Insect Chitin: Metabolism, Genomics and Pest ...

Provided for non-commercial research and educational use only. Not for reproduction, distribution or commercial use. This chapter was originally published in the book Advances in Insect Physiology, Vol. 43, published by Elsevier, and the attached copy is provided by Elsevier for the author's benefit and for the benefit of the author's institution, for non-commercial research and educational use including without limitation use in instruction at your institution, sending it to specific colleagues who know you, and providing a copy to your institution’s administrator.

All other uses, reproduction and distribution, including without limitation commercial reprints, selling or licensing copies or access, or posting on open internet sites, your personal or institution’s website or repository, are prohibited. For exceptions, permission may be sought for such use through Elsevier's permissions site at: http://www.elsevier.com/locate/permissionusematerial From: Daniel Doucet, Arthur Retnakaran, Insect Chitin: Metabolism, Genomics and Pest Management. In Tarlochan S. Dhadialla, editor: Advances in Insect Physiology, Vol. 43, Burlington: Academic Press, 2012, pp. 437-511. ISBN: 978-0-12-391500-9 © Copyright 2012 Elsevier Ltd. Academic Press

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CHAPTER SIX

Insect Chitin: Metabolism, Genomics and Pest Management Daniel Doucet, Arthur Retnakaran Great Lakes Forestry Centre, Canadian Forest Service, Sault Ste. Marie, Canada

Contents 1. Introduction 1.1 Chitin structure, types, distribution in fungi and animals 1.2 Evolution of chitin from prehistoric to present; physiology and function in arthropods 1.3 Chitin in insects, distribution, architecture and sculpturing 2. Chitin Metabolism and Potential Targets for its Disruption 2.1 Chitin biosynthesis, enzymes in the pathway and chitin synthase 2.2 Chitin degradation 2.3 b-N-acetylglucosaminidases 3. Chitin Genomics 4. Benzoylphenyl Ureas as CSIs 4.1 History of development 4.2 Classification and structure of benzylphenyl ureas, structure–activity relationships 4.3 MOA and receptor 4.4 Effects on pests 4.5 Effects on non-target species 4.6 Environmental degradation 5. Non-Benzoylphenylurea Chitin Synthesis Inhibitors 5.1 Structure and properties 5.2 Environmental fate and effects 6. Chitinases and Chitinase-Inhibiting Chemicals for Pest Management 6.1 Fungi and microorganisms with chitinase activity 6.2 Baculoviruses with chitinase gene 6.3 Chemical inhibitors of chitinase 7. Resistance and Resistance Management 8. Conclusions and Future Development Acknowledgements References

Advances in Insect Physiology, Volume 43 ISBN 978-0-12-391500-9 http://dx.doi.org/10.1016/B978-0-12-391500-9.00006-1

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2012 Elsevier Ltd. All rights reserved.

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Daniel Doucet and Arthur Retnakaran

Abstract Chitin is a polymer of N-acetyl glucosamine that forms the protective exoskeleton of all arthropods and is replaced periodically during growth and development. Chitin biosynthesis starts with the disaccharide trehalose, culminating in the polymerization of the N-acetyl glucosamine subunits by chitin synthase to produce chitin microfibrils. Chitin in the old exoskeleton is degraded by chitinases, deacetylases and hexosaminidases and recycled. Chitin synthesis has been used as a target for developing biorational insecticides such as benzoylphenyl ureas, diflubenzuron being the original such compound. Several benzoylphenyl ureas with diverse activity spectra have since been synthesized and widely used for pest control. Newer pesticides targeting not only chitinase and chitin synthase but also other novel sites are being developed. Understanding the various nuances of chitin metabolism and regulation with all the genomic resources on hand will undoubtedly pave the way for developing more target-oriented softer control agents that have minimal impact on the environment.

1. INTRODUCTION In 1811, Henri Braconnot, a French biochemist, described a polysaccharide from mushrooms and called it “fungine” which was later found to be present in insects by Odier (1823) who gave it a more descriptive name, “chitin”, based on the Greek word “chiton” due to its functional resemblance to an envelope or tunic, a name which has received widespread acceptance (Braconnot, 1811; Muzzarelli et al., 2012). Next to cellulose, chitin is the most abundant organic substance present on earth. It acts as a scaffold on which various extracellular matrices are built. Cuticular body walls of arthropods, peritrophic membrane lining of the midgut of insects and cell walls of fungi are some of the major locales of chitin. Chitin has been detected across the board from the simplest algae such as the diatoms to the complex vertebrates such as the blenny fish (Wagner et al., 1993). It is present in sponges, corals, nematodes, molluscs, tubes of worms and even some rhizobial bacteria that secrete a fatty acid-linked chitin oligomer (Ehrlich et al., 2007a; Gaill et al., 1992; Jua´rez-de la Rosa et al., 2012; Merzendorfer, 2011; Spaink, 1994). Only plants, echinoderms and higher chordates appear completely devoid of chitin. In arthropods, chitin is extensively used to construct an exoskeleton that functions as a shield of armour. Even though chitin has been exhaustively studied, certain aspects still remain unresolved. In this chapter, we will try to address various aspects of chitin physiology, biochemistry and genomics as we understand it today. We believe such knowledge will not only help


Insect Chitin: Metabolism, Genomics and Pest Management

understanding how chitin synthesis inhibiting (CSI) insecticides manifest their action but also help explore new targets for intervention with new chemistries and new biotechnology approaches. The physiology of chitin has been elegantly reviewed in the past by many; chief among them are Cohen (1987, 2010), Muzzarelli (1977, 2010), Khoushab and Yamabhai (2010), Lee et al. (2011), Merzendorfer (2006, 2011) and Muthukrishnan et al. (2012).

1.1. Chitin structure, types, distribution in fungi and animals Chitin is a linear amino polysaccharide polymer made up of b-1,4-N-acetylD-glucosamine (GlcNAc) units and exists as two different crystalline forms, a-chitin and b-chitin, while a third form, g-chitin, is a combination of the a and b forms (Lotmar and Picken, 1950; Rudall and Kenchington, 1973). a-Chitin chains are the most abundant form that are arranged in an antiparallel fashion, are very stable and are present in insect cuticles, shells of crabs, lobsters, shrimp and fungal cell walls (Carlstrom, 1957). b-Chitin occurs in diatoms, the pens of squid, the chaetae of annelids and the tubes secreted by tubeworms of the Siboglinidae family (Annelida). The chains are parallel and are less stable than a-chitin (Gardner and Blackwell, 1975). g-Chitin is rare and is present in the stomach of squid and in the cocoons of two genera of beetles. It is a combination of both a- and b-chitin with the two parallel chains arranged in one direction and the third in the opposite direction (Figs. 6.1 and 6.2; Jang et al., 2004; Muzzarelli, 1977; Rudall and Kenchington, 1973). The cell walls of many fungi such as Mucor are made up of not only a-chitin but also chitosan which is a deacetylated form of chitin made up of glucosamine units (Araki and Ito, 1974; Fig. 6.3).

a-Chitin Anti-parallel chains b-Chitin Parallel chains g-Chitin Two parallel and one anti-parallel chains

Figure 6.1 The three forms of chitin chains.



(c) (b) (a) 1620 cm-1

1600

(c) (b) (a) Amide I band

-OH

4500

4000

3500

3000

2500

Amide III

Amide II

2000

1500

1000

500

Wavenumber (cm-1)

Figure 6.2 Fourier Transform Infrared (FT-IR) Spectra of a, b and g chitin chains (a, b and c) isolated from crab shells, squid pens and lucanid beetle cocoons, respectively.

O

CH2OH O H OH

CH2OH O H OH

CH2OH O

O

O

OH

NH2

O

NH2 NHCOCH3

Chitosan CH2OH

CH2OH

O O

O

CH2OH

H OH

O

O

O OH

OH

O

H OH

CH2OH O

O NHCOCH3

OH

O

OH NHCOCH3

Cellulose

Chitin

Figure 6.3 Structure of chitin in relation to chitosan and cellulose.

1.2. Evolution of chitin from prehistoric to present; physiology and function in arthropods The origin of chitin can be traced to prehistoric times dating back 550 million years ago. Among the earliest living organisms, chitin has been shown to be present in diatoms and protozoans from which it has left its fossil imprint



Era

Period

Millions Diatoms of years Fungi ago (0.0)

Quartenary

1.6

Tertiary

66

Cretaceous

138

Jurassic

205

Triassic

240

Palaeozoic Permian

290

Cenozoic Mesozoic

Protozoa Cnidaria Porifera Mollusca

Arthropoda Annelida

Chitinozoa (Flask-like animals)

Pennsylvani 330 an Mississipian 360

Precambrian

Devonian

410

Silurian

435

Ordovician

500

Cambrian

570

2500

Figure 6.4 Fossil history of chitin-containing organisms (after Miller, 1991).

since pre-Cambrian times. During subsequent evolution, the ability to synthesize chitin was split along two lines, one leading to the fungi and the other to the major animal groups such as Porifera, Cnidaria, Arthropoda, Mollusca, Annelida, Nematoda, Rotifera and Brachiopoda among others (Durkin et al., 2009; Flanner et al., 2001; Miller, 1991). In many species of the latter taxa, the exoskeleton has seen the incorporation of minerals to reinforce the chitinous scaffolding. This is well illustrated in diatoms where the cell wall is strengthened by mineralization with silicon (Durkin et al., 2009). Likewise, glass sponges (class Hexactinellida) have a silica–chitin composite in their skeleton (Ehrlich et al., 2007b; Falini and Fermani, 2004). In many crustaceans and molluscs, the chitin in the exoskeleton is impregnated with calcium carbonate. The shells of molluscs are mineralized with calcium carbonate on a b-chitin scaffold and have been well preserved as fossils (Fig. 6.4; Scho¨nitzer and Weiss, 2007). Treating a developing mollusc with a CSI like nikkomycin results in a soft shell with total loss of rigidity (Falini and Fermani, 2004; Scho¨nitzer and Weiss, 2007). In the American Lobster Homarus americanus, amorphous calcium carbonate as well as crystalline calcite is incorporated in the exoskeleton, but in two different layers. The calcium carbonate fraction is



incorporated in chitin fibres oriented in a perpendicular fashion to the cuticle, while the calcite crystals are concentrated in the epicuticle. This arrangement is hypothesized to bestow impact and wear resistance to the exoskeleton (Al-Sawalmih et al., 2008). Fungal cell walls contain, along with chitin, various types of polysaccharides such as cellulose-like glycans to strengthen the cell wall (Adams, 2004; Durkin et al., 2009). In the case of insects, the crystalline chitin nanofibres are embedded in a matrix of proteins and polyphenols to provide strength and flexibility for protection yet allow for flight (Vincent and Wegst, 2004). The hardness of the exoskeleton, especially in beetles, is possible because of sclerotization, where several cuticular proteins are cross linked with orthodiphenols and their quinones (Sugumaran, 2011).

