Content uploaded by Arthur Retnakaran
Author content
All content in this area was uploaded by Arthur Retnakaran on Nov 21, 2017
Content may be subject to copyright.
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 438
1.1 Chitin structure, types, distribution in fungi and animals 439
1.2 Evolution of chitin from prehistoric to present; physiology and
function in arthropods 440
1.3 Chitin in insects, distribution, architecture and sculpturing 442
2. Chitin Metabolism and Potential Targets for its Disruption 444
2.1 Chitin biosynthesis, enzymes in the pathway and chitin synthase 444
2.2 Chitin degradation 456
2.3 b-N-acetylglucosaminidases 459
3. Chitin Genomics 461
4. Benzoylphenyl Ureas as CSIs 465
4.1 History of development 465
4.2 Classification and structure of benzylphenyl ureas, structure–activity
relationships 466
4.3 MOA and receptor 466
4.4 Effects on pests 472
4.5 Effects on non-target species 479
4.6 Environmental degradation 480
5. Non-Benzoylphenylurea Chitin Synthesis Inhibitors 481
5.1 Structure and properties 481
5.2 Environmental fate and effects 484
6. Chitinases and Chitinase-Inhibiting Chemicals for Pest Management 485
6.1 Fungi and microorganisms with chitinase activity 486
6.2 Baculoviruses with chitinase gene 487
6.3 Chemical inhibitors of chitinase 488
7. Resistance and Resistance Management 489
8. Conclusions and Future Development 495
Acknowledgements 496
References 496
Advances in Insect Physiology, Volume 43 #2012 Elsevier Ltd.
ISBN 978-0-12-391500-9 All rights reserved.
http://dx.doi.org/10.1016/B978-0-12-391500-9.00006-1
437
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 biosyn-
thesis 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 insecti-
cides 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 chi-
tin 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 polysac-
charide 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 resem-
blance 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
438 Daniel Doucet and Arthur Retnakaran
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-acetyl-
D-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
aand bforms (Lotmar and Picken, 1950; Rudall and Kenchington, 1973).
a-Chitin chains are the most abundant form that are arranged in an anti-
parallel 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).
Anti-parallel chains
Parallel chains
b-Chitin
g-Chitin
a-Chitin
Two parallel and one anti-parallel chains
Figure 6.1 The three forms of chitin chains.
439Insect Chitin: Metabolism, Genomics and Pest Management
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 mil-
lion 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
4500 4000 3500
-OH Amide I band Amide II Amide III
(c)
(b)
(a)
(c)
(b)
(a)
3000 2500
1600 1620 cm-1
Wavenumber
(
cm-1
)
2000 1500 1000 500
Figure 6.2 Fourier Transform Infrared (FT-IR) Spectra of a,band gchitin chains (a, b and c)
isolated from crab shells, squid pens and lucanid beetle cocoons, respectively.
Chitosan
Cellulose Chitin
CH2OH
NHCOCH3
O
OO
OO
H
CH2OH
CH2OH
OH
NH2
O
O
H
OH
NH2
OH
O
OO
O
O
H
CH2OH
CH2OH
OH
OH
OH
OH
O
OO
O
O
H
CH2OH
CH2OH
OH
OH
NHCOCH3
NHCOCH3
Figure 6.3 Structure of chitin in relation to chitosan and cellulose.
440 Daniel Doucet and Arthur Retnakaran
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 (Ehrlichetal.,2007b;FaliniandFermani,
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
Era
Cenozoic Quartenary
Tertiary
Cretaceous
Jurassic
Triassic
Permian
Pennsylvani
an
Mississipian
Devonian
Silurian
Ordovician
Cambrian
2500
570
500
435
410
360
330
290
240
205
138
66
1.6
Mesozoic
Palaeozoic
Pre-
cambrian
Period Millions
of years
ago (0.0)
Diatoms Protozoa Cnidaria Arthropoda Chitinozoa
(Flask-like animals)AnnelidaMolluscaPoriferaFungi
Figure 6.4 Fossil history of chitin-containing organisms (after Miller, 1991).
441Insect Chitin: Metabolism, Genomics and Pest Management
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 polysaccha-
rides 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 ev-
ery situation, moulding the cuticle to every need. The body wall of the in-
sect 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
pmp
Lc
mv
mv
mf
1 mm
0.1 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).
442 Daniel Doucet and Arthur Retnakaran
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
Epidermal feet
Microvillus
Synthesis and transport
of cuticle precursors
including chitin
Lamellate cuticle
secretory zone
Filopodia
Basement
membrane
N
Actin filaments
Plasma membrane
plaque
Chitin lamellae
-Procuticle
Epicuticle
Cuticulin
Figure 6.6 Schematic representation of the physiology of the epidermal cell. See text
for details. (Based on Locke, 1991, 2001; Moussian, 2010.)
443Insect Chitin: Metabolism, Genomics and Pest Management
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 spe-
cies (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 be-
hind chitin production is warranted, and it implies that intricate induction
and termination controls exist, with ecdysone orchestrating the entire pro-
cess 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, tre-
halose 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
444 Daniel Doucet and Arthur Retnakaran
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 ac-
count for the majority of synthesized chitin (Zaluska, 1959). Glucose then
briefly enters glycolysis to generate fructose-6-phosphate (F6P). F6P is sub-
sequently 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 Neu-
rospora crassa. Observations that the silk worm moth, Bombyx mori, accumu-
lates large amounts of both GlcNac and UDP-GlcNac in the moulting fluid
Glycolysis
Hexosamine
pathway
Glucose
Glucose-6P
Fructose-6P
Trehalose
GlutamineGlutamate
Glutamate
Acetyl CoA
CoASH
Glucosamine-6P
N-acetyl glucosamine-6P
N-acetyl glucosamine-1P
UTP UDP
N-acetyl
glucosamine
Chitin
PPi
Hexokinase
Gluc-6p
isomerase
Glutamine
synthase
Trehalase
GFAT
GNA
AGM
UAP 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 con-
version 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 UDP-
N-acetylglucosamine pyrophosphorylase (UAP). UDP-N-acetyl glucosamine units are
finally converted into chitin polymers by chitin synthase.
445Insect Chitin: Metabolism, Genomics and Pest Management
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 he-
molymph sugar in insects. In addition to being the primary precursor of chi-
tin, 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 cryo-
protectant in insects for winter survival in temperate zones. A comprehen-
sive 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 integ-
ument (epidermis). During the larval development of S.exigua, the expres-
sion 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 wan-
dering 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 asso-
ciation between Tre-1 expression and pupal moulting has been observed in
446 Daniel Doucet and Arthur Retnakaran
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, accom-
panied 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 caus-
ing a slightly more potent effect. This difference was also mirrored at the
phenotypic level, with a high proportion of the SeTre-1-RNAi-linked mor-
tality 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 chi-
tin 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 tran-
scripts encoding UDP-N-acetylglucosamine pyrophosphorylase (UAP) and
CHSB. Direct injection of trehalose in the hemocoel was also found to stim-
ulate 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).
447Insect Chitin: Metabolism, Genomics and Pest Management
Hexokinase seems to accomplish double duty in the specific context of chi-
tin 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 glucos-
amine 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 poly-
saccharides, including chitin. Glutamine:fructose-6-phosphate aminotrans-
ferase (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 gluta-
mine aminotransferase (GAT2) and two sugar isomerase (SIS) motifs. A sub-
stantial 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 yeast-
to-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 abun-
dance 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
448 Daniel Doucet and Arthur Retnakaran
mosquito genes induced by blood meal, the upstream region of AeGFAT1
is populated with binding sites for transcription factors of the ecdysone-
response 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 phos-
phate 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 path-
way 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 ner-
vous 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 pres-
sure for distinct UDP-GlcNAc production patterns has been at play in either
449Insect Chitin: Metabolism, Genomics and Pest Management
this species or its ancestor. Arakane et al. (2011) have characterized the two
genes, TcUAP1 and TcUAP2 and teased apart their functions using RNAi-
mediated 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 de-
pleted 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 me-
tabolite 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 in-
crease in GS production to respond to chitin synthesis demand. GS expres-
sion 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).
450 Daniel Doucet and Arthur Retnakaran
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 ac-
tivity 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 fam-
ily, 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 ho-
mologies 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” de-
notes 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
451Insect Chitin: Metabolism, Genomics and Pest Management
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 trans-
membrane 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 struc-
tures 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
Domain A Domain B Domain C
NC
BD
C
AE
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 asred 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 se-
quence 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.
452 Daniel Doucet and Arthur Retnakaran
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
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.
453Insect Chitin: Metabolism, Genomics and Pest Management
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 devel-
opment. 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 den-
ticle 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 tech-
nology 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 func-
tional 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 rela-
tion 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
454 Daniel Doucet and Arthur Retnakaran
almost all insect CHSAs. The amino acid alignment of the two alternative
exons, obtained from the genomic sequence data from 13 species of hol-
ometabola, is presented in Fig. 6.9B.Arakane et al. (2005) were able to
knock-down either or both TcCHSA isoforms, using specific dsRNA mol-
ecules, 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.
Embryo
Treh
00–02h
02–04h
04–06h
06–08h
08–10h
10–12h
12–14h
14–16h
16–18h
18–20h
20–22h
22–24h
12h
12h
24h
2c post WPP
3c post WPP
4c post WPP
Male 1d
Male 5d
Male 30d
Female 1d
Female 5d
Female 30d
Puff.st. 1–2
Puff.st. 3–6
Puff.st. 7–9
New
L1 L2
CG6262
Hex-A
Hex-C
Pgi
Gfat1
Gfat2
Gs1
Gs2
CG1969
CG10627
mmy
kkv
CS-2
-2.1 4.6
L3
White
prepupa Pupa Adult
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 ¼0
and a variance ¼1. Values were collected for the genes encoding the following
enzymes: trehalases—Treh and CG6262; hexokinases—HEX-A and HEX-C; glucose-
6-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.
455Insect Chitin: Metabolism, Genomics and Pest Management
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 mono-
saccharide acts as the acceptor, the donor (chitin) extends by addition at its re-
ducing end. In both Mesorhizobium chitooligosaccharide synthase (NodC) and
zebrafish chitin synthase, evidence points to polymer elongation at the non-
reducing end (Kamst et al., 1997, 1999). However, hyaluronan synthases
class I, enzymes that are closely related to ChSs, appear to catalyse reducing-
end 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 arthro-
pod 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 pro-
vided 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 enzy-
matic systems are required to carefully dismantle the chitin architecture that
surrounds the epidermis and lines the gut. Our discussion will centre on en-
zymatic systems important in the latter context.
