Major insect and mite pests that occur on tea in India.
Polyphagous insect herbivores encounter numerous toxins (xenobiotics) as they pass through their life cycle; some toxins are produced naturally by the host plants (allelochemicals) and others by humans (insecticides) to manage these insects having pest status. The host plants have evolved defensive mechanisms for protection from herbivory, including chemical repellents and toxins (secondary metabolites). Many classes of insect repellents and toxic substances, such as isoflavonoids, furanocoumarins, terpenoids, alkaloids and cyanogenic glycosides are synthesized in plants. The biosynthetic pathways leading to these allelochemicals are continually evolving to generate new secondary metabolites. Similarly, to control the herbivorous insect pests, numerous chemicals of synthetic origin are used continuously against them. In response, the attacking organisms also evolve mechanisms that enable them to resist the defensive chemicals of their hosts and those toxins of synthetic origin applied for their control. A variety of defence mechanisms, including enzymatic detoxification systems, physiological tolerance and behavioural avoidance, protect insect herbivores from these xenobiotic compounds. Insect pests have evolved the mechanisms to degrade metabolically (enzymatically) or otherwise circumvent the toxic effect of many types of chemicals that we have synthesized as modern insecticides. The extent to which insects can metabolize and thereby degrade these antibiotics or toxins is of considerable importance for their survival in hostile chemical environment. These mechanisms continue to evolve as insects attempt to colonize new plant species or encounter newer molecules of synthetic insecticides. Generally, three main enzymes, general esterases (GEs), glutathione S-transferases (GSTs) and cytochrome P450-mediated monooxygenases (CYPs), are involved in the process of metabolic detoxification of insecticides. During the past 70 years, following the discovery and extensive use of synthetic insecticides, resistance of insects to insecticides has registered the greatest increase and strongest impact. The evolution of resistance to insecticides is an example of evolutionary process. An insecticide is the selection pressure, which results in a very strong but differential fitness of the individual in a population having susceptible and resistant genotypes. The survival and subsequent reproduction of resistant individuals lead to a change in the frequency of alleles conferring resistance in the population over time. While selection pressure acts to change allele frequencies within pest populations, the phenotype upon which selection operates is a function of both genotype and the environment. Recent studies in insect detoxifying enzymes have revealed further versatility in the adaptation of insects to their environment by the phenomenon of induction. This is the process in which a chemical stimulus enhances the activity of the detoxification enzyme systems by the production of additional enzymes that metabolize toxic chemical substances. Hence, the influence of environmental factors such as continuous usage of insecticides and the chemical constituents (allelochemicals) of host plants on phytophagous insects can have a great impact to induce the enzymatic detoxification systems of insects, thereby promoting the insecticide resistance mechanisms. While all insects do possess detoxification ability, its magnitude is expected to vary among the species with the nature of its recent environment and feeding ecology. The level and type of detoxifying mechanisms differ greatly, which therefore result in varying toxicity among different developmental stages, species and populations. Variation in detoxifying enzyme activity is responsible in part for the selective toxicity of different insecticides, the development of resistance to insecticides and selective adaptation to host plants. Over-expression of these detoxifying enzymes, capable of metabolizing insecticides, can result in a high level of metabolic tolerance/resistance to synthetic insecticides. Increased expressions of genes encoding the major xenobiotic metabolizing enzymes are the most common cause of insecticide resistance in insects.
- Insecticide resistance
- insect pests
- detoxifying enzymes
- cytochrome p450
- glutathione S-transferases
- Helopeltis theivora
- Scirtothrips dorsalis
- Empoasca flavescens
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,
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,
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 . Among the insect pests, 32% are from order Lepidoptera, followed by 27% pest species from Hemiptera . In India, only 300 species of arthropods are recorded as pest and about 167 species are from tea-growing Northeast India (Table 1) . 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 , causing on average a 5%–55% yield loss [4–6]. Insect pests alone can cause on an average 11%–55% yield loss, if left unrestrained .
