7 Chitin Metabolism in Insects
Subbaratnam Muthukrishnan
Kansas State University, Manhattan, KS, USA
Hans Merzendorfer
University of Osnabrueck, Osnabrueck, Germany
Yasuyuki Arakane
Chonnam National University, Gwangju, South Korea
Karl J Kramer
Kansas State University, and USDA-ARS,
Manhattan, KS, USA
© 2012 Elsevier B.V. All Rights Reserved
7.1. Introduction
7.2. Chitin Structure and Occurrence
7.3. Chitin Synthesis
7.3.1. Sites of Chitin Biosynthesis
7.3.2. Chitin Biosynthetic Pathway
7.3.3. Chitin Synthases: Organization of Genes and Biochemical Properties
7.3.4. Chitin Synthases: Regulation and Function
7.4. Chitin Degradation and Modification
7.4.1. Insect Chitinases
7.4.2. Insect N-Acetylglucosaminidases
7.4.3. Insect Chitin Deacetylases
7.5. Chitin-Binding Proteins
7.5.1. Chitin-Binding Proteins with the R&R Consensus
7.5.2. Peritrophic Matrix Proteins
7.5.3. Cuticular Proteins Analogous to Peritrophins (CPAPs)
7.5.4. Enzymes of Chitin Metabolism
7.5.5. Role of Secondary Structure of ChtBD2 Motif in Binding to Chitin
7.6. Chitin-Organizing Proteins
7.7. Hormonal Regulation of Chitin Metabolism
7.8. Chitin Metabolism and Insect Control
7.8.1. Inhibition of Chitin Synthesis
7.8.2. Exploiting Chitinases for Insect Control
7.9. Future Studies and Concluding Remarks
7.1. Introduction
“Chitin Metabolism in Insects” was the title of chapters
in both the original edition of the Comprehensive Insect
Physiology, Biochemistry and Pharmacology series published
in 1985 and the follow-up Comprehensive Molecular Insect
Science series in 2005 (Kramer et al., 1985; Kramer and
Muthukrishnan, 2005). Since 2005 substantial progress
in gaining additional understanding of this topic has continued to take place, primarily through the application of
the techniques of molecular genetics, functional genomics, proteomics, transcriptomics, metabolomics, and biotechnology to an assortment of studies focused on insect
chitin metabolism. Several other reviews have also been
published that have reported on some of the advances that
have taken place (Dahiya et al., 2006; Merzendorfer, 2006,
2009; Arakane and Muthukrishnan, 2010). Most interestingly, the list of genes and gene products found to be
involved in insect chitin metabolism has been lengthened
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significantly. In this chapter we will highlight some of the
more recent and important findings, with emphasis on
results obtained from studies conducted on the synthesis,
structure, physical state, modification, organization, and
degradation of chitin in insect tissues, as well as the interplay of chitin with chitin-binding proteins, the regulation
of genes responsible for chitin metabolism, and, finally, the
targeting of chitin metabolism for insect-control purposes.
7.2. Chitin Structure and Occurrence
Chitin is the major polysaccharide present in insects and
many other invertebrates as well as in several microbes,
including fungi. Structurally, it is the simplest of the glycosaminoglycans, being a β(1→4) linked linear homopolymer
of N-acetylglucosamine (GlcNAc, [C8H13O5N]n, where
n >> 1). It serves as the skeletal polysaccharide of several
animal phyla, such as the Arthropoda, Annelida, Molluska,
194 7: Chitin Metabolism in Insects
and Coelenterata. In several groups of fungi, chitin replaces
cellulose as the structural polysaccharide. In insects, it is
found in the body wall or cuticle, gut lining or peritrophic
matrix (PM), salivary gland, trachea, eggshells, and muscle
attachment points. In the course of evolution, insects have
made excellent use of the rigidity and chemical stability of
the polymeric chitin to assemble both hard and soft extracellular structures such as the cuticle (exoskeleton) and PM
respectively, both of which enable insects to be protected
from the environment while allowing for growth, mobility, respiration, and communication. All of these structures
are primarily composites of chitin fibers and proteins with
varying degrees of hydration and trace materials distributed along the structures. The insolubility and structural
complexity of the cuticle has limited its study. However,
sclerotized cuticle can be modeled as an interpenetrating network of chitin fibers with embedded cross-linked
protein and pigments. Both synthesis and degradation of
chitin take place at multiple developmental stages in the
cuticle and the PM. It is usually synthesized as portions
of the old endocuticle and PM and trachaea are resorbed,
and the digested materials are recycled. Although primarily composed of poly-GlcNAc, chitin also can contain a
small percentage of unsubstituted (or N-deacetylated) glucosamine (GlcNAc) residues, making it a GlcNAc-GlcN
heteropolymer (Muzzarelli, 1973; Fukamizo et al., 1986).
When the epidermal and gut cells synthesize and secrete a
particular form of chitin consisting of antiparallel chains or
alpha-chitin, the chains are assembled into microfibrils and
then into sheets. As layers of chitin are added, the sheets
are cross-oriented relative to one another at a constant
angle to form a helicoidal bundle (known as the Bouligand
structure), which can contribute to the formation of an
extremely strong, plywood-like material.
Although there is no doubt that there are strong noncovalent interactions between chitin and chitin-binding
proteins, there is only weak indirect evidence that there
are covalent interactions between them. The evidence so
far for direct involvement of chitin in cross-links to proteins has been inconclusive. Results of solid state NMR
and chemical analyses have indicated the presence of trace
levels of aromatic amino acids in chitin preparations,
suggesting that those amino acids were there because
they were involved in protein cross-links with chitin
(Schaefer et al., 1987). Additional spectroscopic evidence
for glucosamine–catecholamine adducts derived from chitin–protein cross-links in cuticle was obtained using electrospray mass spectrometry and tandem mass spectrometry
(Kerwin et al., 1999). However, those observations have
not been investigated further. More direct evidence for
chitin–protein cross-links from studies of intact cuticle
instead of degraded or digested samples is needed before
the precise nature of the covalent interactions of cuticular proteins with chitin fibers can be resolved (Demolliens
et al., 2008).
Alpha-chitin fibers, because of their hydrophilic nature,
are generally highly hydrated. Chitin dehydration via
impregnation of hydrophobic proteins probably contributes
to tissue stiffening and deplasticization (Vincent, 2009). In
addition, the formation of a cross-linked and interpenetrating protein network in the dehydrated composite leads to
additional hardening (Andersen, 2010); thus, chemical
bonds surely play a crucial role in cuticle mechanics by
increasing the load carried by the proteins and by providing a hydrophobic “coating” around the chitin nanofibers,
thus preventing softening of the latter by water adsorption.
Chitin nanofibrils probably form the initial template, similar to glass or carbon fiber mats in composite processing.
Filler proteins and catechols are then secreted through the
chitinous procuticle. Once oxidation of catechols to quinones and quinone methides has occurred, cross-linking
and hardening of the extracellular matrix ensues. As sclerotization proceeds, water is progressively expelled. The precise role of water removal on the structural properties of the
cuticle is not fully understood, in part because the effect of
water on individual components of the composite is poorly
understood, but some progress is starting to take place.
Also, the individual contributions of chitin and protein to
the mechanical properties are unknown. In the hydrated
state, there is considerable variation in moduli reported for
chitosan/chitin scaffolds (Wu et al., 2006). There is a difference of several orders of magnitude in the stiffness of
chitin/chitosan between the fully hydrated state, where it
is present as a porous, water-saturated scaffold, and the dry
state. To mimic the action of catechols to stiffen chitosan
scaffolds, Wu et al. (2005) achieved a two-fold increase in
stiffness after treatment of chitosan films with oxidized catechols. Although there was a significant increase in stiffness, it was less than the increase observed from insect
cuticle tanning. Recently, dynamic mechanical analysis of
insect cuticle during maturation revealed that while the
water content has an important role in determining cuticle
mechanical properties, the tanning reactions themselves
contribute substantially to these properties beyond simply inducing dehydration (Lomakin et al., 2011). Cuticle,
whether tanned or untanned, increases in hardness while
drying, but the increase is generally less than that observed
from tanning alone.
7.3. Chitin Synthesis
Although extensive knowledge on the precise molecular
mechanism of chitin synthesis is lacking, substantial progress has been made regarding the function and regulation
of several genes involved in the chitin biosynthetic pathway.
In the past 10 years, many genes coding for key enzymes of
this pathway have been isolated and sequenced from various
insect species. Analyses of their expression in different tissues during development have provided the first clues about
their function. The availability of Drosophila melanogaster
7: Chitin Metabolism in Insects 195
(fruit fly) mutants defective in some of these genes, together
with the ability to specifically silence their expression by
RNAi in the fly and other species, has boosted our understanding of this process. Most progress has been made on
chitin synthases (CHSs), which have been identified in a
variety of organisms, including fungi, nematodes, mollusks,
and insects. Amino acid sequence similarities have been the
principal tools used for identifying CHSs, which form a
subfamily within a larger group of the glycosyltransferases
(family GT2) that catalyze the transfer of a sugar moiety
from an activated sugar donor onto saccharide or nonsaccharide acceptors (Coutinho et al., 2003; Cantarel et al.,
2009). CHS has not been an easy enzyme to assay, which
has made its study rather difficult. Traditionally, CHS
activity is measured by a radioactive assay using [14C]- or
[3H]-labeled UDP-GlcNAc as the precursor followed by
quantification of insoluble radiolabeled chitin after acid
precipitation. Alternately, a high-throughput non-radioactive assay is available, which involves binding of synthesized
chitin to a wheat germ agglutinin (WGA)-coated surface,
followed by detection of the polymer with a horseradish
peroxidase–WGA conjugate (Lucero et al., 2002). Also,
the direct incorporation of fluorescently labeled substrates,
such as certain dansyl-UDP-GlcNAc analogs, may prove to
be useful for developing fluorescence-based enzyme assays
(Yeager and Finney, 2005). The paucity of information
concerning the enzyme’s biochemical and kinetic properties
was mainly due to the inability to obtain active soluble CHS
preparations. Recently, however, a purification and solubilization protocol has been developed, which allowed purifying CHS-B from the midgut of Manduca sexta (tobacco
hornworm) as an active, oligomeric complex (Maue et al.,
2009). In addition, first attempts to heterologously express
CHSs from protists and fungi in yeast systems turned out
to be successful (Van Dellen et al., 2006; Martinez-Rucobo
et al., 2009; Barreto et al., 2010). These purification and
expression protocols should facilitate greater progress in
insect CHS studies in the future.
7.3.1. Sites of Chitin Biosynthesis
The epidermis and the midgut are two major tissues where
chitin synthesis occurs in insects. Epidermal cells are
responsible for the deposition of new cuticle during each
molt, and the midgut cells are generally associated with the
formation of the PM during feeding. Both the cuticle and
the PM contain chitin microfibrils, which function as a
matrix that binds numerous cuticle and PM proteins. However, chitin is associated with other tissues as well, including the head skeleton, foregut, hindgut, trachea, wing
hinges, salivary glands, and mouthparts of adults and/or
larvae. In early development, chitin is additionally found
in the cuticle of the developing larva within the embryo,
as well as in the extra-embryonic serosal cuticle and the
eggshells (Wilson and Cryan, 1997; Moreira et al., 2007;
Rezende et al. 2008). In general, it is assumed that the cells
closest to the site where chitin is found are responsible for
its biosynthesis. However, this interpretation is somewhat
complicated by the fact that assembly of chitin microfibrils
occurs in the extracellular space and is influenced by proteins that organize their deposition (see section 7.6).
7.3.1.1. Chitin synthesis in the epidermis and tracheal
system Chitin is a major constituent of the cuticle, the
outermost layer of insects, which serves as an exoskeleton
and protects against various harming agents. Within the
cuticle, chitin is mainly found in the procuticle, with
higher amounts in the endocuticle than in the exocuticle,
but is absent from the epicuticle (Sass et al., 1994). Chitin
deposition in the cuticle was recently reinvestigated in
Drosophila embryos in an ultrastructural study using
electron microscopy and gold-conjugated wheat germ
agglutinin (gold-WGA), which binds to GlcNAc residues
in chitin and glycoproteins (Schwarz and Moussian, 2007).
In agreement with previous findings, gold particles could
only be detected in the procuticle but not in the epicuticle.
The gross architecture of the procuticle is established mainly
by consecutive layers of chitin bundles of microfibrils
embedded in a matrix of cuticle proteins. The orientation
of a single lamina of chitin microfibrils can be twisted in
relation to the neighboring layers above and below it by
different angles in different insect species, giving rise to
helicoidal or pseudo-orthogonal textures. Much of what we
know on cuticle differentiation derives from ultrastructural
studies of cuticle renewal during insect molting (Locke,
2001; Moussian, 2010). The classical concept of cuticle
formation is based on three sequential phases. First,
the envelope is laid down at the plasma membrane
surface, usually above electron-dense plaques at the tips
of microvilli, which were postulated to carry the chitinsynthesizing machinery (Locke, 1991). Then, the epicuticle
is assembled beneath the envelope. Finally, the procuticle,
which is considerably thicker than the other two layers,
is assembled and oriented at the cell surface. However, a
recent ultrastructural study of cuticle differentiation in
Drosophila embryos revealed a slightly different picture,
as envelope, epicuticle, and procuticle are partially formed
in parallel in the first phase, then the cuticle thickens in
the second phase, and in a third phase the chitin laminae
acquire their final orientation (reviewed in Moussian et al.,
2006). Interestingly, the apical membrane of the embryonic
epidermis does not form microvilli-like protrusions.
Instead, it exhibits longitudinal microtubule-stabilized
furrows, which were called apical undulae and are oriented
perpendicular to the first layers of chitin microfibrils
(Schwarz and Moussian, 2007). These apical undulae may
have a crucial role in determining the orientation of chitin
microfibrils, at least in the embryonic cuticle. Factors that
affect the shape of the apical membrane, such as syntaxin
1A, indirectly affect chitin orientation, presumably by
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interfering with the transport of proteins involved in cuticle
or chitin assembly (Moussian et al., 2007).
During embryogenesis, chitin synthesis also plays a role
for tracheal morphogenesis. Chitin is also found in the tracheal cuticle, which has been thought to have a composition similar to that of the epidermal cuticle. This point
needs clarification by direct chemical analysis of tracheal
cuticle. However, it came as a surprise when two research
groups reported independently that chitin forms a transient lumenal matrix during tracheal development in
Drosophila embryos (Devine et al., 2005; Tonning et al.,
2005). The lumenal chitin appears to be necessary to control tube size, diameter, and shape by orchestrating the
function of surrounding tracheal cells. Drosophila genetics, in combination with different microscopic techniques,
have proven most valuable in dissecting cuticle differentiation, and yielded a number of factors that are involved in
controlling this process. Some of these factors will be discussed in more detail later in this chapter (see section 7.6.).
In addition to the histochemical detection of chitin with colored or fluorescent compounds that bind to
chitin with different specificities, the expression of CHS
genes has been used to identify chitin-synthesizing tissues.
CHS gene expression was analyzed in various insects by
RT-PCR, Northern blots, and in situ hybridization. These
studies clearly demonstrated that epidermal and tracheal
cells express CHS genes, and hence confirmed that these
epithelia are sites of chitin biosynthesis. The first cDNA
encoding an insect chitin synthase was identified by Tellam
and colleagues (2000) in Lucilia cuprina (sheep blow fly),
and termed LcCHS1. RT-PCR using total RNA preparations from the carcass and trachea indicated expression of
LcCHS1 in these tissues. In situ hybridization revealed a
strong signal for the LcCHS1 mRNA in a single layer of
epidermal cells immediately underneath the procuticle.
Similar results were obtained for the expression of homologous CHS (also referred to as CHS-A) genes from other
insect sources, including D. melanogaster, M. sexta, Spodoptera frugiperda (fall armyworm), and T. castaneum (red flour
beetle) (Ibrahim et al., 2000; Gagou et al., 2002; Zhu et al.,
2002; Arakane et al., 2004; Bolognesi et al., 2005; Hogenkamp et al., 2005; Zimoch et al., 2005). In agreement with
the detection of chitin in eggs, CHS gene expression was
reported during embryogenesis by RT-PCR using RNA
from Lucilia sericata and Aedes aegypti eggs (Moreira et al.,
2007; Tarone et al., 2007; Rezende et al., 2008).
7.3.1.2. Chitin synthesis in the midgut Chitin is
a component of the insect PM, and accounts for about
3–13% (w/w) of its dry weight. There are two patterns of
PM production in insects. Type I PMs are synthesized and
delaminated throughout the entire midgut epithelium. Type
II PMs are formed as a continuous lining of the gut, which is
produced by a specialized region of the anterior midgut called
the cardia (Lehane, 1997). The most detailed picture of chitin
synthesis and its association with PM proteins has emerged
from observations using transmission, scanning electron,
light, and fluorescence microscopy (TEM, SEM, LM, and
FM, respectively) in three lepidopteran species; namely,
Ostrinia nubilalis (European corn borer), Trichoplusia ni
(cabbage looper), and M. sexta (Harper and Hopkins, 1997;
Harper et al., 1998; Harper and Granados, 1999; Wang
and Granados, 2000; Hopkins and Harper, 2001; Zimoch
and Merzendorfer, 2002). TEM in combination with goldWGA staining demonstrated that the PM of O. nubilalis
contains a fibrous, chitin-containing matrix that appears
first at the tips of the microvilli of the midgut epithelial cells
just past the stomadeal valves, and is rapidly assimilated
into a thin PM surrounding the food bolus (Harper and
Hopkins, 1997). The PM becomes thicker and multilayered
in the middle and posterior regions of the midgut. The
orthogonal lattice of chitin meshwork is slightly larger than
the diameter of the microvilli. SEM and LM studies revealed
that the PM delaminates from the tips of the microvilli.
This observation suggests that microvilli serve as sites (and
possibly as templates) for the organization of the PM by
laying down a matrix of chitin microfibrils, which associate
with PM proteins. A similar pattern of delamination of
the PM containing both chitin and intestinal mucins was
demonstrated in larvae of T. ni (Harper and Granados,
1999; Wang and Granados, 2000). Incorporating WGA
into the diet can interrupt formation of the PM. WGA-fed
O. nubilalis larvae exhibited an unorganized PM, which was
multilayered and thicker than the normal PM (Hopkins
and Harper, 2001). WGA was actually associated with the
PM as well as with the microvillar surface, as revealed by
immunostaining with antibodies specific for WGA. Because
there was very little WGA within the epithelial cells, the
interaction of WGA appears to be extracellular. Presumably,
WGA interferes with the formation of the organized chitin
network and/or the association of PM proteins with the
chitin network, leading to a reduced protein association
with the PM (Harper et al., 1998). There was also extensive
disintegration of the microvilli, and the appearance of dark
inclusion bodies, as well as apparent microvillar fragments
within the thickened multilayered PM. Species such as M.
sexta, which secrete multiple and thickened PMs that are
somewhat randomly organized, tolerated WGA better, and
sequestered larger amounts of WGA within the multilayered
PM (Hopkins and Harper, 2001).
As in the case of epidermal chitin synthesis, RT-PCR,
Northern blots and in situ hybridization demonstrated
the expression of a gene encoding a midgut specific CHS
form. This gene was originally identified in D. melanogaster, but its expression and function were characterized in
Aedes aegypti, M. sexta, and T. castaneum (Ibrahim et al.,
2000; Zimoch and Merzendorfer, 2002; Arakane et al.,
2004), as well as more recently in S. exigua, S. frugiperda,
and O. nubilalis (Bolognesi et al., 2005; Kumar et al.,
2008; Khajuria et al., 2010).
7: Chitin Metabolism in Insects 197
The first evidence that midgut cells express a CHS gene
was provided by Ibrahim et al. (2000) for female Ae. aegypti
mosquitoes dissected several hours after a blood meal.