2.2.1 Chitinases
Chitinases are among the most abundant enzymes involved in chitin metab-
olism. They are glycosyl hydrolases that sever the b-1,4 glycosidic bond of
chitin with a retention of the anomeric configuration (Arakane and
456 Daniel Doucet and Arthur Retnakaran
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 re-
solved; however, the model of the Drosophila IDGF2 protein, a growth fac-
tor 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 pre-
ceded 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 Tri-
bolium, appears to play an important role in the pupal to adult moult. Finally,
knock-down of TcCHT7 demonstrated its essential function in wing and
457Insect Chitin: Metabolism, Genomics and Pest Management
elytral expansion and abdominal expansion after pupation. The physiolog-
ical 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 bio-
medical 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 stud-
ied 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 dupli-
cation 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 sim-
ilarity. Group I and group II CDAs encode a CBD followed by a low-
density 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 en-
code 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
458 Daniel Doucet and Arthur Retnakaran
knock-downs adversely affected the completion of every type of moult (lar-
val, pupal or pupal–adult). RNAi-treated individuals appear unable to break
from the old exuviae (Dixit et al., 2008). The function of the single repre-
sentative 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 un-
derstood. 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 Tri-
bolium). The expression of these CDAs has been tracked with great precision
by Arakane et al. (2009),using RT-PCR and in situ hybridization. Their re-
sults 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 in-
triguing, 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 infec-
tious. These data indicate that regulation of gut CDA gene expression consti-
tutes 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 hy-
drolysis 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 glucosamine-
containing polymers as a carbon source or in the deglycosylation of proteins.
Insect NAGs have been cloned from various species belonging to the Lep-
idoptera, Coleoptera and Diptera and classified into four groups (NAGs
I–IV), based on their phylogenetic relationships and substrate affinity.
459Insect Chitin: Metabolism, Genomics and Pest Management
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 anal-
ysis 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% mor-
tality 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 chi-
tin hydrolysis.
Group III and group IV NAGs are referred as N-glycan-processing en-
zymes and hexosaminidases, respectively. Group III includes the well-
characterized 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 hydro-
lytic 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
460 Daniel Doucet and Arthur Retnakaran
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 dis-
tantly 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 evolution-
ary 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 nar-
row down hypotheses that address gene function. The modENCODE con-
sortium 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),
461Insect Chitin: Metabolism, Genomics and Pest Management
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 interro-
gate 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 co-
regulated. CG1969, the gene encoding glucosamine-6-phosphate-
acetyltransferase (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, al-
though 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.
462 Daniel Doucet and Arthur Retnakaran
Embryo
00–02h
Cht2
Cht3
Cht4
Cht5
Cht7
Cht8
Cht9
Cht11
Cht12
Hexo1
Hexo2
fdl
Verm
Serp
ChLD3
CDa4
CDa5
CDa9
-1.4 5
02–04h
04–06h
06–08h
08–10h
10–12h
12–14h
14–16h
16–18h
18–20h
20–22h
22–24h
12h
12h
24h
2c post WPP
3c post WPP
4c post WPP
Male 1d
Male 5d
Male 30d
Female 1d
Female 5d
Female 30d
Puff.st. 1–2
Puff.st. 3–6
Puff.st. 7–9
New
L1 L2
L3
White
prepupa Pupa Adult
Cht2
CNS
Mg
Hg
MT
FB
SG
Tr
Car
AH
AE
Br
TAG
Cr
Mg
Hg
MT
FB
SG
He
Car
Ov
Tes
VF St
MF St
MAG
Cht3
Cht4
Cht5
Cht7
Cht8
Cht9
Cht11
Cht12
Hexo2
fdl
Verm
Serp
ChLD3
CDa4
CDa5
CDa9
-1.3 4.8
AdultLarva
B
A
Figure 6.11 Developmental and tissue-specific expression patterns of Drosophila mela-
nogaster genes-encoding enzymes involved or putatively involved in chitin breakdown.
463Insect Chitin: Metabolism, Genomics and Pest Management
Intriguingly, the expression of other members of this family (ChLD3, CDA4
and CDA5) follows an almost identical pattern. The functional characteri-
zation 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 ex-
pression 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 lar-
vae 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 re-
lease #29) promise to refine even more the picture of chitin-related gene ex-
pression 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 immunopre-
cipitation at the genomic scale, to identify regulatory elements important in
chitin-metabolism genes, will enable a better understanding of the transcrip-
tion factor networks that turn on and off these genes (Ne
`gre et al., 2011).
(A) Gene expression across embryonic, larval, pupal and adult development. Expres-
sion 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 ¼0 and a variance ¼1. (B) Tissue-specific expression patterns. Data were recov-
ered 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, hind-
gut; 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,Hexo-
2and 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.
464 Daniel Doucet and Arthur Retnakaran
4. BENZOYLPHENYL UREAS AS CSIs
Targeting a biochemical compartment that is important for the sur-
vival of an insect pest is an attractive proposition for pest management. Chi-
tin 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
Dimilin
Ò
(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 com-
bining 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 Dimilin
Ò
;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
465Insect Chitin: Metabolism, Genomics and Pest Management
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 para-
chlorine substitution on the phenyl end. None of the BPUs have a para sub-
stitution 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 rel-
atively 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 synthe-
sized BPUs with larvicidal activity on the larvae of the rice stem borer, Chilo
suppressalis. They found that on the benzoyl end, simple mono- or di-
substitutions 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 hy-
pothesized 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 inthe 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
466 Daniel Doucet and Arthur Retnakaran
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
Diflubenzuron (Dimilin)— Philips-Duphar BV 1972
Fluxyloxuron (PH 60-23)— Philips-Duphar BV 1988
Lufenuron — Novartis A.G. 1977
Fluazuron — Novartis A.G. 1990
Hexaflumuron — Dow Elanco Ltd. 1984
Noviflumuron — Dow Agro Sciences LLC. 2001
Novaluron — Makkhteshim Agan Industries
1990
Flufenoxuron — Shell International
Co.Ltd. 1987
Chlorfluazuron— Ishihara Sangyo
Kaisha Ltd. 1983
Triflumuron (Alsystin) — Bayer Crop
Science 1982
Teflubenzuron — Celamerck GmBH.
1982
FFOO O
F
F
F
F
F
F
Cl O O O
NN
H
F
F
FO
O
F
F
F
F
FF
F
O
OO
O
O
F
F
O
OCl
Cl
F
F
FF
F
FF
F
Cl
Cl
Cl
F
F
FF
FF
O
Cl
ClFFFF
F
F
F
OO
F
O
N
H
F
FFFF
N
H
O
FF
Cl
Cl
H
OO O
H
Cl Cl
HCl NCF3
NN
FFF
F
Cl
N
HN
H
O
N
H
F
N
H
Cl
O
N
HN
HO
Cl
F
N
HN
H
O
N
H
N
H
O
Cl
N
HN
HO
N
N
HN
H
O
N
H
N
H
O
Cl
N
O
Figure 6.12 Chemical structures of a representative group of benzoylphenyl ureas that
have been commercialized.
467Insect Chitin: Metabolism, Genomics and Pest Management
delayed effect at the time of the moult. During larval-to-larval or larval-to-
pupal 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 inhibi-
tion of chitin formation by these compounds became readily apparent (Palli
and Retnakaran, 1998; Retnakaran, 1995). Radioactively labelled UDP-
GlcNAc 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) in-
hibit chitin synthase in an in vitro tissue preparation from the Australian sheep
blowfly, Lucilia cuprina. Since these crude preparations contained other ma-
terials 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 sys-
tems 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 epi-
dermal 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 retic-
ulum 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
468 Daniel Doucet and Arthur Retnakaran
in chitin synthesis is chitin synthase which is a glycosyl transferase that po-
lymerizes 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
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).
469Insect Chitin: Metabolism, Genomics and Pest Management
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
3
H-GlcNAc.
Diflubenzuron completely inhibited chitin synthase activity in this system.
Various ionophores for K
þ
,Ca
þþ
and H
þ
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 ho-
mogenate 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 activ-
ity in these vesicle preparations. They hypothesized that a vesicular ABC
transporter with ATP- or GTP-dependent Ca
þþ
or K
þ
transport was prob-
ably involved. The most likely candidate was an ATP-sensitive K
þ
channel,
a transporter which combines four sulfonylurea receptor (SUR) subunits
along with four K
ir
6.2 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 K
ATP
channel closes
and the K
þ
level inside the b-cell is upset. This change in intracellular K
þ
content depolarizes the cell membrane, which in turn provokes the opening
of Ca
þþ
channels and allows the influx of Ca
þþ
. In turn, the increase in
intracellular Ca
þþ
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 de-
veloped moult deformities strikingly similar to the ones caused by
diflubenzuron. The two compounds share a common urea bridge with sub-
stitutions 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 K
d
values were 44.9 nM for glibenclamide and
64.9 nM for diflubenzuron in Drosophila vesicles. RT-PCR confirmed the
470 Daniel Doucet and Arthur Retnakaran
b-Cell
Ca
++
influx
Depolarization
Glibenclamide
SUR1
K
ir
6.2
K
+
K
ATP
channel
Ca
++
Exocytosis of
vesicle releasing
insulin
K
ATP
channel(K
ir
6.2) is shut–
intracellular K
+
level upset
Insulin
granules
Ca channel
Figure 6.14 Glibenclamide, a sulfonylurea, upon binding to the sulfonylurea receptor
(SUR1) results in the closure of the K
ATP
channel (K
ir
6.2) which depolarizes the
membrane and allows the influx of Ca
þþ
into the bcell resulting in the exocytosis of
the vesicle releasing the insulin granules. (Based on Aittoniemi et al., 2009;
Matsumura, 2010.)
O
OO O
S
N
HN
H
N
H
OCH
3
Glibenclamide
Diflubenzuron
Cl
O
N
H
F
F
N
H
Cl
O
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.
