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First published online November 10, 2003
Review Article |
Chitin metabolism in insects: structure, function and regulation of chitin synthases and chitinases
Department of Biology/Chemistry, University of Osnabrück, 49069 Osnabrück, Germany
* Author for correspondence (e-mail: merzendorfer{at}biologie.uni-osnabrueck.de)
Accepted 9 September 2003
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Summary |
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 TOP Summary IntroductionChitin structureChitin formationInsect chitin synthasesRegulation of chitin synthasesChitin degradationInsect chitinasesRegulation of chitin degradationInhibition of chitin metabolismConclusion and outlookReferences |
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Key words: chitin, chitin synthesis, chitin synthase, chitinase, cuticle, peritrophic matrix, insect
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Introduction |
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TOPSummaryIntroductionChitin structureChitin formationInsect chitin synthasesRegulation of chitin synthasesChitin degradationInsect chitinasesRegulation of chitin degradationInhibition of chitin metabolismConclusion and outlookReferences |
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). Shortly before ecdysis, the molting fluid, which has
accumulated in the apolysial space, is reabsorbed, allowing the recycling of
old cuticle components. Formation of the new cuticle starts after the ecdysial
space opens as a result of the secretion of cuticle proteins and chitin fibers
through the apical membranes of epidermal cells. Initially, patches of
cuticullin, forming later on the outer epicuticle, are secreted, followed by
an unsclerotized, chitinous cuticle referred to as procuticle. Afterwards,
formation of the epicuticle seals the epidermis and protects it against the
digestive enzymes of the molting fluid. Before sclerotization is completed,
the insects expand their new cuticle and shed their old envelope, now called
exuvia, by performing distinct motor programs and increasing body pressure
(Carlson and Bentley, 1977).
Pre-ecdysis behavior and ecdysis are controlled by the action of molting
hormones such as eclosion hormone, which is secreted in response to falling
ecdysteroid titers and causes the release of pre-ecdysis-triggering hormone
and ecdysis-triggering hormone (Truman
and Riddiford, 1970; Zitnan
et al., 1999; Kingan and
Adams, 2000).
Chitin, a polymer of N-acetyl-ß-D-glucosamine, is
a major component of the insect cuticle. Solids NMR and gravimetric analysis
revealed that the chitin content constitutes up to 40% of the exuvial dry mass
depending on the insect species and varies considerably with the different
cuticle types even in a single organism
(Kramer et al., 1995). Chitin
is found in the exo- and endocuticle or in the newly secreted, unsclerotized
procuticle but not in the epicuticle, the outermost part of the integument
(Andersen, 1979). It functions
as light but mechanically strong scaffold material and is always associated
with cuticle proteins that mainly determine the mechanical properties of the
cuticle. In the migratory locust Locusta migratoria, more than a
hundred different cuticle proteins have been observed in 2-D electrophoresis
(Hojrup et al., 1986). Some of
them are highly conserved in various insect orders, some of them are
restricted to specific body regions and others contain repeats of hydrophobic
residues that seem to be linked with cuticle rigidity
(Andersen et al., 1995). One of
the best understood cuticle proteins is resilin, a glycine- and proline-rich
protein that confers high elasticity to the cuticle of hinge regions
(Andersen and Weis-Fogh,
1964).
Chitin is also an integral part of insect peritrophic matrices, which
function as a permeability barrier between the food bolus and the midgut
epithelium, enhance digestive processes and protect the brush border from
mechanical disruption as well as from attack by toxins and pathogens
(Tellam, 1996). Insect
peritrophic matrices have been categorized into two classes, based on their
mode of synthesis (Wigglesworth,
1930; Peters,
1992). Type I peritrophic matrices are synthesized along the whole
midgut and thus form a continuous delamination product. By contrast, type II
peritrophic matrices are exclusively produced by specialized cells in the area
of the cardia, which is located between the esophagus and the anterior midgut.
Peritrophic matrices usually exhibit a chitin content of between 3% and 13%
(Peters, 1992). For the
peritrophic matrix of the tobacco hornworm Manduca sexta, a chitin
content of even 40% has been reported
(Kramer et al., 1995). The
remainder of the peritrophic matrix consists of a complex mixture of proteins,
glycoproteins and proteoglycans. The peritrophic matrix is created when the
chitin microfibrils associate with the highly hydrated proteoglycan matrix
secreted by the gut cells. Further components of the peritrophic matrix, such
as peritrophins, may be added during the gelling process. Peritrophins appear
to link chitin microfibrils via their multiple chitin-binding domains
and additionally mediate binding to other glycoproteins. Consequently, they
may contribute significantly to the tensile strength of the peritrophic matrix
(Lehane, 1997). Variation of
peritrophic matrix formation rate is observed frequently in insects, depending
on the physiological condition (Locke,
1991). Some insects even completely cease peritrophic matrix
production during periods of starvation or molt. The old peritrophic matrix
then gets expelled or reabsorbed and regenerates when the animal starts
feeding again.
Thus, insect growth and development is strictly dependent on the capability to remodel chitinous structures. Therefore, insects consistently synthesize and degrade chitin in a highly controlled manner to allow ecdysis and regeneration of the peritrophic matrices. Chemical compounds that interfere with chitin metabolism, such as diflubenzuron, have been of special interest for the control of agricultural pests. Moreover, due to its unique properties, chitin itself is attracting more and more interest as a basic material for the chemical and pharmaceutical industry. In this review, we will focus on recent advances in understanding biosynthesis and degradation of chitin in cuticles and peritrophic matrices. In particular, we will address the substantial progress that has been made on chitin synthases and chitinases as a result of identification and sequencing of the insect genes encoding these enzymes.
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Chitin structure |
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TOPSummaryIntroductionChitin structureChitin formationInsect chitin synthasesRegulation of chitin synthasesChitin degradationInsect chitinasesRegulation of chitin degradationInhibition of chitin metabolismConclusion and outlookReferences |
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).