1.3. Chitin in insects, distribution, architecture and sculpturing Insects have virtually conquered almost all the ecological niches on earth, except perhaps the ocean. They have adapted themselves remarkably to every situation, moulding the cuticle to every need. The body wall of the insect is called the integument and consists of a non-cellular cuticle which is multilayered with the cellular epidermis underneath. The epidermis is made up of a single layer of cells (Fig. 6.5). Chitin fibrils are arranged in the form of mf

mv 0.1 mm

Lc pmp

mv

1 mm

Figure 6.5 Apical region of an epidermal cell of Calpodes larva showing microvilli (mv) with plasma membrane plaques (pmp) synthesizing lamellate cuticle (Lc) that form the chitin lamellae in the endocuticle. The inset shows a cross section of one microvillus (mv) with the cut ends of actin microfilaments (mf) (from Locke, 1991).



lamellae in the endocuticular region. The cuticle and the epidermis undergo a remarkable series of developmental changes during metamorphosis (Locke, 1991; Moussian, 2010; Vincent and Wegst, 2004). The structural and architectural differences of the cuticle are controlled and precisely regulated by the epidermal cells. These cells communicate with each other by fine filopodia, and the differences in the epidermal feet determine the size and shape of the cuticle they secrete above (Fig. 6.6; Locke, 2001). The regulation of the design and architecture of the cuticle, not only within an insect species but also between insects, rests with the underlying epidermal cell. Chitin synthesis, the synthesis of cuticular proteins, the sclerotization and formation of setae on the surface and pigmentation are some of the major processes that need to be precisely regulated from one end of the body plan to the other as well as between life stages. A polarized pattern of the surface design and sculpture must reside within the epidermal layer. How exactly this pattern is interpreted and quantitatively coordinated is not very well Cuticulin Epicuticle Chitin lamellae -Procuticle

Lamellate cuticle secretory zone

Plasma membrane plaque Actin filaments Microvillus

Synthesis and transport of cuticle precursors including chitin

Filopodia

N

Basement membrane

Epidermal feet

Figure 6.6 Schematic representation of the physiology of the epidermal cell. See text for details. (Based on Locke, 1991, 2001; Moussian, 2010.)



understood. For example, the cuticle in the head has to be thick and hard requiring a thick layer of chitin fibrils embedded in protein with heavy sclerotization and pigmentation. On the other hand, the wings should be thin and flexible with very little sclerotization. The elasticity or stiffness of various cuticular structures vary from being very stiff, such as in the elytra of beetles, to extremely elastic such as the wings, as reflected by the Young’s modulus being either high or low. The tensile strength of various cuticular structures ranges from being either rigid like the elytra of beetles, or elastic like the membranous wings. The degree of stiffness as measured by their Young’s modulus is very high for elytra, 90 Gpa (gigapascals), whereas for larval cuticle, it is about 0.01 Gpa (Vincent and Wegst, 2004). The manifestation of the surface sculpture design from the epidermal cell is transduced through the apical cell membrane which assumes different brush borders according to the signal it receives. The actin cytoskeleton immediately below the apical cell membrane assumes different forms based on the design signal. How exactly these changes transform into the type, shape and size of cuticle that is secreted is unclear. It appears intuitive to assume that polarity genes in the epidermis play an important role in the determination of cuticle morphology (Uv and Moussian, 2010).

2. CHITIN METABOLISM AND POTENTIAL TARGETS FOR ITS DISRUPTION Chitin can represent up to 60% of the dry weight in some insect species (Richards, 1978). This single fact illustrates the importance of this insect component for its survival as well as the huge demand in precursors required for chitin synthesis, mainly glucose, but also glutamine and UTP, along with massive amount of energy needed to set the biosynthetic process in motion. The bulk of chitin synthesis occurs for the most part in short spurts during each moult. Therefore, a rapid mobilization of the metabolic machinery behind chitin production is warranted, and it implies that intricate induction and termination controls exist, with ecdysone orchestrating the entire process all of which are vulnerable points for insect growth disruption.

2.1. Chitin biosynthesis, enzymes in the pathway and chitin synthase Eight enzymatic steps are needed to convert the disaccharide precursor, trehalose into chitin (Kramer et al., 1985). These steps, along with the inclusion of glutamine synthesis, are illustrated in Fig. 6.7. Chitin synthesis starts with



Trehalase

Glycolysis

Glucose

Trehalose

Hexokinase

Glucose-6P Gluc-6p isomerase

Fructose-6P

Glutamine synthase

Glutamate

Glutamine

Hexosamine pathway

GFAT

Glutamate Glucosamine-6P Acetyl CoA GNA

CoASH N-acetyl glucosamine-6P AGM

UTP

N-acetyl glucosamine-1P UAP

Chitin

PPi UDP N-acetyl glucosamine

Chitin synthase

Figure 6.7 Diagram of de novo biosynthesis of chitin. Trehalose serves as the primary carbon source for UDP-GlcNAc. Hemolymph trehalose is converted into alpha- and beta-D-glucose units by trehalase. Glucose enters the glycolytic pathway until its conversion into fructose-6-phosphate (fructose-6P) via the successive actions of hexokinase and glucose-6-phosphate isomerase (gluc-6P isomerase). Fructose-6P is then diverted towards the hexosamine pathway by the action of glutamine: fructose-6-phosphate aminotransferase (GFAT) to produce glucosamine-6-phosphate (Glc-6P). Glc-6P is acetylated by glucosamine-6-phosphate acetyltransferase (GNA), isomerised by N-acetylglucosamine phosphate mutase (AGM) and finally activated by UDPN-acetylglucosamine pyrophosphorylase (UAP). UDP-N-acetyl glucosamine units are finally converted into chitin polymers by chitin synthase.

the hydrolysis of trehalose into two units of D-glucose. Glycogen can also be a precursor for glucose, but pools are often considered too shallow to account for the majority of synthesized chitin (Zaluska, 1959). Glucose then briefly enters glycolysis to generate fructose-6-phosphate (F6P). F6P is subsequently diverted towards the hexosamine pathway to produce, via four steps, UDP-N-acetyl glucosamine (UDP-GlcNac), which will be ultimately polymerized into chitin. Glaser and Brown (1957) first demonstrated the link between the hexosamine pathway and chitin synthesis by showing that UDP-GlcNac acts as the glycosyl donor in cell-free preparations from Neurospora crassa. Observations that the silk worm moth, Bombyx mori, accumulates large amounts of both GlcNac and UDP-GlcNac in the moulting fluid



also suggested that these compounds serve as chitin precursors in insects (Zaluska, 1959; Zielinska and Laskowska, 1958). Candy and Kilby (1962) elegantly showed the presence of glycolytic and hexosamine pathway enzymes in locust wings but could not demonstrate the final transfer of GlcNac from UDP-GlcNac into chitin. Fristrom (1968) obtained similar results from Drosophila wing discs. The pathway was fully deciphered by Cohen and Casida (1980), Mayer et al. (1980), Turnbull and Howells (1983) and others, who confirmed that cell-free insect extracts are indeed able to catalyse chitin synthase activity. 2.1.1 Trehalose hydrolysis and trehalase Trehalose (a-1-D-glucopyranosyl-a-1-D-glucopyranoside) is the major hemolymph sugar in insects. In addition to being the primary precursor of chitin, it fulfils other key functional roles in the life of the insect, perhaps the most important one being an energy source for flight. It also serves as a cryoprotectant in insects for winter survival in temperate zones. A comprehensive review of trehalose biochemistry and functions in insects has been written by Thompson (2003). Consistent with its role as a carbon source for chitin, the hemolymph levels of trehalose become depleted soon after moulting (Howden and Kilby, 1960; Schmidt and Mathur, 1967). Trehalose hydrolysis is catalysed by a,a-trehalase (EC 3.2.1.28), which exists both as a soluble form and as a membrane-bound form (Forcella et al., 2010; Mori et al., 2009). Each enzyme form is encoded by a single gene, and the cDNAs for both the soluble trehalase (also called acid trehalase or Tre-1) and the membrane-bound trehalase (also called neutral trehalase or Tre-2) have been cloned in a few insect species (Chen et al., 2010; Lee et al., 2007; Mitsumasu et al., 2005; Silva et al., 2009; Su et al., 1993, 1994; Takiguchi et al., 1992; Tang et al., 2008; Tatun et al., 2008). Tre-1 and Tre-2 are typically highly expressed in the midgut. In the beet armyworm, Spodoptera exigua, expression of SeTre-1 and SeTre-2 was also observed in other chitin-producing tissues such as the trachea and the integument (epidermis). During the larval development of S. exigua, the expression level of the SeTre-1 transcript shows a distinct peak just before pupation. SeTre-2 expression is more complex, with peaks appearing in the midgut during the early part of the fourth larval instar and also just before the wandering stage. SeTre-2 also shows distinct pulses of expression in pupal fat body at d4 and d7 after pupation (Tang et al., 2008). The same tight association between Tre-1 expression and pupal moulting has been observed in