471Insect Chitin: Metabolism, Genomics and Pest Management
presence of DSUR in Drosophila larvae. The vesicle preparations treated with
diflubenzuron or glibenclamide showed
45
Ca
þþ
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 ques-
tion that remains to be answered. Nevertheless, SUR being the target of
diflubenzuron, based on the studies with glibenclamide, is an attractive hy-
pothesis and when taken together with the recent chitin synthase model, it ap-
pears 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 ma-
chinery is probably formed in the endoplasmic reticulum and the Golgi
where it matures. The vesicle migrates to the plasma membrane and exocy-
toses 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 up-
or 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 genomics-
enabled methods.
4.4. Effects on pests
Introduction of the organochlorine insecticide DDT in 1939 revolutionized
pest control. It was soon followed by organophosphates, carbamates, pyre-
throids, 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
472 Daniel Doucet and Arthur Retnakaran
way or another, target the nervous system of not only insects but also other
animals including humans which share a common neurophysiology. Indis-
criminate and excessive use of neurotoxic insecticides soon manifested their
adverse effects in ecological situations and human health which was dramat-
ically brought into focus by Rachel Carson in her book, “The Silent Spring”
in 1962 alluding to the disappearance of songbird species by DDT. Contin-
uous use at high levels often resulted in the pest populations developing re-
sistance 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 con-
trol, often developed from natural products, made its appearance, and prod-
ucts developed under this new trend were called “biorational pesticides”.
A
C
Chitin
CO2
Plasma membrane
Extracellular
cuticular region
Intracellular
region of the
epidermal cell
UDP + PPi
UDP-GlcNAc
Vesicle
A
BC
DFB / glibenclamide
SUR
CHS
Gol
g
i
B
NH2CA
ES
Figure 6.16 Mode of action of diflubenzuron (DFB) through the ABC transporter, sulfo-
nylurea receptor (SUR), by interfering with the exocytosis of chitin synthase (CHS) of
Drosophila. Normally, the CHS vesicle binds with the plasma membrane where it initi-
ates chitin synthesis and SUR which includes the K
ATP
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 se-
quences are not shown. (Based on Matsumura, 2010; Merzendorfer, 2006; Merzendorfer
et al., 2012; Moussian, 2010; Muthukrishnan et al., 2012.)
473Insect Chitin: Metabolism, Genomics and Pest Management
“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 arthro-
pods 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 re-
place the use of IGRs, as the CSIs like the bisacylhydrazine (ecdysone ag-
onists) 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 commer-
cialized 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 Dimilin
Ò
)
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 effec-
tive 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
474 Daniel Doucet and Arthur Retnakaran
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 Colo-
rado potato beetle, Leptinotarsa decemlineata, than lufenuron or hexaflumuron
(Karimzadeh et al., 2007).
Diflubenzuron is relatively nontoxic to mammals and birds. The LD
50
for rats and mice is >4640 mg/kg. The LC
50
for zebra fish (96 h) is
>0.2 mg/L, and it is nontoxic to bees (LD
50
>100 mg/bee). It is adsorbed
by clay soil and has a half-life of <7 days being degraded by microorganisms.
Crustaceans, in general, are sensitive to diflubenzuron and care has to be
exercised to prevent the material going into the water (Gartenstein et al.,
2006). A suitable buffer zone around the fresh water lakes and coastal areas
needs to be maintained when diflubenzuron is used. This pattern of activity
against pests and toxicology can be extended to most of the BPUs.
4.4.2 Flucycloxuron
This is a second generation BPU that has been optimized as an acaricide.
A dispersible concentrate with special surfactants for topical contact activity
was developed to target a wide spectrum of mites. It has been shown to have
transovarial–ovicidal, ovicidal and ovo-larvicidal activities. Unlike
diflubenzuron, which was effective on only Eriophyid mites, Flucycloxuron
has a wider spectrum of activity and controls both Eriophyid and Tetra-
nychid mites. On apple leaves, it was effective on the two spotted spider
mite, T.urticae, as well as the European red mite, Panonychus ulmi
(Grosscurt, 1993). It appears to be more active than diflubenzuron and pen-
etrates the leaf cuticle. Its toxicology profile is similar to that of
diflubenzuron but might be marginally more toxic in certain respects such
as toxicity to rainbow trout, Oncorhynchus mykiss, and the water flea, Daphnia
(Darvas and Polgar, 1998).
4.4.3 Lufenuron
This BPU is in many respects more active than diflubenzuron and is exten-
sively used in controlling the mushroom sciarid fly, Lycoriella ingenua, which
is one of the most common fly pests affecting the cultivation of the common
475Insect Chitin: Metabolism, Genomics and Pest Management
mushroom Agaricus bisporus (Erler et al., 2011). Lufenuron has been tested
against termites, Reticulitermes hesperus, using baited wood, and the colony
dispersion was monitored by microsatellite analysis. In a period of
10–16 months, wood consumption in the baited sites ceased (Haverty
et al., 2010). Lufenuron has been successfully used against many lepidopteran
pests due to its classic larvicidal effect along with transovarial–ovicidal and
ovicidal effects. It has low toxicity against many parasitoids and has adequate
persistence making it effective on many pests. In New Zealand, both pre-
and post-harvest control against the Tortricid, the light brown apple moth,
Epiphyas postvittana, was initially achieved by tebufenozide (an ecdysone ag-
onist; Chapter 2) and an organophosphate. But the insects soon developed
cross resistance to the organophosphate, which has been since then success-
fully replaced by lufenuron (Whiting et al., 2000).
4.4.4 Fluazuron
Fluazuron has been shown to be effective against ticks and mites. It is effec-
tive against the hard tick, Rhipicephalus sanguineus, and the sarcoptic mange
mite on pigs, Sarcoptes scabiei. Normal treatment for scabies has been per-
methrin or ivermectin, but these do not have any effect on eggs. Combining
permethrin or ivermectin with fluazuron has been shown to be more effec-
tive (de Oliveira et al., 2012; Pasay et al., 2012). Plague epizootics caused by
the bacterium, Yersinia pestis, has been traced to wild populations of rodents
and their fleas in parks and outdoor areas. Since controlling the rodents
would only force the fleas to jump on to other mammals, a programme
to control fleas on wild animals using food cubes impregnated with
lufenuron or fluazuron and a fluorescent-tracking dye was initiated in a
California park. A 6-year trial showed that the flea population was
successfully lowered in all the squirrels and mice but not on chipmunks
(Davis et al., 2008).
4.4.5 Hexaflumuron
The typical use pattern for hexaflumuron has been against the larval stages of
Lepidoptera, Coleoptera and Diptera. Another major use has been in bait
incorporation against the subterranean termites. Field evaluation against
the eastern subterranean termite, Reticulitermes flavipes, showed a reduction
in termites over a 15 m radius from the point of the bait station and was one
of the best among the various insecticides tested (Ripa et al., 2007).
Hexaflumuron baits were used to control the Formosan subterranean ter-
mite, Coptotermes formosanus, over a 12.5-ha plot in an area near New
476 Daniel Doucet and Arthur Retnakaran
Orleans over a 3-month period. Reinvasion took over 7 months, and the
invaders were distinct from the colonies eliminated and neighbouring col-
onies as shown by DNA fingerprinting (Messenger et al., 2005).
4.4.6 Noviflumuron
This is one of the newest (2001) BPUs to enter the fray. It has been found to
be especially active against cockroaches and termites and relatively fast act-
ing. Against the German cockroach, B.germanica, noviflumuron acts faster
than hexaflumuron, and the activity is only by ingestion. The high toxicity
is accentuated by the grooming of spray deposits from the antennae and tarsi
(Ameen et al., 2005). The Formosan subterranean termite, C. formosanus,is
refractory to most control measures because of the way they spread, but
noviflumuron is particularly effective against this insect. Tracking
C.formosanus families is complicated by the fact that simple families are headed
by a pair of reproductive individuals, but extended families are headed by mul-
tiple reproductives. Husseneder et al. (2007) used microsatellite genotyping
before and after treatment to follow the various colonies. A year after treat-
ment, all the colonies vanished excepting for one colony which reappeared,
but the latter was found to be a colony from an untreated area.
4.4.7 Flufenoxuron
Flufenoxuron has been traditionally used against Lepidopteran larvae on
fruits, vegetables, cotton and grain crops with various degrees of success.
For instance, against the cotton leafworm, Spodoptera littoralis, this product
came second best after lufenuron (El-Sheikh and Aamir, 2011). Against
the mushroom sciarid fly, L.ingenua, flufenoxuron fared better than
novaluron, diflubenzuron and teflubenzuron (Erler et al., 2011).
4.4.8 Chlorfluazuron
Chlorfuazuron is truly a broad-spectrum IGR, being active against thrips,
white flies, most lepidopterans, coleopterans, hymenopterans and dipterans.
It has to be ingested in order to be effective and has low environmentalimpact.
All BPUs, in general, and chlorfluazuron, in particular, have an extremely low
impact on adults of the egg parasitoid, Trichogramma nubilale, compared to
neonicotinoids, avermectins, pyrethroids, organophosphates and carbamates
(Wang et al., 2012b). Chlorfluazuron fared well in controlling the Formosan
subterranean termite, C.formosanus, and the eastern subterranean termite,
R.flavipes, as compared to diflubenzuron and hexaflumuron (Osbrink
et al., 2011). When bees were exposed to spinosad, oxymatrine or
477Insect Chitin: Metabolism, Genomics and Pest Management
chlorfluazuron, the LC
50
was 7.34, 10.68 and 2526 mg/L, respectively, show-
ing clearly the safety of chlorfluazuron (Rabea et al., 2010).
4.4.9 Triflumuron
The pest control spectrum of triflumuron covers a whole assemblage of in-
sect groups that include the apple leafminer, boll worm, cabbage moth, cod-
ling moth, cotton leafworm, psyllids, summer fruit moth, tortrix moth
among others. Several BPUs were tested against the mushroom sciarid,
L.ingenua, as a soil drench and triflumuron fared better than novaluron,
diflubenzuron and teflubenzuron (Erler et al., 2011). A combination treatment
using a pyrethroid, cyfluthrin, against the adults and triflumuron against the
larvae was successfully used to control the mealworm, Alphitobius diaperinus,
in broiler and turkey houses (Salin et al., 2003). The sheep blowfly, Lucilia
sericata, has been effectively controlled using odour-baited triflumuron-
impregnated targets. Aluminium sheets covered with a white cloth dipped
in a solution containing sugar, liver homogenate, sodium sulphide and a
10% suspension of triflumuron served as target stations for the flies. Flies walk-
ing and feeding on the mixture wereeffectively coated and controlled through
a combination of ovicidal and larvicidal activities (Smith and Wall, 1998).