In situ hybridizations with an antisense RNA probe for
AeCHS2 (CHS-B) in blood-fed mosquitoes localized the
mRNA at the apical site of midgut epithelial cells. Likewise, in situ hybridization with an antisense RNA probe to
MsCHS2 (CHS-B) from M. sexta revealed that high levels
of transcripts for this gene are present in apical regions of
the columnar cells of the anterior midgut but completely
absent in the epidermis or tracheal system of M. sexta larvae
(Zimoch and Merzendorfer, 2002). An antibody to the catalytic domain of the M. sexta, CHS was used to detect the
enzyme in midgut brush border membranes at the extreme
apical ends of microvilli, a result suggestive of some special compartment or possibly apical membrane-associated
vesicles. In line with its assumed role in PM formation during feeding stages, MsCHS2 mRNA was detected in the
midgut of feeding but not of starving or molting larvae
(Zimoch et al., 2005). Similar expression patterns were
reported for S. exigua, S. frugiperda, and O. nubilalis by RTPCR (Bolognesi et al., 2005; Kumar et al., 2008; Khajuria
et al., 2010). In S. frugiperda, chitin could be stained in the
PM only when SfCHS2 (CHS-B) expression was detectable
(Bolognesi et al., 2005). From the finding that TcCHS2
(encoding CHS-B) expression was observed in T. castaneum only in late larvae and adults, but not in pupal stages,
where chitin is synthesized during cuticle formation, it was
concluded that TcCHS-B functions in the course of PM
formation in the midgut (Arakane et al., 2004), a hypothesis further substantiated by RNAi experiments (Arakane
et al., 2005; see also section 7.3.4.3).
7.3.2. Chitin Biosynthetic Pathway
It has been assumed that most parts of the chitin biosynthetic pathway of insects would be similar or identical to
the Leloir pathway, which has been worked out extensively in fungi and other microbes (Figure 1). This appears
to be the case except for some minor details (Palli and
­Retnakaran, 1999). The source of the sugar residues for
chitin synthesis can be traced to fat body glycogen, which
is acted upon by glycogen phosphorylase. Glucose-1-P
produced by this reaction is converted to trehalose, which
is released into the hemolymph. Trehalose, the extracellular source of sugar in many insect species, is acted upon
by a trehalase, which is widely distributed in insect tissues, including the epidermis and gut, to yield intracellular glucose (Becker et al., 1996). This view was recently
substantiated by Chen et al. (2010), who showed that the
RNAi-induced knockdown of the expression of two trehalase-encoding genes, SeTre1 and SeTre2, caused downregulation of the CHS-encoding genes SeCHS1 and SeCHS2,
respectively, and led to reduced chitin levels in the cuticle
and the PM. The conversion of glucose to fructose-6-P
needed for chitin synthesis involves two glycolytic enzymes
present in the cytosol. These enzymes are hexokinase and
glucose-6-P isomerase, which convert glucose to fructose6-P. From the latter, the chitin biosynthetic pathway
branches off, with the first enzyme catalyzing this branch
being glutamine fructose-6-phosphate amidotransferase
(GFAT, E. C. 2.6.1.16), which might be thought of as
the first committed step in amino sugar biosynthesis. The
conversion of fructose-6-P to GlcNAc phosphate involves
amination, acetyl transfer, and an isomerization step,
which moves the phosphate from C-6 to C-1 (catalyzed
by a phospho-N-acetylglucosamine mutase). The conversion of this compound to the nucleotide sugar derivative
follows the standard pathway and leads to the formation
of an UDP-derivative of GlcNAc, which serves as the substrate for CHS. The entire chitin biosynthetic pathway is
outlined in Figure 1. The involvement of dolichol-linked
GlcNAc as a precursor for chitin was proposed quite some
time ago (Horst, 1983), but that hypothesis has received
very limited experimental support (Quesada-Allue, 1982).
At this point, this possibility remains unproven. Similarly,
the requirement for a primer to which the GlcNAc residues can be transferred also remains speculative. Based on
the model for glycogen biosynthesis, which requires glycogenin as the primer (Gibbons et al., 2002), CHS or an associated protein may fulfill this priming function. Because
each sugar residue in chitin is rotated 180° relative to the
preceding sugar, which requires CHS to accommodate a
alternating “up/down” configuration, another precursor,
UDP-chitobiose, has been proposed to be a disaccharide
donor during biosynthesis (Chang et al., 2003). Evaluation of radiolabeled UDP-chitobiose as a CHS substrate in
yeast, however, revealed that it was not incorporated into
chitin. Nevertheless, by testing monomeric and dimeric
uridine-derived nucleoside inhibitors as mechanistic
probes Yeager and Finney (2004) found a 10-fold greater
inhibition for the dimeric inhibitor than the corresponding monomeric inhibitor. However, both inhibitors bound
with low affinities in the millimolar range. The stereochemical problem in chitin synthesis of adding GlcNAc to
the growing chain in two opposite orientations resembles
the situation with hyaluronan synthases (HAS), which produce the hyaluronan polymer from two different monosaccharides, UDP-GlcNAc and UDP-glucuronic acid. HASs
are “dual action” glycosyltransferases that accomplish hyaluronan biosynthesis by two substrate-binding and active
sites (Weigel and DeAngelis, 2007). As class I HASs are
related to chitin synthase, two binding sites for alternating
GlcNAc orientations may also occur in CHSs.
7.3.2.1. Key enzymes The biosynthetic pathway
of chitin can be thought of as consisting of three
subreactions. The first set leads to the formation of the
amino sugar, GlcNAc, the second to its activated form
UDP-GlcNAc, and the last yields the polymeric chitin
198 7: Chitin Metabolism in Insects
Figure 1 Biosynthetic pathway for chitin in insects starting from glycogen, trehalose, and recycled chitin.
from the amino sugar. The rate-limiting enzyme in the first
subreaction appears to be glutamine-fructose-6-phosphate
aminotransferase (GFAT, EC 2.6.1.16), which is found in
the cytosol. The critical enzyme in the second subreaction
is UDP-N-acetylglucosamine pyrophosphorylase (UAP,
EC 2.7.7.23), which is also found in the cytosol, and that
in the last subreaction is CHS (EC 2.4.1.16), which is
localized in its active form at the plasma membrane. Not
surprisingly, these three enzymes appear to be major sites
of regulation of chitin synthesis.
7.3.2.2. Function and regulation of GFATs In Droso­
phila, two genes encoding GFAT (Gfat1 and Gfat2)
have been identified (Adams et al., 2000; Graack et al.,
2001). Both of these genes are on chromosome 3, but
they are present at different locations. Their intron–
exon organizations are different, as are the amino acid
sequences of the encoded proteins. GFAT consists of two
separate domains: an N-terminal domain that has both
glutamine-binding and aminotransferase motifs identified
in GFATs from other sources; and a C-terminal domain
with both fructose-6-phosphate binding and isomerase
motifs. Gfat1 is expressed in embryos in the developing
tracheal system, cuticle-forming tissues, and corpora cells
of larval salivary glands (Graack et al., 2001). The major
regulation of GFAT1 appears to be post-translational.
When Gfat1 was expressed in yeast cells, the resulting
enzyme was feedback-inhibited by UDP-GlcNAc, and
7: Chitin Metabolism in Insects 199
was stimulated by protein kinase A (PKA). Even though it
has not been demonstrated that there is a phosphorylated
form of GFAT1 which is susceptible to feedback
inhibition by UDP-GlcNAc, this possibility remains
viable. However, the situation may be complicated by
overlapping kinase activities, as recently a novel, highly
conserved phosphorylation site was identified, which
accounts for in vivo phosphorylation of human GFAT1
overexpressed in insect cells by protein kinases other than
PKA (Li et al., 2007). Examination of a mutant that
mimics phosphorylation at this site demonstrated that
the modification stimulates glucosamine-6-phosphatesynthesizing activity, but has no effect on UDP-GlcNAc
inhibition.
Another insect species in which Gfat1 function and regulation has been analyzed in more detail is Aedes aegypti.
The mosquito gene has no introns, and the promoter
appears to contain sequence elements related to ecdysteroid response elements (EcRE) as well as E74 and Broad
complex Z4 elements. E74 and Broad complex Z4 proteins are transcription factors known to be upregulated by
ecdysone (Thummel, 1996). Two Gfat1 transcripts with
different sizes were observed in Northern blot analyses of
RNA from adult females, and their levels increased further
after blood feeding (Kato et al., 2002). Since ecdysteroid
titers increase following blood feeding, it is possible that
this gene is under the control of ecdysteroids, either directly
or indirectly. Silencing of gene expression by dsRNA injection additionally revealed that GFAT1 is necessary for chitin synthesis in the course of PM formation in the midgut,
which occurs in female mosquitoes in response to a blood
meal (Kato et al., 2006). Feedback inhibition of chitin
synthesis by UDP-GlcNAc has also been reported in this
study, indicating that the mosquito enzyme is likely to be
regulated in a manner similar to the Drosophila enzyme.
7.3.2.3. Function and regulation of UAPs Insect
genomes usually possess only one gene encoding UDPGlcNAc pyrophosphorylase (UAP). The known exception
is T. castaneum, which has two UAP genes. The first
phenotypes for defects in the UAP gene were described in
Drosophila, where this gene was alternately termed mummy,
cabrio, or cystic, according to three phenotypes that were
identified in independent genetic screens for genes involved
in tracheal, epidermal, and CNS development (NüssleinVollhard et al., 1984; Hummel et al., 1999; Beitel and
Krasnow, 2000). While cystic was originally recognized to be
important for tracheal morphogenesis and tube size control,
mummy and cabrio mutants were reported to exhibit severe
defects in cuticle formation and CNS development of the
embryo. Eventually, all of these genes were shown to be
allelic by Araújo et al. (2005) and Schimmepfeng et al.
(2006), and the gene encoding UAP is now consistently
named mummy (mmy). Interestingly, the mmy mutant
phenotype is similar to that of the so-called “halloween”
mutants, which fail to produce the morphogenetic
hormone 20-hydroxyecdysone (Gilbert, 2004). UAP
functions in apical extracellular matrix formation by
producing UDP-GlcNAc needed for chitin synthesis and
for protein glycosylation. Consequently, deletion or defects
in mmy can lead to the complete absence of chitin in the
cuticle and tracheal lumen, as evidenced by a lack of WGA
staining in mutant embryos or larvae carrying a single
nucleotide substitution leading to the exchange of glycine
to valine at position 261 (Tonning et al., 2006). Moreover,
the epithelial organization is affected in mmy mutants, as
adherens junctions between epidermal cells appear wider
than in wild type embryos, and the characteristic ladder-like
structure of the septate junctions is missing. Additionally,
a membrane-integral septate junction component (Fas3) is
delocalized in the mutant, indicating that mmy may have
an additional function in proper localization of membranebound septate junction components (Tonning et al.,
2006). Expression of mmy is hormonally regulated in apical
extracellular matrix-differentiating tissues, and selectively
upregulated when chitinous material is deposited during
development. It is possible that the enzyme is also regulated
at the post-translational level by uridine, as this nucleic acid
base was shown to be an effective inhibitor for the yeast
enzyme (Yamamoto et al., 1980). In Ae. aegypti the gene
encoding UAP is constitutively expressed throughout all
life stages, and blood feeding does not significantly alter
mRNA levels (Kato et al., 2005). The cDNA was cloned
and the enzyme expressed as a recombinant enzyme,
allowing determination of substrate specificity. The enzyme
uses GlcNAc-1-P as a substrate, but it also exhibited low
activity when incubated with Glc-1-P. In T. castaneum two
UAP isoforms were identified, which share 60% identical
amino acids but differ significantly in their developmental
and tissue-specific expression patterns, as well as in function,
as revealed by RNAi studies (Arakane et al., 2010). While
the knockdown of TcUAP1 transcripts caused arrested
development at the larval–larval, larval–pupal, and pupal–
adult molts, knockdown of TcUAP2 retarded larval growth
or resulted in pupal paralysis. Results of chitin-staining
experiments in cuticle and PM indicated that chitin
deposition is prevented only when TcUAP1, but not when
TcUAP2, expression was blocked. However, both genes
are essential for beetle development and survival. TcUAP1
obviously is required for chitin synthesis in the course of
cuticle and PM formation, whereas TcUAP2 appears to
have other critical roles, presumably in glycosylation of
proteins.
7.3.3. Chitin Synthases: Organization
of Genes and Biochemical Properties
7.3.3.1. Number and organization of CHS-encoding
genes CHS genes from numerous unicellular and
filamentous species of fungi have been isolated and
200 7: Chitin Metabolism in Insects
characterized (reviewed in Roncero, 2002; Horiuchi,
2009). Genome sequencing revealed three to nine
CHS genes per individual fungal species, which were
categorized into seven gene classes. In contrast, nematode,
mollusk, crustacean, and insect genomes contain only
one or two CHS genes per species (Figure 2A). Since
Tellam et al. (2000) published the first cDNA sequence
for a CHS from Lucilia cuprina (sheep blowfly), cDNA
sequences for CHSs have been reported from numerous
invertebrates, and the availability of an increasing number
of genome sequences has provided additional information
on CHS genes. Nematode CHSs were from two filarial
pathogens, Brugia malayi, and Dirofilaria immitis, the
plant parasite Meloidogyne artiellia and Caenorhabditis
elegans (Harris et al., 2000; Harris and Fuhrman,
2002; Veronico et al., 2001). In both D. immitis and
M. artiella, there is currently only evidence for a single
gene, but in B. malyai and C. elegans, two genes were
identified. CeCHS1 is required for eggshell formation,
whereas CeCHS2 is needed to form the grinder in the
ectodermal pharynx (Zhang et al., 2005). CHS sequences
from crustaceans and chelicerates were deduced from
the Daphnia pulex and Ixodes scapularis genome projects,
both of which have two CHS genes. Likewise, all insect
genomes available so far harbor two CHS genes, which
have been divided into class A and class B genes, with the
latter appearing to be the more ancient form (Figure 2A).
The insect species from which complete cDNAs for CHSs
have been isolated are L. cuprina (Tellam et al., 2000),
D. melanogaster (Gagou et al., 2002), Ae. aegypti (Ibrahim
et al., 2000), Anopheles quadrimaculatus (Zhang and
Zhu, 2006), M. sexta (Zhu et al., 2002), S. frugiperda
(Bolognesi et al., 2005), Spodoptera exigua (Chen at al.,
2007; Kumar et al., 2008) and T. castaneum (Arakane
et al., 2004). Genomic sequences from Anopheles gambiae,
T. castaneum, D. melanogaster and M. sexta, which were
deduced from available genome projects or obtained by
individual nucleotide sequencing, were used to determine
the organization of CHS genes in these species (Figure 3).
The overall structure of CHS genes varies among different insect species and gene classes. The numbers of exons
range from 8 to 24, with lengths from 46 bp to more than
3000 bp. While most genes contain at least some exons
that contribute longer ORFs, the lepidopteran CHS genes
appear more fragmentized, because they contain a higher
number of shorter exons (Zhu et al., 2002; Kumar et al.,
2008). Insect CHS-A genes have two mutually exclusive
exons, resulting in two mRNA splice variants. Both exons
code for 59 amino acids comprising extracellular, transmembrane and intracellular domains, the latter being located
near the carboxyl terminus of the protein. One major difference between the two exons that are alternately spliced is
that all of the b forms code for segments that have a site for
N-linked glycosylation just before the transmembrane helix,
Figure 2 Phylogenetic trees of CHS proteins and conserved exons. The trees are based on ClustalW alignments and were
performed with the neighbor joining method. Bootstrap tests of phylogeny were performed with 10,000 replications., (A) Bootstrap
consensus tree of CHS proteins from fungi, nematodes, mollusks and arthropods., (B) Bootstrap consensus tree of exons a
and b found in class A CHS genes, and the corresponding region of class B CHS genes. Aa, Aedes aegypti (XP_001662200.1,
XP_001651163.1); Af, Aspergillus fumigatus (XP_749322.1, XP_746604.1, XP_748263.1, XP_752630.1, CAA70736.1,
XP_747364.1, XP_754184.1, XP_755676.1); Ar, Atrina rigida (AAY86556.1); Bm, Brugia malayi (XP_001898491.1, AAS77206.1);
Ce, Caenorhabditis elegans (NP_492113.2, NP_493682.2); Dm, Drosophila melanogaster (AAG22215.3, AAF51798.2);
Di, Dirofilaria immitis (AAG39382.1); Dp, Daphnia pulex (NCBI_GNO_134384, NCBI_GNO_326244); Is, Ixodes scapularis
(XP_002405234.1; XP_002405231.1); Lm, Locusta migratoria (ACY38589.1); Ms, Manduca sexta (AAL38051.2, AAX20091.1);
Ma, Meloidogyne artiellia (AAG40111.1); Mg, Mytilus galloprovincialis (ABQ08059.1); Nv, Nasonia vitripennis (XP_001602290.1,
XP_001602181.1); Pf, Pinctada fucata (BAF73720.1); Ph, Pediculus humanus corporis (XP_002423597.1), XP_002423604.1); Sc,
Saccharomyces cerevisiae (NP_014207.1, NP_009594.1, NP_009579.1); Tc, Tribolium castaneum (AAQ55059.1, AAQ55061.1).
7: Chitin Metabolism in Insects 201
whereas none of the a forms do. The precise physiological
significance of alternate exon usage and potential glycosylation in CHS expression and function is still unknown,
even though it is clear that there is developmental regulation of alternate exon usage (see section 7.3.4.2.).
7.3.3.2. Modular structure of chitin synthases CHSs
are members of family GT2 of the glycosyltransferases
(Coutinho et al., 2003), which generally utilize a
mechanism where inversion of the anomeric configuration
of the sugar donor occurs. The protein fold (termed
GT-A) for this family is considered to be two associated
β/α/β domains that form a continuous central sheet of at
least eight β-strands. The GT-A enzymes share a common
ribose/metal ion-coordinating motif (termed DxD
motif ) as well as another carboxylate residue that acts as a
catalytic base. The general organization of CHSs has been
deduced from a comparison of amino acid sequences of
these enzymes from several species of insects, nematodes,
and yeasts (Merzendorfer, 2006). These enzymes have
three distinguishable domains: an N-terminal domain
with moderate sequence conservation among different
species and containing several transmembrane segments;
a central catalytic domain that is believed to be orientated
toward the cytoplasm; and a C-terminal domain with
multiple transmembrane segments (Figure 4). The
catalytic domain contains several highly conserved
stretches including GT2 consensus sequences, which
have been suggested to be involved in binding of UDP,
the donor and acceptor saccharides, and the product.
They include sequences similar to the Walker A and B
motifs for binding of the nucleotide moiety (Walker et al.,
1982), sequences similar to the DXD and G(X)4(Y/F)
R motifs likely involved in substrate binding, the
GEDRxx(T/S) motif at the acceptor binding site, and
the (Q/R)XXRW motif involved in product binding.
The latter motif is present only in processive GTs.
While the transmembrane segments in the N-terminal
domain show different patterns among different insect
species, the transmembrane segments in the C-terminal
domain are remarkably conserved both with respect
to their location and the spacing between adjacent
transmembrane segments. Particularly striking is the fact
that five such transmembrane segments are found in a
cluster immediately following the catalytic domain, and
two more segments are located closer to the C-terminus.