Chitin is composed largely of alternating N-acetylglucosamine
residues, which are linked by ß-(1-4)glycosidic bonds. Since hydrolysis
of chitin by chitinase treatment leads to the release of glucosamine in
addition to N-acetylglucosamine, it was concluded that glucosamine
might be a significant portion of the polymer. However, solids NMR analysis of
tobacco hornworm cuticle preparations suggested that little or no glucosamine
is present (Kramer et al.,
1995). Chitin polymers tend to form microfibrils (also referred to
as rods or crystallites) of 
3 nm in diameter that are stabilized by
hydrogen bonds formed between the amine and carbonyl groups. Chitin
microfibrils of peritrophic matrices may even exceed 0.5µm in length and
frequently associate in bundles containing parallel groups of 10 or more
single microfibrils (Peters et al.,
1979; Lehane,
1997). X-ray diffraction analysis suggested that chitin is a
polymorphic substance that occurs in three different crystalline
modifications, termed 
-, ß- and 
-chitin. They mainly differ
in the degree of hydration, in the size of the unit cell and in the number of
chitin chains per unit cell (Rudall and
Kenchington, 1973; Kramer and
Koga, 1986). In the form, all chains exhibit an
anti-parallel orientation; in the ß form the chains are arranged in a
parallel manner; in the form sets of two parallel strands alternate
with single anti-parallel strands. In addition, non-crystalline, transient
states have also been reported in a fungal system
(Vermeulen and Wessels,
1986). All three crystalline modifications are actually found in
chitinous structures of insects. The form is most prevalent in
chitinous cuticles, whereas the ß and forms are frequently found
in cocoons (Kenchington,
1976; Peters,
1992). Peritrophic matrices usually consist of - and
ß-chitin. Sometimes the presence of ß-chitin in cocoons is traced
back to the fact that some cocoons are formed from peritrophic matrices; for
example, those of Australian spider beetle Ptinus tectus, a
specialized beetle (Rudall and
Kenchington, 1973).
The anti-parallel arrangement of chitin molecules in the form
allows tight packaging into chitin microfibrils, consisting of 20 single
chitin chains that are stabilized by a high number of hydrogen bonds formed
within and between the molecules. This arrangement may contribute
significantly to the physicochemical properties of the cuticle such as
mechanical strength and stability
(Giraud-Guille and Bouligand,
1986). By contrast, in the ß- and -chains, packing
tightness and numbers of inter-chain hydrogen bonds are reduced, resulting in
an increased number of hydrogen bonds with water. The high degree of hydration
and reduced packaging tightness result in more flexible and soft chitinous
structures, as are found in peritrophic matrices or cocoons. The picture drawn
above is certainly oversimplified and does not explain the physicochemical
properties of cuticles and peritrophic matrices adequately because it is
reduced to only one component of a complex structure. However, differences in
the arrangement of chitin microfibrils between cuticles and peritrophic
matrices may help to understand their function. The cuticle is secreted in the
form of thin layers by the apical microvilli of epidermal cells. The chitin
microfibrils are embedded into the protein matrix and stabilize it in a way
that resembles constructions of steel-reinforced concrete. Since horizontal
microfibrils, in parallel with the cuticle plane, rotate either progressively
or abruptly from one level to another, complex patterns (e.g. helicoidal) and
textures (e.g. plywood-like structures) arise, depending on the degree of
rotational displacement (Bouligand,
1972). By contrast, in peritrophic matrices, the microfibrils are
normally arranged as a network of randomly organized, felt-like structures
embedded in an amorphous matrix, and only in a few cases have higher ordered
configurations been reported (Lehane,
1997).
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Chitin formation |
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TOPSummaryIntroductionChitin structureChitin formationInsect chitin synthasesRegulation of chitin synthasesChitin degradationInsect chitinasesRegulation of chitin degradationInhibition of chitin metabolismConclusion and outlookReferences |
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). Candy and Kilby
(1962) were the first to
propose a biosynthetic pathway for insect chitin synthesis starting with
glucose and ending with UDP-GlcNAc. The pathway from UDP-GlcNAc to chitin was
finally established in insects by Jaworski et al.
(1963) using cell-free
extracts from the southern armyworm Spodoptera eridania. Many
subsequent studies conducted with preparations from various insects supported
this general pathway, which is shown in
Fig. 1. Little is known about
the final steps in chitin synthesis in any organism. One intermediate step
might be the transfer of GlcNAc onto a lipid to form
dolichyldiphospho-N-acetylglucosamine, as was deduced from studies
with microsomal fractions of bugs and brine shrimps
(Quesada-Allue et al., 1976;
Horst, 1983). However, the
current evidence for the occurrence of lipid-GlcNAc intermediates or acceptors
during chitin assembly is weak and therefore not included in the depicted
pathway. In fungal systems, at least, an acceptor other than UDP-GlcNAc seems
not to be required to initiate polymerization
(McMurrough and Bartnicki-Garcia,
1971; Orlean,
1987; Merz et al.,
1999a; Chang et al.,
2003). Chitin synthase is definitely the key enzyme within the
biosynthetic pathway, but detailed understanding of its mode of action seems
to be a distant prospect, in particular because the enzyme has not yet been
purified to homogeneity.