4.4.10 Teflubenzuron
When newly emerged females of the migratory locust, Locusta migratoria,
were fed with either a methanolic extract of the plant, Haplophyllum
tuberculatum, or teflubenzuron, the latter treatment completely blocked
egg hatch (Acheuk et al., 2012). Atlantic salmon in fish farms have been
plagued by sea lice (a copepod ectoparasite), Lepeophtheirus salmonis. The lice
have 10 developmental stages starting with two free-swimming nauplius
stages (nauplius 1 and 2), copepodit stages 3 when it attaches itself to the fish,
the chalimus stages 4–7 when it starts feeding on the salmon and, finally,
8 and 9 pre-adult mobile stages before reaching the 10th which is the adult
stage which is also mobile. Teflubenzuron was administered along with the
feed at the rate of 10 mg/kg for 7 days to salmon in cages. The chalmis, pre-
adult and adult stages were counted at weekly intervals, and they were sig-
nificantly low compared to the controls (Campbell et al., 2006).
4.4.11 Novaluron
Among the BPUs, it is one of the late entries, making its debut in 1990. It has
been shown to be an effective IGR against several lepidopteran, coleop-
teran, homopteran and dipteran pests. In the United States, it has been
478 Daniel Doucet and Arthur Retnakaran
formulated for controlling pests of apples, potatoes, brassicas, ornamentals
and cotton. It has also been designated as a reduced risk alternative to organ-
ophosphate insecticides. It exhibits low acute mammalian toxicity, no sig-
nificant sub-chronic effects on mammals and poses low risk to the
environment and non-target organisms. It is well suited for IPM and inte-
grated resistant management (IRM) programmes (Cutler and Scott-Dupree,
2007). Ishaaya has done considerable work on novaluron and demonstrated
its effectiveness against many pests such as white flies, leafminers, the beet
armyworm and many other important pests (Ishaaya and Horowitz, 1998;
Ishaaya et al., 1996, 1998). In Brazil, the dengue vector mosquito,
A.aegypti, has been traditionally controlled by neurotoxic insecticides,
but repeated use over many years has made these pesticides ineffective. As
an alternative, novaluron has been successfully used to keep the
population down. Various physiological parameters, including the
inhibition of chitin synthesis, have been well studied after novaluron
treatment (Farnesi et al., 2012).
4.5. Effects on non-target species
It was described earlier that BPUs act on organisms that actively synthesize
chitin with the exception being the fungi. These compounds are relatively
safe to vertebrates, in general, and mammals, in particular. The real environ-
mental impact is on non-target arthropods, typically aquatic crustaceans.
The effect on many such organisms has been described in earlier reviews
(Dhadialla et al., 2005; Retnakaran et al., 1985). One of the most
sensitive crustaceans to diflubenzuron is the freshwater flea, Daphnia, and
has been used as an indicator species in environmental toxicity studies.
During the moulting process, the old cuticle is degraded by enzymes in the
moulting fluid and a new cuticle is synthesized. The moulting fluid which
is released into the surrounding water during ecdysis contains, among other
chemicals, the enzymes chitinase, which hydrolyses the chitin into
oligomers and trimers and chitobiase which hydrolyses these into
monomers (GlcNAc). The chitobiase activity in the water is an indicator
of the health of the aquatic population and is directly correlated to the
robustness of the organisms (Hanson and Lagadic, 2005). For Daphnia,the
LC
50
at 48 h is 7.1 mg/L of diflubenzuron. The effect of diflubenzuron on
D.pulex and D.magna was studied using the chitobiase activity assay. As
the concentration of Diflubenzuron increased, the chitobiase level
decreased indicating the negative impact of diflubenzuron. D.magna was
479Insect Chitin: Metabolism, Genomics and Pest Management
more sensitive than D.pulex (Duchet et al., 2011). Utilizing a suitable buffer
zone between the spray area and the water body will lessen the impact of
BPUs on aquatic crustaceans.
BPUs are generally not deleterious to adult insects, due to lack of contact
activity, unless they are specially formulated. If high concentrations are con-
sumed (which is unlikely), then there is a risk of a transovarial effect on eggs,
leading to the failure of eggs to hatch. Pollinators in the spray zones are gen-
erally not affected because of the low level of exposure. Fields treated with
35–400 g/ha of diflubenzuron are safe for honey bees, but anything above
300 g/ha is harmful to bumble bees (Tasei, 2001). Larvae of the Caribbean
fruit fly, Anastrepha suspensa, were parasitized by the wasp, Biosteres
longicaudatus, and they were treated topically by three different concentra-
tions, 100, 500 and 1000 ppm of diflubenzuron. At the higher two levels,
the parasites had difficulty in locating the hosts. Fewer eggs were laid, but
they were all viable (Lawrence, 1981).
The egg parasitoid commonly known as the “stingless wasp”,
Trichogramma, has been used extensively around the world as a biological
control agent by itself or in combination with a soft pesticide in IPM. Over
230 species in the family Trichogrammatidae are known, and they are often
used as indicator species for testing the adverse effects of insecticides on par-
asitoids. The toxicity of 30 insecticides belonging to seven chemical classes
was tested on Trichogramma japonicum, an egg parasitoid of lepidopteran pests
of rice. Organophosphates (e.g. fenitrothion) and carbamates (e.g. carbaryl)
were the most toxic with an LC
50
that ranged from 0.035 to 0.49 mg AI/L
followed by antibiotics (e.g. abamectin), phenylpyrazoles (e.g. fipronil), py-
rethroids (e.g. cypermethrin) and the neonicotinoids (e.g. imidacloprid).
The IGDs, chlorfluazuron, fufenozide, hexaflumuron and tebufenozide
were the least toxic with an LC
50
ranging from 3383 to 30,206 mg AI/L
(Zhao et al., 2012). Adult predators such as coccinellids (ladybird beetles),
predatory mites and spiders are unaffected by novaluron, but the juveniles
are typically susceptible (Cutler and Scott-Dupree, 2007).
4.6. Environmental degradation
BPUs tend to be adsorbed strongly to soil and cuticular waxes which tends to
extend their half-life as well as protect them from degradation. While this
may extend their insecticidal activity, it raises environmental concerns about
persistence. It can be argued that when tightly absorbed to the soil, BPUs
become unavailable to organisms, thus unlikely to cause any toxic effects.
480 Daniel Doucet and Arthur Retnakaran
These compounds are degraded by conjugation, hydrolysis or both. The
urea bridge is split at the benzoyl end, resulting in a benzamide and a phen-
ylurea followed by further degradation. The fate of diflubenzuron has been
well studied and has been summarized by Retnakaran (1995, 1985).
Degradation of novaluron depends on the soil type and the presence of
microorganisms. In unsterilized alluvial soil, its half-life is 17.0–17.8 days,
whereas in sterilized alluvial soil, it is prolonged to 53.7–59.0 days. In coastal
saline unsterilized soil, its half-life ranges from 11.4 to 12.7 days, but in ster-
ilized soil, it is 28.9 to 29.8 days. It is evident that both abiotic and biotic
factors play a role in the degradation of this compound. These results can
probably be extended to all BPUs (Das et al., 2008).
In recent years, liquid chromatography with tandem mass spectrometry
detection (LC–MS/MS) has made analysis of pesticide residues fast, sensitive
and reliable. There is minimal clean-up required and a large number of com-
pounds can be simultaneously analysed. Nine BPUs were simultaneously
detected from fruits, vegetables, cereals and animal products. Samples were
homogenized with diatomaceous earth and extracted with ethyl acetate at
80 C and 1500 psi (pressurized liquid extraction), concentrated and
analysed in an LC–MS/MS (Brutti et al., 2010).
5. NON-BENZOYLPHENYLUREA CHITIN SYNTHESIS
INHIBITORS
Several compounds other than BPUs that inhibit chitin synthesis have
been reported. These products do not bear any resemblance to BPUs and do
not share any structural similarity among themselves but appear to inhibit
chitin formation in different ways. They have been successfully used as con-
trol agents, and their environmental impact has been investigated.
5.1. Structure and properties
Etoxazole, buprofezin and cyromazine are three of the non-BPU CSIs that
have been successfully used as insecticides (Fig. 6.17). They have been as-
sembled together because of their common characteristic of interfering with
moulting and chitin formation.
5.1.1 Etoxazole (TetraSan
Ò
)
Etoxazole was developed in 1994 by Yashima Chemical Industry Co. in
Japan. It is classified as a 2-diphenyl-1,3-oxazoline and was developed as
an acaricide against tetranychid spider mites such as Panonychus and
481Insect Chitin: Metabolism, Genomics and Pest Management
Tetranychus species. It also has been shown to have insecticidal activity against
aphids, leafhoppers, the fall armyworm and the diamond back moth (Naun
and Smagghe, 2006). On spider mites, it is active against eggs, larvae and nymphs
but not on adults. The rat LD
50
is >5000 mg/kg and it has some effect on the
liver and the prostate in the dog. Degradation of etoxazole in the soil is slow and
also undergoes partial photolysis. Its half-lifeinclayloamis52.0days,butinall
other types of soil, it is <30.0 days (EPA petition for Etaxazole, 2010).
5.1.2 Buprofezin (Applaud
Ò
)
This compound, developed by Hoechst, not only inhibits chitin synthesis
but also acts on cholinesterase and has been used extensively against the
whitefly, Bemisia tabaci (Cottage and Gunning, 2006). It has good activity
Etoxazole
Buprofezin
C
y
romazine
F
H
3
C
CH
3
CH
3
CH
3
CH
3
CH
3
CH
3
CH
3
H
3
C
N
O
FO
O
O
N
NH
NH
2
H
2
N
N
N
N
N
N
Figure 6.17 Non-benzoylphenylurea chitin synthesis inhibitors.
482 Daniel Doucet and Arthur Retnakaran
against homopterans such as mealy bugs, scale insects and whiteflies.