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B. mori (Mitsumasu et al., 2008). Tatun et al. (2008) recently demonstrated that in larvae of the bamboo borer Omphisa fuscidentalis soluble and membrane-bound gut trehalase activities differ markedly in their response to hormones. JH acid promotes diapause termination and pupation, accompanied by a tripling of soluble trehalase activity. 20E injection of the insect moulting hormone, 20-hydroxyecdysone (20E), causes a similar effect. Consistent with this increase in soluble trehalase activity, the expression of the OfTreh-1 gene increases after JH acid or 20E treatment. In contrast, membrane-bound trehalase activity and OfTreh-2 expression levels were completely unresponsive to both hormones. The localized functions of Tre-1 and Tre-2 in relation to chitin synthesis were also investigated further by gene knock-down experiments in S. exigua (Chen et al., 2010). Injection of SeTre-1 or SeTre-2 dsRNA into d1 fifth instar larvae resulted in 50–60% mortality, with SeTre-1 knock-down causing a slightly more potent effect. This difference was also mirrored at the phenotypic level, with a high proportion of the SeTre-1-RNAi-linked mortality occurring as larval–pupal intermediates, while the effect of SeTre-2 RNAi was delayed, the latter generating pupal–adult intermediates and adults that failed to eclose. The depression of epidermal chitin content was also stronger after SeTre-1 RNAi than SeTre-2 RNAi treatment, while SeTre-2 RNAi had a higher impact on midgut chitin content. These experiments highlight the critical role of Tre-1 in integumental chitin synthesis. The study by Chen et al. (2010) also provided hints that trehalose and/or glucose levels control the expression of genes further downstream in the chitin biosynthetic pathway. For instance, knocking down SeTre-1 reduces the transcripts of the glucose-6-phosphate isomerase (G6PI) and chitin synthase A (CHSA) genes. In contrast, SeTre-2 mRNA depletion reduces the transcripts encoding UDP-N-acetylglucosamine pyrophosphorylase (UAP) and CHSB. Direct injection of trehalose in the hemocoel was also found to stimulate UAP expression. Thus, the authors postulate that a complex crosstalk exists between glycometabolism and 20E to regulate chitin synthesis. 2.1.2 Glycolytic and hexosamine pathways Glycolytic enzymes are ubiquitous among living organisms. Hexokinase (EC 2.7.1.1) is the first of the two glycolytic enzymes by which glucose is directed towards chitin synthesis, the next one being G6PI (EC 5.3.1.9). Beyond early confirmations of their presence in insect tissue extracts, scant attention has been paid to these two enzymes (Candy and Kilby, 1962).



Hexokinase seems to accomplish double duty in the specific context of chitin production. It can first utilize de novo glucose and, albeit with a lower affinity, use glucosamine and convert it into glucosamine-6-phosphate (Moser et al., 1980). This would be important in the recycling of glucosamine pools generated by chitin degradation at moulting. In S. exigua, the transcription of the SeG6PI gene is upregulated by 20E, indicative of an increased need for chitin production. Double-stranded RNA-mediated inhibition of the S. exigua ecdysone receptor gene (SeEcR) in larvae decreases SeG6PI expression observed at pupation and, conversely, 20E injection boosts its expression (Yao et al., 2010). The hexosamine pathway is the route by which sugar units are converted into sugar nucleotides, before their final combination into structural polysaccharides, including chitin. Glutamine:fructose-6-phosphate aminotransferase (GFAT, EC 2.6.1.16) is the first and rate-limiting enzyme in the hexosamine biosynthetic pathway. It catalyses the conversion of F6P into glucosamine-6-phosphate and structurally is composed of a class II glutamine aminotransferase (GAT2) and two sugar isomerase (SIS) motifs. A substantial body of knowledge exists on the importance of this enzyme in fungal chitin production (reviewed in Durand et al., 2008; Milewski, 2002; Milewski et al., 2006). Candida albicans GFAT activity increases in line with chitin demand during germ tube formation occurring at the yeastto-mycelium transformation (Chiew et al., 1980). This increase can be attributed to both an upregulation of GFAT gene transcription and a post-transcriptional control of the enzyme (Milewski et al., 2006; Smith et al., 1996). It has long been known that eukaryotic GFAT is inhibited by the end product of the hexosamine pathway, UDP-GlcNac (Endo et al., 1970; Kornfeld, 1967; Watzele and Tanner, 1989). The inhibition is allosteric, and in C. albicans GFAT, UDP-GlcNac inhibition is modulated (or reduced) by glucose-6-P (Milewski et al., 1999). Invertebrate GFATs have been characterized from Drosophila, mosquito and tick. Drosophila GFAT1 has been found in embryonic tissues associated with cuticle deposition (dorsal side and trachea at stages 16 and 17), but its role changes later in development, being restricted to the salivary glands in late third instar larvae (Graack et al., 2001). Aedes aegypti GFAT1 (AeGFAT1) is closely related to DmGFAT1 but has a different expression pattern. The gene transcript is present in the midgut and increases in abundance after a blood meal. This observation is consistent with the demand in chitin triggered by the production of the peritrophic matrix (PM) lining the midgut (Kato et al., 2006; Perrone and Spielman, 1988). As with a few other


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mosquito genes induced by blood meal, the upstream region of AeGFAT1 is populated with binding sites for transcription factors of the ecdysoneresponse pathway. Four Broad-, two E74- and one ecdysone-response elements have been discovered (Kato et al., 2002). Post-blood feeding induction of GFAT has also been observed in the ixodid tick, Haemaphysalis longicornis. The gene is ubiquitously expressed, but expression rises only in the gut and the epidermis after a blood meal (Huang et al., 2007). Knocking down HlGFAT by RNAi inhibits blood feeding, likely caused by the inability of individuals to synthesize chitin and extend the surface of the PM and the cuticle during the short feeding span. Phosphorylated glucosamine produced by GFAT is further N-acetylated by glucosamine-6-phosphate acetyltransferase (GNA), and the phosphate group moves from position 6 to position 1 via the action of N-acetylglucosamine phosphate mutase (AGM). Very little is known about the regulation of these two enzymes in the context of chitin synthesis. In yeast, deletion of the genes encoding GNA and AGM are lethal. However, the lethal phenotypes are not specifically linked to any deficiency in chitin. They are rather a consequence of the essential role that the hexosamine pathway plays in the synthesis of glycosylphosphatidylinositol anchors and mannoproteins, in addition to chitin (Milewski et al., 2006). The last step before the polymerization of UDP-GlcNAc units by chitin synthase consists of the condensation of GlcNAc-1-P with UTP. This step is catalysed by UDP-GlcNAc pyrophosphorylase (UAP, EC 2.7.7.23). Along with GFAT, UAP constitutes an important control point in the provision of precursors to chitin synthase. Most insects encode a single UAP gene, but the flour beetle, Tribolium castaneum, is known to harbour a second copy (Arakane et al., 2011). Mutations in the Drosophila UAP gene (named mummy or cystic) have revealed its essential role in a range of developmental processes that depend on UDP-GlcNAc, including but not limited to chitin synthesis. Mutants of mummy show defects in cuticle synthesis, central nervous system morphogenesis and tracheal tube elongation (Arau´jo et al., 2005; Beitel and Krasnow, 2000). The cystic mutants also fail to deposit any chitin in the lumen of embryonic tracheal tubes. Interestingly, chitin not only serves a protective role in trachea but also stabilizes the expanding epithelium and organizes the behaviour of the surrounding tracheal cells (Devine et al., 2005). The presence of two UAP genes in Tribolium implies that selection pressure for distinct UDP-GlcNAc production patterns has been at play in either



this species or its ancestor. Arakane et al. (2011) have characterized the two genes, TcUAP1 and TcUAP2 and teased apart their functions using RNAimediated knock-down in both adults and larvae. TcUAP1 fulfils a clear role in chitin synthesis in both the epidermis and the midgut. Injection of TcUAP1-dsRNA in larvae triggers developmental arrest at the larval–larval, larval–pupal or pupal–adult moults. The cuticle in the elytra becomes depleted of chitin and displays a loss of integrity, and the PM chitin content is likewise decreased. Injection of TcUAP2-dsRNA is also lethal when injected into larvae, leading to pupal paralysis, but chitin content in either the midgut or the epidermis appears normal. The authors indicate that while both genes seem to be required to fulfil a range of vital functions dependent on UDP-GlcNAc production (e.g. protein glycosylation and secondary metabolite production), only TcUAP1 appears to have retained a significant role in chitin synthesis. Consistent with its proximity to chitin synthase in the chitin metabolic pathway, the UAP gene expression also appears to be regulated by 20E. Injection of 20E in S. exigua larvae greatly increases SeUAP transcripts 12 h later (Yao et al., 2010), along with the transcription of SeG6PI and SeTre-1 noted above. The delay in transcription and the sensitivity of SeUAP, SeG6PI and SeTre-1 to the protein synthesis inhibitor, cycloheximide, warrants their positioning as “late gene” in the ecdysone-response cascade. It is quite likely therefore that the metabolic steps spanning trehalose breakdown to chitin polymerization are controlled in unison during the larval to pupal transition. 2.1.3 Glutamine synthase Given that equimolar amounts of glutamine and fructose-6-P enter in the production of glucosamine-6-P by GFAT, it is not surprising that glutamine availability is critical to chitin synthesis. De novo glutamine synthesis from intracellular glutamate pools is catalysed by glutamine synthase (GS, EC 6.3.1.2). Smartt et al. (1998) demonstrated that the inhibition of A. aegypti GS with L-methionine-S-suffoximine disrupts PM formation, presumably as a result of a deficiency in chitin content. The transcriptional control of the AeGS gene is complex, with a core promoter and regulatory elements that likely allow differential modulation of expression between the midgut and other tissues (Niu et al., 2003). At least one other arthropod displays an increase in GS production to respond to chitin synthesis demand. GS expression in the Antarctic krill, Euphausia superba, increases threefold between the late intermoult stage and apolysis, a time that corresponds to chitin synthesis (Seear et al., 2010).


