficult. For all such difficulties and with the concurrent advancement in biochemical methodologies, protein markers then became more popular. Protein markers made a significant contribution in the early periods when DNA technologies were not so much advanced, as it is now [223]. A diverse range of novel molecular (DNA) markers are now available for entomological investigations. Currently, both DNA and protein markers have revolutionized the biological sciences and have enhanced many fields of insect study, especially agricultural entomology [224].

Insecticide resistance is the result of an increase in the ability of individuals of an insect species to survive insecticide application and is an important example of man-driven evolution [225]. Alleles conferring resistance may arise and spread in populations and to other populations with variable success, depending on factors such as selective forces, genetic variability, gene flow, population size and environmental conditions [220]. Studies that map the population structure of pest insects, as well as the potential for gene flow between populations, are needed to understand the development of resistance and prevention of its spread [226, 227]. Development of resistance is often rapid in isolated populations that have been treated by insecticides [228]. The rate of development of insecticide resistance may, however, be influenced by gene flow between treated and untreated populations by maintaining the frequency of resistance alleles at a low level [229]. Contaminant exposure was a poor predictor of population structure and the level of gene flow was a better predictor of relatedness [230]. Gene flow may balance divergence by opposing the effect of selection pressures [229]. Population genetic patterns should therefore be investigated with reference to geographical variability, as well as selection pressure. Detoxification resistance occurs when enhanced levels or modified activities of biotransformation enzymes prevent the insecticide from acting on its site of action because the metabolites produced have little or no activity compared with the original substance [231]. These changes may be due to mutations resulting in a protein with slightly different properties or altered expression.

As chemical control is frequently used to avoid economic damage, the sucking insects have been subjected to major selection pressure. Insecticides will probably continue to be the main control method in the near future and therefore it is important to study the structure of sucking insect population and change in insecticide susceptibility. There are several techniques for estimating the genetic diversity such as randomly amplified polymorphic DNA analysis, microsatellites, minisatellites, restriction fragment length polymorphism analysis and amplified fragment length polymorphism (AFLP) analysis. DNA markers are also suitable for use with small amounts of insect material and can be used with stored, dry or old samples. Some have complex multi-locus banding patterns, which may be of a non-Mendelian nature (e.g. randomly amplified polymorphic DNAs (RAPDs)). They have an expanding range of applications, many involving intra- and interspecific discriminations.


The author expresses his sincere thanks to the Head, Department of Zoology, University of North Bengal, for providing laboratory space. The author also expresses sincere appreciation to those scientists and authors based on whose concept, hypothesis and work this chapter has been developed. The author also expresses his thanks to the University of North Bengal for providing uninterrupted local area network that has immensely helped in searching and collecting information. The author also expresses his sincere thanks to InTech Open Access Publishing Group, editor of the book ‘Insecticide Resistance’ Prof. Stanislav Trdan (University of Ljubljana, Biotechnical Faculty, Dept. of Agronomy, Chair for Phytomedicine, Agricultural Engineering, Crop Production, Pasture and Grassland Management, Slovenia) and Ms Sndra Bakic (InTech Europe, Rijeka, Croatia).


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