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Many basic studies have been performed with fungal systems, and some of the
results seem to be valid for the insect enzymes as well. Common features of
most chitin synthases are that enzyme activity is dependent on the presence of
divalent cations such as Mg2+ or Mn2+ and that it is
increased by mild proteolysis, suggesting the existence of a zymogenic form
(Duran et al., 1975;
Mayer et al., 1980;
Hardy and Gooday, 1983;
Kramer and Koga, 1986;
Merz et al., 1999a). Usually,
chitin synthase activity can be inhibited by structural UDP-GlcNAc analogues
such as polyoxins and nikkomycin (Gooday,
1972; Dahn et al.,
1976). Enzyme activity seems to be restricted exclusively to
membrane-containing fractions
(Ruiz-Herrera and Martinez-Espinoza,
1999). Since chitin synthase has been localized in the membranes
of Golgi complexes (Horst and Walker,
1993) and intracellular vesicles
(Sentandreu et al., 1984), as
well as in plasma membranes (Duran et al.,
1975; Vardanis,
1979), it may be concluded that the enzyme follows an exocytotic
pathway, accumulating in cytoplasmic vesicles during its transport to the cell
surface. This view is supported by studies performed with imaginal discs of
Indian mealmoth Plodia interpunctella, which showed that chitin
synthesis is inhibited when microtubules are disrupted by cytoskeletal poisons
such as colchicine or vinblastine
(Oberlander et al.,
1983).
In fungal systems, substantial data have accumulated indicating that chitin
synthase activity of at least one chitin synthase isoform (CHS3p) is
associated with specialized intracellular microvesicles, known as chitosomes,
which exhibit a special lipid and protein composition
(Bracker et al., 1976;
Hernandez et al., 1981;
Florez-Martinez et al., 1990).
Electron microscopy has revealed that, in the presence of UDP-GlcNAc and
activators, purified chitosomes synthesize microfibrils that crystallize in
the lumen of the vesicles (Bracker et al.,
1976). Similar results were obtained when cell-free precipitates
resulting from chitin synthase activity in crude extracts of red flour beetle
Tribolium castaneum were examined. Electron micrographs of the chitin
synthase products showed a network of long, parallel-aligned microfibrils that
varied in thickness from 10 nm to 80 nm. The microfibrils were associated with
particles ranging from approximately 50 nm to 250 nm indiameter, which may be
interpreted as `insect chitosomes' (Cohen,
1982). However, final proof for direct involvement of `insect
chitosomes' in chitin synthesis is missing. Interestingly, chitosome-like
structures do not seem to occur in insect epidermal cells from Brazilian
skipper butterfly Calpodes ethlius and Australian sheep blowfly
Lucilia cuprina. Instead, electron microscope studies showed densely
stained areas at the tips of microvilli from epidermal cells, referred to as
plasma membrane plaques, which were considered as clusters of
chitin-synthesizing enzymes. During cuticle formation, these areas undergo
hormonally controlled cyclic turnovers
(Binnington, 1985;
Locke, 1991;
Locke and Huie, 1979). In
accordance with the predicted site of chitin synthesis, immunohistochemistry
using polyclonal antibodies raised against a conserved region of the chitin
synthase showed strong labeling within the apical region of the epidermis from
the epiproct of the American cockroach Periplaneta americana
(Fig. 2; H. Merzendorfer and L.
Zimoch, unpublished).
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Similar results have been obtained for the chitin synthase found in insect
intestinal systems to produce chitin for peritrophic membranes, which are
thought to be secreted by the microvilli of gut epithelial cells, since in
electron microscopy secreted material appears as more or less electron-dense
aggregation on top of or in between the microvilli
(Peters, 1992). By secreting
the peritrophic matrix, the microvilli act as a mold that causes microfibril
spacing and, in doing so, contribute to the formation of regular patterns that
are sometimes found in peritrophic matrices. Recently, Hopkins and Harper
(2001) used transmission
electron microscopy and wheat germ agglutinin (WGA)-gold staining to visualize
newly secreted chitinous fibers in lepidopteran midgut sections. They found
them on the microvillar surface but also within the apical region of
microvilli. In line with this view, immunohistochemistry conducted with
polyclonal antibodies raised against a conserved polypeptide of the
Manduca sexta chitin synthase demonstrated that the enzyme is
restricted to the apical tips of microvilli from columnar cells, one major
cell type found in larval midgut (Zimoch
and Merzendorfer, 2002). However, as may also be the case for
epidermal cells, it is not yet clear whether chitin synthase is actually
integrated into the plasma membrane or resides in vesicles enriched underneath
the plasma membrane. Confocal laser scanning microscopy, at least, unveiled
vesicular structures within the cytoplasm of columnar cells that immunoreacted
with the anti-chitin synthase antibodies and, hence, may represent `insect
chitosomes' on their way from the Golgi complex to the apical tips of
microvilli (Zimoch and Merzendorfer,
2002).
The specific mechanism by which chitin is produced is still unknown.
However, evidence suggests that chitin is synthesized through an asymmetric
mechanism, accepting GlcNAc units from the cytosolic UDP-GlcNAc pool and
releasing the nascent chain into the extraplasmic phase
(Ruiz-Herrera and Martinez-Espinoza,
1999). Indeed, from predictive analysis it seems likely that the
catalytic site of the chitin synthase that binds UDP-GlcNAc faces the
cytoplasm (Tellam et al.,
2000). On the basis of the presented data, one can propose two
alternative models for insect chitin synthesis
(Fig. 3). In one model,
intracellular vesicles merely function as exocytotic conveyors responsible for
the transport of chitin synthase to the plasma membrane. After membrane
fusion, the chitin synthase may be activated and subsequently secretes chitin
into the extracellular space. This model requires some regulatory step, which
controls enzyme activity, keeping the enzyme switched off until the vesicles
fuse with the plasma membrane. Since proteolytic activation of chitin
synthesis is observed in microsomal preparations from stable fly Stomoxys
calcitrans pupae (Mayer et al.,
1980), onset of chitin synthase activity upon vesicle fusion might
be achieved by extracellular proteases present in the midgut or in the molting
fluid (Law et al., 1977;
Reynolds and Samuels, 1996;
Terra et al., 1996).
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In a more speculative model, chitin is secreted into the lumen of specialized vesicles, which accumulate underneath the terminal web and fuse with the plasma membrane when chitin needs to be released. This model allows storage of chitin polymers and their rapid release, which may be important for peritrophic matrix secretion upon feeding of blood-sucking mosquitoes. However, the length of chitin polymers may be restricted due to the limited volume of the vesicles.