The cluster of five transmembrane helices spanning the
membrane, known as 5-TMS (5-transmembrane spans),
has been suggested to be involved in the extrusion of the
polymerized chitin chains across the plasma membrane
to the exterior of the cell, as has been proposed for the
extrusion of cellulose (Richmond, 2000). Following
the last transmembrane helix of the 5-TMS, a sequence
similar to the (S/T)WGT(R/K) motif found in fungal
chitin synthases is located at the extracellular site. The
CHSs derived from class A genes were predicted to have
a coiled-coil region following the 5-TMS region (Zhu
et al., 2002; Arakane et al., 2004). Also, all of the genes
encoding the class A CHSs have two alternate exons
(corresponding to alternate exon 7 of D. melanogaster,
exon 8 of T. castaneum, exon 6 of A. gambiae, and the
exon 20 homolog of M. sexta). The alternate exons are
located on the C-terminal side of the 5-TMS region, and
encode the next transmembrane segment and flanking
Figure 3 Schematic diagram of the organization of insect CHS-A and CHS-B genes. The exon–intron organization was
deduced from comparisons of available cDNA and genomic sequences. Boxes indicate exons; lines indicate introns. The
second of the two alternative exons (8b) of TcCHS1, DmCHS1 (7b), AgCHS2 (7b), and MsCHS1 (homolog of exon 20 from
MsCHS2, 20b) are indicated as closed boxes (modified according to Arakane et al., 2004, and Hogenkamp et al., 2005). Ag,
Anopheles gambiae; Dm, Drosophila melanogaster; Ms, Manduca sexta; Tc, Tribolium castaneum.
202 7: Chitin Metabolism in Insects
sequences (Figure 4). The alternate exon-encoded regions
of the CHS proteins differ in sequence by as much as 30%,
and most of these differences are in the regions flanking
the transmembrane segment. This finding suggests that
the proteins may differ in their ability to interact with
cytosolic or extracellular proteins, which might regulate
chitin synthesis, transport, and/or organization. An
attractive hypothesis is that these flanking sequences
may influence the plasma membrane location of a CHS
by interacting with cytoskeletal elements, or perhaps
by generation of extracellular vesicles involved in chitin
assembly.
7.3.3.3. Zymogenic properties of chitin synthase In
numerous fungal and insect systems, chitin synthesis
is activated by trypsin and other serine proteases,
suggesting that CHS is produced as a zymogen (reviewed
in Merzendorfer, 2006). However, there is very little
knowledge on the significance of this phenomenon
in arthropods. In yeast, which has three CHS genes,
proteolytic activation by trypsin has been reported
for Chs1 and Chs2 (Cabib and Farkas, 1971; Sburlati
and Cabib, 1986). With Chs3, the situation is more
complicated, as the zymogenic properties appear to
depend on UDP-GlcNAc and additional proteins, such
as the regulatory subunit Chs4 (Choi et al., 1994; Ono
et al., 2000). However, no endogenous proteinase has
been identified that would cleave the CHS zymogen. The
zymogenic properties of yeast Chs2 and Chs3 have been
reinvestigated recently. For Chs2 it was demonstrated that
trypsin acts on a soluble protease that, once activated,
stimulates Chs2 activity (Martínez-Rucobo et al., 2009).
Another study reports a role of the CaaX proteinase
Ste24 in chitin synthesis (Meissner et al., 2010). Ste24
is a membrane-integral protease of the endoplasmic
reticulum, which is known to be involved in proteolytic
maturation of the yeast mating factor a. Yeast two-hybrid
studies have indicated, however, that Ste24 interacts
with Chs3. The interacting domain was mapped to a
cytosolic region that immediately precedes the catalytic
domain of Chs3. Deletion of ste24 led to Calcofluor
white (CFW) resistance and decreased chitin levels,
whereas overexpression led to CWF hypersensitivity and
increased chitin levels. The CFW phenotype of wild type
cells could be rescued by expressing the homologous gene
from T. castaneum in ste24Δ cells, indicating orthologous
functions. Although Ste24 directly binds to Chs3, it
appears not to be a substrate of the protease. Instead,
genetic experiments indicate that Chs4 is cleaved by Ste24
in a prenylation-dependent manner at its C-terminal
CaaX motif, and that this processing is required for
intracellular transport of Chs3 to the plasma membrane
(Meissner et al., 2010). Addition of trypsin to cell-free
extracts obtained from different insect species such as
Diaprepes abbreviatus, M. sexta, T. castaneum, and Stomoxys
calcitrans leads to the stimulation of chitin synthesis by
Figure 4 Structural model of the tripartite domain organization of Drosophila DmCHS1. The N-terminal domain A of Drosophila
contains 8 transmembrane helices (TMHs), but this number varies, between different insect species, from 7 to 10. The central
domain B is facing the cytoplasm, and forms the catalytic site. The ensuing domain C contains 5 + 2 TMHs, and the C-terminus
is located at the extraplasmatic site. Generally, all TMHs are highly conserved in insects. Putative motifs involved in nucleotide,
donor, acceptor, and product binding are indicated. The polymer is synthesized in the cytosol and the chitin chain needs to be
translocated across the membrane, a process that might require the 5TMS cluster and the extrusion motif SWGTR.
7: Chitin Metabolism in Insects 203
30–50% (Cohen and Casida 1980b; Mayer et al., 1980;
Ward et al., 1991; Zimoch et al., 2005). In Manduca,
trypsin-dependent stimulation of chitin synthesis was
observed in crude midgut extracts, but not in membrane
fractions of the midgut. However, it could be restored by
re-adding a soluble fraction, suggesting that trypsin does
not directly act on CHS but on a soluble protein that in
turn stimulates chitin synthesis, which is similar to the
recent finding of a soluble factor that activates yeast Chs2
(Zimoch et al. 2005; Martínez-Rucobo et al., 2009).
Attempts to directly purify and identify the soluble factor
from M. sexta have failed. However, a chympotrypsinlike peptidase (MsCTLP-1) was identified in the midgut,
which binds to the extracellular C-terminal domain of
MsCHS2 (Broehan et al., 2007). MsCTLP-1 is secreted
into the gut lumen when the larvae start to feed, and it
stimulates chitin synthesis after being proteolytically
activated by trypsin (Broehan et al., 2008). In line with the
assumption that CHS enzymes are produced as zymogens,
denaturing gel electrophoresis and immunoblotting of
oligomeric CHS complexes purified from the midgut of
M. sexta yielded a distinct pattern of CHS fragments,
which is consistent with the assumption that the CHS
monomer is cleaved twice during maturation, and that
the resulting three fragments are part of the active enzyme
(Maue et al., 2009).
7.3.4. Chitin Synthases: Regulation and Function
7.3.4.1. Regulation of chitin synthase gene expression Insect class CHS1and CHS2 genes encoding CHS-A and
CHS-B enzymes, respectively, are expressed in different
tissues and exhibit different patterns of expression during
development. Although technical difficulties associated
with the isolation of specific tissues free of other
contaminating tissues (mainly trachea) initially hampered
their unambiguous assignment, some general conclusions
can be drawn from studies investigating CHS gene
expression in different species and stages of development.
CHS genes are expressed at all stages of growth, including
embryonic, larval, pupal, and adult stages. CHS1 genes
are expressed over a wider range of developmental stages
(Tellam et al., 2000; Gagou et al., 2002; Zhu et al.,
2002). CHS2 genes are not expressed in the embryonic
or pupal stages but are expressed in the larval stages,
especially during feeding in the last instar and in adults,
including blood-fed mosquitoes (Ibrahim et al., 2000;
Zimoch and Merzendorfer, 2002; Arakane et al., 2004).
These developmental differences in CHS1 and CHS2
expression prompted the assumption that insect CHS
genes have specialized functions in different tissues or
at different developmental stages. Accordingly, LcCHS1
is expressed only in the carcass (larva minus internal
tissues) and trachea of L. cuprina, but not in salivary
gland, crop, cardia, midgut, or hindgut (Tellam et al.,
2000). In blood-fed female mosquitoes, a gene encoding
a CHS-B enzyme is expressed in epithelial cells of the
midgut (Ibrahim et al., 2000). In T. castaneum, TcCHS1
is expressed in embryos, larvae, pupae, and young adults,
but not in mature adults (more than a month old), while
TcCHS2 is expressed at early and late larval stages as well
as in adult stages, but not in embryos and pupae (Arakane
et al., 2004). Similar expression profiles were reported in
two lepidopteran insects for the CHS genes SeCHS1 in
S. exigua and SfCHS2 in S. frugiperda (Bolognesi et al.,
2005; Chen et al., 2007). Tissue-specific expression was
also investigated systematically in M. sexta (Zhu et al.,
2002; Hogenkamp et al., 2005; Zimoch et al., 2005).
These studies demonstrated that MsCHS1 is expressed in
epidermal cells and in the tracheal system of larvae and
pupae, whereas MsCHS2 is expressed only in midgut
tissue. Transcriptional regulation of CHS expression has
been suggested to be mediated by ecdysone-responsive
elements in the upstream regions of both Drosophila genes
(Merzendorfer and Zimoch, 2003; also see section 7.7).
For krotzkopf verkehrt (kkv), the gene encoding DmCHS1,
another mode of transcriptional control appears to exist,
as it is strongly upregulated in epidermal cells surrounding
wounds caused by microinjection needles (Pearson et al.,
2009). What is remarkable, however, is that kkv uses a
fundamentally different signaling pathway for wound
activation than other genes involved in wound healing,
such as ddc and ple coding for dopa decarboxylase and
tyrosine hydroxylase, respectively. While the latter two
genes require the JUN/FOS and grainy head (GRH)
transcription factors to induce the wound response,
transcriptional activities of the identified wound enhancer
in the kkv upstream region was not affected by these
transcription factors (Pearson et al., 2009).
A more recent finding is that a chitinous serosal cuticle
containing chitin is produced very early in development
of Aedes aegypti (Rezende et al., 2008). The serosal cuticle
was shown to contain chitin and to be responsible for the
development of desiccation tolerance of mosquito eggs.
The serosal chitin is apparently the product of a class A
CHS derived from the CHS1 gene. This burst of chitin
synthesis occurs long before organogenesis and before formation of the larval cuticle. Chitin has also been detected
in eggs, eggshells, and ovaries of Aedes aegypti (Moreira
et al., 2007). In ovaries and eggs of T. castaneum, we have
detected transcripts of TcCHS-A (our unpublished data).
To summarize, the analysis of expression patterns of
the two CHS genes in different tissues and periods of
development of several insects suggests that class A CHS
enzymes are synthesized by epidermal cells when cuticle
deposition occurs in embryos, larvae, pupae, and young
adults, whereas class B enzymes are produced by the midgut epithelial cells in the course of PM formation in the
larval and adult stages and is probably limited to these
feeding stages.
204 7: Chitin Metabolism in Insects
7.3.4.2. Tissue-specific expression of alternate exons The genes encoding class A CHSs from D. melanogaster,
A. gambiae, Ae. aegypti, T. castaneum, and M. sexta,
but not the genes encoding class B CHSs, exhibit two
alternately spliced exons, which are highly conserved
between different insect species (Figure 2B). Each exon
encodes a 59-amino acid segment following the 5-TMS
region. This segment contains a 20-aa transmembrane
region and flanking sequences. In addition, the presence
of a predicted coiled–coil region immediately following
the 5-TMS region in the CHSs encoded by those genes
that have the alternate exons suggests a link between these
two structural features, and the possibility of regulation
of alternate exon usage. In agreement with this idea,
transcripts containing either one of these exons have been
detected in T. castaneum, M. sexta, and, more recently, in
Ae. aegypti (Arakane et al., 2004; Hogenkamp et al., 2005;
Zimoch et al., 2005; Chen et al., 2007; Rezende et al.,
2008). In T. castaneum embryos, transcripts with either
exon 8a or 8b were detected, whereas in last instar larvae
and prepupae, only exon 8a transcripts were present. In
the pupal stage, however, transcripts with exon 8a or
exon 8b were abundant, along with trace amounts of a
transcript with both exons. In mature adults none of these
transcripts were detected, whereas TcCHS2 transcripts
were easily detected (Arakane et al., 2004). Injection of
dsRNA specific to either one of both alternately spliced
mRNAs revealed that splice variant 8a of TcCHS1 is
required for both the larval–pupal and pupal–adult molts,
whereas splice variant 8b is required only for the latter.
This finding, together with the relative amounts of these
mRNAs, suggested that the splice variant with exon 8a
contributes mostly to pupal cuticular chitin synthesis.
Nevertheless, the variant with exon 8b appears to have
a vital role in the emergence of the adult from the pupal
cuticle, which obviously cannot be fulfilled by the exon
8a isoform alone. With regard to relative amounts of both
splice variants, similar results were observed in fifth instar
larvae of M. sexta (Hogenkamp et al., 2005; Zimoch
et al., 2005). RT-PCR based detection of the alternately
spliced transcripts at different developmental stages in
the epidermis revealed that the ratio of mRNA levels
for both splice variants varies during development, with
MsCHS1 exon 20a being more predominant generally
than that with exon 20b (Hogenkamp et al., 2005).
Tracheal cells also express both variants of MsCHS1, but,
in this tissue, MsCHS1 with exon 20b is more abundant
(Zimoch et al., 2005). The latter finding was confirmed
also in L. migratoria (Zhang et al., 2010a). When
LmCHS1 expression was silenced by dsRNA injection
into second instar nymphs, the locusts developed three
distinct phenotypes exhibiting severe molting defects
and eventually died. While the knockdown of LmCHS1a
expression revealed phenotypes similar to those for
LmCHS1, the knockdown of LmCHS1 transcripts with
alternate exon b, which is more abundant than the one
with alternate exon a in tracheal tissue, led to crimped
cuticles. The major finding of that study, however, was
that the function of insect CHSs and their alternate exons
are conserved in both holo- and hemimetabolous insects.
As discussed above, AaCHS1 accounts for chitin
synthesis in the course of serosal cuticle formation in
Ae. aegypti embryos, and two splice variants containing
either exon 6a or exon 6b have been identified. Quantitative PCR showed that at the moment of serosal cuticle
formation, splice variant 6a is predominantly expressed.
The biochemical basis for a specific function, however,
remains unknown.
7.3.4.3. Knockout mutants and RNAi reveal differen­
tial functions of CHS genes Drosophila mutants and
RNAi experiments were extremely helpful in analyzing
the differential functions of the two CHS genes. EMS
mutagenesis and screening of the resultant mutant
embryos for defects in epidermal differentiation and
cuticular patterning helped to identify genes involved
in controlling cuticle morphology (Jüergens et al.,
1984; Nüsslein-Volhard et al., 1984; Wieschaus et al.,
1984; Ostrowski et al., 2002). These genes include kkv,
knickkopf (knk), grainy head (grh), retroactive (rtv), and
zepellin (zep), some of which will be discussed later, in
section 7.6. Mutations in these genes resulted in poor
cuticle integrity and reversal of embryonic orientation
in the egg to varying degrees. Generally, homozygous
mutant embryos failed to hatch. When these mutant
embryos were mechanically devitellinized, the cuticles
became grossly enlarged, yielding the “blimp” phenotype.
Interestingly, embryos derived from wild type females
treated with diflubenzuron or lufenuron displayed a
similar “blimp-like” phenotype when devitellinized,
indicat­ing that either genetic or chemical disruption of
chitin deposition leads to this phenotype (Ostrowski
et al., 2002; Gangishetti et al., 2009). Also, inhibition of
chitin synthesis in D. melanogaster embryos induced by
the CHS-specific inhibitor nikkomycine Z leads to cuticle
defects, as they are similar to those observed in Drosophila
kkv mutants (Tonning et al., 2006; Gangishetti et al.,
2009). Ostrowski et al. (2002) characterized the kkv gene
and identified it as a CHS-like gene, and Moussian et al.
(2005a) finally showed that this class A CHS is essential
for chitin synthesis in epidermal and tracheal cuticles.
Careful analysis of the ultrastructure of the embryonic
cuticle of kkv mutants confirmed that chitin synthesis
by a class A CHS is essential for procuticle formation.
Another interesting finding was that in kkv mutants the
cuticle frequently detaches from underlying epidermal or
tracheal cells, suggesting that chitin is also required for
anchoring the cuticle (Moussian et al., 2005a). In addition,
the head skeleton of kkv mutant embryos is undersized
and deformed, and sclerotization and pigmentation
7: Chitin Metabolism in Insects 205
are impaired. An unexpected finding was that kkv is
required for tracheal tube expansion, which starts before
chitin is actually deposited in the tracheal cuticle during
embryogenesis (Devine et al., 2005; Tonning et al., 2005).
This finding suggested that chitin has an additional
function in early tracheal morphogenesis. Histological
stainings with Congo red, WGA, or a fluorescence-labeled
chitin-binding domain revealed that the tracheal lumen
contains chitin cables before the tracheal cuticle is formed.
The loss of lumenal chitin evidently affects subapical
cytoskeletal organization of tracheal cells. Therefore, it was
hypothesized that the chitinous lumenal matrix is sensed
by tracheal cells to coordinate cytoskeletal organization,
which controls the diameter size of the tracheae.
RNAi experiments to interfere with CHS expression
and investigate CHS function have been performed in
T. castaneum, A. aegypti, A. gambiae, S. exigua, O. nubilalis, and Locusta migratoria. In T. castaneum, injection
of dsRNA for either TcCHS1 or TcCHS2 into young
larvae, penultimate instar larvae, and prepupae resulted
in a substantial knockdown of TcCHS1 and TcCHS2
mRNA levels. TcCHS1-specific RNAi disrupted larval–
larval, larval–pupal, and pupal–adult molts, and caused
a significant reduction in total chitin content (Arakane
et al., 2005). Interestingly, the phenotypes differed significantly depending on whether the insects were injected
in the penultimate larval, last larval, or prepupal instar.
The first of these groups failed to pupate and died without any splitting of the old larval cuticle, while the second group initiated the larval–pupal molt, but the pupae
died without shedding their larval exuviae, although
splitting of the old cuticle had occurred. The third group
failed to carry out the pupal–adult molt, and died as pharate adults trapped in their pupal exuviae. In contrast,
TcCHS2 dsRNA injection into last instar larvae or prepupae had no effect on pupal or adult development, but
when injected into penultimate instar, the larvae shrank
in size and died without molting to the last instar. As the
knockdown affected only immature or penultimate larvae,
it was suggested that TcCHS2 knockdown impairs chitin
synthesis necessary for PM formation. Indeed, when midguts prepared from last instar larvae treated with dsRNA
to TcCHS1 and TcCHS2 were stained with a fluoresceinconjugated chitin-binding domain, a fluorescent PM was
detected in larvae treated with dsRNA for TcCHS1, but
not following RNAi for TcCHS2 (Arakane et al., 2005).
These experiments provided strong evidence that the Tribolium chitin synthase genes, TcCHS1 and TcCHS2, have
different functions, as they are involved in the synthesis
of chitin in epidermal/tracheal cuticles and midgut PM,
respectively. In a succeeding study, dsRNA for either
one of the two CHS genes was injected into young and
old female adults to investigate effects on egg-laying and
embryogenesis (Arakane et al., 2008). When dsRNA for
TcCHS1 was injected into young female adults (less than
10 days old), the beetles died without laying any eggs.
When older female adults were injected, the beetles developed normally and laid eggs that were drastically reduced
in chitin content and failed to hatch. The embryos had
a twisted and enlarged blimp-like phenotype (Figure 5).
Hence, TcCHS1 appears to have roles in the development
of embryos and adults, in addition to its role in cuticle
formation. Interestingly, injection of dsRNA for TcCHS2
into adults led to a significant reduction in chitin content
of the PM, and caused death after 2 weeks. The female
beetles treated with dsRNA for TcCHS2 also failed to lay
eggs, presumably due to starvation, because the fat body
was significantly depleted due to autophagy (Arakane
et al., 2008). Similar to the situation in the beetle, RNAi
experiments to knock down the AaCHS2 transcripts of Ae.
aegypti showed that it is required in female mosquitoes for
the de novo synthesis of the PM after a blood meal (Kato
et al., 2006).