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2.1.4 Chitin synthase The final step in chitin synthesis is one that is exceedingly complex. One has to be reminded that both the substrate and the product differ radically in physicochemical properties, UDP-GlcNAc being small and soluble while chitin is insoluble, and can reach micrometers in length. Furthermore, the polymerization occurs at the boundary of two topological spaces, from the cytosol to the lumen of vesicles or outside the cell. This final step is catalysed by chitin synthase. Needless to say, the biochemistry and molecular biology of chitin synthase has been the subject of intense scrutiny, yet the full mechanistic details of its activity remain elusive. Because of its intricate activity and absolute requirement, chitin synthesis has long been considered an “Achilles’ heel” of arthropod pests and fungal diseases (Cohen, 1993; Georgopapadakou and Tkacz, 1995; Rogg et al., 2012). Chitosan, a useful derivative of chitin, has also spurred interest in understanding the chitin synthase machinery so that its production can be better harnessed (de Assis et al., 2010; Je and Kim, 2012). Excellent reviews on chitin synthase enzymatic properties, regulation and taxonomic distribution have been published recently (Cohen, 2010; Merzendorfer, 2006, 2011; Muthukrishnan et al., 2012). Chitin synthases (ChSs, EC 2.4.1.16) are inverting glycosyltransferases belonging to the “GT2” family (Lairson et al., 2008). Members of this family, which also include hyaluronan- and cellulose-synthases, catalyse glycosyl group transfer with an inversion of the anomeric stereochemistry relative to the donor sugar. ChSs are integral transmembrane proteins in which three major domains (A–C) have been recognized based on primary sequence homologies between insect and fungal ChSs. The A domain comprises the N-terminal part of the protein, and in insect ChSs, it is occupied by 7–10 transmembrane helices, a feature that distinguishes them from ChSs of other taxa where at most two such helices can be found (Merzendorfer, 2011). The central B domain is entirely cytosolic and displays conserved amino acid motifs important for the activity/processivity of the enzyme. Common to all GT2 enzymes, the B domain harbours Walker A and B motifs (A: GXXXXGK(T/S), B: (R/K)XXXXGXXXXLhhhhD, where “h” denotes an hydrophobic amino acid) that bind to the nucleotide moiety and a “D(I/V)D” motif that coordinates divalent cation and facilitates UDP departure (Lairson et al., 2008). Donor sugar binding is presumed to take place through the latter motif and another conserved motif (sequence GCF(A/S)LFR) 63 amino acids downstream. Completing the B domain motifs are a putative acceptor sugar donor site (sequence GEDRW) and a



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Figure 6.8 Domain organization of the insect chitin synthase 1 protein. Chitin synthase 1 is a transmembrane protein divided into three structural domains, denoted A, B and C above the polypeptide chain. N- and C-termini are labelled on the left and right side of the polypeptide chain, respectively. Transmembrane helices are indicated as red cylinders spanning the lipid bilayer. Domain A contains nine transmembrane helices. Domain B is entirely cytoplasmic and contains five conserved motifs among chitin synthase, indicated by dark blue circled letters: Motifs “A” and “B” denote the Walker A and B motifs that bind to the nucleotide moiety. Motif “C” denotes the donor saccharide-binding site, while motif “D” indicates the acceptor saccharide-binding site. Motif “E” indicates a conserved sequence involved in product binding. Domain C contains two clusters of transmembrane helices: the first one is a bundle of five helices putatively involved in chitin extrusion. The second cluster, containing two helices, is located closer to the C-terminus of the protein.

highly conserved sequence [(Q/R)RRW] that is thought to be involved in binding the nascent chitin chain. The C-domain is located at the C-terminal part of the protein. Typical for insect ChSs C-domain, a cluster of five transmembrane helices can be found close to the boundary of domain B, and a further two closer to the C-terminal end (Fig. 6.8). The function of this domain remains enigmatic, though it is suggested to help the translocation of the chitin polymer across the cell membrane (Cohen, 2010). Chitin synthase cDNA sequences and their corresponding gene structures have been characterized in several arthropod species (Fig. 6.9). As a rule, most species encode two genes (ChS1 and 2, encoding ChS-A and -B proteins, respectively), a fact supported by the recent release of fully sequenced genomes. These include crustacean (Daphnia pulex), chelicerates (Tetranychus urticae, Ixodes scapularis) and all insects with the exception of the pea aphid, Acyrthosiphon pisum, where only one ChS gene has been found. In theory, all chitin synthesis in the latter species could be accomplished by the


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Figure 6.9 (A) Structure of the chitin synthase 1 gene of Tribolium castaneum. Exons are indicated by large rectangles linked by lines. The white section at the 50 - and 30 - most extremities indicate the untranslated regions, while black sections represent the coding portion of the TmChS1 gene. The alternatively spliced exons 8a and 8b are labelled. The length of the bar on the upper right corner is 1 kb. (B) Multiple amino acid sequence alignment of TmChS1 exons 8a and 8b polypeptides and homologous exon sequences from 12 other insect species. All “a” exons are grouped in the upper alignment, while “b” exons are in the lower alignment, with their respective cytoplasmic, transmembrane and extracellular regions boxed and labelled. Consensus residues within each “a” and “b” exon alignments are indicated as stars (*) to denote identity, or by a colon (:) or a dot (.) to denote similarity. A global consensus denoting identical residues for all exons “a” and “b” sequences is indicated in red, at the bottom of the figure. The naming of the ChS1 exons polypeptides is as follows: two letters for the species name, followed by “CHSA”, the exon number and the exon type (a or b). The two-letter species are the following: Dm, Drosophila melanogaster; Ag, Anopheles gambiae; Ae, Aedes aegypti; Cq, Culex quinquefasciatus; Tc, Tribolium castaneum; Nv, Nasonia vitripennis; Cf, Camponotus floridanus; Pb, Pogonomyrmex barbatus; Hs, Harpegnathos saltator; Lh, Linepithema humile; Am, Apis mellifera; Dp, Danaus plexippus; Of, Ostrinia furnacalis.

product of a single gene, although it is also possible that the “missing” copy lies on an unsequenced portion of the genome. The Drosophila CHS-A and CHS-B proteins display 47% identity at the amino acid level. This low percentage of identity is typical of most ChS-A



and ChS-B pairs across insect species and is suggestive of an ancient gene duplication event. Studies on the expression of ChS-A and -B enzymes also indicate that the two forms accomplish different roles during insect development. Mutations in the Drosophila ChS1 gene were first identified under the name krotzkopf verkehrt (kkv), in a classical saturating mutagenesis screen that uncovered embryonic lethal mutants on the third chromosome (Jurgens et al., 1984). kkv mutants display a crumbled head skeleton and narrow denticle bands phenotype and even present deficiencies in cuticle sclerotization and pigmentation (Moussian et al., 2005). Accordingly, the DmCHS1 gene is strongly expressed in stage 14–16 embryos at the time of peak embryonic cuticle deposition. It is also expressed at high levels later in development in white prepupae (Gagou et al., 2002; also see Fig. 6.10). The ChS2 gene (DmCHS2) is for its part expressed at much lower levels in embryos and peaks at an earlier time (8–10 h embryos). Combined with an absence of embryonic cuticle defect alleles mapping to DmChS2, all available evidence indicates that DmChSB is dispensable for embryonic cuticle synthesis. According to the FlyAtlas tissular gene expression database, the highest levels of DmChS-2 expression can be found in both larval and adult hindguts and trachea, with very little transcript detectable in the carcasses (which include the epidermis) or the trachea. By contrast, DmChS1 is abundantly expressed in the two latter tissues, in larvae (Chintapalli et al., 2007). Although Drosophila would be an ideal system to decipher the exact roles of ChS-A and ChS-B, due to the ease of generating mutants in this insect, more progress has been accomplished of late by applying gene knock-down technology in Tribolium. Using RNAi-mediated gene knock-down, Arakane et al. (2005) demonstrated convincingly that TcChSA (encoded by TcCHS1) is involved in the synthesis of chitin in the epidermis and the trachea, while TcCHSB (encoded by TcCHS2) functions in the synthesis of chitin embedded within the midgut peritrophic membrane. The functional specialization of CHS-A and CHS-B between the epidermis and the midgut is probably a feature conserved in several other insect species. Alternative splicing is known to increase the diversity of ChS1 gene products. Two alternatively spliced exons, 8a and 8b, can be found in the TcCHS1 gene (Arakane et al., 2005; Fig. 6.9A). These exons encode two slightly differing version of a C-domain transmembrane helix, flanked by an extracellular segment at its N-terminus and a cytoplasmic fragment at its C-terminus. The stretch of amino acid (59–60 aa) is rather small in relation to the total length of CHS-B, but the capacity to switch between the two isoforms appears critical, as these alternative exons are conserved in



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Figure 6.10 Developmental mRNA expression profiles for 14 Drosophila melanogaster genes involved or putatively involved in chitin biosynthesis. Expression data were obtained from the modENCODE consortium dataset (http://modencode.org; Celniker et al., 2009) and converted into a heatmap format by using the matrix2png program version 1.2.2 (http://www.chibi.ubc.ca/matrix2png/; Pavlidis and Noble, 2003). Bright yellow squares indicate high expression levels, while black squares indicates no expression. The 30 developmental time points are organized in columns, while genes are in rows. Values were normalized within each row to obtain a mean ¼ 0 and a variance ¼ 1. Values were collected for the genes encoding the following enzymes: trehalases—Treh and CG6262; hexokinases—HEX-A and HEX-C; glucose6-phosphate isomerase—Pgi; glutamine-fructose-6-phosphate-amidotransferase— GFAT1 and GFAT2; glutamine synthase—Gs1 and Gs2; glucosamine-6-phosphate acetyltransferase—CG1969; N-acetylglucosamine phosphate mutase—CG10627; UDP-N-acetylglucosamine pyrophosphorylase—mmy and chitin synthases—kkv and CS-2.

almost all insect CHSAs. The amino acid alignment of the two alternative exons, obtained from the genomic sequence data from 13 species of holometabola, is presented in Fig. 6.9B. Arakane et al. (2005) were able to knock-down either or both TcCHSA isoforms, using specific dsRNA molecules, and observe the phenotype of injected individuals, in terms of their ability to complete larval and pupal moults and adult emergence. The data generated so far indicate that the TcChSA-8a splice variant is required for proper larval–pupal and pupal–adult moulting, while TcChSA-8b is required for adult emergence.