If the catalytic site really faces the cytoplasm, UDP-GlcNAc could directly
bind from the cytoplasmic pool. Consequently, in both presented models,
nascent chitin has to be transported across the membrane, possibly involving
transmembrane regions of the chitin synthase. If the catalytic domain should,
contrary to the predictions, face the extraplasmic site, UDP-GlcNAc would need
to be transported either into the extracellular environment or into the lumen
of the vesicles. Substrate transport might be achieved either by the chitin
synthase itself or by transmembrane proteins similar to the UDP-GlcNAc
transporters that reside in the endoplasmic reticulum or the Golgi vesicles
(Perez and Hirschberg, 1985;
Cecchelli et al., 1986;
Segawa et al., 2002).
Although no biochemical data that support intravesicular catalysis are
currently available, it would cleverly circumvent the unsolved problem of how
to translocate the nascent chitin polymer across the membrane, because chitin
would already be synthesized on the side of its subsequent release.
Chitin synthase can be assayed readily and some progress has been made in
purifying active components in fungal systems
(Duran and Cabib, 1978;
Kang et al., 1984;
Machida and Saito, 1993;
Uchida et al., 1996).
However, despite all efforts that have been made during the past decades, the
enzyme has still not been purified to homogeneity. Therefore, we have only a
vague image of the molecular mechanism of chitin synthesis. In contrast to
fungi, only few studies have been conducted using chitin synthase-containing
preparations from insects. In vivo studies, as well as in
vitro studies using insect organ and cell cultures, first provided
insights into insect chitin synthesis
(Candy and Kilby, 1962;
Marks and Leopold, 1971;
Marks, 1972;
Surholt, 1975;
Vardanis, 1976). More
detailed knowledge emerged from investigations performed in cell-free systems,
although preservation of enzyme activity turned out to be difficult.
Quesada-Allue et al. (1976)
were among the first to measure chitin synthase activity in cell-free extracts
of insects. For this purpose, they used crude extracts from the kissing bug
Triatoma infestans integument and monitored
[14C]N-acetylglucosamine incorporation into the polymer.
Chitin synthase activity exhibited a pH optimum of about 7.2 and was dependent
on the presence of Mg2+ and GlcNAc. Interestingly, radioactivity
was also found concomitantly with chitin synthesis in a liposoluble fraction.
Chromatographic analysis of this fraction suggested the involvement of
N-acetylglucosaminyl-phospholipid in insect chitin synthesis, which
was supported by the finding that chitin synthesis was blocked by tunicamycin,
an inhibitor of UDP-N-acetylglucosamine:dolichyl-phosphate
N-acetylglucosaminephosphotransferase
(Heifetz et al., 1979;
Quesada-Allue, 1982).
Supporting evidence came from studies performed with microsomes from brine
shrimps (Artemia salina), which catalyzed the transfer of
N-acetylglucosamine from UDP-N-acetylglucosamine to a lipid
acceptor. The resulting dolichyldiphosphate-linked chito-oligomer may act as a
GlcNAc acceptor for chitin synthesis
(Horst, 1983). By contrast,
from kinetic studies it was concluded that chitin synthesis generally occurs
without the need for soluble or lipid GlcNAc acceptors functioning as primers
for chain assembly (Horsch et al.,
1996; Merz et al.,
1999a). In line with this interpretation, some groups have
reported that chitin synthesis was not affected significantly by tunicamycin
in several insect systems (Mayer et al.,
1981; Fristrom et al.,
1982; Bade, 1983).
The inconsistency regarding the published data, together with the fact that
chain assembly occurs without the need of an initial acceptor other than
UDP-GlcNAc in fungal systems, however, raises doubt about the significance of
lipid intermediates or primers in arthropod systems.
Chitin synthesis is influenced in different ways by other effectors as
well, depending on the particular enzyme source. For instance, GlcNAc has been
reported to stimulate chitin synthesis in fungi and also in some insects
(Keller and Cabib, 1971;
Quesada-Allue et al., 1976;
Cohen and Casida,
1980a,b,
1982). By contrast, studies
with microsomal fractions from Stomoxys showed almost complete
inhibition of chitin synthesis with 1 mmol l-1 GlcNAc
(Mayer et al., 1980). Even
more confusing, the activity of classical inhibitors of chitin synthesis such
as polyoxin, nikkomycin and diflubenzuron also seems to depend on the insect
system used for the particular study. Cohen and Casida
(1982), for instance, reported
different effects of polyoxins and nikkomycin on chitin synthesis in cecropia
moth Hyalophora cecropia and cabbage looper Trichoplusia ni.
Mayer et al. (1980,
1981) observed polyoxin D
inhibition in microsomal preparations from Stomoxys only at high
concentrations but no inhibitory effect for diflubenzuron, whereas Turnbull
and Howells (1983) showed for
crude homogenates of larval integuments from Lucilia that chitin
synthesis was inhibited by both polyoxin D and diflubenzuron. However, due to
the crude character of the investigated preparations, care has to be taken not
to jump to conclusions. Besides cell-free extracts, chitin synthesis has also
been reported for several insect cell lines. For instance, Marks et al.
(1984) demonstrated chitin
synthase activity in MRRL-CH cells, a continuous cell line from
Manduca embryos. Londershausen and colleagues showed chitin synthesis
in an epithelial-like cell line from the non-biting midge, Triatoma
infestans as well as in Kc cell lines from Drosophila
melanogaster. In cell cultures from Chironomus, incorporation of
radiolabeled glucosamine was partially inhibited by the acyl urea SIR 8514,
polyoxin D and nikkomycin (Londershausen
et al., 1988).