Buprofezin suppresses oviposition in adult homopterans and reduces viabil-
ity of eggs while at the same time inhibiting chitin synthesis during the
growth stages. Generally, it takes 3–7 days for it to act but feeding damage
is low and by and large not disruptive to beneficial insects and mites. Typ-
ically, it is used at a rate of 0.25–0.38 lb AI/acre, applied twice a year per
season on almonds, bananas, citrus, cotton, cucumbers, grapes, lettuce,
melons, pumpkins, squash and tomatoes. It is nontoxic to birds but mildly
toxic to mammals and is not acutely toxic to aquatic organisms (summarized
in Palli and Retnakaran, 1998).
5.1.3 Cyromazine (Larvadex
Ò
)
It is an aminotriazine and a cyclopropyl derivative of melamine developed as
a pesticide by Ciba-Geigy (now Novartis). It acts both as an insecticide and
as an acaricide and has contact activity that interferes with moulting and pu-
pation similar to BPUs (Fig. 6.18). It is also systemic and has translaminar
activity. It has many interesting applications, one of which is on beans
(Phaseolus vulgaris) grown hydroponically where it is added to the nutrient
solution at the rate of 20 mg/plant to control mite and insect pests
(Patakioutas et al., 2007). It has been successfully used against insect
pests of vegetables, mushrooms, potatoes and ornamentals. It provides
excellent control of stable fly maggots in winter hay (Taylor et al., 2012).
A
B
Figure 6.18 Two second instars of the Colorado potato beetle, Leptinotarsa
decemlineata, 72 h after treatment with cyromazine. A blister filled with hemolymph
protruding from prothorax (A) and the hindgut everted from the anus (B) can be
observed. (After Karimzadeh et al., 2007, with the permission of Professor Hejazi).
483Insect Chitin: Metabolism, Genomics and Pest Management
The MOA was thought to be due to an inhibition of either phenylalanine
hydroxylase during sclerotization or inhibition of dihydrofolate reductase
(DHFR), but both have been disputed and the question remains unresolved
(Bel et al., 2000).
5.2. Environmental fate and effects
All three compounds persist in the environment to various degrees and are
broken down or conjugated in the soil over time. This process is accentuated
by soil microorganisms and affected by pH. Etoxazole poses minimal risk to
fresh- and saltwater fish. It is highly risky, however, to fresh- and saltwater
invertebrates but still has a substantially lower impact than most competitive
products. It also poses a minimal risk to avian species, but again lower than sim-
ilar products. Bioconcentration is minimal, and the material is rapidly cleared
from the system. The LD
50
for mallard ducks is >200 mg/kg and no reproduc-
tive toxicity has been recorded. Contact and oral toxicity to honeybees was
>200 mg/bee. It persists in water in the dark with a half-life of 161 days at
pH 7 but photodegrades with an average half-life of 17.4 days. Soil half-life
is from 9 to 52 days. Bioaccumulation is minimal with a low octanol/water par-
tition coefficient, log P
ow
¼5.52 at 20 C. Bioaccumulation is only marginally
above the level of concentration (EPA petition for Etaxazole, 2010). It has no
inhibitory effect on brain acetylcholinesterase (U
¨ner et al., 2006).
Buprofezin has been found to be moderately toxic to mammalian systems.
Some dermal and liver effects were noticed in rabbits at 1000 mg/kg. It is
slightly persistent in the environment compared to other insecticides with a
half-life of 51 days. Minimal photolysis occurs with this chemical. Field dis-
sipation in sandy loam had a half-life of 38.1 days (EPA Buprofezin, 2003).
Buprofezin degradation in rice fields in Japan was simulated in the laboratory,
and it indicated that in flooded soils the half-life was 104 days, whereas in up-
land soils it was 80 days. The half-life was extended when sterile soils were
used (Funayama et al., 1986).
Cyromazine is rapidly metabolized in rats with 97% of the administered
dose being excreted in the urine within 24 h. Methylation, hydroxylation
and N-dealkylation are the metabolic processes involved. Approximately
7% is converted into melamine, 11% into hydroxycyromazine and
methylcyromazine and the rest is excreted.
14
C-labelled cyromazine metab-
olism was studied in rats, hens, goats and sheep and >95% was recovered
(Fig. 6.19). Environmental fate of cyromazine in the soil showed that mel-
amine was the major degradation product formed. The half-life of
484 Daniel Doucet and Arthur Retnakaran
cyromazine in the soil ranged from 2.9 to 107 days depending on the type of
soil. One of the key ways in which cyromazine gets degraded is photolysis.
The calculated half-life after photolytic degradation in sunlight is 3.5 days.
The overall metabolic and environmental performance is similar to other
comparable insecticides (Dorne et al., 2012). Judicious use of this insecticide
with caution is the conclusion one can draw from the environmental impact
studies.
6. CHITINASES AND CHITINASE-INHIBITING CHEMICALS
FOR PEST MANAGEMENT
The activity displayed by chitinases makes them in theory very attrac-
tive as pest management tools. The large number of chitinase genes
expressed by insects reflects the fundamental need to carefully control chitin
R, G, S, H
R, G GCyromazine
Hydroxy-cyromazineMethyl-cyromazine
Melamine
R = Rat
G = Goat
H = Hen
S = Sheep
NH
N
N
N
NH
2
H
2
N
NH
N
N
N
NH
2
HO
N
N
N
NH
2
NH
2
H
2
N
NH
N
+
N
N
NH
2
H
2
N
Figure 6.19 Cyromazine metabolism in selected vertebrates.
485Insect Chitin: Metabolism, Genomics and Pest Management
degradation in various tissues and as part of the normal ontogenetic pro-
gramme. As the destruction of chitin is deleterious in any other circum-
stance, chitinases are prime candidates for effective and targeted insect
pest control. Numerous microorganisms secrete chitinases to breach the
insect cuticle and accelerate the penetration of their hosts. This activity
contributes to the success of several arthropod biocontrol agents currently
used in the field (Fernandes et al., 2012).
6.1. Fungi and microorganisms with chitinase activity
The two most successful fungal entomopathogens, Beauveria bassiana and
Metarhizium anisopliae, are known to secrete chitinases during the process
of host infection (Bogo et al., 1998; Fang et al., 2005; Kang et al., 1999;
Krieger de Moraes et al., 2003; St Leger et al., 1996). The B.bassiana
BbChit1 gene and its upstream regulatory sequence were cloned by Fang
et al. (2005), and the chitinase enzyme was characterized both in vitro and
in vivo. Bbchit1 is a 33-kDa protein with homology to both Aspergillus
and Trichoderma family 18 (endo-) chitinases. Transformation and
overproduction of the protein in B.bassiana, using a constitutive
Aspergillus gene promoter were successfully achieved, and the new strain
used against aphids in bioassays. The recombinant strain displayed
significantly lower values of LC
50
and LT
50
(lethal time necessary for 50%
mortality), when compared to the non-transformed (wild-type) strain
and, therefore, Bbchit1 provided increased virulence. By contrast,
manipulating the expression of a similar chitinase gene in M.anisopliae
did not result in increased virulence in the host Manduca sexta (Screen
et al., 2001). The M.anisopliae Chit1 chitinase gene was cloned from two
different isolates and found to encode 44 kDa proteins differing in
isoelectric points. Overexpression of the basic Chit1 enzyme in a wild-
type M.anisopliae using the constitutive Aspergillus promoter failed to
increase virulence, for reasons that are not entirely clear.
In an interesting twist, Fan et al. (2010) conducted further genetic engi-
neering involving a chitinase, this time by joining the CBD, BmChBD from
aB.mori chitinase to the subtilisin-like protease CDEP-1 from B.bassiana
and expressing the fusion protein either in vitro or in vivo. The fusion protein
showed greater insecticidal activity than the wild-type CDEP-1, possibly by
altering the interaction of the enzyme with host chitin. The synergy
between proteases and chitinases is known to be important in the fungal in-
fection process. Mixing B.bassiana CDEP-1 and BbChit1 results in greater
486 Daniel Doucet and Arthur Retnakaran
insect cuticle degradation. Fusing CDEP-1 and BbChit1 together as a hybrid
enzyme and transforming the corresponding construct in a B.bassiana
recipient strain (along with a constitutive Aspergillus promoter) conferred
hypervirulence towards insect hosts (Fang et al., 2009). Further manipula-
tion of the chitinases and associated proteases during the infection and col-
onization process should be able to modulate the B.bassiana and M.anisopliae
virulence phenotypes.
The chitinases of entomopathogenic bacteria have also been important
to kill or otherwise successfully parasitize their hosts. For example, the bac-
terium, Yersinia entomophaga, produces a toxin complex, comprising TcA,
TcB and TcC, that is instrumental in conferring entomopathogenicity
(Hurst et al., 2011). In addition, the Yen-Tc toxin produced by
Y.entomophaga strain MH96 confers a broad spectrum of insecticidal activity
by ingestion. Recent 3D structure and enzymatic activity determination
revealed that the Yen-Tc complex is in fact an endochitinase, and these sub-
units may decorate the scaffold of the TcA toxin complex (Landsberg et al.,
2011). This chitinase, along with other secreted chitinases, is hypothesized
to confer the extremely rapid (less than 72 h) speed of kill and broad
spectrum of Y.entomophaga (Busby et al., 2012).
6.2. Baculoviruses with chitinase gene
Baculoviruses are important viral control agents of insects, especially those in
the order Lepidoptera. Many of these viruses require the production of
chitinases in the final stage of infection so that infectious particles (occluded
viruses) can be released successfully in the environment. The Autographa
californica Nucleopolyhedrosis virus (AcMNPV) chitinase gene chiA was
the first such gene cloned and characterized from a baculovirus (Hawtin
et al., 1995). The gene encodes a 551-amino acid protein of 58 kDa that
localizes in the cytoplasm of infected cells. The gene appears late in the
AcMNPV infection cycle, consistent with a need of chitinase activity for
the rupturing of the cuticle of individuals liquefied by the infection. A care-
ful comparison of chiA with chitinases from various organisms led the
authors to conclude that the gene was acquired by horizontal transfer from
a bacterium, rather than from an insect.
The chiA enzymes have also been shown to work in concert with a pro-
tease, the cathepsins V-CATH, to destroy the cuticle of infected cadavers.