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 (
||Bunch caterpillar||Folivores||Bombycidae: Lep.|
||Red slug caterpillar||Folivores||Zygaenidae: Lep.|
||Looper caterpillar||Folivores||Geometridae: Lep.|
||Black inch worm||Folivores||Geometridae: Lep.|
||Sungma caterpillar||Folivores||Lymantridae: Lep.|
||Tea leaf roller||Folivores||Gracilariidae: Lep.|
||Cockchafer grubs||Root of young tea||Scarabaeidae: Col.|
||Leaf-eating cockchafer||Leaf eater||Scarabaeidae: Col.|
||Large green weevil||Leaf eater||Curculionidae: Col.|
||Tea leaf miner||Leaf eater||Agromyzidae: Dip.|
||Live wood-eating termite||Stem and root eater||Termitidae: Iso.|
||Scavenging termite||Stem and root eater||Termitidae: Iso.|
||Tea mosquito bug||Leaf sucker||Miridae: Hem.|
||Tea greenfly/tea jassid||Leaf sucker||Jassidae: Hem.|
||Tea aphid||Leaf sucker||Aphididae: Hem.|
||Yellow tea thrips||Leaf and bud sucker||Thripidae: Thy.|
||Common thrips||Leaf and bud sucker||Thripidae: Thy.|
||Red spider mite||Leaf sucker||Tetranychidae: Aca.|
||Scarlet mite||Leaf sucker||Tenuipalpidae: Aca.|
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) . 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 , outbreak of secondary pests , development of insecticide resistance [10, 11], including undesirable residues in made tea . 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
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 . 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, 19–21]. A major concern in managing the major insect pests of tea is its high potential to develop resistance rapidly to regularly used insecticides . 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 . Recently, there are reports on the development of resistance to many commonly used synthetic insecticides and consequent failure in controlling many tea pests [10, 22–26]. 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 [19–21, 26]. The development of resistance in
For the management of other sucking insects such as yellow tea thrips,
Similarly, in another emerging sucking insect pest of tea, tea greenfly,
A high level of insecticide resistance in folivores, such as black hairy caterpillar, bunch caterpillar, looper pest complex (
Detoxification of insecticides is an important toxicokinetic mechanism for insect pests to tolerate regularly applied insecticides [8, 40–42]. 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 [43–45].
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 . 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  These mechanisms also cross-protect insect pests when they are exposed to synthetic insecticides . Herbivorous insect groups (agricultural pests) are significantly more diverse than their non-herbivorous sister groups . The role of plant in promoting diversification in insects has occurred through co-evolutionary ‘defence strategies’ among them . 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 . 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 .
Among sucking insect herbivores, the actual food used, i.e. digested whole tissue particularly parenchyma (as in
Insecticide resistance is a genetic change in response to selection pressure of toxicants that impair pest control in the field . 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 .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:
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 ‘
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 . 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,
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 . 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 18.104.22.168) 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) . Alteration at the target site, to less a sensitive target for neurotoxic insecticides, is an important toxicodynamic resistance mechanism in insects .
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 . 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 . 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 . The presence of insensitive AChE conferring resistance was first noticed in OP-resistant mites,
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).
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 .
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
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
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 . 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).
O2 to H2O, H2O2, O2 −
|CYP6A1 (and probably most P450 enzymes)|
|CYP4C7, CYP6A1, CYP6A2, CYP6A8, CYP6G1, CYP6M2, CYP6CM1vQ, CYP9T2, CYP12A1, CYP18A1, CYP302A1 CYP306A1, CYP312A1, CYP314A1, CYP315A1|
|O-dealkylation||CYP6A1, CYP6D1, CYP6A5, CYP6B4, CYP6B17, CYP6B21, CYP6G1, CYP6Z2, CYP6CM1vQ, CYP9A12, CYP9A14, CYP12A1, CYP321A1,|
|Epoxidation||CYP6A1, CYP6A2, CYP6B8, CYP6B27, CYP6AB3, CYP6 CYP6AB11, CYP9E1, CYP12A1, CYP15A1, CYP321A1|
|Aromatic hydroxylation||CYP6D1, CYP6G1, CYP6M2|
|Heteroatom oxidation and dealkylation
Phosphorothioate ester oxidation
|CYP6A1, CYP6A2, CYP6D1, CYP12A1|
|Complex and atypical reactions
Cyanogenic glucoside biosynthesis:
Val/Ile to oximes
Oximes to cyanohydrins
Aryl ether cleavage
Decarbonylation with C–C cleavage
The first insect P450 cloned and sequenced was CYP6A1 from
5.1.2. Carboxylesterases (EC 22.214.171.124)
Carboxylesterase or esterase is a collective term for the enzymes that hydrolyse carboxylic esters . Classification of these enzymes is difficult because of their overlapping substrate specificity . However, the esterase classification of Aldridge  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 . 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 .