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Insect chitin synthases |
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TOPSummaryIntroductionChitin structureChitin formationInsect chitin synthasesRegulation of chitin synthasesChitin degradationInsect chitinasesRegulation of chitin degradationInhibition of chitin metabolismConclusion and outlookReferences |
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). The
cDNA was derived from Lucilia, the Australian sheep blowfly. It
encoded a presumed protein with a molecular mass of approximately 180 kDa, and
predictive analysis suggested the existence of 15-18 transmembrane helices,
indicating that the enzyme is an integral membrane protein. Analysis of the
LsCHS-1 expression pattern by RT-PCR and in situ
hybridization revealed that the transcripts are detectable in all
developmental stages of the fly and that they localize in the epidermal cells
underlying the procuticle.
Meanwhile, progress has been made in investigating insect chitin synthases
due to the availability of an increasing number of gene and cDNA sequences
deposited in sequence databases or published within the past three years,
although final proof that the deduced proteins synthesize chitin is pending
(Ibrahim et al., 2000;
Tellam et al., 2000;
Gagou et al., 2002;
Zhu et al., 2002).
In contrast to fungi, which possess multiple genes encoding chitin synthase
isoforms (Munro and Gow,
2001), molecular analysis of nematode and insect chitin synthase
genes (CHS) has so far revealed a limited number of gene copies.
Genome sequencing projects have shown that Caenorhabditis elegans,
Drosophila and Anopheles gambiae possess two different
CHS genes, and recently cDNA sequencing or genomic Southern blotting
also provided evidence for two gene copies in Lucilia, Manduca and
Tribolium (Gagou et al.,
2002; Tellam et al.,
2000; Zhu et al.,
2002; Zimoch and
Merzendorfer, 2002; Y. Arakane, D. Hogenkamp, Y. C. Thu, C. A.
Specht, R. W. Beeman, K. J. Kramer, M. Kanost and S. Muthukrishnan,
unpublished results). Comparison of amino acid sequences from fungal, nematode
and insect chitin synthases has revealed that insect enzymes are more closely
related to those of nematodes than those of fungi.
Insect chitin synthases are large transmembrane proteins with theoretical
molecular masses ranging from 160 kDa to 180 kDa and exhibit a slightly acidic
isoelectric point between 6.1 and 6.7. Alignments of the amino acid sequences
from Lucilia, Drosophila and Caenorhabditis revealed a
tripartite domain structure (Tellam et
al., 2000; see also Fig.
4A). Domain A is found in the N-terminal region, has
varying numbers of transmembrane helices and shows the least sequence
similarity among any of the species. Depending on the number of predicted
transmembrane helices in the A domain, the N-terminus appears to be
located at different sides of the membrane, facing either the extracellular
environment or the cytoplasm. However, this may also reflect shortcomings
regarding the computer-based prediction of transmembrane helices.
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Domain B is found in the center of chitin synthases, comprises 400
amino acids and contains the catalytic center of the protein. The B domain is
highly conserved and contains two unique motifs, EDR and
QRRRW, that are present in all types of chitin synthases and
therefore can be regarded as signature sequences. Some of the conserved
residues have been implicated to be essential for the catalytic mechanism,
since they may be involved in protonation of the substrate
(Sinnott, 1990;
Breton et al., 2001). In
particular, even conservative substitutions of those residues that have been
highlighted in bold above drastically decrease chitin synthase activity in
yeast, although they do not significantly affect the apparent
Km values for the substrate
(Nagahashi et al., 1995).
Similar sequences have been found in bacterial and vertebrate hyaluron
synthases (Rosa et al., 1988;
DeAngelis et al., 1994;
Pummill et al., 1998),
cellulose synthases (Saxena et al.,
2001) and N-actelyglucosaminyltransferases such as the
NodC protein (Geremia et al.,
1994). An aspartic acid residue at position 441 of the yeast
chitin synthase 2 protein (CHS2p) was also suggested to be conserved in all
chitin synthases. Its substitution by glutamic acid led to a severe loss of
chitin synthase activity in the resulting CHS2 mutant
(Nagahashi et al., 1995).
This aspartic acid residue is nevertheless replaced by glutamate in some
insect chitin synthases at a corresponding position, supporting the necessity
of at least an acidic residue at this position. However, a highly conserved
aspartic acid is also found at position 344 of the yeast CHS2p. Unfortunately,
this position has not been addressed by in vitro mutagenesis so far.
Zhu et al. (2002) described
three additional highly conserved blocks in insect chitin synthases, CATMWHXT,
QXFEY and WGTRE (at positions 583-590, 794-798 and 1076-1080 of the
Drosophila CHS-1 protein, respectively; see
Fig. 4A), with most of the
amino acids also conserved in fungal or nematode chitin synthases.
Domain C comprises the C-terminal part of the enzyme and contains
two amino acids that might also be involved in catalysis, since site-directed
mutagenesis performed with CHS2p of yeast showed that enzyme activity was
diminished when W803 or T805 were exchanged for alanine
(Yabe et al., 1998). Both
residues are conserved in insects at positions comparable to those of the
yeast enzyme, immediately following transmembrane helix five of the C domain.
Although this domain is far less conserved than the catalytic domain, it
exhibits seven transmembrane helices as a common feature.
As has been reported for several fungal chitin synthases, insect enzymes
may also be glycosylated because they exhibit several putative
N-glycosylation sites of which one is conserved in every insect
chitin synthase (Table 1). In
fungal systems, the affinity of lectins such as concanavalin A or wheat germ
agglutinin to the sugar portion of N-glycosylated residues has
already been used for the purification of active components of chitin
synthases (Machida and Saito,
1993; Merz et al.,
1999a,b).
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Based on relative sequence differences, chitin synthases have been grouped
into two classes, class CHS-A and class CHS-B enzymes. So far, most insects
seem to have one gene copy for each enzyme. Since both genes are located at
one chromosome in both Drosophila and Anopheles, it is
likely that they have evolved from a common ancestor by gene duplication
(Gagou et al., 2002; Y.