Hom and Volkman (2000) provided evidence that pro-V-CATH requires
the presence of chiA in order to be processed into the active form V-CATH.
487Insect Chitin: Metabolism, Genomics and Pest Management
AcMNPV chiA thus appears to display a chaperone-like activity towards
proV-CATH and given its intracellular location, the direct interaction be-
tween the two proteins contributes to the retention of pro-V-CATH in the
cytoplasm (Hodgson et al., 2011). Manipulating the expression of the
AcMNPV chiA gene appears to be a viable avenue in the improvement of
the efficacy of the wild-type virus. Studies on the modification of the chiA
promoter have already been initiated (Hodgson et al., 2007).
6.3. Chemical inhibitors of chitinase
The search for small-molecular-weight chitinase inhibitors has been on-
going for several years, yielding numerous lead compounds such as
allosamidin-, agrifin-, argadin-, purine-based compounds and several others
(Cohen, 2010). However, due to their broad spectrum of activity (with their
possible concomitant adverse affects on non-target organisms), none of the
compounds have reached the market as insecticides yet.
Allosamidin is a trisaccharide-like molecule that competitively inhibits
chitinases. Its basic structure presents two successive b-1,4-N-GlcNac units
joined to an allosamizoline group (Sakuda et al., 1986). Several further
modifications of the allosamidin basic plan have been attempted, with vary-
ing impacts on chitinase activity (Huang, 2012; Terayama et al., 1993).
Allosamidin was shown to be active against chitinase from the aphid,
Myzus persicae (Francis et al., 2012), the moth, Ostrinia furnacalis (Wang
et al., 2012a) and several other insect species (see Cohen, 2010) as well as
the hard tick, H.longicornis (You et al., 2003). The molecular mechanisms
by which allosmidin binds to O.furnacalis chitinases have been studied by
way of 3D docking and cluster analysis. Wang et al. (2012a) proposed
that the compound binds to the 2/3 sugar-binding site in chitinase.
In doing so, the enzyme is arrested in a catalytic transition state that
precludes completion of the hydrolysis reaction.
Cyclopeptides are a second group of promising chitinase inhibitors.
These compounds were isolated from fungal sources and include argifin
and argadin (Hirose et al., 2010).
Like allosamidine, they are competitive inhibitors, but their wide spec-
trum of binding activity beyond fungal and insect chitinases, including hu-
man acidic mammalian chitinase, makes it unlikely that they will be used in
pest management. Other compounds in the early stages of development
might prove more promising in achieving a high specific efficacy against
insects. In this regard, the psammaplins and styloguanidin are cyclic peptide
488 Daniel Doucet and Arthur Retnakaran
metabolites derived from marine sponges. In contrast to allosamidin,
argifin and argadin, psammaplin is a non-competitive inhibitor, implying
that a more insect-specific binding pocket could potentially be targeted
(Cohen, 2010).
7. RESISTANCE AND RESISTANCE MANAGEMENT
In 1914, Melander observed that the San Jose scale, Aspidiotus
perniciosus, infesting apple trees in an orchard in Clarkston valley
(Washington State) could no longer be controlled by the prevailing
inorganic insecticide, sulphur-lime, and this was the first recorded case of
insecticide resistance. According to the Michigan State University database,
as of 2008, there are 7747 cases of resistance against 331 insecticides
(Mota-Sanchez et al., 2008).
Resistance originates because of the excessive application of selection
pressure by the repeated use of an insecticide over time. At least three factors
are critically linked to the development of resistance to insecticides. (1) An-
cestral detoxification pathways: Insects and plants have co-evolved over a
long geological time period. The oldest recorded plant, Cooksonia species
dates back to the Silurian, some 425 million years ago. Insects originated
during the Devonian period, about 400 million years ago according to
the discovery of the first springtail fossil, Rhyniognatha hirsti. Just to place
it in perspective, humanoid fossils are <1 million years old. The 400 million
years of co-evolution between insects and plants has been a battlefield of
constant chemical warfare between the two. Every time a host plant makes
a compound to ward off an insect herbivore, the insect comes up with a way
to detoxify the defence chemical. This constant arms race has resulted in in-
sects having a library of genetic schemes to thwart multiple forms of plant
defences. Much of these detoxifying protocols can lie dormant when they
are not needed only to be called into action when an insecticide is intro-
duced. Only this time it is a human intervention and not a plant creation.
(2) Reproductive capacity: Insects, in general, have a great propensity to
procreate in large numbers, increasing the probability of random mutations.
This in turn increases the probability of beneficial mutations, for example,
those conferring the ability to neutralize insecticides, and the mutation to
arise in the general susceptible insect population. (3) Repeated exposure:
Constant heavy use of a pesticide over a time period increases the selection
pressure exponentially which leads to the development of new resistance
mechanisms. Resistance is often quantified and reported as resistant ratios
489Insect Chitin: Metabolism, Genomics and Pest Management
(RR
50
) which is the LD
50
of the field population divided by the LD
50
of the
susceptible population (Rodrı
´guez et al., 2007). It takes ordinarily several
generations to build resistance to any insecticide, and it is usually an auto-
somal recessive. Recently, a novel way to acquire resistance within one gen-
eration was reported (Kikuchi et al., 2012). The bean bug, Riptortus pedestris,
harbours a symbiotic bacterium, Burkholderia species, in the intestine which
can degrade fenitrothion. This bacterium is acquired from the soil by the
nymph of the bean bug at every generation (Fig. 6.20).
Sooner or later, all insecticides have the potential to generate resistance,
and this phenomenon is dependent inter alia on the usage pattern. In this re-
gard, it is interesting to follow the development of BPUs and the evolution
of resistance. Diflubenzuron, which was introduced in the 1970s, has devel-
oped widespread resistance. The repeated use of this BPU against the Egyp-
tian leafworm, S.littoralis, in cotton fields in Egypt resulted in resistance
(Elghar et al., 2005). But the most recent entry of BPU for field
use, noviflumuron, which was introduced in 2001 by Dow Agro Sciences,
has so far not showed any resistance. Fluazuron, hexaflumuron,
noviflumuron and novaluron are the four BPUs that are resistance
free, an outcome possibly due to a careful usage extending the life of these
pesticides (Whalon et al., 2008). Diflubenzuron resistance has been reported
in the codling moth, Cydia pomonella, infesting fruit trees and walnut and also
the Australian sheep blow fly, L.cuprina, on sheep; the house fly, M.dome-
stica, on various foods; the diamond back moth, Plutella xylostella, on cruci-
fers and Nasturtium; the beet armyworm, S.exigua, on cotton, tomato,
AB
Figure 6.20 The bean bug, Riptortus pedestris, (A) and its digestive system (B) illustrating
a new type of resistance mechanism utilizing a symbiotic bacterium. Arrow points to the
part of the intestine where the symbiotic bacterium, Burkholderia, resides and that de-
grades fenitrothion resides. (Kikuchi et al., 2012 with permission from PNAS).
490 Daniel Doucet and Arthur Retnakaran
celery, lettuce, cabbage and alfalfa; and the tomato leafminer, Tuta absoluta,
on tomatoes and potatoes. Flucycloxuron resistance has been found in the
two spotted spider mite, T.urticae, on cotton, fruit trees, vegetables, walnut
and ornamentals. Lufenuron resistance is seen on the fruit fly, D.melano-
gaster, on fruits and the beet army worm, S.exigua. Flufenoxuron resistance
has been reported on the codling moth, C.pomonella. Chlorfluazuron resis-
tance is seen in the notorious diamond back moth, P.xylostella, which shows
resistance to a whole list of insecticides. Triflumuron has been found to be
resistant in C.pomonella,P.xylostella and T.absoluta. The three non-BPU
CSIs, etoxazole, buprofezin and cyromazine show resistance as well
(Fig. 6.17). Etoxazole has been shown to have resistance mutations by seg-
regant mapping (Van Leeuwen et al., 2012). The two spotted spider mite in
a limited region of Japan has been shown to have developed resistance to
Etoxazole (Masanobu et al., 2001). Buprofezin is resistant to the sweet potato
whitefly, B.tabaci, on cotton; brown leafhopper, Nilaparvata lugens,onriceand
the green house whitefly, Trialeurodes vaporariorum, on cucumber, tomato and
ornamentals. Cyromazine resistance has been reported in M.domestica,
L.cuprina,D.melanogaster and the serpentine leafminer, Iriomyza trifolii,on
chrysanthemum and celery (Bell et al., 2010). It must beemphasized that these
insecticides are not resistant to all populations globally but only to certain
populations in defined areas (Kotze and Sales, 2001; Whalon et al., 2008).
The mechanism of resistance follows the principles of natural selection.
When a pest is constantly bombarded by a chemical insecticide, it kills almost
all the insects except a few that have a mechanism to neutralize the chemical.
Repeated applications of the pesticide over several seasons and generations
kills almost all the susceptible insects but concomitantly there is an increase
in the population of the resistant variety. Over several years, the susceptible
forms are completely replaced by the resistant ones. It is a classic case of nat-
ural selection where the selection pressure applied by the pesticide leads to
the survival of the fittest, namely, the resistant insects. The resistance con-
ferred on the insect is due inter alia to the detoxification of the MOA or the
effect on the target site of the insecticide. An insect that is resistant to an
insecticide with a particular MOA can become resistant to another insecti-
cide with a similar MOA, a phenomenon called cross resistance. A resistant
species to one insecticide can become resistant to another insecticide with a
different MOA which is called multiple resistance. One should bear in
mind that all insects have a basic detoxifying system, and only when the in-
secticide overwhelms the system, the insect succumbs to the treatment.
Sometimes insects are not affected by a pesticide because of their size,
491Insect Chitin: Metabolism, Genomics and Pest Management
or the thickness of the exoskeleton, or innate metabolism and not due to
selection pressure in which case it is referred to as tolerance and not resis-
tance (Whalon et al., 2008).
Metabolic resistance mechanisms in insects are primarily based on detox-
ification enzymes such as esterases, glutathione-S-transferases (GSTs) and
monooxygenases. The role of esterases in pyrethroid and organophosphate re-
sistance has been well documented. Carboxylesterase inhibitor induces suscep-
tibility in malathion-resistant insects and has been used to study resistance.