Several questions that surround the mechanistic properties of ChSs, and in fact many inverting glycosyltransferases, remain unanswered. A first, uncertainty concerns the identity of the donor and acceptor sugars in the b-1,4 glycosidic bond formation. If UDP-GlcNAc acts as the donor, then the growing chitin polymer extends by addition to its non-reducing end. Conversely, if the monosaccharide acts as the acceptor, the donor (chitin) extends by addition at its reducing end. In both Mesorhizobium chitooligosaccharide synthase (NodC) and zebrafish chitin synthase, evidence points to polymer elongation at the nonreducing end (Kamst et al., 1997, 1999). However, hyaluronan synthases class I, enzymes that are closely related to ChSs, appear to catalyse reducingend elongation (Lairson et al., 2008). The initiation of chitin polymerization is likewise contentious. It has been proposed that arthropod ChSs require a soluble or covalently bound primer to start chain elongation. The identity of the primer has been variously proposed to be a chitooligomer, a glycolipid, or a dolichol derivative (Horst, 1983; Palli and Retnakaran, 1998), but this hypothesis lacks strong foundations (Merzendorfer, 2011). Support for a primer-induced polymerization traces its origins to one of the first cell-free assay of insect chitin synthase (Quesada-Allue et al., 1976), but further experimental evidence will be needed to infirm or confirm it.

2.2. Chitin degradation The catabolism of chitin is important in two fundamental contexts of arthropod biology. Firstly, chitin can be a significant barrier or a source of energy for a large number of microorganisms, parasites and predators that consume arthropods. To breach the chitin barrier, or unlock its basic building blocks and release energy, almost all such organisms depend on extremely efficient chitin degradation enzymes. Secondly, the very structural advantage provided by chitin, in the form of a rigid exoskeleton, is incompatible with the linear and gradual growth of body size. Arthropods have resolved this conundrum by punctually destroying and rebuilding the exoskeleton through the process of moulting. In this context as well, a number of enzymatic systems are required to carefully dismantle the chitin architecture that surrounds the epidermis and lines the gut. Our discussion will centre on enzymatic systems important in the latter context. 2.2.1 Chitinases Chitinases are among the most abundant enzymes involved in chitin metabolism. They are glycosyl hydrolases that sever the b-1,4 glycosidic bond of chitin with a retention of the anomeric configuration (Arakane and


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Muthukrishnan, 2010). Insect chitinases are divided in numerous “groups” based on the number and type of domains they encode. At a minimum, a catalytic domain that confers the chitinase activity must be present. The 3D structure of catalytically active chitinase domain has not yet been resolved; however, the model of the Drosophila IDGF2 protein, a growth factor closely related to insect chitinases, has been published (Varela et al., 2002). The catalytic domain presents the TIM barrel scaffold present in all glycosyl hydrolases family 18 so far studied, to which all insect chitinases belong (Terwisscha van Scheltinga et al., 1996). The vast majority of insect chitinases comprise the catalytic domain preceded by a signal peptide, implying that they are secreted. In the beetle, T. castaneum, fully 13 of the 20 chitinase genes, belonging to groups IV, V and VII, encode proteins that display this arrangement of the two domains (Arakane and Muthukrishnan, 2010; Zhu et al., 2008a). In chitinases of group VIII, a single transmembrane domain takes the place of the signal peptide, suggesting that these proteins are membrane bound rather than secreted (e.g. TcCHT11). A third distinct domain, the chitin-binding domain (CBD), is present at the C-terminal end of group I chitinases and in some chitinases of group IV. The CBDs of insect chitinases are closely related to the carbohydrate motif 14 found in peritrophins (Jasrapuria et al., 2010). A single CBD is also present in group VI chitinases, but it is followed by an extremely long region (up to 2500 residues) rich in serine and threonine residues. A yet more complex arrangement of domains is displayed in group II and III chitinases. Group III chitinases (represented by TcCHT7) present two catalytic domains in tandem, flanked by an N-terminal transmembrane domain and a C-terminal CBD (Arakane and Muthukrishnan, 2010). Group II are extremely long proteins ( 2700 amino acids) with multiple catalytic domains and CBDs. Group II chitinases from dipterans display a domain order with the formula “CatCBD 3Cat 2CBDCat” (where “Cat” indicates the catalytic domain). In all other insect species, group II chitinases are longer still and present an extra “Cat” and CBD at the N-terminus, giving the formula “CatCBDCatCBD 3Cat 2CBDCat”. The distinct functional roles of this bevy of chitinases are starting to be understood in Tribolium. Knocking down the gene expression of the most complex (group II) chitinase in this insect, TcCHT10, blocks moulting at all stages (Zhu et al., 2008b). TcCHT5, the only group I chitinase in Tribolium, appears to play an important role in the pupal to adult moult. Finally, knock-down of TcCHT7 demonstrated its essential function in wing and



elytral expansion and abdominal expansion after pupation. The physiological and/or developmental roles of the numerous group IV chitinases (encoded by 14 different genes) are still unclear at this point, likely because of functional redundancy between them. 2.2.2 Chitin deacetylases Chitin deacetylases (CDAs, EC 3.5.1.41) are secreted metalloenzymes that catalyse the removal of acetyl groups from the chitin polymer. Deacetylated chitin is also known as chitosan, a biomaterial with useful biomedical applications. The cloning of insect CDAs is recent, with the first sequence isolated from the lepidopteran Trichoplusia ni by Guo et al. (2005). The characterization of CDA functions and expression patterns has been brisk since then, especially in the more genetically tractable Drosophila and Tribolium species. CDA genes have been systematically studied by Dixit et al. (2008) and Arakane et al. (2009) in these two genomes, along with those from Anopheles gambiae and the honeybee. The number of CDA genes varies between a minimum of five (in the mosquito and the honeybee) up to nine in Tribolium, again suggestive of ancestral gene duplication events followed by specialization in the latter species. CDA proteins are modular, presenting at a minimum a catalytic (CDA) domain at the C-terminus. Five groups of CDAs are recognized, depending on the presence and order of other domains and their overall degree of similarity. Group I and group II CDAs encode a CBD followed by a lowdensity lipoprotein receptor class A (LDLa) domain and the CDA domain. Although similar in modular architecture, members of group I and group II CDAs display overall low levels of identity. Insects encode two genes from group I CDA and a single gene from group II. Mutations affecting two group I CDA genes have been recovered in Drosophila and have been shown to affect embryonic tracheal tube morphogenesis (Luschnig et al., 2006). Named Serpentine (Serp) and Vermiform (Verm), these two genes encode DmDCA1 and DmDCA2 enzymes, respectively. Their peculiar names stem from the extremely long and convoluted appearance of tracheal tubes displayed by mutants at stage 15 and 16 of embryonic development. It has been suggested that by converting chitin into chitosan, DmCDA1 and DmCDA2 increase the rigidity of tracheal chitin cylinders. Through direct and indirect influence on the tracheal epidermis, this change restricts the longitudinal growth of tracheal tubes. In Tribolium, the function of the two DmCDA1 and DmCDA2 orthologues, TcCDA1 and TcDCA2, were investigated through RNAi injections. Both TcCDA1 and TcCDA2


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knock-downs adversely affected the completion of every type of moult (larval, pupal or pupal–adult). RNAi-treated individuals appear unable to break from the old exuviae (Dixit et al., 2008). The function of the single representative of group II CDAs remains largely unknown, except for the restricted expression of TcCDA3 in Tribolium thoracic muscles. The roles of the other three groups of insect CDAs are also not fully understood. Group III and group IV CDAs display a CBD and a CDA domain but are devoid of the central LDLa domain. Group V CDAs seem to have retained only the CDA domain, with no recognizable CBD or LDLa domains. Typically, insects encode one gene of each group III and IV CDAs, while a more variable number of group V CDAs can be present (up to four in Tribolium). The expression of these CDAs has been tracked with great precision by Arakane et al. (2009), using RT-PCR and in situ hybridization. Their results indicate that transcripts from TcDCA4, the only group III CDA gene, can be detected in the epidermal cells of imaginal appendages. TcCDA5 (group IV) is for its part expressed in the carcass and at all stages of development. Finally, members of the group V CDAs (TcCDA6–TcCDA9) show strong expression in the midgut. The variable number of group V CDA genes is particularly intriguing, and their strong expression in the gut indicates a possibly important role in modifying the chitin-to-chitosan ratio in the peritrophic membrane. Interestingly, Jakubowska et al. (2010) have observed that infection of Spodoptera frugiperda by the Helicoverpa armigera single nucleopolyhedrovirus (HearNPV) baculovirus increases the accumulation of the HaCDA5a gene. Mutant baculoviruses expressing the HaCDA5a gene were also more infectious. These data indicate that regulation of gut CDA gene expression constitutes an important strategy in breaking host defence barriers, presumably by modifying PM permeability. The larger number of gut-specific CDAs might be a reflection of the host–pathogen arms race taking place in this tissue.