Arakane, D. Hogenkamp, Y. C. Thu, C. A. Specht, R. W. Beeman, K. J. Kramer, M.
Kanost and S. Muthukrishnan, unpublished results). Gene expression studies
performed in Lucilia, Tribolium and Manduca indicated that
class A chitin synthases are specifically expressed in the epidermis and
related ectodermal cells such as tracheal cells, while expression of class B
chitin synthases may be restricted to gut epithelial cells that produce
peritrophic matrices (Y. Arakane, D. Hogenkamp, Y. C. Thu, C. A. Specht, R. W.
Beeman, K. J. Kramer, M. Kanost and S. Muthukrishnan, unpublished results). In
Lucilia, LcCHS-1, a class A chitin synthase, was found in the
carcass, which is free of internal tissues, but not in the midgut
(Tellam et al., 2000). In
Aedes aegypti, RT-PCR with a probe to AaCHS-1, a class B chitin
synthase, resulted in products that were detectable in midgut or whole
mosquitoes but not in the carcass
(Ibrahim et al., 2000).
Moreover, RT-PCR that was conducted with mRNA preparations from
Manduca using isoform-specific primers suggests that expression of
class B chitin synthases is restricted, since MsCHS-2-specific products can
only be observed in the midgut but not in other tissues (D. Hogenkamp and S.
Muthukrishnan, personal communication; K. Gerdemann and H. Merzendorfer,
unpublished). Besides homology criteria, class A insect chitin synthases are
characterized by the presence of a coiled-coil region immediately following
the five transmembrane helices of the C domain
(Tellam et al., 2000; Y.
Arakane, D. Hogenkamp, Y. C. Thu, C. A. Specht, R. W. Beeman, K. J. Kramer, M.
Kanost and S. Muthukrishnan, unpublished results;
Fig. 4A; Table 1). The coiled-coil
region is predicted to face the extracellular space and may be involved in
protein-protein interaction, vesicle fusion or oligomerization
(Skehel and Wiley, 1998;
Burkhard et al., 2001).
Interestingly, cellulose synthases from mosses, ferns, algae and vascular
plants, which have some similarities with chitin synthases, are organized in
rosettes consisting of six subunits, which in turn may each contain six single
polypeptides (Doblin et al.,
2002). Rosette assembly may involve oxidative dimerization between
single cellulose synthase polypeptide subunits via zinc finger
domains (Kurek et al., 2002).
It is therefore tempting to speculate that oligomerization may be important
for chitin synthases too, possibly mediated by the coiled-coil region.
It seems that class A chitin synthases are encoded by a gene that is
differentially spliced, resulting in the expression of an alternate exon
comprising 59 amino acids and encoding transmembrane helix six and adjacent
regions of the C domain (Tellam et al.,
2000; Y. Arakane, D. Hogenkamp, Y. C. Thu, C. A. Specht, R. W.
Beeman, K. J. Kramer, M. Kanost and S. Muthukrishnan, unpublished results).
The alternate exons share 70%, 72% and 78% identical amino acids in TcCHS-1,
DmCHS-1 and MsCHS-1, respectively. Recently, Arakane and colleagues
demonstrated that both exons are actually expressed in Tribolium (Y.
Arakane, D. Hogenkamp, Y. C. Thu, C. A. Specht, R. W. Beeman, K. J. Kramer, M.
Kanost and S. Muthukrishnan, unpublished results). Although their expression
pattern differs to some extent during development, the functional significance
of alternate exon usage is not yet clear.
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Regulation of chitin synthases |
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).
Therefore, the expression of the involved genes has to be precisely controlled
during each molt. Insect metamorphosis is known to be regulated by the release
of ecdysone, a steroid hormone secreted into the hemolymph by the prothoracic
gland, which could be involved in the regulation of chitin synthesis. For
Drosophila imaginal disks, it has been shown that the presence of
ecdysone inhibits chitin synthesis as well as the expression of procuticle
proteins (Apple and Fristrom,
1991; Hiruma et al.,
1991). Converse data were obtained for cultured wing discs from
Plodia, where ecdysone stimulated uptake and incorporation of
radiolabeled glucosamine (Oberlander,
1976). These opposite effects on chitin synthesis are not
necessarily conflicting; rather, they may reflect a dual effect of
ecdysteroids, since these steroids are known to act as both positive and
negative regulators (Apple and Fristrom,
1991; Spindler et al.,
2001). Upregulation of transcriptional activities by ecdysteroids
during insect development was found in a number of cases. The
Drosophila genes Eips 28 and Eips 29, for instance,
are controlled tissue- and stage-specifically by ecdysone-responsive elements
present in the upstream and downstream flanking regions
(Andres and Cherbas, 1994).
Further examples of this mode of ecdysteroid-mediated transcriptional
regulation are the Drosophila genes encoding the yolk protein
(Bownes et al., 1996), the
heat-shock proteins hsp23 and hsp27 (Luo
et al., 1991), the caspase DRONC
(Dorstyn et al., 1999;
Hawkins et al., 2000) and the
Manduca genes EcR-A and EcR-B1, which encode two
ecdysone receptor (EcR) isoforms (Jindra
et al., 1996). Downregulation of transcriptional activities by
ecdysteroids was observed too, but these effects may be due to indirect
actions. For instance, the ecdysteroid-regulated gene esr20, which is
expressed in the trachea of Manduca, has been suggested to be
downregulated at ecdysis. In this case, downregulation may be caused by a
decline in transcript stability triggered indirectly by 20-hydroxyecdysone
(Meszaros and Morton, 1997).
Moreover, transcription of the dopa decarboxylase-encoding gene from
Manduca may be indirectly suppressed by 20-hydroxyecdysone
via an ecdysteroid-induced transcription factor that itself
suppresses dopa decarboxylase transcription
(Hiruma et al., 1995).