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 . Phosphatases are A-esterases that detoxify many organophosphorous insecticides especially phosphates in insects. In houseflies, paraoxon can be hydrolysed to diethyl phosphoric acid and
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 126.96.36.199.)
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 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 . 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 , 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
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 . On the contrary, parathion can be de-arylated by glutathione S-transferases to produce diethyl phosphorothioic acid and S-(
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 . 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 . 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).
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 . Similarly, permethrin resistance in horn flies is suppressible by piperonyl butoxide, and negatively related to diazinon toxicity . In
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 . 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  and diafenthiuron by S-oxidation . 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).
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,
||CYP6A1||OP, carbametes |
|IGR [123, 124]|
|CYP6A5v2, CYP6A36||Pyrethroids [125–127]|
|CYP6D1, CYP6D3||Pyrethroids [128–130]|
|CYP6D1, CYP6D3v2||Pyrethroids |
||CYP6A2||DDT, malathion [135–137]|
|CYP6A8||Malathion [137, 138]|
|CYP6G1||DDT [140, 141]|
|Lufenuron, propoxur |
||CYP6G1||DDT, imidacloprid, Malathion |
|CYP6M2, CYP6P3||Pyrethroids |
|CYP6M2, CYP6Z2||Permethrin |
|CYP4C27, CYP4H15||DDT |
|CYP6Z1,2, CYP12F1, CYP314A1||DDT |
||CYP6P4, CYP6P9||Pyrethroids [153, 154]|
|CYP4H34, CYP6Z10, CYP9M10||Permethrin |
|CYP4H21, H22, H23, CYP4J4, CYP4J6||Deltamethrin |
|CYP6B7, CYP9A12, CYP9A14||Pyrethroids |
|CYP4S1, CYP337B1||Fenvalerate |
|CYP4L5,11, CYP4M6,7, CYP6AE11, CYP9A14, CYP332A1, CYP337B1||Deltamethrin |
||CYP6BQ8,9,10, CYP436B1, B2||Deltamethrin |
In Western Flower Thrips,
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 . 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.
|Alkaloids||Neuroreceptors (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||
|Cardenolides||Nervous system (depressing activity); Na+, K+-ATPase (specific inhibitor)||Canal trenching behaviour,
target site mutation
|Cyanogenic glycosides||Electron transport (inhibition of mitochondrial cytochrome oxidase)||Ingestion avoidance,
sequestration and detoxification
|Glucosinolates||Respiration (inhibition)||Detoxification by GSTs,
detoxification by glucosinolate sulphatase, formation of nitriles instead of isothiocyanate detoxification by P450s, detoxification by N-oxidation and sequestration
|Flavonoids and phenolic acids||Respiration (inhibition),
decrease of toxin levels in gall tissue, glycosylation by UDP-glycosyl transferase, sequestration and/or excretion
|Iridoid glycosides||Feeding (deterrent owing to bitterness), nucleic acids (inhibition of DNA polymerase), proteins (denaturant and cross-linking activities)||Sequestration||
|Coumarins and furanocoumarins||Nucleic acids (photoactivie DNA bonding),
|Detoxification by P450s,
detoxification by GSTs
|Protease inhibitors||Digestive system (inhibition of protease)||Over-expression of insensitive protease||
|Terpinoids||Nervous 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||
|Tannins||Feeding (complexation of salivary and gut proteins); pro-oxidant activity||Synthesis of anti-oxidant compounds||
Biotransformation of plant toxins is one of the major weapons that insects have evolved in their co-evolutionary arms race with plants . To date, metabolic resistance to plant chemicals has been identified not only in herbivorous insects  but also in detrivorous insects such as mosquito larvae feeding plant debris . 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
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
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.
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 . 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 .
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 . 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 . 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 . 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 . Contaminant exposure was a poor predictor of population structure and the level of gene flow was a better predictor of relatedness . Gene flow may balance divergence by opposing the effect of selection pressures . 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 . 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 ‘
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