2.3. b-N-acetylglucosaminidases b-N-acetylglucosaminidases (NAG, EC 3.2.1.52) are responsible for the hydrolysis of terminal N-acetylglucosamine residues from the non-reducing end of oligosaccharides (Cohen, 2010). This family of enzymes is extremely well represented in various taxa of microorganism that use glucosaminecontaining polymers as a carbon source or in the deglycosylation of proteins. Insect NAGs have been cloned from various species belonging to the Lepidoptera, Coleoptera and Diptera and classified into four groups (NAGs I–IV), based on their phylogenetic relationships and substrate affinity.



Group I and group II NAGs (NAG1 and NAG2) are bona fide chitinolytic enzymes, and insects generally encode one gene of each group. The only exceptions are the three group I NAGs present in mosquitoes of the genus Culex (C. pipiens, C. fasciatus; Muthukrishnan et al., 2012). Functional analysis of NAG1 and NAG2 enzymes has been performed in Drosophila. DmHEXO1 and DmHEXO2 (now renamed DmNAG1 and DmNAG2) cloned and expressed in a heterologous yeast expression system could digest chitotriose (Le´onard et al., 2006). Similarly, a BmNAG purified from B. mori could catalyse the hydrolysis of chitooligosaccharides into GlcNac (Nagamatsu et al., 1995). Using RNAi, Hogenkamp et al. (2008) tested the effect of knocking down the expression of the TcNAG1 and TcNAG2 genes in T. castaneum. TcNAG1 is the most abundantly expressed NAG in this species and its mRNA levels peak at the late pupal stage. Consistent with a requirement of this enzyme during moulting, TcNAG1 RNAi induces up to 90% mortality at the pupal–adult moult. RNAi-mediated knock-down of TcNAG1 is also effective in blocking larval–larval and larval–pupal moults. In contrast, the dsRNA knock-down of TcNAG2 produced a weaker lethal phenotype: while pupal–adult moults were effectively blocked, TcNAG2 RNAi could not arrest larval–larval and larval–pupal moults completely. This weaker phenotype might be linked to the more restricted expression pattern of TcNAG2. TcNAG2 is expressed at very low levels in the epidermis, but at high levels in the midgut, while TcNAG1 is expressed in roughly equal amounts in each tissue. Hence, TcNAG2 appears to play a more specialized role in the hydrolysis of midgut chitooligosaccharides than in epidermal chitin hydrolysis. Group III and group IV NAGs are referred as N-glycan-processing enzymes and hexosaminidases, respectively. Group III includes the wellcharacterized fused lobes (fdl) gene product of Drosophila (Boquet et al., 2000). Mutations in the fdl gene are associated with the fusion of mushroom bodies in the adult brain. The FDL enzyme has been purified and its hydrolytic activity towards a range of different N-glycans and oligosaccharides has been studied. While FDL was completely unable to release GlcNAc units from chitin oligomers, it was able to do so with GlcNAc units attached to the a-1,3-linked mannose of the core pentasaccharide of N-glycans (Le´onard et al., 2006). Hogenkamp et al. (2008) have hypothesized that the orthologue of DmFDL, TcFDL, is likewise involved in N-glycan processing but has retained some of the chitin-degrading activity of group I and group II NAGs. This was suggested by knock-down experiments on


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TcFDL, which caused a significant failure (80% of injected individuals) to properly complete the pupal to adult moult, along with an inability of pharate adults to shed the pupal cuticle. The insect hexosaminidases that form the group IV NAGs are only distantly related to NAGs of the other three groups. These enzymes are closely related to mammalian hexosaminidases that are active on the sugar chain of mammalian monosialic ganglioside 2 (GM2) (Kolter and Sandhoff, 1998). These enzymes are unlikely to play a significant role in chitin degradation.

3. CHITIN GENOMICS The public release of complete insect genomes in the past decade is an opportunity to better understand the activity, regulation and evolution of enzymes involved in the chitin metabolism and use this understanding to discover new chemotypes for new target sites that disrupt cuticle synthesis for pest control. At present, the genomes of 26 insect species have been sequenced, annotated and published. Slightly more than half of those are from dipterans that serve as model species in developmental and evolutionary genetics or are vectors of viral or parasitic diseases [Drosophila melanogaster and 11 congeneric species (Drosophila 12 Genomes Consortium, 2007); A. aegypti, Culex quinquefasciatus and A. gambiae (Arensburger et al., 2010; Holt et al., 2002; Nene et al., 2007)]. Hymenopterans (bees, wasps, ants) are represented by eight genomes (Smith et al., 2011a,b; Werren et al., 2010), while the genome of two species within the Lepidoptera (moths and butterflies) and one each in the Coleoptera (beetles) and Homoptera (aphids) have also been released (International Aphid Genomics Consortium, 2010; Tribolium Genome Sequencing Consortium, 2008; Xia et al., 2004; Zhan et al., 2011). Last, the recent completion of the water flea (D. pulex) genome will also be instrumental in identifying the conserved arthropod proteins involved in chitin metabolism (Colbourne et al., 2011). The community of Drosophila genomicists has pioneered numerous “-omics” tools that allow one to investigate gene function, or at least to narrow down hypotheses that address gene function. The modENCODE consortium is particularly active in this area, with a stated goal of “identify(ing) all of the sequence-based functional elements in the Caenorhabditis elegans and D. melanogaster genomes” (http://www.modencode.org; Celniker et al., 2009). The consortium has so far generated and curated an extremely large amount of data on gene profiling (e.g. coding and noncoding RNAs),



the binding sites of transcription factors, chromatin structure and the results of other related experiments. When extended to other arthropod species, this type of project could fundamentally affect the orientation of hypothesis-driven studies on the evolution of chitin metabolism. Gathering and analysing the complete modENCODE dataset pertaining to chitin-metabolism genes and their transcriptional control elements in Drosophila are beyond the scope of this review. However, a glimpse of the utility of the modENCODE dataset can be obtained from key studies; in particular, the one authored by Graveley et al. (2011) on the precise determination of all gene transcript levels across development by next-generation sequencing. The authors of this study used a combination of RNAseq, tiling oligonucleotide arrays and cDNA sequencing to interrogate gene expression at 30 times during development (Mockler et al., 2005; Wang et al., 2009). This increase in sampling density covers the important events in Drosophila cuticle synthesis, particularly at the embryonic and pupal stage. Figure 6.10 illustrates the modENCODE gene expression profiles obtained for 14 Drosophila genes that have a putative or confirmed role in chitin synthesis. While the upregulation of the terminal genes of the chitin synthesis pathway during pupal stage was expected (i.e. mmy and the two ChS genes), a few other genes are partially coregulated. CG1969, the gene encoding glucosamine-6-phosphateacetyltransferase (GNA), appears also strongly upregulated in pupae. Similarly, the GFAT1 and Treh genes follow a strong or a slight increase in transcription at the same stage in 12 h white prepupae. The expression profiles of these genes are similar at the time of embryonic cuticle deposition (14–18 h) with some key differences, such as in the low expression of the ChS-2 gene. Likewise, an increase in Treh expression is not apparent at that stage, and the increase in GFAT1 is much more muted than in pupae. Differences in the expression levels from different chitin synthesis pathway genes are indicative that complex levels of control are at play. Figure 6.11A illustrates the expression of Drosophila chitinolytic genes. It is clearly apparent that most chitinase genes are expressed in pupae, although the increase in expression is over a broader time span than observed for the ChS-1 and mmy genes. While chitinases might not be expected to play a major role at the time of the first (embryonic) cuticle deposition, the Cht7 gene is, nonetheless, very transcriptionally active at this time (14–20 h after egg laying). As noted previously, the CDA genes that control tracheal elongation (Verm and Serp) are highly expressed in embryo.



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Figure 6.11 Developmental and tissue-specific expression patterns of Drosophila melanogaster genes-encoding enzymes involved or putatively involved in chitin breakdown.



Intriguingly, the expression of other members of this family (ChLD3, CDA4 and CDA5) follows an almost identical pattern. The functional characterization of ChLD3, CDA4 and CDA5 in the context of embryonic cuticle would be worthy of future investigations. The chitinolytic gene expression presented in Fig. 6.11B was constructed from the dataset of the FlyAtlas project, which collected genome-wide expression data from multiple larval and adult tissues (Chintapalli et al., 2007). This figure demonstrates the striking degree of tissue specificity of some Drosophila chitinolytic enzymes. Cht8 and Cht9 appear highly specific to the midgut, but the former is restricted to adults, while the latter is found only in larvae. Cht4 is also midgut specific but expressed approximately equally in larvae and adults. Other restricted expression patterns include the Malpighian tubule-specific Cht11 gene and the adult testis-specific Cht12. Future releases of the modENCODE dataset (which is currently at release #29) promise to refine even more the picture of chitin-related gene expression and how it affects chitin metabolism. Evaluating the impact of long intergenic noncoding RNA on neighbouring loci will be possible (Young et al., 2012). Additionally, analysing data obtained by chromatin immunoprecipitation at the genomic scale, to identify regulatory elements important in chitin-metabolism genes, will enable a better understanding of the transcription factor networks that turn on and off these genes (Ne`gre et al., 2011). (A) Gene expression across embryonic, larval, pupal and adult development. Expression data were obtained from the modENCODE consortium dataset (http:// modencode.org; Celniker et al., 2009) and converted into a heatmap format by using the matrix2png program version 1.2.2 (http://www.chibi.ubc.ca/matrix2png/; Pavlidis and Noble, 2003). Bright yellow squares indicate high expression levels, while black squares indicates no expression. The 30 developmental time points are organized in columns, while genes are in rows. Values normalized within each row to obtain a mean ¼ 0 and a variance ¼ 1. (B) Tissue-specific expression patterns. Data were recovered from the FlyAtlas database (http://flyatlas.org/; Chintapalli et al., 2007) and converted into a heatmap as in (A) above. Values for the following larval and adult tissues, in columns, were included: CNS, central nervous system; Mg, midgut; Hg, hindgut; MT, Malpighian tubules; FB, fat body; SG, salivary glands; Tr, trachea; Car, carcass; AH, adult head; AE, adult eye; Br, brain; TAG, thoracicoabdominal ganglion; Cr, crop; He, heart; Ov, ovaries; Tes, testis; VF St, virgin female spermatheca; MF St, mated female spermatheca. Values were collected for the genes encoding the following enzymes: chitinases—Cht2-5, Cht7-9 and Cht11-12; hexosaminidases—Hexo-1, Hexo2 and fdl; chitin deacetylases—Verm, Serp, ChLD3, CDa4-5 and CDa9. Expression data could not be retrieved for Cht6 and Cht10 genes for either the developmental or tissue-specific dataset. FlyAtlas expression data was also unavailable for the Hexo-1 gene.