Analysis of chitin synthase expression during Drosophila
metamorphosis indicates that ecdysone has a regulatory role on CHS-1 (DmeChSB)
and CHS-2 (DmeChSA) transcript levels
(Gagou et al., 2002). In third
instar larvae and shortly after pupariation CHS transcripts were barley
detectable. However, in response to the first ecdysone pulse, both transcripts
were drastically upregulated, although at different points in time. CHS-1
transcripts were upregulated first, coinciding with the formation of pupal
inner epicuticle, whereas CHS-2 transcripts were upregulated a few hours
later, concurrent with pupal procuticle formation. The progression of
transcript upregulation may suggest that ecdysone activates transcription of
the CHS genes by activating a nuclear receptor heterodimer consisting
of the EcR and the Drosophila retinoid X receptor homologue USP, the
ultraspiracle protein (Yao et al.,
1993). Indeed, computational scanning of the `transfac database'
revealed that both genes contain putative ecdysone responsive elements (EcREs)
in their upstream regions. The regulatory elements correspond with the
consensus sequences (G/T)NTCANTNN(A/C)(A/C) and (A/G)G(G/T)T(G/C)
ANTG(A/C)(A/C)(C/T)(C/T), deduced from promoters of hsp23, hsp27 and
Fbp1, which encode two Drosophila heat-shock proteins and a
fat body protein, respectively (Luo et
al., 1991; Antoniewski et al.,
1993; Wingender et al.,
1997; Tellam et al.,
2000). Somewhat different results were obtained when
MsCHS-1 expression was investigated in Manduca 5th instar
larvae and pupae (Zhu et al.,
2002). During feeding, transcript levels were observed to be
relatively constant, but dropped drastically when feeding ceased and gradually
increased again in the wandering stage to a maximum at pupal molt. Correlation
with ecdysteroid titers in the Manduca hemolymph suggests that the
MsCHS-1 gene is negatively controlled by ecdysteroids, because
ecdysteroid titers increase prior to wandering and decrease before pupation
(Bollenbacher et al., 1981;
Baker et al., 1987).
Transcriptional or post-transcriptional regulation also seems to occur for
the midgut-specific chitin synthase isoform encoded by class B genes. In
situ hybridization performed with midgut sections from the mosquito
Aedes showed that the amount of transcripts was upregulated in
response to a bloodmeal (Ibrahim et al.,
2000). Interestingly, transcripts were localized to the apical
region of epithelial cells. Similar results were obtained by in situ
hybridization of cryosections from the anterior midgut of Manduca 5th
instar larvae (Zimoch and Merzendorfer,
2002). The observed apical localization may reflect the site of
CHS-2 biosynthesis because, in Manduca, columnar cell apical regions
with large whorls of rough endoplasmic reticulum and Golgi complexes are found
beneath the terminal web (Cioffi,
1979). This interpretation is also supported by the observation
that in the basal region of the anterior midgut both rough endoplasmic
reticulum and Golgi complexes are missing but are present in the basal region
of the median and posterior midgut. Correspondingly, CHS-2 transcripts are
evenly spread throughout the cytoplasm of the columnar cells in the median and
posterior midgut (Zimoch and
Merzendorfer, 2002). The only cell organelles that have been
observed in the region of the Manduca columnar cells' terminal web
were interpreted as small Golgi vesicles with electron-dense contents that
appeared to be collected at the apical border of the cell
(Cioffi, 1979). Are these
vesicles loaded with chitin that will be released upon a secretory signal? In
any case, inactive chitin synthases also have to be transported to the apical
plasma membrane, and vesicle transport may be regulated as well. This notion
may be supported by the finding that microtubule disruptans interfere with
chitin synthesis (Oberlander et al.,
1983).
Since insect chitin synthase activity is increased by limited proteolysis,
it is tempting to speculate about the existence of a cellular pool of inactive
proenzymes being activated by specific signals. However, even in fungal
systems, the significance of this phenomenon has not yet been elucidated
(Merz et al., 1999a). Besides
proteases, further regulatory factors that affect chitin synthase activity may
exist in insects. In yeast, several proteins that are involved in the
regulation of chitin synthesis have been described. Yeast CHS4p, for instance,
seems to stimulate chitin synthase III (CHS3p) activity by a direct
protein-protein interaction and may be needed for septin-dependent, localized
chitin deposition in the yeast cell wall
(Ono et al., 2000). SHC1p is
a protein homologous to CHS4p and functions in cell wall ascospore assembly
but regulates CHS3p activity exclusively during the sporulation process
(Sanz et al., 2002). Another
protein that is required for fusion and mating, CHS5p, has been implicated in
regulation of chitin synthase, since chitin synthase III targeting to cortical
sites in yeast is dependent on both CHS5p and the actin cytoskeleton/Myo2p
(Santos and Snyder, 1997).
Further proteins have been discovered by genetic screens, including CHS6p,
which is necessary for the anterograde transport of CHS3p from the chitosome
to the plasma membrane (Ziman et al.,
1998), and CHS7p, which regulates CHS3p export from the
endoplasmic reticulum (Trilla et al.,
1999). So far, no orthologs have been described in insects.
However, future experiments with two- or three-hybrid systems may reveal
interaction partners that regulate chitin synthase activity in insects.
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Chitin degradation |
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; Fukamizo,
2000; Kondo et al.,
2002). Since chitin-degrading enzymes can be used to convert
chitin-containing raw material into biotechnologically utilizable components,
they are of significant interest for the chemical and pharmaceutical industry.