As the function of CHSs is vital for insect development
and survival, RNAi-mediated knockdown of CHS genes
could be a powerful approach in pest control. Based on
the observation that chitin synthesis can be blocked by
dsRNA injection in mosquitoes, Zhang et al. (2010b)
developed a method to generate a systemic knockdown
of CHS gene expression in A. gambiae larvae by feeding
nanoparticles consisting of chitosan and dsRNA specific
for the target gene. In line with the presumed function
of both CHSs in cuticle and PM syntheses, the larvae
became more susceptible to diflubenzuron and to Calcofluor White (CFW), when AgCHS1 or AgCHS2 expression, respectively, was inhibited. Another promising
approach would be to feed bacteria expressing dsRNA to
target genes, as was originally performed with C. elegans
(Timmons and Fire, 1998). Indeed, when E. coli bacteria
expressing dsRNA to SeCHS1 were fed to larvae of the
lepidopteran pest S. exigua, the survival rate was decreased
as they advanced in development (Tian et al., 2009).
7.4. Chitin Degradation and
Modification
Insects must periodically replace their old cuticle with a
new one because it is too rigid to allow for growth. Key
to this process is the elaboration of the molting fluid with
an assortment of chtitinases and proteases. Chitinases
are among a group of proteins that insects use to digest
the structural polysaccharide in their exoskeletons and
gut linings during the molting process (Kramer et al.,
1985; Kramer and Koga, 1986; Kramer and Muthukrishnan, 1997; Fukamizo, 2000). Precise regulation of chitin metabolism is a complex and intricate process that is
critical for insect growth, metamorphosis, organogenesis,
and survival (Arakane and Muthukrishnan, 2010). Chitin
content, which fluctuates throughout the life cycle of the
insect, is directly influenced not only by chitin synthases
206 7: Chitin Metabolism in Insects
Figure 5 Phenotypes of T. castaneum larvae after RNAi for genes of chitin metabolism. dsRNAs for the indicated genes (200 ng
per insect, n = 20) were injected into penultimate instar larvae (young larvae), last instar larvae, pharate pupae as indicated
above each panel. All animals injected with dsRNA for CHS-A, TcCHT10, TcNAG1, and TcCDA1 died at the ensuing molt.
Unlike RNAi of TcCHT10, injection of dsRNA (200 ng per insect) for TcCHT5 into penultimate instar and last instar larvae as well
as pharate pupae prevented only adult molt. When dsRNA for TcCHT7 (200 ng per insect) was injected into pharate pupae,
normal phenotypes were observed in the pupal stage. However, unlike buffer-injected controls, TcCHT7 dsRNA-treated insects
failed to expand their adult elytra and their wings did not fold properly (modified from Zhu et al., 2008c). Animals injected with
control dsRNA for EGFP developed in a normal fashion, and had no mortality or abnormal phenotype.
(CHSs), but also by chitinases (CHTs, EC 3.2.1.14) and
β-N-acetylglucosaminidases (NAGs, EC 3.2.1.52). Chitin
is digested in the cuticle and PM to GlcNAc by a binary
enzyme system composed of CHT and NAG (Fukamizo
and Kramer, 1985a, 1985b; Filho et al., 2002). The former enzyme from molting fluid hydrolyzes chitin into
oligosaccharides, whereas the latter, which is also found
in the molting fluid, further degrades the oligomers to
the monomer from the non-reducing end. In some cases,
additional unrelated proteins that possess one or more
chitin-binding domains (CBD), but are devoid of chitinolytic activity, enhance degradation of chitin (VaajeKolstad et al., 2005). This system also probably operates
in the gut during degradation of PM, and increases the
porosity of the PM. It may also help in the digestion of
chitin-containing prey (Bolognesi et al., 2005; Khajuria
et al., 2010).
The precise control of chitin content is critical not
only for the survival of the insect, but also for optimal
function of individual anatomical structures such as
wings and other appendages. In addition, modulation
of the physical properties of chitin-containing structures
of insects is accomplished, in part, by the deacetylation
of the polysaccharide by chitin deacetylases (CDAs, EC
3.5.1.41). Partially deacetylated chitin may have different protein-binding and physical properties than those of
chitin. The process of partially deacetylating chitin and
the importance of this modification for insect growth
7: Chitin Metabolism in Insects 207
and development have emerged as new areas of research
in insect molecular science (Luschnig et al., 2006; Wang
et al., 2006; Arakane et al., 2009).
7.4.1. Insect Chitinases
7.4.1.1. Cloning of genes encoding insect chitinases
and chitinase-like proteins Since the first report of
an insect chitinase, its cDNA and its corresponding
gene from M. sexta (MsCHT5) (Koga et al., 1987;
Kramer et al., 1993; Choi et al., 1997; Kramer and
Muthukrishnan, 1997), numerous insect CHT genes and
cDNAs have been cloned and characterized from several
insect species belonging to different orders, including
dipterans, lepidopterans, coleopterans, hemipterans, and
hymenopterans (Kramer and Muthukrishnan, 2005). The
organization of most of these genes is very similar to that
of MsCHT5, and most of the proteins display a domain
architecture consisting of catalytic, linker, and/or chitinbinding domains (CBD) similar to MsCHT5. These genes/
enzymes include epidermal chitinases from the silkworm
Bombyx mori (Kim et al., 1998; Abdel-Banat and Koga,
2001), the fall webworm Hyphantria cunea (Kim et al.,
1998), wasp venom from Chelonus sp. (Krishnan et al.,
1994), the common cutworm Spodoptera litura (Shinoda
et al., 2001), the fall armyworm Spodoptera frugiperda
(Bolognesi et al., 2005), a molt-associated chitinase from
the spruce budworm Choristoneura fumiferana (Zheng
et al., 2002), and midgut-associated chitinases from the
malaria mosquito A. gambiae (Shen and Jacobs-Lorena,
1997), yellow fever mosquito Ae. aegypti (de la Vega et al.,
1998; Khajuria et al., 2010), the beetle Phaedon cochleariae
(Girard and Jouanin, 1999), and the sand fly Lutzomyia
longipalpis (Ramalho-Ortigao and Traub-Cseko, 2003),
as well as several deduced from Drosophila genome data.
A smaller linkerless fat body-specific chitinase from the
tsetse fly Glossina morsitans (Yan et al., 2002), and a very
large epidermal chitinase with five copies of the catalytic
domain and multiple chitin-binding domain from the
yellow mealworm Tenebrio molitor (Royer et al., 2002),
have also been described.
Daimon et al. (2003) described a gene encoding
another type of chitinase from the silkworm, BmCHT-h.
The encoded chitinase shared extensive similarities with
microbial and baculoviral chitinases (73% amino acid
sequence identity to Serratia marcescens chitinase, and
63% identity to Autographa californica nuclear polyhedrosis virus chitinase). Even though this enzyme had the
signature sequence characteristic of a family 18 chitinase,
it had a rather low percentage of sequence identity with
the family of insect chitinases. It was suggested that an
ancestral species of B. mori acquired this chitinase gene
via horizontal gene transfer from Serratia or a baculovirus.
A gene encoding a CHT-like protein that is highly related to
BmCHT-h was also found in the pea aphid, Acyrthosiphon
pisum (Nakabachi et al., 2010).
Only after the complete genome sequences became
available was it recognized that insect genomes contain
a large number of genes encoding CHT-like proteins
widely divergent not only in their DNA and amino acid
sequences, but also in the organization of their domains
(Zhu et al., 2004, 2008a; Arakane and Muthukrishnan,
2010). The number of CHT genes per insect genome is
in the range of 7 to 24 for D. melanogaster, A. gambiae,
Ae. aegypti, B. mori, A. pisum, and T. castaneum. This
range excludes genes encoding CHT-like proteins whose
consensus sequences are poorly conserved (see section
7.1.2.2; Khajuria et al., 2010; Nakabachi et al., 2010;
Zhu et al., 2004, 2008a). The 22 genes that encode CHTs
or chitinase-like proteins (CHLPs) in T. castaneum have
been divided into eight subgroups, based on sequence
similarity and domain organization (Figure 6) (Arakane
Figure 6 Domain organization of T. castaneum chitinase gene family. The program SMART was used to analyze the identified
domains. TcCHT7 and TcCHT11 have a single transmembrane span at the N-terminal region. Blue boxes, signal peptide; pink
boxes, catalytic domain; green boxes, chitin binding domain; red boxes, transmembrane span; lines, linker regions.
208 7: Chitin Metabolism in Insects
and Muthukrishnan, 2010). The chitinases in all insect
species can be similarly classified into multiple groups
(Figure 7). There is only one copy of the gene encoding a group I chitinase (CHT5) in all species except for
A. gambiae, Ae. aegypti, and the human body louse Pediculus humanus corporis, in which obvious gene duplications
have occurred, resulting in one to four additional copies
(Khajuria et al., 2010). To date, only one gene representing each of the groups II, III, VI, VII, and VIII (CHT10,
7, 6, and 11, respectively) has been found in various insect
species. Interestingly, in addition to the group III CHT
genes (CHT7s with two catalytic domains) identified
in fully sequenced insect genomes such as T. castaneum,
D. melanogaster, A. gambiae, Ae. aegypti, C. pipiens, A. mellifera, N. vitripennis, A. pisum, and P. corporis, orthologs
have also been found in non-insect arthropod genomes,
including those of the crustacean water flea Daphnia
pulex, and the arachnid deer tick Ixodes scapularis, indicating an ancient origin of CHT7 that predates separation of the class Chelicerata more than five million years
ago. Group IV appears to be the largest group in all insect
species studied, containing 5, 8, 10, and 14 genes in D.
melanogaster, A. gambiae, Ae. aegypti, and T. castaneum,
respectively. The sole exception so far is A. pisum. No chitinase gene encoding a protein that belongs to this group
was identified in A. pisum (Nakabachi et al., 2010). In
T. castaneum, most group IV CHT genes form a large cluster within a small region of the genome, suggesting the
occurrence of a recent gene duplication event. Group V is
composed of the genes encoding CHLPs such as imaginal
disc growth factors (IDGFs). The number of genes for this
group ranges from one in B. mori and A. pisum to as many
as six in D. melanogaster (Arakane and Muthukrishnan,
2010; Nakabachi et al., 2010).
7.4.1.2. Domain organization of insect chiti­
nases Insect CHTs belong to family-18 glycosylhydro­
lases (the GH-18 super family) and function in hydrolysis
of chitin in the exoskeleton and PM-associated chitin in
the midgut, utilizing an endo-type cleavage mechanism
during the molting process (Kramer and Muthukrishnan,
1997, 2005). Members of the CHT family contain a
multidomain structural organization that includes a
leader peptide and/or a transmembrane span, one to five
catalytic domains (GH-18), multiple Ser/Thr-rich linker
regions that are usually heavily glycosylated, and zero to
seven six-cysteine-containing chitin-binding domains
(CBDs) related to the peritrophin A domain (Figure 6;
Royer et al., 2002; Arakane et al., 2003; Zhu et al.,
2008b). The catalytic domains of all insect CHTs, which
are comprised of about 370 amino acids, assume a β8α8barrel structure and possess signature motifs of family
18 glycosylhydrolases (Kramer et al., 1993; Perrakis
et al., 1994, Terwisscha van Scheltinga et al., 1994; de
la Vega et al., 1998; Fusetti et al., 2002; Varela et al.,
2002; Tsai et al., 2004; Arakane and Muthukrishnan,
2010). The consensus sequence for conserved motif I
is KXX(V/L/I)A(V/L)GGW in the β3-strand, where X is
a non-conserved amino acid. The conserved motif II is
FDG(L/F)DLDWE(Y/F)P, which is known to be located
in or near the catalytic site (β4-strand) of the enzyme,
with a glutamate residue (E) being the most critical
residue in this motif as the putative proton donor in the
catalytic mechanism (Watanabe et al., 1993; Lu et al.,
2002; Zhang et al., 2002). Conserved motifs III and IV
are MXYDL(R/H)G in the β6-strand and GAM(T/V)
WA(I/L)DMDD in the β8-strand.
CBDs found in insect CHTs all belong to carbohydratebinding module 14 (CBM-14, pfam 01607; ChtBD2
family = SMART family 00494, Boraston et al., 2004).
Insect CBDs are only about 60 amino acids long and have
less conserved amino acid sequences, with the exception
of the six cysteines and several aromatic residues whose
relative locations are highly conserved (Jasrapuria et al.,
2010). The proposed function(s) of the CBD is to help
anchor the enzyme onto the insoluble chitin to enhance
chitin degradation efficiency (Linder et al., 1996; Arakane
et al., 2003). As described in section 7.4.1.1, based on the
amino acid sequence similarity and domain architecture,
insect CHTs can be classified into eight groups (Figures
6 and 7). Group I CHTs (CHT5s) represent the prototypical and enzymatically characterized CHTs purified
from molting fluid and/or integument of M. sexta and
B. mori (Koga et al., 1983, 1997). All of these group
members contain a signal peptide, one catalytic domain, a
Ser/Thr-rich linker region, and one CBD. Group II CHTs
(CHT10s) are rather diverse in their domain architecture,
and have four or five catalytic domains, together with
four to seven CBDs. Dipterans and A. pisum (hemiptera)
appear to be unique in having only four catalytic domains
and four CBDs. The domain corresponding to the most
N-terminal catalytic domain and one CBD found in
group II chitinases from other species appear to be missing in the dipteran CHT10s (Zhu et al., 2008b; Arakane
and Muthukrishnan, 2010; Nakabachi et al., 2010). The
second catalytic unit of all CHT10s (the first catalytic
unit in the case of the dipteran and A. pisum proteins) is
predicted to lack chitinolytic activity due to a substitution
of the most critical amino acid residue glutamate (E) with
asparagine (N) in conserved motif II. Group III CHTs
(CHT7s) possess two catalytic domains and one C-terminal CBD. The first catalytic domains of the group III proteins from all insect species studied share greater sequence
similarity with each other than they do to the second catalytic domain, suggesting a unique function and/or evolutionary origin for each of the catalytic domains. Unlike
most insect CHTs, CHT7s are predicted to have an
N-terminal transmembrane segment, and are likely to be
membrane-bound proteins. Indeed, recombinant T. castaneum CHT7 (TcCHT7) that was expressed in Hi-5 insect
7: Chitin Metabolism in Insects 209
Figure 7 Phylogenetic analysis of putative chitinases and chitinase-like proteins (IDGFs) in insects. ClustalW software was
used to perform multiple sequence alignments prior to phylogenetic analysis. The phylogenetic tree was constructed by
MEGA 4.0 software using UPGMA (Tamura et al., 2007). Protein sequences obtained from GenBank as follows: Tribolium
castaneum, TcCHT2 (AY873913); TcCHT4 (EF125543); TcCHT5 (AY675073); TcCHT6 (AY873916); TcCHT7 (DQ659247); TcCHT8
(DQ659248); TcCHT9 (DQ659249); TcCHT10 (DQ659250); TcCHT11 (DQ659251);TcCHT12 (XM_967709); TcCHT13 (DQ659252);
TcCHT14 (XM_967912); TcCHT15 (XM_967984); TcCHT16 (AY873915); TcCHT17 (XP_972719); TcCHT18 (XP_973161); TcCHT19
(XP_973119); TcCHT20 (NP_001034516); TcCHT21 (NP_001034517); TcCHT22 (NP_001038095); TcIDGF2 (DQ659253);
TcIDGF4 (DQ659254); Aedes aegypti, AaCHT1 (XP_001656232); AaCHT2 (XP_001662520); AaCHT3 (XP_001663568);
AaCHT4 (XP_001663099); AaCHT5 (XP_001656234); AaCHT6 (XP_001662588); AaCHT7 (XP_001650020); AaCHT8
(XP_001663098); AaCHT9 (XP_001663099); AaCHT10 (XP_001655973); AaCHT11 (XP_001654045); AaCHT12 (XP_001658836);
AaCHT13 (XP_001656231); AaCHT14 (XP_001656233); AaBR1 (XP_001660745); AaBR2 (XP_001660748); Apis mellifera,
AmCHT2 (XP_623744); AmCHT5 (XP_623995); AmCHT6 (XP_393252); AmCHT7 (XP_396925); AmCHT10 (XP_395734);
AmCHT11 (XP_395707); AmIDGF (XP_396769); Drosophila melanogaster, DmCHT2 (NP_477298); DmCHT5 (NP_650314);
DmCHT6 (NP_572598); DmCHT7 (NP_647768); DmCHT10 (NP_001036422); DmCHT11 (NP_572361); DmIDGF1 (NP_477258);
DmIDGF2 (NP_477257); DmIDGF3 (NP_723967); DmIDGF4 (NP_727374); DmIDGF5 (NP_611321); DmDS47 (NM_057733);
Bombyx mori, BmCHT2 (BGIBMGA009695); BmCHT5 (BGIBMGA010240); BmCHT6 (BGIBMGA009890); BmCHT7
(BGIBMGA005539); BmCHT10 (BGIBMGA006874); BmCHT11 (BGIBMGA005859); BmIDGF (BGIBMGA000648);
Anopheles gambiae, AgCHT2 (XP_315650); AgCHT5 (XP_001237469); AgCHT6 (AGAP000198); AgCHT7 (XP_308858);
AgCHT10 (XP_001238192); AgCHT11 (XP_310662); AgBR1 (AAS80137); AgBR2 (AY496421); Nasonia vitripennis, NvCHT2
(XP_001601416); NvCHT5 (NP_001155084); NvCHT6 (); NvCHT7 (XP_001604515); NvCHT10 (XR_036825); NvCHT11
(XP_001604954); NvIDGF (XP_001599305.
210 7: Chitin Metabolism in Insects
cells using the baculovirus protein expression system was
found to be in the cell pellet rather than in the medium,
as expected for secreted proteins. The washed cell pellet
containing recombinant TcCHT7 could hydrolyze chitin
added to the culture medium, suggesting that the catalytic domains of this putative membrane-bound protein
face the extracellular space (Arakane, unpublished data).
Group IV CHTs comprise the largest and most divergent
group of proteins. CHTs in this group have a signal peptide and one catalytic domain. Most (but not all) of the
members lack a CBD (Figure 6). Group V chitinase-like
proteins (CHLPs) include the imaginal disc growth factors
(IDGFs) and the hemocyte aggregation inhibitor protein
(HAIP, Kanost et al., 1994; Pan et al., 2010). CHLPs have
a signal peptide, one catalytic domain, and no CBDs. Like
other family-18 proteins, the crystal structure of D. melanogaster IDGF2 and homology modeling of all proteins in
this group revealed the β8α8-TIM barrel structure (Varela
et al., 2002). However, members of this group have an
additional loop sequence located between the β4-strand
and the α4-helix immediately after conserved region II.
Although these proteins possess all four of the family-18
conserved motifs, the glutamate residue in conserved
motif II is substituted by a glutamine in all members of
the group, with the exception of two T. castaneum IDGFs
(TcIDGF2 and TcIDGF4; Zhu et al., 2008b). TcIDGF2
and TcIDGF4 retain the glutamate residue in conserved
region II but lack chitinase activity, either due to a D to A
substitution in the conserved motif II, or to an extra loop
stretching between the β4-strand and the α4-helix that
possibly interferes with a productive substrate–enzyme
interaction (Zhu et al., 2008a), or both. Group VI CHTs
(CHT6s) exhibit a domain architecture similar to that of
group I (a signal peptide, one catalytic domain, and one
CBD), but they have a very long C-terminal stretch (e.g.,
1819 amino acids in length after the CBD in TcCHT6)
that has no predicted conserved domain (Figure 6) except
for the A. pisum enzyme, which possesses an additional
CBD at the C-terminal region (Nakabachi et al., 2010).