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4. BENZOYLPHENYL UREAS AS CSIs Targeting a biochemical compartment that is important for the survival of an insect pest is an attractive proposition for pest management. Chitin is an integral part of the exoskeleton of insects and is essential for the protection of the insect against dehydration, microbial infection and physical injury. A serendipitous discovery at the Philips–Duphar laboratory in Holland led to the development of a benzoylphenyl urea (BPU), diflubenzuron or DimilinÒ (Maas et al., 1981; Verloop and Ferrell, 1977). Since most of the commonly used pesticides such as organophosphates, carbamates, pyrethroids and neonicotinoids are neurotoxins that target neurotransmission by interacting with acetyl cholinesterase or its receptor, or sodium channel in the nervous system whereas BPUs inhibit chitin synthesis in the epidermis, it was felt that cross resistance was less likely to occur. Also chitin synthesis is primarily an arthropodan feature that is absent in vertebrates, thus narrowing the activity spectrum.

4.1. History of development One of the first compounds tested at the Philips–Duphar laboratory in 1970 was designated as DU-19.111 and consisted of a substituted benzoyl group attached to a substituted phenyl group by a urea bridge. It was made by combining two herbicides, dichlorobenil and diuron. It was routinely tested for herbicidal and insecticidal activity. When tested on insect larvae, there was no immediate knock-down effect, but treated individuals showed various degrees of moult deformities. This discovery sparked the research on moult inhibiting BPUs as a new class of insecticides-targeting chitin synthesis. One of the earliest compound synthesized had a difluorobenzoyl group on one end of the urea bridge and a chlorophenyl group on the other side and came to be known as PH 60-40 or diflubenzuron (commercialized under the name DimilinÒ; Maas et al., 1981; Verloop and Ferrell, 1977). Diflubenzuron became the harbinger of the vast array of “insect growth regulators (IGRs)” or “CSIs” or “BPUs” that were developed by various groups. Since it had to be ingested in order to be effective and interfered with cuticle formation, it had a narrow spectrum of activity. It became especially attractive because it was effective only on organisms that had a chitinous exoskeleton. BPUs have been reviewed several times, and in this review, we will provide the major conclusions from the past and



update them to their current status (Dhadialla et al., 2005; Matsumura, 2010; Retnakaran et al., 1985; Wright and Retnakaran, 1987).

4.2. Classification and structure of benzylphenyl ureas, structure–activity relationships BPUs are a class of compounds with a central urea moiety, with a benzoyl group attached to a nitrogen on one side of the urea bridge and a phenyl group attached to the nitrogen on the other side. Diflubenzuron has two ortho-fluorine substitutions on the benzoyl moiety and a parachlorine substitution on the phenyl end. None of the BPUs have a para substitution on the benzoyl part of the molecule (Fig. 6.12). Most of the complex substitutions occur on the phenyl end, whereas the benzoyl part remains relatively simple. The basic skeleton remains constant within all the BPUs. Nakagawa and his group (Nakagawa et al., 1991) conducted an in-depth study of the quantitative structure–activity relationship (QSAR) of various synthesized BPUs with larvicidal activity on the larvae of the rice stem borer, Chilo suppressalis. They found that on the benzoyl end, simple mono- or disubstitutions at the ortho-position with either single chlorine or two fluorines were optimal, making it strongly electron withdrawing and hydrophobic. Very little tinkering at this end was possible to increase the activity. They hypothesized that this end was the one that attached itself to the unidentified receptor resulting in the inhibition of chitin synthesis. The phenyl (or the anilide) end permitted profound substitutions so long as they maintained high hydrophobicity and electron-withdrawing ability. Chlorfluazuron with an unoccupied ortho-position in the phenyl end and a dichloro pyridyloxy group in the para-position made it highly hydrophobic and electron withdrawing, making it very active. Similarly, teflubenzuron has four halogen substituents in the phenyl end, making it both hydrophobic and electron withdrawing. The biological spectrum of activity, however, defies accurate prediction by QSAR studies making us rely on trial-and-error assays on various insects.

4.3. MOA and receptor Upon ingestion by larvae, BPUs induce overt deformities during moulting, which can be traced to the process of chitin synthesis and assembly in the cuticle, and this is the basis of their insecticidal action (Khan and Qamar, 2011; Retnakaran et al., 1976). There are also subtle and covert effects on reproduction and egg hatch (Lo´pez et al., 2011). Delayed effects on pupae and adults have also been observed (Eisa et al., 1991). One of the



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Figure 6.12 Chemical structures of a representative group of benzoylphenyl ureas that have been commercialized.

earliest explanations for these moult deformities was hormone imbalance. The adverse effect on cuticle morphogenesis, which was widely observed, indicated that chitin formation was probably involved. When actively feeding larvae ingest BPU, the effect is manifested during the moult. There is no immediate knock-down effect, but instead there is a



delayed effect at the time of the moult. During larval-to-larval or larval-topupal moulting, among other things, a new cuticle is secreted and the old cuticle is digested. The adverse effect of BPUs at these stages reinforces the idea that chitin formation was probably the target of these compounds (Gangishetti et al., 2009). Chitin inhibition has been tracked histochemically with selective stains such as calcofluor white or fluoroscein isothiocyanate labelled with wheat germ agglutinin (FITC-WGA) (Meyberg, 1988). Using the FITC-WGA stain, the synthesis of chitin during moulting and the inhibition by BPUs were studied on the spruce budworm, Choristoneura fumiferana. The inhibition of chitin formation by these compounds became readily apparent (Palli and Retnakaran, 1998; Retnakaran, 1995). Radioactively labelled UDPGlcNAc was used to track chitin synthesis in BPU-treated and untreated spruce budworm larvae (Retnakaran et al., 1989). Histochemical and radioactive labelling studies confirmed that BPUs inhibit chitin formation during moulting. BPU-induced chitin inhibition has also been examined at the ultrastructural level (Retnakaran et al., 1989; Unsal et al., 2004). The results clearly show that chitin deposition is inhibited, and instead of chitin lamellae, a fibrous zone is observed (Fig. 6.13). In vitro assays using tissue preparations have demonstrated the activity of chitin synthase, catalysing the polymerization of UDP-GlcNAc to chitin (Cohen and Casida, 1980). Turnbull and Howells (1983) showed that diflubenzuron and Polyoxin-D (a structural analogue of UDP-GlcNAc) inhibit chitin synthase in an in vitro tissue preparation from the Australian sheep blowfly, Lucilia cuprina. Since these crude preparations contained other materials besides the enzyme, it was unclear whether the BPUs directly inhibited chitin synthase or the effect was indirect. Leighton et al. (1981) proposed that chitin synthase existed as a zymogen, and the benzoylurea inhibited the proteolytic enzyme required for activation. Since chitin synthase is a membrane-bound enzyme, it is difficult to obtain a cell-free preparation without losing some of its activity. Nevertheless, cell-free systems have been used to show chitin synthase inhibition by BPUs (Merzendorfer and Zimoch, 2003). Chitin synthesis is a complex process that takes place in the polarized epidermal cells where the precursors enter through the basal lamina from the hemolymph and the polymerization occurs at the apical plasma membrane. The biosynthesis takes place within the cytoplasm in the endoplasmic reticulum and the Golgi, and the chitin fibril inside the vacuole is exocytosed and assembled on the surface to a preset architecture. The key enzyme involved


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Figure 6.13 Ultrastructural changes in the sixth instar spruce budworm that were force fed with 1 ng of chlorfluazuron per larva (24-h old) and examined when they were 48 h (A), 60 h (B) and 72 h (C) old. de, dense epicuticle; dv, dense vesicle; f, amorphous layer denoting fibrous zone; G, golgi bodies; l, endocuticular lamellae; m, mitochondria; mv, microvilli; n, nucleus; pc, pore canal; rer, rough endoplasmic reticulum; s, smooth apical membrane; t, transitional layer of disrupted endocuticle; vl, lucent vesicle. (From Retnakaran et al., 1989, with the permission of Pesticide Biochemistry and Physiology).