Controlled atmosphere in warehouses induces esterase activity in stored prod-
uct insects promoting resistance to fumigants such as methyl bromide. GST is
responsible for organophosphate, organochlorine and cyclodiene resistance.
Microsomal P-450-dependent monooxygenases are of importance in
imparting resistance (Boyer et al., 2012). Synergists can often render a resistant
population susceptible or prevent the development of resistance. Synergists
such as piperonyl butoxide (PBO), sesamex and sulfoxide inhibit mixed func-
tion oxidases and increase the effectiveness of carbaryl, permethrin, malathion
and DDT. The synergists S,S,S-tributylphosphorotrithioate (TBBT),
O,O,O-triphenylphosphate (TPP) and 5-benzyl-O,O-diisopropyl pho-
sphorothioate (IBP) inhibit carboxylesterase and phosphotriesterase and
augment parathion, permethrin and dimethoate activity (see Raffa and
Priester, 1985). CH
3
I, t-phenylbutanone and diethylmaleate inhibit GST
and make organophosphates active (Raffa and Priester, 1985). ABC
transporters (ATP-binding cassette transporter) have been shown to be
involved in manifestation of resistance for insecticides such as thiodicarb in
Heliothis virescens and cypermethrin, fenvalerate and methylparathion in
H.armigera as well as Bt resistance (Gahan et al., 2010).
Diflubenzuron resistance in L.cuprina has been shown to be due to en-
hanced monooxygenase enzyme levels and significant synergism with PBO
suggesting a role for P-450-mediated metabolism. Autosomal inheritance of
this trait has been demonstrated, but the actual resistance appears to be poly-
genic involving mechanisms additional to monooxygenase (Kotze and Sales,
2001). Recently, it has been convincingly suggested that BPUs block chitin
synthesis by binding to the SUR, which is an ABC transporter, and inhibit
chitin synthase activity (Matsumura, 2010). It is quite conceivable that over
expression of this ABC transporter cum receptor might play a role in the
resistance mechanism of BPUs.
Resistance in the three non-BPU CSIs, buprofezin, cyromazine and
etoxazole, has been reported, and some work on resistance management
in the case of buprofezin has been initiated. Buprofezin resistance in the
492 Daniel Doucet and Arthur Retnakaran
brown plant hopper, N.lugens, was reported after 11 years of usage on rice.
This particular population had a resistance ratio of 3599. Use of synergists
such as O,O,-diethyl-O-phenyl phophorothioate (SV1), PBO and diethyl
maleate (DEM) reduced the resistance only by 0.5- to 1.6-fold indicating
that esterases, P-450 monooxygenases and GSTs are not involved in the re-
sistance mechanism (Wang et al., 2008). The exact mechanism of resistance
has not been resolved. Dennehy and his team at the University of Arizona
have developed a three regimen resistance management against the white fly,
B.tabaci, on cotton which has been widely adopted. They used three differ-
ent insecticides with different MOA in sequence, pyriproxifen (juvenile
hormone analogue), buprofezin (CSI) and synergized pyrethroid (Na
þ
channel modulator in the nervous system) and have successfully avoided re-
sistance from appearing (Dennehy and Williams, 1997). The brown leafhop-
per, N.lugens, on rice is controlled by buprofezin, imidacloprid (nicotinic
acetylcholine receptor (nAChR) agonist) and fipronil (GABA-gated chlo-
ride channel antagonist) (Denholm et al., 1999; Ling and Zhang, 2011).
Alternating insecticide treatments with compounds with unrelated
modes of action (Table 6.1)has been the method of choice for IRM. Incor-
porating biorational and biological pesticides would make it much more en-
vironmentally friendly. One has to balance economics with the
environment. Careful application from the onset of introduction will no
doubt prolong the life of an insecticide and the ideal way would be to com-
bine IPM with IRM.
Resistance management for the future entails the discovery of new target
sites and many have been suggested in recent years; (i) with the advent of
new immunofluorescent imaging technologies, the polymeric proteins that
provide a multifunctional scaffold within the plasma membrane in the cy-
toskeleton have been identified as a possible target sites. These include
spectrins, integrins and ankyrins. Any disruption will be lethal as has been
shown in Drosophila. (ii) The membrane-spanning protein receptor, the gua-
nosine protein-coupled receptor that depends on crosstalk with different sig-
nalling pathways can have drastic pleiotropic consequences if disrupted.
(iii) Odour-binding proteins are very selective and recognize specific pher-
omones and any blockage will have severe effects. (iv) Calcium channel is
gated with a Ca
2þ
–ATPase pump that acts as a critical intracellular messen-
ger, and if kept frozen at the open state, it can have dire consequences.
(v) Yet another potential target site is the nitric oxide signalling pathway
controlling mitochondrial apoptosis and DNA repair critical to the insect.
These are but a few of the major target sites that can be exploited to create
493Insect Chitin: Metabolism, Genomics and Pest Management
Table 6.1 Mode of action of pest control agents (based on various sources)
Mode of action Chemical group Example of insecticide
1. Acetylcholinesterase
(AChE) inhibitors
Carbamates,
organophosphates
Carbaryl, carbofuran,
Acephate, diazinon
2. GABA-gated chloride
channel antagonist
Cyclodienes,
phenylpyrazoles
Chlordane, endosulfan,
Fipronil
3. Na channel modulator Pyrethroids,
DDT group
Pyrethrin, cypermethrin,
DDT, methoxychlor
4. Nicotinic acetylcholine
receptor (nAChR)
agonists
Neonicotinoids Imidacloprid, acetamiprid
5. Nicotinic acetylcholine
receptor (nAChR)
allosteric activator
Spinosyns Spinosad
6. Chloride channel
activator
Avermectins Abamectin, emamectin
7. Juvenile hormone
mimic
Juvenile hormone
analogues
Methoprene, pyriproxifen
8. Midgut membrane
disruptor
Bacillus
thuringiensis
Cry toxins of Bt.
9. Inhibitor of
mitochondrial ATP
synthase
– Diafenthiuron
10. Uncoupler of oxidative
phosphorylation
– Chlorfenapyr
11. Nicotine acetylcholine
receptor (nAChR)
channel blocker
– Thiocyclam
12. Inhibitors of chitin
synthesis
Benzoylphenyl
ureas
Diflubenzuron, novaluron
13. Moult disruptor Non-
benzoylphenyl
urea
Buprofezin, cyromazine,
etoxazole
14. Ecdysone agonist Bisacyhydrazines Tebufenozide,
methoxyfenozide,
halofenozide,
chromafenozide, fufenozide
494 Daniel Doucet and Arthur Retnakaran
a new set of pesticides with new and different modes of action (MOA) crit-
ical to combat resistance (Hollomon, 2012; Mercer et al., 2007). For millions
of years, plants and insects have co-evolved, and now armed with the new
technologies, humans have entered the fray on the side of plants. Immediate
knock-down effects were the results operators were looking for in the early
days of pest management. With a better understanding of the physiology of
the insect and the MOA of an insecticidal compound, one can look for
targeted effects. New approaches such as the use of RNA interference with
dsRNA and hairpin RNA have already started making inroads into pest
management. Knockouts generated through genomic screening can identify
potential new target sites. Predicting and overcoming resistance mutations
through computational biochemistry incorporating our latest knowledge of
protein modelling will soon be a reality. The potential for this approach is
illustrated by how methicillin-resistant Staphylococcus aureus (MRSA) can be
overcome by using novel inhibitors of DHFR. A protein algorithm defined
the binding site for DHFR and much of the analyses were done in silico.In
the coming years, molecular diagnostics will be used in bioassays and
resistance monitoring will greatly aid in our quest to find new target sites,
new control methods (Hollomon, 2012).
8. CONCLUSIONS AND FUTURE DEVELOPMENT
Moulting is a critical phase in the life of an insect and the insect is ex-
tremely vulnerable during this state. The exoskeleton is replaced during this
process and inter alia the old chitin is digested and replaced with a newly
Table 6.1 Mode of action of pest control agents (based on various sources)—cont'd
Mode of action Chemical group Example of insecticide
15. Octopamine receptor
agonist
– Amitraz
16. Electron transport
inhibitor
– Hydramethylnon, rotenone
17. Voltage-dependent Na
channel blocker
– Indoxacarb
18. Inhibitors of acetyl
CoA carboxylase
– Spirodiclofen
19. Ryanodine receptor
modulator
– Flubendiamide
495Insect Chitin: Metabolism, Genomics and Pest Management
synthesized chitin. The carefully orchestrated sequence of events provides
many opportunities for pest control, chitin synthesis being the primary
one. While we understand the basic biological and biochemical events leading
to the dissolution and replacement of chitin, the molecular aspects of regula-
tion remain incomplete. With the advent of genomics and its exponential
growth of genomic data from many species of arthropods, our understanding
of tissue transcription, proteomes and interactomes has greatly advanced, and
this will probably result in answering the unresolved questions in the near fu-
ture. The genomics of the diversity in exoskeleton morphology such as inter-
specific differences (e.g. coleopterans vs. lepidopterans), ontogenic differences
(embryonic, larval, pupal and adult cuticle) and the basic body plan including
appendages (intersegmental, segmental, wing and elytra regions) afford nu-
merous opportunities for identifying new target sites that can be exploited
in one form or another for pest control. All these morphological differences
have evolved over 400 million years as adaptations for the insects to colonize
diverse ecological niches and may have vulnerable target sites.
Genomic analyses are being applied to explain many of the molecular
aspects of chitin formation and BPU MOA, which will eventually lead to
new protocols for pest management. The regulatory interrelationships be-
tween the various enzymes involved in chitin synthesis as well as the ultimate
step of polymerization can all be better understood using genomic resources.
Unravelling the mechanism of chitin polymerization by chitin synthase may
not only help us optimize the control protocols but also assist in understand-
ing other related inverting glycosyltransferases such as cellulose synthase and
hyaluron synthase. New technologies in sequencing, liquid chromatography
and tandem mass spectrometry will greatly aid in not only developing new
pesticides but also monitoring their non-target roles in the environment.
With the genomic resources growing at an astronomical rate along with
our understanding and concern for the environment, the quest for new con-
trol measures based on chitin physiology appears propitious.
ACKNOWLEDGEMENTS
We thank the authors who were kind enough to provide us with their manuscripts in print.
This work was supported by a grant from the Genomics Research and Development
Initiative (GRDI) of the Canadian Forest Service.