Group VII CHTs (CHT2s) possess a domain architecture
similar to that of group IV CHTs, which have a signal
peptide, one catalytic domain, and no CBDs. They are
classified as a separate group because phylogenetic analysis
clearly indicates that these CHTs form a different clade
near group II CHT10s. Group VIII CHTs (CHT11s)
have one catalytic domain and no CBD. Interestingly,
they have a predicted transmembrane segment instead of
a signal peptide at the N-terminus, and they fall into a
branch next to group III (CHT7s), all of which are predicted to be membrane-bound proteins.
7.4.1.3. Gene expression and functions of insect
chitinases The redundancy of genes for CHTs raises
important questions about their functions. Several
insect CHT cDNAs have been obtained from epidermis,
gut, and fat body, and extensively characterized
(Kramer and Muthukrishnan, 2005). The epidermal
endochitinases presumably function in turnover of the
old cuticle, as these enzymes are found in the molting
fluid along with N-acetylglucosaminidases, whereas the
gut CHTs are thought to participate in the breakdown
of chitin in the PM. In T. castaneum, tissue specificity
and developmental patterns of expression of the 22
TcCHT and TcCHLP genes were analyzed by RT-PCR
using cDNAs prepared from RNAs isolated at different
developmental stages, such as embryo, larva, pharate
pupa, pupa, and adult (Zhu et al., 2008c; Arakane and
Muthukrishnan, 2010). The group I gene TcCHT5,
group II gene TcCHT10, group III gene TcCHT7,
group V genes TcIDGF2 and TcIDGF4, group VI gene
TcCHT6, group VII gene TcCHT2, and group VIII
gene TcCHT11 are expressed at all stages analyzed, with
some variation, whereas all group IV genes (TcCHTs
2, 4, 8, 9, and 12 to 22) were predominantly expressed
in the feeding stages (larva and adult). In addition, all
chitinase genes belonging to group IV were expressed
in larval gut tissue but not in the carcass (whole body
minus gut), suggesting a possible function of these
TcCHTs in PM-associated chitin turnover or digestion
of dietary chitin (Zhu et al., 2008c). Khajuria et al.
(2010) recently reported that orally feeding dsRNA for
a midgut-specific chitinase gene (encoding a group IV
CHT) from larvae of O. nubilalis (OnCHT) significantly
reduced the transcript levels of this gene and led to a
significant increase of chitin content in the PM. The
body weight of dsRNA OnCHT-fed larvae was decreased
by 54% as compared with that of control dsRNA GFPfed larvae, suggesting that some group IV CHTs are
critical for regulating PM-chitin content, insect growth,
and development. Interestingly, A. pisum appears to
have no group IV CHT genes (Nakabachi et al., 2010).
A. pisum (hemipteran) possesses a perimicrovillar
membrane (PMM) that is devoid of chitin, suggesting
that group IV CHTs may not play a role in the PM
turnover. Instead, one CHT gene, ApCHT6 (encoding
a group VIII CHT), was highly expressed in the midgut
of A. pisum. Similarly, TcCHT11 (encoding a group VIII
CHT) was expressed in larval midgut, but not in the
carcass (Arakane and Muthukrishnan, 2010). Group
VIII CHTs, as well as group VI CHTs, may play critical
roles in PM/PMM chitin degradation and turnover.
RNAi for group IV chitinases in T. castaneum for individual chitinases (and some combinations of chitinases)
failed to produce any visible phenotypes, perhaps reflecting
the redundant functions of this large group of chitinolytic
enzymes. In contrast, injection of dsRNA for all chitinases
belonging to groups I, II, III, and V resulted in unique
lethal phenotypes. The most severe molting defect was
observed after injection of dsRNA for TcCHT10 (encoding a group II CHT). Injections of dsRNA for TcCHT10
7: Chitin Metabolism in Insects 211
prevented the embryo from hatching and also averted all
types of molts, including larval–larval, larval–pupal, and
pupal–adult, depending on the timing of administration
of the dsRNA (Figure 5; Zhu et al., 2008c). These results
suggest a critical role for group II CHTs at every molt
and developmental stage. Other CHTs (e.g., CHT5, also
expressed in the epidermis) could not compensate for the
loss of function of a group II CHT.
Unlike RNAi for TcCHT10, injection of dsTcCHT5
(encoding a group I CHT) prevented only the pupal–
adult molt (Figure 5). Although the gene encoding this
prototypical CHT was expressed throughout all developmental stages, and the corresponding enzymes from several other insect species have been found in larval molting
fluid, the failure to obtain a larval–larval or larval–pupal
molting arrest probably indicates that one or more of
the other CHTs (e.g., group II CHT, TcCHT10) could
compensate for TcCHT5 at all molts except during adult
eclosion. Group III CHTs, which appear to encode membrane-bound enzymes with two catalytic domains and
one CBD at the C-terminus, appear to be critical for tissue differentiation, rather than chitin degradation associated with molting. Indeed, in D. melanogaster, expression
of the DmCHT7 (CG1869) gene increased more than
40-fold in the wing during the 32- to 40-h pupal wing
differentiation period (Ren et al., 2005). In T. castaneum,
injection of dsRNA for CHT7 resulted in a defective elytral and hindwing expansion without affecting molting
(Figure 5; Zhu et al., 2008c). Group V is composed of
IDGFs that are known to be involved in cell proliferation and differentiation (Kawamura et al., 1999; Zhang
et al., 2006). It is worthy of note that although group V
CHTs have no chitinolytic activity (Zhu et al., 2008b),
they appear to be important for the adult molt. Injection of dsRNA for one of these CHLPs in T. castaneum,
TcIDGF4, prevented adult eclosion (Zhu et al., 2008c).
It is possible that TcIDGF4 may be required for tracheal
proliferation during adult metamorphosis. Two A. gambiae proteins, AgBR1 and AgBR2, which belong to this
group, were induced specifically in the hemolymph by
bacterial challenge (Shi and Paskewitz, 2004), suggesting
that some members of the CHLP group (and/or members
of other CHT groups) may have a role in the immune
response.
7.4.2. Insect N-Acetylglucosaminidases
7.4.2.1. Phylogenetic analysis of insect N-acetylglu­
cosaminidases Beta-N-acetylglucosaminidases (NAGs;
EC 3.2.1.30) have been defined as enzymes that release –
acetylglucosamine residues from the non-reducing
end of chitooligosaccharides and from glycoproteins
with terminal N-acetylglucosamines. Insect NAGs are
members of the family-20 hexosaminidase super-family
of the glycosylhydrolases of the Carbohydrate Active
Enzymes database, CAZY (Coutinho and Henrissat,
1999; Cantarel., et al., 2009). These enzymes have been
detected in the molting fluid, hemolymph, integument,
and gut tissues of several species of insects (Kramer and
Koga, 1986; Hogenkamp et al., 2008), and cooperate
with CHTs to hydrolyze chitin to generate monomers
of N-acetylglucosamine (Fukamizo and Kramer, 1985a,
1985b). Insect CHTs are unable to convert the chitin
substrate completely to GlcNAc monomers. Therefore,
NAG is the enzyme primarily responsible for the
production of the monomer from chitooligosaccharides
for recycling. Kinetic studies with M. sexta CHT
(MsCHT5, group I CHT) have revealed that this enzyme
is subject to substrate and/or product inhibition when
chitooligosaccharides and/or colloidal chitin are utilized as
substrates (Koga et al., 1982, 1983; Arakane et al., 2003).
Therefore, one of the potential functions of NAGs may
be to prevent the accumulation of chitooligosaccharides
at concentrations that are high enough to interfere with
efficient degradation of chitin by CHT (Kramer and
Muthukrishnan, 2005).
cDNAs for epidermal β-N-acetylglucosaminidases
of B. mori, B. mandarina, T. ni, and M. sexta have been
isolated and characterized (Nagamatsu et al., 1995; Zen
et al., 1996; Goo et al., 1999; Hogenkamp et al., 2008).
A NAG also has been detected in the gut of Ae. aegypti,
where its activity increased dramatically upon blood feeding (Filho et al., 2002). A search of the D. melanogaster,
A. gambiae, Ae. aegypti, Culex pipiens, A. mellifera, N.
vitripennis, B. mori, and T. castaneum genome databases
revealed the presence of multiple NAG genes, as well as
the genes encoding β-N-acetylhexosaminidases (HEXs)
in these species (Hogenkamp et al., 2008). Phylogenetic
analysis of NAGs from these insects indicates that NAGs
can be classified into four distinct groups – NAG group
I (NAG1), NAG group II (NAG2), N-glycan processing NAGs (FDL) (group III, Leonard et al., 2006), and
HEX group IV – according to their amino acid sequences
­(Figure 8). To date, only a single gene representing each
of the groups I, II, and III has been found in the various insect species, with the exception of C. pipiens, which
appears to have three genes encoding NAG-like proteins
closely related to group I NAGs. Group I is composed
of the enzymatically well-characterized NAGs, including
NAGs from M. sexta (MsNAG1) and B. mori (BmNAG1).
DmHEXO2, which has been shown to have NAG activity (Mark et al., 2003; Leonard et al., 2006), was placed
in group II. Group III is composed of the D. melanogaster fused lobes protein (DmFDL), along with the fused
lobes (fdl) homologs of other insect species (Leonard
et al., 2006). All of the proteins belonging to this group
possess a predicted transmembrane anchor and a signal
anchor, except for a signal peptide that can be found in
NAGs belonging to groups I, II, and IV. In T. castaneum,
TcNAG3 could not be unambiguously assigned to any of
212 7: Chitin Metabolism in Insects
7: Chitin Metabolism in Insects 213
Figure 8 Phylogenetic analysis of NAGs and hexoaminidases in Tribolium, other insects and metazoans. MEGA4.0 (Tamura
et al., 2007) was used to construct the consensus phylogenetic tree using UPGMA. Bootstrap analyses of 1000 replications
are shown. Protein sequences extracted from GenBank include: MsNAG, Manduca sexta (AY368703); BmNAG, Bombyx mori
(genbank: AF326597); TnNAG, Trichoplusia ni (AY078172); AmNAG1, Apis mellifera (XM_624790); TcNAG1, Tribolium castaneum
(EF592536); DmNAG1 (DmHEXO1), Drosophila melanogaster (NM_079200); AgNAG1, Anopheles gambiae (XP_315391);
CqNAG1a, Culex quinquefasciatus (XP_001864406); AaNAG1, Aedes aegypti (EAT43909); CqNAG1b, Culex quinquefasciatus
(XP_001864407); CqNAG1c, Culex quinquefasciatus (XP_001866097); TcNAG3, Tribolium castaneum (EF592538); TcFDL,
Tribolium castaneum (EF592539); AmFDL, Apis mellifera (XP_394963); DmFDL, Drosophila melanogaster (NP_725178); AgFDL,
Anopheles gambiae XP_308677); CqFDL, Culex quinquefasciatus (XP_001850423); AaFDL, Aedes aegypti (EAT36388); TcNAG2,
Tribolium castaneum (EF592537); DmNAG2 (DmHEXO2), Drosophila melanogaster (NM_080342); AgNAG2, Anopheles gambiae
(XM_307483); CqNAG2, Culex quinquefasciatus (XP_001842710); AaNAG2, Aedes aegypti (EAT40440); HsHEXA, Homo
sapiens (NM_000520); HsHEXB, Homo sapiens (NM_000521); MsHEXA, Mus musculus (NM_010421); MsHEXB, Mus musculus
(NM_010422); SfHEX1, Spodoptera frugiperda (DQ183187); SfHEX2, Spodoptera frugiperda (DQ249307); BmHEX, Bombyx
mori (AY601817); TcHEX3, Tribolium castaneum (XM_970565); AmHEX, Apis mellifera (XM_001122538); TcHEX1, Tribolium
castaneum (XM_970563); TcHEX2, Tribolium castaneum (XM_970567); CqHEX2, Culex quinquefasciatus (XP_001867058);
AgHEX, Anopheles gambiae (XM_319210); CqFEX1, Culex quinquefasciatus (XP_001867057); and AaHEX, Aedes aegypti
(EAT43655.
the three subgroups. TcNAG3 is more closely related to
TcFDL than to TcNAG1 and TcNAG2, but the TcFDL
and TcNAG3 genes are present on different linkage
groups (Figure 8, Hogenkamp et al., 2008).
7.4.2.2. Expression and functional analysis of insect
N-acetylglucosaminidases Hogenkamp and colleagues
(2008) performed dsRNA-mediated post-transcriptional
downregulation (RNAi) of transcripts for all four NAG
genes from a single insect species (T. castaneum) to study
the functions of insect NAGs. Injection of a dsRNA
corresponding to any one TcNAG gene resulted in
substantial downregulation of the target transcript without
significantly affecting the levels of the other TcNAG
transcripts. Depletion of transcripts for any one of the
targeted genes produced lethal molting arrest phenotypes.
However, some of the injected insects did succeed in
completing each type of molt (larval–larval, larval–pupal,
and pupal–adult). TcNAG1 appeared to be most critical
in chitin catabolism during molting. Administration of
dsRNA for TcNAG1 resulted in developmental arrest,
and more than 80% of the insects died at the time of
the next molt (Figure 5). During each type of molt,
larval–larval, larval–pupal, and pupal–adult, the insects
were unable to completely shed their exoskeleton. The
pupa–adult molting phenotype produced by injection of
dsRNA for TcNAG1 is strikingly similar to that obtained
in RNAi studies with dsTcCHT5 (Figure 5; see section
7.4.1.3). Insects injected with dsRNA for TcCHT5 also
failed to shed their old cuticle, and the new cuticle was
visible underneath the old cuticle (Zhu et al., 2008c;
Arakane and Muthukrishnan, 2010). It has been shown
that in M. sexta, CHT is susceptible to oligosaccharide
inhibition (Koga et al., 1982, 1983; Arakane et al.,
2003). Injection of dsRNA for TcNAG1 may result in
the accumulation of chitiooligosaccharides in the molting
fluid, and therefore it may cause inhibition of TcCHT5
activity, resulting in a phenotype similar to that observed
in dsRNA for TcCHT5-treated insects. The high level
of expression of TcNAG1, its phylogenetic relationship
to other well-characterized molting-associated insect
NAGs (Figure 8), and the phenotypic effect of knocking
down TcNAG1 transcripts suggest that, among all of
the TcNAGs, TcNAG1 (group I NAG) is the enzyme
primarily responsible for the efficient degradation of
cuticular chitin, in concert with TcCHT5 (group I
CHT), in T. castaneum, and that this may be the case in
other insect species as well.
Although TcNAG1 is most likely to be the principal
NAG for catabolism of cuticle-associated chitin, the other
three NAGs identified in T. castaneum also appear to play
important and perhaps indispensable roles in cuticle turnover and development. Injection of dsRNA for TcNAG2
(encoding a group II NAG orthologous to DmHEXO2)
prevents all types of molts, especially the pupal–adult
molt. Like the phenotype produced by injection of
dsRNA for TcNAG1 (Figure 5), more than 75% of the
animals treated with dsRNA for TcNAG2 were unable to
fully shed the old pupal cuticle. Since injection of dsRNA
for TcNAG2 did not change the level of TcNAG1 transcripts, TcNAG1 could not compensate for the lack of
TcNAG2 in adult eclosion in T. castaneum. In addition,
TcNAG2 transcript level in the midgut is relatively higher
than that in the carcass (whole body minus midgut), suggesting TcNAG2 as well as TcNAG1, which are highly
expressed in both tissues, also play critical roles in the PMassociated chitin turnover.
Group III consists of the insect orthologs of the
D. melanogaster fused lobes gene, DmFDL. The FDL
proteins are predicted to be membrane-bound, with a
single transmembrane helix located near the N-terminus.
Furthermore, ultracentrifugation experiments on a lepidopteran protein from the culture media of Sf 9 and Sf 21
cells indicated that a major portion of the NAG activity
214 7: Chitin Metabolism in Insects
resided in the membrane fraction (Altmann et al., 1995;
Tomiya et al., 2006). This lepidopteran NAG was capable
of effectively hydrolyzing chitotriose-PA (pyridylamino),
while the recombinant DmFDL was unable to digest
chitotriose (Leonard et al., 2006). The latter hydrolyzed
only the GlcNAc residue attached to the α-1,3-linked
mannose of the core pentasaccharide of N-glycans.
No cleavage activity of any other GlcNAc residues was
observed, including the GlcNAc residue attached to
the α-1,6-linked mannose of the core pentasaccharide.
Furthermore, DmFDL did not catalyze the endo-type
hydrolysis of the N,N′-diacetylchitobiosyl unit in the
high-mannose pentasaccharide core. A similar N-glycan
substrate specificity for the terminal GlcNAc attached
to the α-1,3-linked mannose was observed in membrane-bound β-N-acetylhexosaminidases from several
lepidopteran insect cell lines, including Sf21, Bm-N, and
Mb-0503 (Altmann et al., 1995; Tomiya et al., 2006).
Taken together, FDLs may play a critical role in N-glycan
processing.
Unlike RNAi for TcNAG1 (group I NAG), injection of
dsRNA for TcFDL exhibits a small percentage (10–20%)
of lethal molting defect phenotypes at the larval–larval
and larval–pupal molts (Hogenkamp et al., 2008). Much
higher mortality (80%), however, was observed at the
pupal–adult molting stage, indicating that TcFDL plays
an essential role for adult eclosion. The transcript level of
TcFDL in the midgut was relatively low compared to that
of the carcass. Therefore, the observed lethal phenotype
at the pharate adult stage may be a direct result of the
knockdown of this transcript in the cuticular epidermal
cells, rather than in the gut lining cells. If TcFDL does
in fact play a role in chitin turnover in the cuticle, then
this protein may be secreted and not membrane-bound.
Indeed, Leonard and colleagues (2006) have observed that
DmFDL is, to a large extent, secreted into the extracellular space. Whether there is another point of regulation at
the level of release of membrane-bound FDLs is an interesting possibility.
Another T. castaneum NAG, TcNAG3, has not been
unambiguously assigned to any of the three NAG groups
(Figure 8). Similar to TcNAG2 (group II NAG), TcNAG3
is also expressed at a significantly higher level in the larval midgut than in the carcass (Hogenkamp et al., 2008).
Furthermore, an analysis of the developmental pattern
of expression of TcNAG3 indicated that it is primarily
expressed during the larval stages. Unlike RNAi for the
other three TcNAGs, injection of dsTcNAG3 did not consistently result in lethal phenotypes, and the majority of
dsRNA-injected insects survived to adults with no visible
phenotypic changes. However, a small number of individuals (approximately 20%) did exhibit a lethal larval
phenotype similar to that of TcNAG1 RNAi (Figure 5).
In addition, a few insects (approximately 10%) exhibited
a lethal pharate adult molting phenotype after dsRNA
TcNAG3 injection. These insects were unable to fully shed
their old pupal cuticle, similar to the phenotypes observed
after dsRNA TcNAG1 and dsRNA TcNAG2 injections.
The TcNAG3 gene is expressed predominantly in the
larval stages, with only trace levels of expression in the
pupal and adult stages (Hogenkamp et al., 2008). In other
insect species analyzed, only genes that can be classified
into groups NAG1, NAG2, and FDL have been identified
(Figure 8). Therefore, TcNAG3 appears to be unique, and
its relatively high expression in the midgut compared to
the carcass suggests that it may be specialized for the turnover of PM-associated chitin rather than cuticular chitin
during larval stages.