in chitin synthesis is chitin synthase which is a glycosyl transferase that polymerizes the GlcNAc to form chitin microfibrils utilizing UDP-GlcNAc as the building block and is also the one that is inhibited by BPUs. As described earlier, chitin synthase has been sequenced from many insects (Ampasala et al., 2011; Merzendorfer, 2006, 2011) and recently an elegant structural



model has been proposed on the basis of several studies by Muthukrishnan et al. (2012). It has been suggested that the phenyl end (also called the anilide end) which permits minimal di-ortho substitutions is probably the region that interacts with the chitin synthase through perhaps a yet to be identified receptor (Nakagawa et al., 1991). Nakagawam and Matsumura (1993) used newly moulted integument preparations from the American cockroach, Periplaneta americana, to study chitin synthase activity by measuring the incorporation of 3H-GlcNAc. Diflubenzuron completely inhibited chitin synthase activity in this system. Various ionophores for Kþ, Caþþ and Hþ inhibited chitin synthase activity as well, suggesting that diflubenzuron inhibition was related to ion transport which in turn is related to intracellular exocytosis of vesicles. Using a homogenate of newly moulted integument, they were able to obtain an active preparation of intracellular vesicles of chitin synthase (Nakagawa and Matsumura, 1994). Diflubenzuron effectively blocked chitin synthase activity in these vesicle preparations. They hypothesized that a vesicular ABC transporter with ATP- or GTP-dependent Caþþ or Kþ transport was probably involved. The most likely candidate was an ATP-sensitive Kþ channel, a transporter which combines four sulfonylurea receptor (SUR) subunits along with four Kir6.2 subunits (Abo-Elghar et al., 2004). In order to test this hypothesis, they used an anti-diabetic sulfonylurea drug, glibenclamide (also known as glyburide), which is known to bind to SUR subunits present in channels of human pancreatic b-cell and help in the release of insulin. Upon binding of glibenclamide to SUR subunits, the KATP channel closes and the Kþ level inside the b-cell is upset. This change in intracellular Kþ content depolarizes the cell membrane, which in turn provokes the opening of Caþþ channels and allows the influx of Caþþ. In turn, the increase in intracellular Caþþ induces the exocytosis of the insulin-carrying vesicles, releasing its contents (Fig. 6.14; Aittoniemi et al., 2009). When nymphs of the German cockroach, Blattella germanica, were topically treated with the anti-diabetic drug glibenclamide, the nymphs developed moult deformities strikingly similar to the ones caused by diflubenzuron. The two compounds share a common urea bridge with substitutions on either end with the phenyl or the cyclohexane end having very few substituents (Fig. 6.15). Competitive-binding assays with tritiated diflubenzuron and glibenclamide using vesicle preparations from Blattella and Drosophila had similar results. The unlabelled ligand was able to completely displace the radioligand. The Kd values were 44.9 nM for glibenclamide and 64.9 nM for diflubenzuron in Drosophila vesicles. RT-PCR confirmed the



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Figure 6.15 Structural similarity between the anti-diabetic drug, glibenclamide and the chitin synthesis inhibitor, diflubenzuron. Both have a central urea bridge with a simple substitution at the phenyl or anilide end in diflubenzuron and the corresponding end in glibenclamide which has been suggested as the receptor-binding end.



presence of DSUR in Drosophila larvae. The vesicle preparations treated with diflubenzuron or glibenclamide showed 45Caþþ uptake which was stimulated by ATP (Matsumura, 2010). When DSUR was first isolated from Drosophila embryos, it was present in only certain areas (Nasonkin et al., 1999). How widespread is SUR expression during insect development is a lingering question that remains to be answered. Nevertheless, SUR being the target of diflubenzuron, based on the studies with glibenclamide, is an attractive hypothesis and when taken together with the recent chitin synthase model, it appears convincing (Matsumura, 2010; Muthukrishnan et al., 2012). The precursors for chitin from the hemolymph enter through the basal lamina of the epidermal cell, and the vesicle with the chitin biosynthetic machinery is probably formed in the endoplasmic reticulum and the Golgi where it matures. The vesicle migrates to the plasma membrane and exocytoses at the tips of microvilli where the nascent chitin microfibril is extruded. It is conceivable that, when treated with diflubenzuron or glibenclamide, the vesicle exocytoses precociously before the chitin is formed and the entire process comes to a halt (Fig. 6.16). It has been suggested many years ago that the transport of chitin synthase is inhibited by diflubenzuron (Eto, 1990). It appears that the enzyme chitin synthase itself is not inhibited by diflubenzuron, but the processing of the enzyme is inhibited. The processing of the enzyme is probably different in fungi and that is perhaps why diflubenzuron has no effect on the fungal synthesis of chitin. Diflubenzuron was fed to Tribolium, and a genomic tiling array was performed to examine the expression levels of 11,000 genes. While many of the genes were upor downregulated, chitin synthase itself was not affected (Merzendorfer et al., 2012). Some exciting work lies ahead, and finally, the MOA of BPUs might be unambiguously resolved in the next few years, using genomicsenabled methods.

4.4. Effects on pests Introduction of the organochlorine insecticide DDT in 1939 revolutionized pest control. It was soon followed by organophosphates, carbamates, pyrethroids, neonicotinoids, avermectins and many others making the synthetic insecticides an attractive option for pest control. The broad spectrum of activity, low cost and rapid kill made them appealing to end users. The use of these pesticides resulted in the phenomenal increase in the production of food and fibre as well as significant reduction in the incidence of arthropod-borne diseases by vector control. All these insecticides, in one



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Figure 6.16 Mode of action of diflubenzuron (DFB) through the ABC transporter, sulfonylurea receptor (SUR), by interfering with the exocytosis of chitin synthase (CHS) of Drosophila. Normally, the CHS vesicle binds with the plasma membrane where it initiates chitin synthesis and SUR which includes the KATP channel and maintains energy homeostasis within the cell. A, B, C—the three domains of CHS, B being the catalytic domain and the other two are transmembrane domains; CA, Catalytic area with the Walker motifs; ES, consensus sequence for extrusion site of chitin. Other consensus sequences are not shown. (Based on Matsumura, 2010; Merzendorfer, 2006; Merzendorfer et al., 2012; Moussian, 2010; Muthukrishnan et al., 2012.)

way or another, target the nervous system of not only insects but also other animals including humans which share a common neurophysiology. Indiscriminate and excessive use of neurotoxic insecticides soon manifested their adverse effects in ecological situations and human health which was dramatically brought into focus by Rachel Carson in her book, “The Silent Spring” in 1962 alluding to the disappearance of songbird species by DDT. Continuous use at high levels often resulted in the pest populations developing resistance that reached crisis proportions in some instances (Carson, 1962). In an attempt to curb their excessive use, a new strategy called “integrated pest management” (IPM) was developed. IPM is the judicious use of chemical control methods compatible with biological control methods, ecologically sensitive and nontoxic to human health. Soon a softer type of chemical control, often developed from natural products, made its appearance, and products developed under this new trend were called “biorational pesticides”.



“Biorational” is a loose term embracing many different compounds that were more specific against a given pest and less toxic to the environment. BPUs have been considered a biorational pesticide because of their low mammalian toxicity as well as their specificity to growing stages of arthropods in the process of actively synthesizing chitin. BPUs have been referred to as IGRs, CSIs and moult inhibitors. In Chapter 1, Pener and Dhadialla propose the more appropriate term, insect growth disruptors (IGDs), to replace the use of IGRs, as the CSIs like the bisacylhydrazine (ecdysone agonists) and juvenile hormone analogues (JHA) insecticides, which mimic the 20E and JH, respectively, actually disrupt, and not regulate, growth and development in susceptible insect pests. BPUs are slow-acting larvicides that manifest their effects at the time of moulting resulting in the mortality of the insect. These compounds also affect other tissues that have chitin such as the peritrophic membrane creating feeding problems and egg shell resulting in the inhibition of egg hatch. A selective list of 11 of the more important BPUs that have been commercialized is shown in Fig. 6.12. The control of various pests, the environmental effects, toxicology and pharmacodynamics have been extensively covered in earlier reviews (Dhadialla et al., 2005; Retnakaran et al., 1985; Wright and Retnakaran, 1987). Here, we will update some of the control measures that are more recent. Upon examination of the 11 structures shown in Fig. 6.12, it becomes apparent that the benzoyl end of the urea bridge is remarkably simple and uniform with di-ortho substitution with fluorine except for triflumuron which has a mono–ortho substitution with chlorine. The phenyl or the anilide end has all the extensive substitutions which probably accounts for the differential effects on pest populations. 4.4.1 Diflubenzuron (Dfb or DimilinÒ) Diflubenzuron is the harbinger of all the BPUs and has been extensively studied and used around the world. It is highly insoluble in water and has to be ingested to be effective. It is not systemic in plants and therefore does not work on sap-sucking insects. Although one would expect that it should be effective on all open feeding lepidopteran larvae it is not uniformly effective and this could be due to detoxification in some species. Species such as the fruit tortrix moths Adoxophyes orana, and Pandemis heparana are relatively insensitive to diflubenzuron (Eck, 1981). In the spruce budworm, C. fumiferana, larvae in the fifth and sixth stadia were more susceptible to diflubenzuron than in the earlier stages (Granett and Retnakaran, 1977). Some species like the forest tent caterpillar, Malacosoma disstria, and the gypsy


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moth, Lymantria dispar, are very sensitive to this compound (Retnakaran et al., 1985). It has been used to control cockroaches, locusts, grasshoppers and most leaf-feeding larvae, in general (Weiland et al., 2002). Insect pests of cotton, soyabean and horticultural crops are all susceptible. Larvae of sciarid flies, phorid flies on mushrooms, mosquitoes and most fly larvae can all be controlled with diflubenzuron. In veterinary applications, feed through trials on cattle led to the control of the house fly larvae (Musca domestica) in the dung. Diflubenzuron is less effective on the Colorado potato beetle, Leptinotarsa decemlineata, than lufenuron or hexaflumuron (Karimzadeh et al., 2007). Diflubenzuron is relatively nontoxic to mammals and birds. The LD50 for rats and mice is > 4640 mg/kg. The LC50 for zebra fish (96 h) is >0.2 mg/L, and it is nontoxic to bees (LD50 > 100 mg/bee). It is adsorbed by clay soil and has a half-life of