REFERENCES
Abo-Elghar, G.E., Fujiyoshi, P., Matsumura, F., 2004. Significance of the sulfonylurea re-
ceptor (SUR) as the target of diflubenzuron in chitin synthesis inhibition in Drosophila
melanogaster and Blattella germanica. Insect Biochem. Mol. Biol. 34, 743–752.
496 Daniel Doucet and Arthur Retnakaran
Acheuk, F., Cusson, M., Doumandji-Mitiche, B., 2012. Effects of a methanolic extract of the
plant Haplophyllum tuberculatum and of teflubenzuron on female reproduction in the
migratory locust, Locusta migratoria (Orthoptera: Oedipodinae). J. Insect Physiol. 58,
335–341.
Adams, J.A., 2004. Fungal cell wall chitinases and glucanases. Microbiology 150, 2029–2035.
Aittoniemi, J., Fotinou, C., Craig, T.J., Wet, H.D., Proks, P., Ashcroft, P.M., 2009. SUR1:
a unique ATP-binding cassette protein that functions as an ion channel regulator. Philos.
Trans. R. Soc. B 364, 257–267.
Al-Sawalmih, A., Li, C., Siegel, S., Fabritius, H., Yi, S., Raabe, D., Fratzl, P., Paris, O., 2008.
Microtexture and chitin/calcite orientation relationship in the mineralized exoskeleton
of the American lobster. Adv. Funct. Mater. 18, 3307–3314.
Ameen, A., Wang, C., Kaakeh, W., Bennett, G.W., King, J.E., Karr, L.L., Xie, J., 2005.
Residual activity and population effects of noviflumuron for German cockroach
(Dictyoptera: Blattellidae) control. J. Econ. Entomol. 98, 899–905.
Ampasala, D.R., Zheng, S., Zhang, D., Ladd, T., Doucet, D., Krell, P.J., Retnakaran, A.,
Feng, Q., 2011. An epidermis-specific chitin synthase CDNA in Choristoneura fumiferana:
cloning, characterization, developmental and hormonal-regulated expression. Arch.
Insect Biochem. Physiol. 76, 83–96.
Arakane, Y., Muthukrishnan, S., 2010. Insect chitinase and chitinase-like proteins. Cell.
Mol. Life Sci. 67, 201–216.
Arakane, Y., Muthukrishnan, S., Kramer, K.J., Specht, C.A., Tomoyasu, Y.,
Lorenzen, M.D., Kanost, M., Beeman, R.W., 2005. The Tribolium chitin synthase genes
TcCHS1 and TcCHS2 are specialized for synthesis of epidermal cuticle and midgut
peritrophic matrix. Insect Mol. Biol. 14, 453–463.
Arakane, Y., Dixit, R., Begum, K., Park, Y., Specht, C.A., Merzendorfer, H., Kramer, K.J.,
Muthukrishnan, S., Beeman, R.W., 2009. Analysis of functions of the chitin deacetylase
gene family in Tribolium castaneum. Insect Biochem. Mol. Biol. 39, 355–365.
Arakane, Y., Baguinon, M.C., Jasrapuria, S., Chaudhari, S., Doyungan, A., Kramer, K.J.,
Muthukrishnan, S., Beeman, R.W., 2011. Both UDP N-acetylglucosamine
pyrophosphorylases of Tribolium castaneum are critical for molting, survival and fecundity.
Insect Biochem. Mol. Biol. 41, 42–50.
Araki, Y., Ito, E., 1974. A pathway of chitosan formation in Mucor rouxii: enzymatic
deacetylation of chitin. Biochem. Biophys. Res. Commun. 56, 669–675.
Arau
´jo, S.J., Aslam, H., Tear, G., Casanova, J., 2005. Mummy/cystic encodes an enzyme
required for chitin and glycan synthesis, involved in trachea, embryonic cuticle and
CNS development—analysis of its role in Drosophila tracheal morphogenesis. Dev. Biol.
288, 179–193.
Arensburger, P., Megy, K., Waterhouse, R.M., Abrudan, J., Amedeo, P., Antelo, B.,
Bartholomay, L., Bidwell, S., Caler, E., Camara, F., Campbell, C.L., Campbell, K.S.,
Casola, C., Castro, M.T., Chandramouliswaran, I., Chapman, S.B., Christley, S.,
Costas, J., Eisenstadt, E., Feschotte, C., Fraser-Liggett, C., Guigo, R., Haas, B.,
Hammond, M., Hansson, B.S., Hemingway, J., Hill, S.R., Howarth, C., Ignell, R.,
Kennedy, R.C., Kodira, C.D., Lobo, N.F., Mao, C., Mayhew, G., Michel, K.,
Mori, A., Liu, N., Naveira, H., Nene, V., Nguyen, N., Pearson, M.D.,
Pritham, E.J., Puiu, D., Qi, Y., Ranson, H., Ribeiro, J.M., Roberston, H.M.,
Severson, D.W., Shumway, M., Stanke, M., Strausberg, R.L., Sun, C., Sutton, G.,
Tu, Z.J., Tubio, J.M., Unger, M.F., Vanlandingham, D.L., Vilella, A.J., White, O.,
White, J.R., Wondji, C.S., Wortman, J., Zdobnov, E.M., Birren, B.,
Christensen, B.M., Collins, F.H., Cornel, A., Dimopoulos, G., Hannick, L.I.,
Higgs, S., Lanzaro, G.C., Lawson, D., Lee, N.H., Muskavitch, M.A., Raikhel, A.S.,
Atkinson, P.W., 2010. Sequencing of Culex quinquefasciatus establishes a platform for
mosquito comparative genomics. Science 330, 86–88.
497Insect Chitin: Metabolism, Genomics and Pest Management
Beitel, G.J., Krasnow, M.A., 2000. Genetic control of epithelial tube size in the Drosophila
tracheal system. Development 127, 3271–3282.
Bel, Y., Wiesner, P., Kayser, H., 2000. Candidate target mechanisms of the growth inhibitor
cyromazine: studies of phenylalanine hydroxylase, puparial amino acids, and
dihydrofolate reductase in dipteran insects. Arch. Insect Biochem. Physiol. 45, 69–78.
Bell, H.A., Robinson, K.A., Weaver, R.J., 2010. First report of cyromazine resistance in a
population of UK house fly (Musca domestica) associated with intensive livestock produc-
tion. Pest Manag. Sci. 66, 693–695.
Bogo, M.R., Rota, C.A., Pinto Jr., H., Ocampos, M., Correa, C.T., Vainstein, M.H.,
Schrank, A., 1998. A chitinase encoding gene (chit1 gene) from the entomopathogen
Metarhizium anisopliae: isolation and of genomic and full-length cDNA. Curr. Microbiol.
37, 221–225.
Boquet, I., Hitier, R., Dumas, M., Chaminade, M., Pre
´at, T., 2000. Central brain
postembryonic development in Drosophila: implication of genes expressed at the inter-
hemispheric junction. J. Neurobiol. 42, 33–48.
Boyer, S., Zhang, H., Lempe
´rie
`re, G., 2012. A review of control methods and resistance
mechanisms in stored-product insects. Bull. Entomol. Res. 102, 213–229.
Braconnot, H., 1811. Recherches analytique sur la nature des champignons. Ann. Chim. 79,
265–304.
Brutti, M., Blasco, C., Pico
´, Y., 2010. Determination of benzoylurea insecticides in food by
pressurized liquid extraction and LC-MS. J. Sep. Sci. 33, 1–10.
Busby, J.N., Landsberg, M.J., Simpson, R.M., Jones, S.A., Hankamer, B., Hurst, M.R.,
Lott, J.S., 2012. Structural analysis of Chi1 Chitinase from Yen-Tc: the
multisubunit insecticidal ABC toxin complex of Yersinia entomophaga. J. Mol. Biol.
415, 359–371.
Campbell, P.J., Hammell, K.L., Dohoo, I.R., Ritchie, G., 2006. Randomized clinical trial to
investigate the effectiveness of teflubenzuron for treating sea lice on Atlantic salmon. Dis.
Aquat. Organ. 70, 101–108.
Candy, D.J., Kilby, B.A., 1962. Studies on chitin synthesis in the desert locust. J. Exp. Biol.
39, 129–140.
Carlstrom, D., 1957. The crystal structure of alpha-chitin (poly-N-acetyl-D-glucosamine).
J. Biophys. Biochem. Cytol. 3, 669–683.
Carson, R., 1962. The Silent Spring. Houghton Mifflin Co., New York, 378 pp.
Celniker, S.E., Dillon, L.A., Gerstein, M.B., Gunsalus, K.C., Henikoff, S., Karpen, G.H.,
Kellis, M., Lai, E.C., Lieb, J.D., MacAlpine, D.M., Micklem, G., Piano, F., Snyder, M.,
Stein, L., White, K.P., Waterston, R.H., modENCODE Consortium, 2009. Unlocking
the secrets of the genome. Nature 459, 927–930.
Chen, J., Tang, B., Chen, H., Yao, Q., Huang, X., Chen, J., Zhang, D., Zhang, W., 2010.
Different functions of the insect soluble and membrane-bound trehalase genes in chitin
biosynthesis revealed by RNA interference. PLoS One 5, 10133–10146.
Chiew, Y.Y., Shepherd, M.G., Sullivan, P.A., 1980. Regulation of chitin synthesis during
germ-tube formation in Candida albicans. Arch. Microbiol. 125, 97–104.
Chintapalli, V.R., Wang, J., Dow, J.A.T., 2007. Using FlyAtlas to identify better Drosophila
melanogaster models of human disease. Nat. Genet. 39, 715–720.
Cohen, E., 1987. Chitin biochemistry: synthesis and inhibition. Annu. Rev. Entomol. 32,
71–93.
Cohen, E., 1993. Chitin synthesis and degradation as targets for pesticide action. Arch. Insect
Biochem. Physiol. 22, 245–261.
Cohen, E., 2010. Chitin biochemistry: synthesis, hydrolysis and inhibition. Adv. Insect Phys.
38, 5–74.
Cohen, E., Casida, J.E., 1980. Properties of Tribolium gut chitin synthetase. Pestic. Biochem.
Physiol. 13, 121–128.
498 Daniel Doucet and Arthur Retnakaran