Moreover, chitinases and their inhibitors may be adopted as insecticides to
combat pests or as fungicides for the treatment of microbial infections
(Kramer and Muthukrishnan,
1997; Herrera-Estrella and
Chet, 1999). In chitin-producing organisms, chitinolytic enzymes
are essential for maintaining normal life cycle functions such as
morphogenesis of arthropods or cell division and sporulation of yeast and
other fungi (Passonneau and Williams,
1953; Elango et al.,
1982; Kuranda and Robbins,
1991). They are also found in organisms that do not contain chitin
themselves but utilize chitin as a nutrient source. Several bacterial genera,
such as Streptomyces spp., convert insoluble chitin into soluble,
metabolizable compounds by means of different chitinolytic enzymes and
chitin-binding proteins that act as a glue for adherence on chitinous
substrates (Charpentier and Percheron,
1983; Schrempf,
2001). Chitinase genes have been identified in the genome of the
Autographa californica nucleopolyhedrovirus, a member of the virus
family Baculoviridae, which is restricted to arthropod hosts
(Hawtin et al., 1995). The
baculoviral chitinase may play a crucial role in viral infectivity
(Thomas et al., 2000;
Saville et al., 2002). For
plant and vertebrate chitinases, including those produced by human
macrophages, it is proposed that they act in defense against chitin-containing
pathogens or pests (Leah et al.,
1991; Boot et al.,
1995,
1998;
Gooday, 1999;
Carlini and Grossi-de-Sa,
2002).
In insects, chitin-degrading enzymes play a crucial role in postembryonic
development, especially during larval molt and pupation. During the molt,
proteases and chitinases are synthesized by epidermal cells and accumulate in
the molting fluid between the epidermis and the old cuticle
(Dziadik-Turner et al., 1981;
Samuels and Reynolds, 1993;
Samuels and Paterson, 1995;
Reynolds and Samuels, 1996).
Most of the digestion products are transported via the molting fluid
to the mouth and anal openings and are subsequently accumulated in the midgut
(Reynolds and Samuels, 1996;
Yarema et al., 2000).
However, direct reabsorption by the epidermis may also occur. In any case, the
reincorporated constituents seem to be recycled and used to produce the new
procuticle (Surholt, 1975;
Reynolds and Samuels, 1996;
Kaznowski et al., 1986). In
addition, some larvae ingest the shed exuvia to regain its constituents. This
behavior coincides with the period of chitinase expression in the gut
(Kramer et al., 1993).
Moreover, the midgut chitinases seem to be involved in the formation,
perforation and degradation of the midgut peritrophic matrix, which protects
the gut epithelium from damaging factors
(Peters, 1992;
Shen and Jacobs-Lorena, 1997;
Filho et al., 2002).
Chitinolytic enzymes are also found in some hymenopteran venoms and in the
digestive fluid of spiders, where they may facilitate the entry of harmful
ingredients through the cuticle of the prey
(Mommsen, 1980;
Krishnan et al., 1994;
Jones et al., 1996).
Recently, a fat body-specific chitinase that is detected in milk gland tissue
and could therefore be important for the development of intrauterine larvae
was characterized in the viviparous tsetse fly Glossina morsitans
(Yan et al., 2002).
Since chitin is hard to break due to its physicochemical properties, its
degradation usually requires the action of more than one enzyme type.
Endo-splitting chitinases produce chitooligomers that are subsequently
converted to monomers by exo-splitting
ß-N-acetylglucosaminidases. The latter enzyme cleaves off
N-acetylglucosamine units from non-reducing ends and prefers smaller
substrates than chitinases (Koga et al.,
1982,
1983,
1997; Fukamizo and Kramer,
1985a,b;
Kramer and Koga, 1986;
Kramer et al., 1993;
Zen et al., 1996;
Filho et al., 2002). As a
consequence of these properties, the overall rate of chitin hydrolysis is
limited by the action of the chitooligomer-producing chitinase, which
drastically increases the effective substrate concentration for the
ß-N-acetylglucosaminidase.
The mechanism of catalysis seems to be quite similar to that postulated for
the cellulase complex and other multi-enzyme systems hydrolyzing linear
polymers (Easterby, 1973;
Klesov and Grigorash, 1982).
The first enzyme of the `cellulosome', a multiple cellulase-containing protein
complex, is an endocellulase that limits monosaccharide formation, because
exocellulases are inefficient in degrading insoluble polysaccharides. In
contrast to the cellulolytic enzymes, however, chitinolytic enzymes are not
believed to assemble into corresponding `chitinosomes', although evidence
excluding their existence is lacking.
Interestingly, the appearance and activity of both chitinolytic enzymes
seem to be in reverse order as they function in chitin degradation. In
Manduca, the silkworm Bombyx mori and Locusta, the
exo-splitting ß-N-acetylglucosaminidase appears earlier in the
molt than the endo-splitting chitinase. This was verified by activity assays
and immunoblot analysis with polyclonal antibodies raised against both enzymes
(Kimura, 1973a,
1977;
Zielkowski and Spindler,
1978; Fukamizo and Kramer,
1987; Koga et al.,
1989). Since the cuticle is a complex matrix of chitin and tightly
bound proteins, enzyme accessibility is restricted, and free non-reducing ends
are limited. Thus, further mechanisms of cuticle degradation exist, including
degradation by proteases that are also present in the molting fluid
(Law et al., 1977).
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Insect chitinases |
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TOPSummaryIntroductionChitin structureChitin formationInsect chitin synthasesRegulation of chitin synthasesChitin degradationInsect chitinasesRegulation of chitin degradationInhibition of chitin metabolismConclusion and outlookReferences |
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; Fig. 4B). The
last two domains, however, do not seem to be necessary for chitinase activity
because naturally occurring chitinases that lack these regions are still
enzymatically active (Girard and Jouanin,
1999; Feix et al.,
2000; Yan et al.,
2002). In agreement with these observations, C-terminus-truncated
versions of the recombinant Manduca chitinase still exhibit catalytic
activity (Wang et al., 1996;
Zhu et al., 2001).
|
In all insect chitinases sequenced so far, a hydrophobic signal peptide is
predicted to precede the N-terminal region of the mature protein
(Kramer et al., 1993;
Koga et al., 1997;
Choi et al., 1997;
Nielsen et al., 1997;
Shen and Jacobs-Lorena, 1997