7.4.3. Insect Chitin Deacetylases
7.4.3.1. Phylogenetic analysis and domain organi­
zation of chitin deacetylases The extracellular matrix
(ECM) of the insect exoskeleton is modified in different
ways to give the cuticle its proper physiological and
mechanical properties – namely, rigidity and thickness,
or flexibility and thinness (Kramer and Muthukrishnan,
2005). Chitin deacetylases (CDAs, EC 3.5.1.41) are
secreted metalloproteins that belong to a family of
extracellular chitin-modifying enzymes that catalyze the
N-deacetylation of chitin to form chitosan, a polymer of
β-1,4-linked D-glucosamine residues with electrostatic
properties very different from chitin. This modification
might contribute to the affinity of chitosan for a variety of
cuticular proteins distinct from those that bind specifically
to chitin. CDAs have been well characterized in various
fungi and bacteria (Caufrier et al., 2003), and belong to
the carbohydrate esterase family 4 (CE4) of the CAZY
database (www.cazy.org; Cantarel et al., 2009). CE4
esterases catalyze deacetylation of different carbohydrate
substrates, such as chitin, acetylxylan, and bacterial
peptidoglycan. Chitooligosaccharide deacetylases and
NodB, a nodulation protein from Rhizobium, belong to
this family, and possess a similar catalytic domain (John
et al., 1993).
The first cDNA encoding an insect CDA-like protein
(TnPM-P42, also referred to as TnCDA9) was characterized from the PM in the cabbage looper, Trichoplusia ni,
only 5 years ago (Guo et al., 2005). Since then, several
genes/cDNAs encoding insect CDAs have been identified
from different species (Luschnig et al., 2006; Wang et al.,
2006; Campbell et al., 2008; Dixit et al., 2008; Toprak
et al., 2008; Jakubowska et al., 2010). A comparative
analy­sis of CDA gene families in several insect species with
fully sequenced genomes, including Diptera, Coleoptera,
Hymenoptera, and Lepidoptera, revealed that the number
of CDA genes varies with species. Based on amino acid
sequence similarity, insect CDAs are classified into five
groups, I to V (Figure 9; Dixit et al., 2008; Jakubowska
et al., 2010).
7: Chitin Metabolism in Insects 215
Figure 9 A phylogenetic tree of putative CDAs from different insects. A consensus phylogenetic tree was constructed using
neighbor-joining method in the software MEGA 4.0 (Tamura et al., 2007). Protein sequences obtained from GenBank as
follows; NvCDA1, Nasonia vitripennis (XP_001604765); AmCDA1, Apis mellifera (XP_391915); TcCDA1, Tribolium castaneum
(ABU2522); HaCDA1, Helicoverpa armigera (ADB43610); BmCDA1, Bombyx mori (BGIBMGA006213); DmCDA1, Drosophila
melanogaster (NP_730444); AgCDA1, Anopheles gambiae (XP_320597); DmCDA2, Drosophila melanogaster (NP_001163469);
AgCDA2, Anopheles gambiae (XP_320596); AmCDA2, Apis mellifera (XP_623723); TcCDA2, Tribolium castaneum (ABU25224);
NvCDA2, Nasonia vitripennis (XP_001604838); BmCDA2, Bombyx mori (BGIBMGA006214); DmCDA3, Drosophila melanogaster
(NP_609806); AgCDA3, Anopheles gambiae (XP_317336); BmCDA3, Bombyx mori (BGIBMGA008988); NvCDA3, Nasonia
vitripennis (XP_001606617); AmCDA3, Apis mellifera (XP_001121246); TcCDA3, Tribolium castaneum (ABW74145); TcCDA4,
Tribolium castaneum (ABW74146); AgCDA4, Anopheles gambiae (XP_310753); DmCDA4, Drosophila melanogaster
(NP_728468); BmCDA4, Bombyx mori (BGIBMGA010573); AmCDA4, Apis mellifera (XP_001120478); NvCDA4, Nasonia
vitripennis (XP_001607989); AmCDA5, Apis mellifera (XP_624655); NvCDA5, Nasonia vitripennis (XP_001603918); TcCDA5,
Tribolium castaneum (ABW74147); BmCDA5, Bombyx mori (BGIBMGA002696); AgCDA5, Anopheles gambiae (XP_316929);
DmCDA5, Drosophila melanogaster (NP_001097044); TcCDA6, Tribolium castaneum (ABW74149); TcCDA7, Tribolium
castaneum (ABW74150); TcCDA8, Tribolium castaneum (ABW74151); TcCDA9, Tribolium castaneum (ABW74152); DmCDA9,
Drosophila melanogaster (NP_611192); TnCDA9, Trichoplusia ni (AAY46199); BmCDA9-3, Bombyx mori (BGIBMGA013758);
HaCDA5a, Helicoverpa armigera (ADB43611); HaCDA5b, Helicoverpa armigera (ADB43612); BmCDA9-1, Bombyx mori
(BGIBMGA013756); BmCDA9-2, Bombyx mori (BGIBMGA013757).
216 7: Chitin Metabolism in Insects
Group I CDAs (CDA1s and CDA2s) consist of
D. melanogaster Serpentine (DmSerp) and Vermiform
(DmVerm) (referred to as DmCDA1 and DmCDA2,
respectively) and their orthologs (CDAs 1 and 2) from
each species. All group I CDAs have a chitin-binding
peritrophin-A domain (CBD), a low-density lipoprotein
receptor class A domain (LDLa), and a CDA catalytic
domain. There are two to four transcript variants produced by alternative splicing and/or exon skipping from
the CDA2 pre-mRNAs (Dixit et al., 2008). Group II, III,
and IV families are represented by only one CDA in each
species, namely CDA3, CDA4, and CDA5, respectively.
Although, like group I CDAs, CDA3s also possess a single copy of each of the three domains, the overall amino
acid sequence identity is only about 38% with CDA1s
and CDA2s (amino acid sequence identity between
CDA1s and CDA2s is about 60%). Group III enzymes
(CDA4s) have a single copy of the CBD and the CDA
catalytic domain, but lack an LDLa domain. Group IV
CDAs (CDA5s), like CDA4s, each possess a single CBD
and a single CDA catalytic domain. These two domains,
however, are connected by a long Ser/Thr/Pro/Gln-rich
linker (e.g., about 2400 amino acids in AgCDA5), which
results in CDA5s being the largest CDA proteins. At least
three insect species, D. melanogaster, A. mellifera, and
T. castaneum, have more than one isoform of CDA5 due to
alternative splicing and/or exon skipping during the processing of pre-mRNA for these genes. Group V consists of
two subgroups. One subgroup includes the CDA9s. Two
CDAs (HaCDA5a and HaCDA5b), identified recently by
proteomic analysis and EST sequence analysis of the PM
of the cotton bollworm Helicoverpa armigera (Campbell
et al., 2008; Jakubowska et al., 2010), also belong to this
CDA9 subgroup of group V (Figure 9). Interestingly, two
lepidopterans, B. mori and H. armigera, appear to have
multiple genes related to CDA9. The other subgroup
of group V consists of paralogs from T. castaneum only
(TcCDAs 6, 7, and 8), and not from other insect species.
All the proteins belonging to this group have only a CDA
catalytic domain, and no CBD or LDLa domains.
7.4.3.2. Functional analysis of insect chitin deacety­
lases Developmental patterns and tissue-specific expres­
sion of different CDA genes in the same species suggest that
the chitin deacetylases may have specific functions. In D.
melanogaster, the two group I genes, DmSerp (DmCDA1)
and DmVerm (DmCDA2), are required for normal
tracheal tube development and morphology (Lusch­ing
et al., 2006; Wang et al., 2006). D. melanogaster mutants
lacking either serp or verm exhibited excessively long
and tortuous embryonic tracheal tubes. In T. castaneum,
injection of dsRNA for TcCDA1 or TcCDA2, which
are predominantly expressed in epidermis and tracheae,
prevented all types of molts, including larval–larval,
larval–pupal, and pupal–adult (Figure 5; Arakane et al.,
2009). Furthermore, alternative exon-specific RNAi for
TcCDA2 (TcCDA2a and TcCDA2b) revealed functional
specialization of the isoforms for this CDA. Unlike exon
non-specific RNAi for TcCDA2, injection of dsRNAs
specific for either one of alternative exons did not prevent
any molts, suggesting that the proteins TcCDA2a and
TcCDA2b could compensate for each other. However, the
resulting adults exhibited different abnormal phenotypes.
RNAi for TcCDA2a affected only femoral–tibial joint
movement, while dsRNA for TcCDA2b resulted in elytra
with crinkled and rough dorsal surfaces (Arakane et al.,
2009). These results suggest that group I CDAs play
critical roles in maintaining the structural integrity of the
cuticlular chitin laminae and chitin fibers of the tracheal
tube. It is possible that there are unique cuticular proteins
that preferentially bind to deacetylated portions of chitin,
whereas others preferentially bind to fully acetylated
chitin. These proteins may help to organize the chitinous
cuticular layers and provide the proper rigidity and/or
flexibility in different regions of the cuticle.
Injection of a mixture of dsRNAs for T. castaneum
group V CDAs, TcCDAs 6, 7, 8, and 9, which are all predominantly expressed in the gut, significantly reduced the
transcript levels of individual CDAs. However, no adverse
effects on the appearance, behavior, or survival of these
dsRNA-treated insects were observed (Arakane et al.,
2009). Interestingly, Jakubowska et al. (2010) observed
that one of the group V (CDA9 subgroup) CDA genes
from H. armigera (HaCDA5a) was downregulated by baculovirus infection in larvae. Like TnCDA9, HaCDA5a
had a strong binding affinity for chitin, although it lacks
any predicted chitin-binding domain. Incubation of the
PM from S. frugiperda with recombinant HaCDA5a
increased PM permeability in a concentration-dependent
manner. Infection of insects with a recombinant baculovirus carrying this gene significantly increased the speed of
kill for S. frugiperda and S. exigua. Together, these observations indicate that the group V CDA, HaCDA5a, may
have a role in determining PM structure/morphology or
permeability. For instance, downregulation of transcripts
for this gene after pathogen attack resulted in reduced PM
permeability, presumably to avoid pathogen infection.
Additional studies in the future may reveal the physiological functions of the many CDAs belonging to groups II,
III, and IV.
7.5. Chitin-Binding Proteins
Chitin is almost always found in association with numerous proteins that influence the overall mechanical and
physicochemical properties of the chitin–protein matrix,
which can range from very rigid (e.g., head capsule and
mouth parts) to fully flexible (e.g., larval body and wing
cuticle). Since chitin is an extracellular matrix polysaccharide, the proteins that have an affinity for chitin are
7: Chitin Metabolism in Insects 217
expected to be extracellularly secreted proteins. This is
generally true, with the constraint that some CBPs can be
in vesicles or storage granules between the time they are
synthesized and when they are secreted or released into
the extracellular space by exocytosis.
There are three broad groups of insect proteins containing sequence motifs that have been associated with
chitin-binding ability. The first group consists of a very
large number of insect cuticular proteins, belonging to the
CPR family, containing a consensus sequence(s) known as
the extended Rebers & Riddiford Consensus (R&R Consensus) of a stretch of about 70 amino acids that defines
pfam 00379 (Willis, 2010; see also Chapter 5). The second
group of proteins contains an amino acid sequence motif
known as the “peritrophin A” motif (Tellam et al., 1999).
To avoid confusion about its biological role(s), this motif
will be referred to as the ChtBD2 domain in this chapter, because it is found not only in the group of proteins
extracted from the peritrophic matrix, but also in proteins
extracted from (or expressed in) cuticle-forming tissues.
Proteins with the ChtBD2 motif are further subdivided
intro three groups: peritrophic matrix proteins (with 1–19
ChtBD2 domains, determined to date); cuticular proteins
analogous to peritrophins-3 (with 3 ChtBD2 domains);
and cuticular proteins analogous to peritrophins-1 (with
1 ChtBD2 domain) (Jasrapuria et al., 2010). This domain
consists of a linear sequence of about 60 amino acids with
6 cysteines and conserved spacings between successive
cysteine residues. The ChtBD2 domain defines family
14 of carbohydrate-binding proteins with chitin-binding
ability (CBM14; pfam01607; SMART 00494). The second group also includes enzymes of chitin metabolism
(chitinases, chitin deacetylases, and a protease) that have
one or more ChtBD2 domains in addition to their catalytic domains. The third group of chitin-binding proteins
consists of the family of antimicrobial peptides related
to tachystatins from horseshoe crab (denoted as A1, A2,
B1, B2, and C subfamilies), as well as the calcium channel antagonists, agatoxins from spider venom. Tachystatins are expressed in hemocytes, where they are stored
in the form of small granules and are released into the
hemolymph upon an immune stimulus. This group of
proteins with six cysteines and a high affinity for chitin
has a triple-stranded β-sheet structure with an inhibitory
cysteine knot motif (Fujitani et al., 2007). This structure
is quite different from the peritrophin A motif and tachystatin (see below), and belongs to pfam 11478. They are
not associated with cuticle or the PM, but they do play a
major role in immune defense against bacteria, fungi, and
other pathogens.
Representative members of each of the three groups of
chitin-binding proteins have been extracted from the cuticle or the PM, or isolated from hemocytes. They have also
been expressed in bacterial or other hosts, and some of the
purified proteins have been shown to have chitin-binding
ability. Several proteins belonging to the first and second
groups of chitin-binding proteins are only predicted from
known cDNA or genomic sequences and have not been
biochemically characterized, largely as a result of difficulties associated with extracting them from highly sclerotized cuticular preparations or exuviae. The following
sections will focus on the proteins of the second group of
proteins with ChtBD2 motifs, and also include a limited
discussion of group 3 chitin-binding proteins. A discussion on the first group of cuticular proteins with the R&R
or other consensus motifs is kept to a minimum, because
it is the subject of Chapter 5 in this book (Willis, 2010).
7.5.1. Chitin-Binding Proteins with the R&R
Consensus
The CPR family of cuticular proteins is generally rich in
histidines and devoid of cysteines. The absence of cysteines has been regarded as a defining characteristic of this
group of proteins, with rare exceptions. The number of
cuticular proteins belonging to the CPR subfamily in different insects varies widely, ranging from 32 in A. mellifera to >150 in A. gambiae (see Chapter 5), indicating a
genus-specific expansion of specific families of cuticular
proteins. Among the many families of cuticular proteins
in insects, only some members of the CPR family with the
R&R Consensus have been unequivocally shown to bind
to chitin (summarized in Chapter 5). A member each of
the Tweedle family from B. mori (Tang et al., 2010) and
one protein of the CPAP family (see below) have also been
shown to possess chitin-binding ability. Modeling studies using the 65-aa long R&R Consensus have led to the
notion that this region assumes a half-barrel structure into
which a liner chain of N-acetylglucosamines can be fitted
using van der Waals interactions between the sugar oligomer and the hydrophobic rings of conserved aromatic
amino acids in this consensus (Iconomidou et al., 2005).
In an interesting study, Rebers and Willis (2001) demonstrated that the addition of this consensus sequence alone
to glutathione-S-transferase resulted in acquisition of an
affinity for chitin by this chimeric protein.
7.5.2. Peritrophic Matrix Proteins
The second group of proteins with the ChtBD2 motif
is the family of proteins known as “peritrophins” that
can be extracted from the PM using strong denaturing/
chaotropic reagents, such as 6-M urea or 6-M guanidine
hydrochloride (Tellam et al., 1999). The extracted PMPs
or recombinantly expressed PMPs have chitin-binding
activity (Elvin et al., 1996; Wijffels et al., 2001; Wang
et al., 2004). This motif was shown to be responsible for
binding to chitin by expressing a single ChtBD2 domain
of Trichoplusia ni peritrophin, CBP1, in an insect cell
line, and demonstrating its chitin-binding ability (Wang
218 7: Chitin Metabolism in Insects
et al., 2004). Proteins with multiple ChtBD2 domains are
commonly found strongly associated with the PM. Not
all of them are actually extractable, even with strong chaotropic agents. Some require extraction with strong organic
solvents, such as anhydrous trifluoromethanesulfonic
acid, which also deglycosylates O-linked glycoproteins
(Campbell et al., 2008).
The number of ChtBD2 domains in insect PMPs varies
from 1 to as many as 19 in the bertha armyworm Mamestra configurata (Shi et al., 2004; Dinglasan et al., 2009;
Venancio et al., 2009; Jasrapuria et al., 2010; Toprak et al.,
2010). Some PMPs have multiple ChtBD2 repeats in a
tandem arrangement with short spacers rich in P, S, and
T residues. Some of these linkers are potential sites of
O-glycosylation. Other PMPs have mucin domains interspersed between ChtBD2 domains in various patterns of
alternating ChtBD2 and mucin domains (Wang et al.,
2004; Venancio et al., 2009). PMPs with only one or two
ChtBD2 domains have also been reported ­( Jasrapuria
et al., 2010; Toprak et al., 2010). The number of PMPs
in different species is variable. Both Ae. aegypti and D.
melanogaster have been predicted to have about 65 PMPs,
though many of these may not be components of the
PM (Venancio et al., 2009). Detailed expression studies
of all proteins with ChtBD2 domains in T. castnaeum
have demonstrated that there are only 11 bona fide PMPs
in this beetle (Jasrapuria et al., 2010). Direct proteomic
analysis of >200 proteins extracted from PMs dissected
from adult A. gambiae females fed a protein-free diet has
revealed the presence of only 12 PMPs, with the number
of ChtBD2 repeats ranging from 1 to 4. It is likely that
the total number of PMPs in insects is in the range of
10–20, although it can’t be ruled out that additional PMP
genes are expressed in the gut. However, their conceptual
protein products were not detected in proteomic analyses
because they were still in the insoluble pellet after extraction with detergents used in an extensive study (Dinglasan
et al., 2009). Interestingly, different PMP genes of T. castaneum were not expressed uniformly through the length
of the midgut, with some PMPs being expressed in the
anterior midgut, whereas others coding for proteins with
multiple ChtBD2 domains were expressed in the posterior midgut (Jasrapuria et al., 2010). Whether this differential spatial expression results in altered permeability
of the PM along the length of the midgut remains to be
investigated.
7.5.3. Cuticular Proteins Analogous
to Peritrophins (CPAPs)
In addition to the PMP genes, which are expressed exclusively in the midgut lining cells, there are other genes
encoding proteins with ChtBD2 domains, which are
expressed in tissues other than the midgut. All of these
proteins are predicted to have a cleavable signal peptide,
and are expected to be capable of interacting with extracellular chitin. These genes are expressed predominantly
in epidermal tissue as well as in other cuticle-forming
tissues, including tracheae, elytra, hindwings, and hindgut. These genes have been subdivided into two groups,
CPAP1 and CPAP3, to reflect the fact that they encode
proteins with one or three ChtBD2 domains, respectively
(Jasrapuria et al., 2010). CPAP3 is the new name given to
the orthologs of the previously characterized D. melanogaster “obstructor” or “gasp” gene family.
Mutants of the D. melanogaster CPAP3-C gene are
embryo-lethal, and have been reported to exhibit cuticular defects (Barry et al., 1999; Behr and Hoch, 2005). In
D. melanogaster there are 10 genes encoding CPAP3 proteins, which can be further subdivided into two groups of
5 genes each. Only orthologs for the first group (CPAP3A, CPAP3-B, CPAP3-C, CPAP3-D, and CPAP-E) are
present in insects other than Drosophila species. There are
significant variations in the expression profiles of these
genes in different cuticle-forming tissues and/or developmental stages, suggesting functional differences among
the CPAP3 proteins. RNA interference studies carried
out in T. castaneum are consistent with such specialized
functions of individual CPAP3 proteins (Jasrapuria et al.,
unpublished data).
While it is expected that the CPAP3 proteins with
three ChtBD2s will bind to chitin strongly, this has been
demonstrated for only one recombinant protein from the
spruce budworm Choristoneura fumiferana, which was
expressed in E. coli (Nisole et al., 2010). However, only
a minor percentage of the His-tagged protein bound to
the chitin, with the major portion appearing in the flowthrough fraction, perhaps indicating that not all molecules of this recombinant protein had folded properly to
exhibit strong chitin-binding activity. So far, there is no
report of expression of this class of proteins in an insect
cell system that may overcome the problem of misfolding
as demonstrated for two PMP proteins with 10 and 12
repeats of ChtBD2 domains (Wang et al., 2004).
A second group of genes encoding proteins with one
ChtBD2 domain, referred to as the CPAP1 family proteins, has been characterized extensively using a bioinformatics analysis of the T. castaneum genome (Jasrapuria
et al., 2010). These proteins vary extensively in size, and
in the location of the ChtBD2 domain. Like CPAP3,
they are also expressed in cuticle-forming tissues and
have putative cleavable signal sequences consistent with
a role involving interactions with chitin. So far, there are
no reports on the chitin-binding ability of these proteins.
Only some of these proteins have orthologs in D. melanogaster, casting doubt on whether these proteins are ubiquitous in insects. However, RNAi studies have produced
lethal phenotypes when transcripts for 3 of the 10 genes
encoding CPAP1 proteins were depleted in T. castaneum
(Jasrapuria, unpublished data).
7: Chitin Metabolism in Insects 219
7.5.4. Enzymes of Chitin Metabolism
Enzymes of chitin metabolism, including some members
of the chitinase and chitin deacetylase families, have the
ChtBD2 motif (Kramer et al., 1993; Campbell et al.,
2008; Dixit et al., 2008; Zhu et al., 2008a). The presence
of one or more copies of this ChtBD2 motif has been suggested to increase the affinity of enzymes of chitin metabolism for the insoluble substrate, chitin, and to increase
the processivity of these extracellular enzymes. Support
for this idea comes from the drastic loss of ability to bind
to insoluble chitin upon removal of the region containing the ChtBD2 motif from the C-terminal region of an
M. sexta chitinase, which follows the catalytic domain.
A C-terminal fragment of only 58 amino acids with this
domain did bind to colloidal chitin, and addition of one
or two copies of this domain to the chitinase catalytic
domain progressively increased the affinity of the chitinase to colloidal chitin (Arakane et al., 2003). While most
of these enzymes have only one ChtBD2 motif, one class
of insect chitinases (group II) has four or five ChtBD2
motifs dispersed among multiple catalytic domains (Royer
et al., 2002; Zhu et al. 2008a). A role for these multiple
ChtBD2s in facilitating the depolymerizing chitin crystallites has been suggested (Arakane and Muthukrishnan,
2010).
7.5.5. Role of Secondary Structure of ChtBD2
Motif in Binding to Chitin
Tertiary structures based on 2D-NMR studies in solution
are available for only two insect proteins with ChtBD2
domains with high affinity for chitin; namely, tachycitin and scarabacin. The antimicrobial peptide tachycitin
from the horseshoe crab, which has a structure different from the tachystatins, is 76 aa long, and has a higher
Km for chitin binding than tachystatin – 19.5 μM versus
4.3 μM, respectively (Kawabata et al., 2003). The second chitin-binding antimicrobial peptide for which an
NMR-deduced structure is available is scarabacin from
the coconut rhinocerous beetle Oryctes rhinoceros, which
is 36 aa long and has a Kd = 1.3 μM (Hemmi et al., 2003).
A comparison of these two structures with that of another
chitin-binding minimal fragment called hevein-32, from
the rubber latex protein hevein, has provided some interesting insights about the role of a part of the ChtBD2
motif in chitin binding.
Tachycitin has 10 cysteines in the form of 5 disulfide
bonds, and has significant similarity to several peritrophins from a wide spectrum of insects, including PMP3
of T. castaneum, with which it shares 51% amino acid
sequence identity. Of these 10 cysteines, 5 are in perfect
register with the linear arrangement of the cysteines in
PMP3 ChtBD2 domains, without introducing gaps in
either sequence, except for the first cysteine in the motif.
More importantly, the amino acid sequence from positions 40 to 60 of tachycitin, which includes one disulfide bond, shares significant similarities to those of several
other peritrophins from a wide range of insect species
(Suetake et al., 2000). Furthermore, the three-dimensional structure of this stretch of 21 amino acids is nearly
identical to that of a hevein-32 from positions 20 to 32
(Aboitiz et al., 2004). Both proteins have two anti-parallel
β-sheets followed by a short α-helix in this region, which
also includes a disulfide bond. Several aromatic amino
acids that have been shown to contact the oligosaccharide ligands (GlcNAc)3–6 are also conserved in the two
sequences.
The 3D structure of scarabacin reveals the presence of
only one disulfide bond between cys18 and cys29. The
C-terminal half of this peptide from cys18 to ser 36 also
has a secondary structure consisting of two anti-parallel
β-sheets and a short α-helical turn super-imposable on
hevein-32 or tachycitin (Hemmi et al., 2003). These data
suggest that only the C-terminal half of the ChtBD2
domain may be critical for chitin binding. Consistent
with this interpretation is the finding that the N-terminal
half of tachycitin has a completely different 3D structure,
consisting of a three-stranded β-sheet while retaining the
hevein/scarabacin-like chitin-binding motif on the C-terminal domain (Suetake et al., 2000; Hemmi et al., 2003).
These data suggest that all three chitin-binding proteins
(hevein, scarabacin, and tachycitin) share a common
chitin-binding secondary/tertiary structure, even though
they do not have extensive amino acid sequence identity.
By extrapolation, we expect that all of the proteins with
ChtBD2 domains will also have this structural motif consisting of two anti-parallel β-sheets and a short α-helical
turn.
A protein from the vestimentiferan Riftia pachyptila has
been shown to bind specifically to β-chitin, but not to
α-chitin or cellulose (Chamoy et al., 2001). The sequence
of this protein includes a cysteine-rich region that resembles the C-terminal region of many mammalian chitinases, and is likely to be a chitin-biding motif. However, it
does not have the consensus sequence or the characteristic
spacing between adjacent cysteines of the ChtBD2 motif,
and may represent yet another type of chitin-binding
domain.
7.6. Chitin-Organizing Proteins
In addition to the CPR proteins with the R&R Consensus
and the CPAP proteins with the ChtBD2 motif, which
are expected to interact with chitin, some additional proteins may be associated with chitin, and help to organize
it into bundles and the laminae that are characteristic
of a mature procuticle. Two proteins encoded by Knickkopf (Knk) and Retroactive (Rtv) genes are known to be
involved in this process in D. melanogaster (Ostrowski
220 7: Chitin Metabolism in Insects
et al., 2002; Moussian et al., 2005b; Tonning et al., 2005).
Mutations in these two genes result in a dilated cuticle
and loss of the fibrillar organization of tracheal chitin, and
death of the developing embryo. Transmission electron
microscopic analyses of the developing embryonic cuticle
in these mutants revealed loss of the laminar architecture
of chitin and the accumulation of electron-dense material
in the procuticle.
How do these two proteins function to organize the
cuticle-associated chitin? The domain organization of these
proteins and their predicted properties offer some hints.
The 75-kDa KNK is a GPI-anchored membrane protein
with a multidomain architecture consisting of two DM13
domains, a dopamine monooxygenase N-terminal domain
(DOMON domain), and a unique C-terminal region
that has not been associated with any well-­characterized
domain. However, this sequence has some similarities to
plastocyanin (Moussian, 2010). Interestingly, this region
also has some sequence similarity to several plant proteins
that possess DM13 and DOMON domains as well as a
cytochrome b561 domain. It is possible that Knickkopf
and its orthologs are extracellular proteins that may have
a role in oxidation–reduction reactions perhaps involving
dopamine. T. castaneum KNK expressed in a baculovirusinsect cell expression system does bind to colloidal chitin
(Chaudhari, unpublished data). RNAi of this KNK gene
results in loss of chitin, and this loss appears to be due to
the protective effect of KNK on chitin against degradation
by chitinolytic enzymes. The distribution of this protein
between the procuticle and plasma membrane is consistent with such a chitin-protective role.
RTV is also a membrane-bound protein with a single
C-terminal transmembrane domain, which localizes this
protein to the apical surface of the plasma membrane
(Schwarz and Moussian, 2007). RTV mutants have a
spindle-shaped body, and often the cuticle separates from
the epidermal layer underneath. This protein, which is
about 150 amino acids long, has 10 cysteines, belongs to
the neurotoxin-like SCOP superfamily of proteins, and
has a β-sandwich structure with 2 and 3 β-strands in the
2 β−sheets. Its ability to bind to chitin has not been demonstrated, but the six aromatic amino acids present in the
loops indicate such a possibility.
7.7. Hormonal Regulation of Chitin
Metabolism
Chitinolytic activity in the molting fluid rises just prior to
each molt and falls shortly thereafter. These changes parallel the increasing and falling ecdysteroid titers prior to
ecdysis, as observed initially by Kimura (1976). A direct
role for ecdysteroids in inducing chitinase expression was
demonstrated using M. sexta larval abdomens that were
precluded from receiving hormonal signals from the brain
by a ligature below the second thoracic segment. Injection
of 20-hydroxyecdysone (20HE) into these ligated abdomens resulted in a sharp and rapid increase in transcripts
for chitinase. This increase was abolished by a simultaneous injection of a juvenile hormone mimic (Fukamizo
& Kramer, 1987). Koga et al. (1992) reported a similar
induction of chitinase by ecdysteroid, utilizing isolated
Bombyx abdomens. Zheng et al. (2003) observed that
injection of an ecdysteroid agonist resulted in induction
of expression of a chitinase gene in epidermal tissue of C.
afumiferana, and demonstrated the accumulation of chitinase in molting fluid. It appears that hormonal regulation of chitinase genes occurs in a broad range of insect
species. However, the presence of multiple genes encoding
chitinases was not appreciated when these early studies
were done, and it was not apparent which class of chitinases was induced by the ecdysteroid treatment. Based
on our present knowledge about the tissue specificity of
expression of different groups of chitinases, it is likely that
these early studies were only focused on the expression of
group I chitinases.
A group II chitinase gene with five catalytic domains
from the beetle Tenebrio molitor has also been shown to
be hormonally regulated (Royer et al., 2002). During
pupal–adult metamorphosis, the abundance of transcripts
for this gene paralleled the changes in ecdysteroid (20HE)
titers during metamorphosis. Interestingly, even topical
application of the JH analog, methoprene, induced transcripts for this chitinase within 8 hours after treatment.
These results are somewhat contradictory to the studies
on M. sexta chitinase, in which JH had no inductive effect
on chitinase transcript levels (Kramer et al., 1993). In B.
mori, another chitinase gene, BmChiR1, required 20HE
for induction, and was suppressed by the simultaneous
application of a JH analog (Takahashi et al., 2002). Even
though this chitinase was reported to have only two inactive catalytic domains and one CBD, our bioinformatics
analysis (Merzendorfer, unpublished data) indicated that
this gene actually encodes a protein with five catalytic
domains and seven CBDs, and appears to be a group II
chitinase. A recent study on the regulation of chitinase
gene expression in a shrimp species demonstrated that it
is induced by ecdysteroids. Hence, ecdysteroids may be
required for induction of chitinases in most arthropods
(Priya et al., 2009). While it is clear that the expression of
more than one chitinase gene is controlled by ecdysteroid
and possibly by JH, it is likely that these effects are mediated through one or more transcription factors induced
by ecdysteroids (Riddiford et al., 2003). However, there
are no published reports on the identification of hormone
response elements in the promoters of any of the insect
chitinase genes.
There is little evidence to support the idea that hormones play a direct role in the control of chitin synthesis.
Instead, chitin synthesis is initiated at about the time of
(or prior to) apolysis, when new cuticle is being deposited.
7: Chitin Metabolism in Insects 221
In general, chitin synthesis reaches peak levels in between
molts when new cuticle is being synthesized at the maximal rate. In larval stages, this is also the period when
PM-associated chitin is synthesized. Thus, both CHS-A
and CHS-B levels are high during feeding periods in
larval stages. In the pupal stage CHS-B transcripts are
undetectable, whereas levels of transcripts for CHS-A
have multiple peaks roughly corresponding to periods of
synthesis of pupal cuticle, adult epidermal cuticle, and
tracheal chitin (Hogenkamp et al., 2005; Arakane et al.,
2008).
7.8. Chitin Metabolism and Insect
Control
7.8.1. Inhibition of Chitin Synthesis
The absence of chitin in animals and plants has led to
the development of insect control strategies that target
enzymes involved in the synthesis, modification, and
degradation of chitin. Several membrane proteins that
are likely to be involved in the assembly of chitin in the
procuticle, or regulation of chitin metabolism, may also
be attractive targets. Compounds that directly or indirectly interfere with chitin biosynthesis include peptidyl
nucleosides, acylureas, thiadiazines, and different kinds of
chitin-binding molecules. The peptidyl nucleosides were
isolated originally from different Streptomyces species, and
include polyoxins and nikkomycins (Hori et al., 1971;
Dahn et al., 1976). They are substrate analogs resembling
the structure of UDP-GlcNAc, and competitively inhibit
chitin synthases of fungal and insect sources, with nikkomycin being the most potent inhibitor (Cohen, 2001). As
peptidyl nucleosides that exhibit low permeability across
the hydrophobic epicuticle are easily degraded in the
intestine and show toxic side effects in vertebrates, they
have not been developed further to control insect pests,
but some of them are in use as fungicides in agriculture
(Zhang and Miller, 1999; Cohen, 2001; Ruiz-Herrera
and San-Blas, 2003). In contrast, since the discovery of
the high insecticidal potential of diflubenzuron in the
early 1970s by Dutch scientists, various acylurea derivatives, such as lufenuron, novaluron, and hexaflumuron,
have been developed commercially for controlling agricultural pests (Palli and Retnakaran, 1999). They have
been shown to inhibit chitin synthesis and to disturb
cuticle formation, causing abortive molting. Ultrastructural analysis revealed defects in chitin synthesis, abnormal deposition of endocuticular layers, and impaired PM
formation. Studies with these “chitin synthesis inhibitors”
have provided some insights concerning the role of chitin
in development, and its biological function. In particular, the use of the acylurea compound lufenuron has provided substantial information on chitin synthesis during
Drosophila development (Wilson and Cryan, 1997). The
effects of this insect growth regulator were complex and
variable, depending on the developmental stage and dose
at which the insects were exposed to this agent. When
newly hatched larvae were reared on a diet containing very
low concentrations of lufenuron, the larvae did not die
until the second or third instar, and some pupariated even
though the pupae were abnormally compressed. Pharate
adults either failed to eclose or died shortly after emergence, and had deformed legs. The flight ability of the
emerged adults was also affected when the larvae were
exposed to very low concentrations of lufenuron. First
and second instar larvae fed higher concentrations of
lufenuron had normal growth and physical activity for
several hours, but the insects died at about the time of the
next ecdysis. Third instar larvae fed high concentrations
of lufenuron underwent pupariation, but the puparia had
an abnormal appearance, and the anterior spiracles failed
to evert. Strikingly, adults showed no mortality and had
no flight disability even when fed high levels of lufenuron,
indicating that once all chitin-containing structures had
been formed, this “chitin inhibitor” had very little effect
on morphology and function. Thus, insect development
is affected by lufenuron at all stages when chitin synthesis
occurs. Another phase of insect development affected by
this compound was egg hatching, even though oviposition was normal. The embryos completed development,
but failed to rupture the vitelline membrane. In an ultrastructural study of acylurea effects on Drosophila embryogenesis, Gangishetti and colleagues have shown recently
that egg hatching is completely abolished after treating
female flies with a high dose of lufenuron and mating
them with untreated males (Gangishetti et al., 2009). In
line with its lower insecticidal activity, the same treatment
performed with diflubenzuron resulted in a constant rate
of larval survival. Overall, the hatching rates depended
on the dosage of the insecticides. The embryonic phenotypes were grouped into five classes: (1) hatching wild
type larvae; (2) non-hatching larvae that appeared slightly
bloated after being released manually from the eggshells;
(3) non-hatching larvae with a strongly melanized head
skeleton and a cuticle detached from the epidermis, which
is similar to knk and rtv phenotypes (see section 7.6); (4)
non-hatching larvae with a crumbled head skeleton and
detached cuticle, which is similar to the kkv phenotype
(see section 7.3.4.3. and Figure 5); and (5) non-hatching larvae with strong segmentation and morphological
defects. The latter phenotypes were indistinguishable from
the effects of the nucleoside peptide antibiotic nikkomycin, which is a competitive inhibitor of chitin synthase.
Electron microscopy revealed that the treatment with
lower doses of the insecticides affected cuticle thickness
and orientation of microfibrils, while higher doses disrupted chitin synthesis completely, as evidenced by the
lack of Calcofluor white fluorescence in the cuticle (Gangishetti et al., 2009). Interestingly, no changes in kkv and
222 7: Chitin Metabolism in Insects
mummy gene expression were observed, but the expression of certain genes encoding cytochrome P450 enzymes
was substantially upregulated, indicating that the respective enzymes are involved in diflubenzuron and lufenuron
detoxification. Similar results were also observed in Tribolium, where diflubenzuron fed to larvae did not significantly influence TcCHS1 or TcCHS2 expression, but did
affect mRNA levels for certain cytochrome P450 enzymes
(merzendorfer, unpublished data). In contrast to Drosophila and Tribolium, RT-PCR and Northern blot analyses
carried out with A. quadrimaculatus revealed a two-fold
upregulation of AqCHS1 mRNA levels in response to a
high dose of diflubenzuron, while the chitin content in
surviving larvae decreased in a dose-dependent manner (Zhang and Zhu, 2006). The observed increase in
AqCHS1 mRNA levels associated with a decrease in chitin content corroborates the common view that acylurea
insecticides affect chitin synthesis at a post-transcriptional
level. Hence, diflubenzuron-induced AqCHS1 expression may serve as a mechanism to compensate for chitin
deficiency.
Several studies have aimed to elucidate the underlying
mechanism of the insecticidal activity of diflubenzuron.
Diflubenzuron efficiently blocks chitin synthesis, as the
incorporation of radiolabeled sugars into the growing
chitin chain is inhibited (Post and Vincent, 1973; Hajjar and Casida, 1978; Mayer et al., 1980; Clarke and
­Jewess, 1990). However, in contrast to peptidyl nucleosides that block chitin polymerization, diflubenzuron
obviously does not affect the catalytic step, because chitin
synthesis is not impaired in cell-free systems (Cohen and
Casida 1980a; Mayer et al., 1980; Kitahara et al., 1983;
Zimoch et al., 2005). It also does not interfere with any
of the metabolic reactions yielding UDP-N-acetylglucosamine, and neither does it affect chitin synthesis in
fungi (­Verloop and Ferrel, 1977; Cohen, 1987). Based on
these and other findings, it was suggested that diflubenzuron acts at a post-­catalytic step of chitin synthesis
(Cohen, 2001). Many other mechanisms for the action of
diflubenzuron have been suggested, including effects on
glycolytic enzymes, chitinases, phenoloxidases, hormonal
sites, and microsomal oxidases (Ishaaya and Cohen, 1974;
Ishaaya and Ascher, 1977; Mitlin et al., 1977; DeLoach
et al., 1981; Soltani, 1984). Studies using imaginal discs
and cell-free systems indicated that benzoylphenylureas
inhibit ecdysteroid-dependent GlcNAc incorporation
into chitin (Mikolajczyk et al., 1994; Oberlander and Silhacek, 1998). These results indicated that acylurea compounds target ecdysone-dependent sites, which eventually
leads to inhibition of chitin formation. However, direct
proof for this hypothesis is lacking. On the basis of competitive binding assays performed with glibenclamide, a
more recent study suggested that a sulfonylurea receptor
might be the target for diflubenzuron (Abo-Elghar et al.,
2004). As the sulfonylurea receptors (SURs) may also
act as regulatory subunits of inward rectifying potassium
channels in insects (Akasaka et al., 2006), inhibition of a
SUR could alter the membrane potential in such a way
that Ca2+ homeostasis and eventually protein secretion
required for cuticle and PM formation is affected. In line
with this assumption, glibenclamide as well as diflubenzuron were found to affect Ca2+ uptake by isolated cuticular vesicles from the German cockroach Blatella germanica
(Abo-Elghar et al., 2004). Although the significance of
this finding remains uncertain, future research following up on this hypothesis may elucidate the target site of
acylureas.
Another chemical group of “chitin synthesis inhibitors” comprises thiadiazine derivatives, such as buprofezin
(Applaud), which is used as an insecticide that specifically
acts on sucking insects such as homopterans and hemipterans (Kanno, 1981). Although quite different in chemical structure, the effect of buprofezin resembles that of
acylureas, as it blocks incorporation of radiolabeled chitin
precursors and interferes with insect development. However, buprofezin may have a different target site in insects,
as it also blocks acetylcholinesterase (AChE) activity. The
activity of AChE in crude homogenates from the whitefly
Bemisia tabaci was significantly inhibited by buprofezin at
a concentration of 0.5 μM (Cottage and Gunning, 2006).
Strikingly, inhibition was not observed in buprofezinresistant flies.
Chitin-binding molecules interfere with the microfibril
assembly, and hence block chitin deposition at its final
step. There are polysaccharide-binding dyes, such as Calcofluor White (CFW), Congo red or primuline, which
interfere with chitin crystallization by disrupting hydrogen
bond formation and hence perturbing microfibril assembly (Vermeulen and Wessels, 1986). Accordingly, these
dyes were reported to impair fungal cell wall morphogenesis (Selitrennikoff, 1984; Roncero and Duran, 1985). In
insects, the process of PM formation appears to be particularly susceptible to CFW, and its effects were studied
in flies, mosquitoes and caterpillars. Injection of as little as
0.05 μg CFW into Calliphora erythrocephala flies led to perturbations of PM formation and increased permeabilities
for FITC-labeled dextrans with molecular masses ranging
between 17 and 32 kDa (Zimmermann and Peters, 1987).
However, in contrast to other PM-disrupting agents such
as dithiothreitol or chitinase, changes in PM permeabilities
for FITC-labeled dextrans with a molecular mass of 2 MDa
were not observed when mosquito larvae were treated with
CFW or Congo red (Edwards and Jacobs-Lorena, 2000).
In L. cuprina the PM structure was not affected, although
the larvae showed growth retardation and a reduction in
lifespan (Tellam and Eisenmann, 2000). In the mite Acarus
siro, combinations of diflubenzuron and CFW were most
effective in reducing chitin content of the PM (Sobotnik
et al., 2008). Hence, combinations of CFW with other
insecticidal compounds affecting chitin synthesis may
7: Chitin Metabolism in Insects 223
prove to be a useful strategy for insect control. Disruption
of the PM structure was consistently reported in various
lepidopteran species (Wang and Granados, 2000; Bolognesi et al., 2001; Zhu et al., 2007). When larvae of T. ni and
S. exigua were fed with a CFW-containing diet, an increase
in PM permeability was observed and the larvae became
more susceptible to baculoviral infections. Interestingly,
a significant amount of proteins was released upon CFW
treatment, which may explain altered permeabilities
(Wang and Granados, 2000; Zhu et al., 2007). Next to
chitin-binding dyes, numerous sugar-binding proteins
(lectins) from animals and plants such as galectins, WGA,
and chitinase-like lectins bind chitin or chitosan because
of their high preference for GlcNAc. Like CFW, they disrupt PM formation in numerous cases, and therefore have
been investigated for their insecticidal potential (Cohen,
2010). The effects of WGA on PM formation are summarized in section 7.3.1.2. However, these types of proteins
also bind to glycoproteins and proteoglycans present in the
PM, and hence their particular mode of action is difficult
to asseess in vivo.
7.8.2. Exploiting Chitinases for Insect Control
Chitinases have been used in a variety of ways for insect
control and other purposes (Kramer and Muthukrishnan,
1997; Gooday, 1999). Several chitinase inhibitors with
biological activity have been identified based on natural
products chemistry (Spindler and Spindler-Barth, 1999),
such as allosamidin, which mimics the carbohydrate substrate (Rao et al., 2003), and cyclic peptides (Houston
et al., 2002). Although useful for biochemical studies,
none of these chitin catabolic inhibitors have been developed for commercial use, primarily because of the high
cost of production and potential side effects. As we learn
more details about chitinase catalysis, it might become
more economically feasible to develop and optimize chitinase inhibitors for insect pest management.
Fungi and plants use chitinases for establishing infection and as a defense against invading pathogens, respectively. Entomopathogens secrete a plethora of extracellular
proteins with potential activity in insect hosts. One of
these proteins is chitinase, which is used by fungi such as
Metarhizium anisopliae to help penetrate the host cuticle
and render host tissues suitable for consumption (St Leger
et al., 1996; Krieger de Moraes et al., 2003). Among the
10 most frequent transcripts in a strain of M. anisopliae
are 3 encoding chitinases and a chitosanase (Freimoser
et al., 2003a). However, when M. anisopliae was transformed to overexpress its native chitinase, the pathogenicity towards the tobacco hornworm was unaltered,
suggesting that wild type levels of chitinase are not limiting for cuticle penetration (Screen et al., 2001). Another
fungal species, Conidiobolus coronatus, also produces both
endo- and exo-acting chitinolytic enzymes during growth
on insect cuticle (Freimoser et al., 2003b). Apparently,
both M. anisopliae and C. coronatus produce a chitinolytic
enzyme system to degrade cuticular components.
Both microbial and insect chitinases have been shown
to enhance the toxicity of the entomopathogenic bacterium Bacillus thuringiensis (Bt) (Regev et al., 1996; Tantimavanich et al., 1997; Ding et al., 1998; Sampson and
Gooday, 1998; Wiwat et al., 2000). For example, when
the chitinolytic activities of several strains of B. thuringiensis were compared with their insecticidal activity, it was
determined that the enzyme could enhance the toxicity of
Bt to S. exigua larvae by more than two-fold (Liu et al.,
2002). Microbial chitinases have been used in mixing
experiments to increase the potency of entomopathogenic
microorganisms (Kramer and Muthukrishnan 1997).
Synergistic effects between chitinolytic enzymes and
microbial insecticides were reported as early as the 1970s.
Bacterial chitinolytic enzymes were first used to enhance
the activity of Bt and a baculovirus. Larvae of C. fumiferana died more rapidly when exposed to chitinase–Bt
mixtures than when exposed to the enzyme or bacterium
alone (Smirnoff and Valero, 1972; Lysenko, 1976; Morris,
1976). Mortality of gypsy moth, Lymantria dispar, larvae
was enhanced when chitinase was mixed with Bt, relative
to treatment with Bt alone, in laboratory experiments
(Dubois, 1977). The toxic effect was correlated positively
with enzyme levels (Gunner et al., 1985). The larvicidal
activity of a nuclear polyhedrosis virus toward L. dispar
larvae was increased about five-fold when it was administered with a bacterial chitinase (Shapiro et al., 1987).
Inducible chitinolytic enzymes from bacteria cause
insect mortality under certain conditions. These enzymes
may compromise the structural integrity of the PM barrier and improve the effectiveness of a Bt toxin by enhancing contact of the toxin molecules with their epithelial
membrane receptors. For example, five chitinolytic bacterial strains isolated from midguts of Spodoptera littoralis
induced a synergistic increase in larval mortality when
combined with Bt spore-crystal suspensions relative to
either an individual bacterial strain or a Bt suspension
alone (Sneh et al., 1983). An enhanced toxic effect toward
S. littoralis also resulted when a combination of low levels
of a truncated recombinant Bt toxin and a bacterial endochitinase was incorporated into a semisynthetic insect diet
(Regev et al., 1996). Crude chitinase preparations from
B. circulans enhanced the toxicity of Bt kurstaki toward
diamondback moth larvae (Wiwat et al., 1996). Liu et al.
(2002) reported that several strains of Bt produced their
own chitinases, which had synergistic larvicidal activity
with the endotoxins.
A family-18 insect chitinase has been used as an
enhancer of baculovirus toxicity and as a host plant
resistance factor in transgenic plants. Introduction of an
M. sexta chitinase cDNA into Autographa californica
multiple nuclear polyhedrosis viral (AcMNPV) DNA
224 7: Chitin Metabolism in Insects
accelerated the rate of killing of fall armyworm compared
to the wild type virus (Gopalakrishnan et al., 1995). Baculoviral chitinases themselves play a role in liquefaction of
insect hosts (Hawtin et al., 1997; Thomas et al., 2000).
A constitutively expressed exochitinase from B. thuringiensis potentiated the insecticidal effect of the vegetative
insecticidal protein Vip when they were fed to neonate
larvae of S. litura (Arora et al., 2003). Mutagenesis of the
AcMNPV chitinase gene resulted in cessation of liquefaction of infected T. ni larvae, supporting a role of chitinase
in viral spreading (Thomas et al., 2000). When diet containing AcMNPV chitinase expressed in E. coli was fed to
B. mori larvae, a dose-dependent increase in loss of integrity of the PM was observed. Even at a dose of 1 mg/g of
larvae, there was 100% mortality (Rao et al., 2004).
Tobacco budworms were killed when reared on transgenic tobacco expressing a truncated, enzymatically active
form of M. sexta class I chitinase (Ding et al., 1998). A
synergistic interaction between insect chitinase expressed
in transgenic tobacco plants and Bt applied as a spray at
sublethal levels occurred when using the tobacco hornworm as the test insect. In contrast to results obtained
with the tobacco budworm, studies with the hornworm
revealed no consistent differences in larval growth or foliar
damage when the insects were reared on first-generation
transgenic chitinase-positive tobacco plants as compared
to chitinase-negative control plants. When Bt toxin was
applied at levels where no growth inhibition was observed
on control plants, chitinase-positive plants had significantly less foliar damage and lower larval biomass production. These results indicated that the insect chitinase
transgene did potentiate the effect of sublethal doses of
Bt toxin, and vice versa (Ding et al., 1998), but chitinase
was not very effective on its own as a biocontrol agent.
Tomato plants have been transformed with fungal chitinase genes with concomitant enhancements in resistance
to insect pests (Gongora et al., 2001). Effects observed
include reduced growth rates and increased mortality, as
well as a decrease in plant height and flowering time, with
an increase in the number of flowers and fruits
(Gongora and Broadway, 2002). Chitinase-secreting bacteria have been used to suppress herbivorous insect pests.
A strain of Enterobacter cloacae transformed with a chitinase gene digested the chitinous membranes of phytophagous ladybird beetles, Epilachna vigintioctopunctata, and
also suppressed leaf feeding and oviposition when the
beetles ingested transformed bacteria entrapped in alginate microbeads sprayed on tomato seedlings (Otsu et al.,
2003). When pure chitinase from tomato moth larvae was
injected into larvae, decreased cuticle thickness and 100%
mortality was observed even at a low dose (2.5 μg/g).
Insects fed this protein exhibited reductions in growth and
food consumption (Fitches et al., 2004). Acaricidal activity of a purified chitinase from a hard tick, Haemophysalis
longicornis, has also been demonstrated (Assenga et al.,
2006). Immunization with this chitinase as the antigen
protected mice from tick infections (You et al., 2009).
Several GlcNAc-specific lectins from plants have been
evaluated for insect toxicity (Harper et al., 1998; Macedo
et al., 2003). These proteins appear to disrupt the integrity
of the PM by binding to chitin or glycan receptors on
the surface of cells lining the insect gut. Moreover, they
may bind to glycosylated digestive enzymes and inhibit
their activity. Another type of plant chitin-binding protein is the seed storage protein vicilin, which is actually
a family of oligomeric proteins with variable degrees of
glycosylation (Macedo et al., 1993; Shutov et al., 1995).
Some vicilins are insecticidal to bruchid beetles and stalk
borers (Sales et al., 2001; Mota et al., 2003). Apparently,
these proteins bind to the PM, causing developmental
abnormalities and reduced survival rates. To date, no
non-enzymatic carbohydrate-binding protein derived
from an insect has been evaluated for biocidal activity.
A novel approach has been proposed to develop strategies
for insect control by utilizing chitin-binding molecules to
specifically target formation of the PM. CFW, a chemical whitener with chitin-binding properties, was used as
a model compound in the diet to inhibit PM formation
in T. ni, and to increase larval susceptibility to baculovirus infection (Wang and Granados, 2000). It was also
effective in suppressing PM formation in S. frugiperda,
and at the same time in preventing the establishment of a
decreasing gradient of proteinases along the midgut tissue
(Bolognesi et al., 2001).
A protease from A. gambiae with a chitin-binding
domain has been described, which may be involved in
insect defense (Danielli et al., 2000). This 147-kDa protein, sp22D, is expressed in a variety of tissues, most
strongly in hemocytes, and is secreted into the hemolymph. Upon bacterial infection, the transcripts for this
protein increase by about two-fold, suggesting a role in
insect defense. This protein has a multidomain organization that includes two copies of an N-terminal ChtBD2
domain, a C-terminal protease domain, and several receptor domains. It binds strongly to chitin, and undergoes
complex proteolytic processing during pupal to adult
metamorphosis. It has been proposed that exposure of this
protease to chitin may regulate its activity during tissue
remodeling or wounding.
Two synthetic peptides were found to inhibit A. gambiae midgut chitinase, and also to block sporogonic
development of the human malaria parasite Plasmodium falciparum and avian malaria parasite P. gallinaceum, when the peptides were fed to infected mosquitoes
(Bhatnagar et al., 2003). The design of these peptides was
based on the putative proregion sequence of mosquito
midgut chitinase. The results indicated that expression
of chitinase inhibitory peptides in transgenic mosquitoes
might alter the vectorial capacity of mosquitoes to transmit malaria.
7: Chitin Metabolism in Insects 225
7.9. Future Studies and Concluding
Remarks
Although substantial progress in studies of insect chitin
metabolism has occurred since the initial edition of Comprehensive Insect Physiology, Biochemistry and Pharmacology was published in 1985, we still do not know much
about how chitin is produced and transported across the
cell membrane so that it can interact perfectly with other
components for assembly of supramolecular extracellular
matrices such as the exoskeleton and PM. These structures
are still very much biochemical puzzles in which we do
not understand well how the various components come
together during morphogenesis, or are digested during
the molting process. Hopefully, this chapter will stimulate
more effort to gain an understanding of how insects utilize
chitin metabolism for growth and development, and also
to facilitate development of materials that may perturb
insect chitin metabolism for pest management purposes.
Since 2005, many questions have been answered about
the biosynthesis of insect chitin, including: why do insects
have two genes for CHS, and at what developmental
stages are the various CHSs produced? However, we know
little about the unique properties and functions of each
CHS. Of particular interest is the role of alternate splicing
in generating different isoforms of CHSs from the same
gene. The developmental cues that control alternate splicing and how they affect chitin synthesis and/or deposition
will be the subject of future studies. Attempts to express
full-length CHS genes in heterologous systems for the
production of active recombinant enzymes or subdomains
has met with very limited success, probably because CHSs
are membrane-bound proteins. The recent finding that
proteolytic processing may be necessary for CHS activation may also have contributed to this lack of success
(Broehan et al., 2007, 2008). In the future, the availability
of pure proteins and molecular probes for specific CHSs
would facilitate a better understanding of chitin biosynthesis and its regulation.
Two other significant questions about the regulation
of insect chitin biosynthesis are: what is the mechanism
of the initiation phase, and is there an autocatalytic initiator molecule? Like glycogen synthesis, chitin synthesis may involve both initiation and elongation phases.
As the initiator of glycogen synthesis, glycogenin transfers glucose from UDP-glucose to itself to generate an
­oligosaccharide–protein primer for elongation (Gibbons
et al., 2002). Like chitin synthase, glycogenin is a glycosyltransferase, which raises the question of whether chitin
synthase has an autocatalytic function similar to glycogenin, and whether there is a separate “chitinogenin”-like
protein. Another possibility is the participation of a lipid
primer for chitin synthesis. Cellulose synthesis in plants
involves the transfer of lipid-linked cellulodextrins to a
growing glucan chain (Read and Bacic, 2002). The lipid
in this case is sitosterol-β-glucoside. No lipid primer has
been identified to date for insect chitin synthesis.
Little is known about the catalytic mechanism of any
insect CHS. Once active insect CHS-related recombinant
proteins can be produced in a cell line, site-directed mutagenesis can be used to probe for essential residues in the
catalytic and regulatory domains. It is likely that acidic
amino acids play critical roles in CHS catalysis in a manner comparable to those identified in other glycosyltransferases (Hefner et al., 2002) and in yeast chitin synthases
(Nagahashi et al., 1995).
One of the major unanswered questions about insect
chitinolytic enzymes and chitin deaceylases is: why are
there so many of these enzymes? Some species have more
than 20 chitinase or chitinase-like genes, and we only
know the function of a few of them. Chitinolytic enzymes
are gaining importance for their biotechnological applications in agriculture and healthcare (Dahiya et al., 2006).
Additional success in using chitinases from both insects
and other organisms for different applications depends
on a better understanding of their biochemistry and regulation so that their useful properties can be optimized
through genetic and biochemical engineering. Reasons for
the rather high number of chitinolytic and chitin deacetylase enzymes with various domain structures are not fully
understood.
So far there has been little success in using chitinase in
pest-control applications, but it may prove more useful
as an enhancer protein in a cocktail with other biopesticides targeted at the cuticle or gut (Fiandra et al., 2010; Di
Maro et al., 2010). Also, only a few catalytic domains or
chitin-binding domains, or various combinations thereof
(domain shuffling and/or swapping), have been evaluated
for biocidal activity, and thus further toxicological experimentation after recombinations is warranted (Zakariassen et al., 2009; Li and Greene, 2010; Neeraja et al.,
2010). With good progress occurring in regard to functional analysis from RNAi studies, the ability to choose an
appropriate target gene or protein associated with insect
chitin metabolism that can be exploited to achieve targeted and selective control of pest insects has improved.
A hypothetical model for chitin-containing extracellular matrices in insects is the following: a fiber-reinforced
composite structure whereby chitin fibers form the initial
scaffold that is subsequently impregnated with a blend
of proteins into which some components of lower abundance, such as water, catechols, lipids, pigments, and minerals, are interspersed. For a soft hydrated material such as
the PM and trachea, chitin/chitosan and protein are the
major components that associate primarily non-covalently
via hydrogen bonding, as well as through hydrophobic
and electrostatic interactions with relatively little protein
cross-linking. The chitin-organizing proteins may have a
role in the precise arrangement of the individual laminar
layers of chitin, as well as their relative orientation with
226 7: Chitin Metabolism in Insects
the layers above and below it. For matrices that become
sclerotized, such as tanned cuticle, catechols are incorporated and oxidized to quinones and quinone methides,
which subsequently cross-link the proteins, and perhaps
chitin/chitosan as well. Future studies are needed to characterize more fully the covalent and non-covalent interactions and reactions of chitin, protein, lipid, mineral salts,
and oxidized catechols (chitin–water, chitin–protein,
chitin–catechol, chitin–lipid, chitin–pigment, chitin–
mineral interactions) from appropriate secretory tissues.
Results from such studies will provide critical insights into
the anabolic and catabolic pathways by which the chitin–
protein composite is formed and recycled, as well as into
the bioinspired fabrication of environmentally sustainable
load-bearing materials whose formulation is based, at least
in part, on insect chitin chemistry and metabolism.
Acknowledgments
We thank Dr. Mi Young Nho for helping with the phylogenetic tree construction and The National Science Foundation for financial support.
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