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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Key words: chitin, chitin synthesis, chitin synthase, chitinase, cuticle, peritrophic matrix, insect
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Introduction |
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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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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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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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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.
![]() |
Regulation of chitin synthases |
---|
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.
![]() |
Chitin degradation |
---|
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
).
![]() |
Insect chitinases |
---|
|
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
;
Kim et al., 1998
;
Mikitani et al., 2000
;
Royer et al., 2002
). The
signal peptide presumably mediates secretion of the enzyme into the
endoplasmic reticulum and it is cleaved off by signal peptidases after the
protein has been transported across the membrane
(von Heijne, 1990
;
Müller, 1992
).
The catalytic domain of family 18 chitinases comprises the N-terminal half
of the enzyme. It was suggested that the N-terminal part of this domain
influences the binding or the hydrolysis of the substrate
(Perrakis et al., 1996).
Sequence alignments revealed two highly conserved regions within the catalytic
domain, the second one including the catalytic center
(Henrissat, 1991
;
Coutinho et al., 2003
; see
also Fig. 4B). The catalytic
domain of family 18 chitinases has a TIM-barrel structure
(Lasters et al., 1988
) that
forms a groove on the enzyme's surface. This groove is considered as the
active center, which binds sugar units of chitin, possibly
(GlcNAc)6 moieties, that are subsequently cleaved by a retaining
mechanism discussed later on (Armand et
al., 1994
; Drouillard et al.,
1997
). The hallmarks of the chitinase structure are eight parallel
ß-strands, forming the barrel's core, which is surrounded by eight
-helices connected to the barrel by linkers of different length and
form. The two consensus sequences lie along ß-strands three and four of
the
/ß barrel and represent the substrate-binding site
(Aronson et al., 1997
). So
far, no crystal structure of an insect chitinase is available, but homology
modeling using crystal structures of bacterial and plant chitinases has
revealed three-dimensional models of the catalytic domain from the
Manduca chitinase showing striking similarities with the
/ß barrel structure described above
(Kramer and Muthukrishnan,
1997
; Huang et al.,
2000
). Although the models lack a well defined
(
/ß)8 folding, they predict eight ß-sheets and
four complete and several incomplete
-helices. In some insects, the
catalytic region is followed by a less conserved domain containing a putative
PEST-like region that is also found near the C-terminus of the yeast chitinase
(Kim et al., 1998
;
Kuranda and Robbins, 1991
;
Kramer et al., 1993
;
Royer et al., 2002
). As
already mentioned, insect chitinases without a PEST-like region have also been
described in the literature (Girard and
Jouanin, 1999
; Feix et al.,
2000
; Yan et al.,
2002
). PEST-like regions presumably increase the susceptibility of
the enzyme to proteolysis by a calcium-dependent protease or to degradation
via the 26S proteasome (Rogers
et al., 1986
; Rechsteiner and
Rogers, 1996
). Therefore, these regions could play a role in
enzyme turnover or activation of zymogenic chitinases.
Like some fungal chitinases, the chitinases found in insect molting fluids
are extensively glycosylated. Thus, insect chitinases can be easily detected
by carbohydrate staining after sodium dodecyl sulfate polyacrylamide gel
electrophoresis (SDS-PAGE) and blotting. Several putative N-linked
glycosylation sites that may be necessary for the secretion of the protein and
maintenance of its stability are found within the deduced amino acid sequences
of insect chitinases (Gopalakrishnan et
al., 1995; Kramer and
Muthukrishnan, 1997
; Kim et
al., 1998
; Fig.
4B). Moreover, the serine/threonine-rich PEST-like region of the
Manduca chitinase is extensively modified by O-glycosylation
(Kramer et al., 1993
;
Arakane et al., 2003
).
Previous determination of the carbohydrate composition of the Manduca
chitinases revealed N-acetylglucosamine and several neutral hexoses
as part of the sugar portion (Koga et al.,
1983
,
1997
). The attachment of
oligosaccharides probably increases solubility and protects the peptide
backbone against proteases.
Insect chitinases are anchored to their substrate through the C-terminal
chitin-binding domain, which is characterized by a six-cysteine motif that is
also found in nematode chitinases
(Venegas et al., 1996). It
functions in targeting of the enzyme to its substrate and thereby facilitates
catalysis. The sixcysteine motif is also found in several peritrophic matrix
proteins, as well as in receptors and other proteins that are involved in
cellular adhesion (Tellam et al.,
1992
; Tellam,
1996
; Kramer and
Muthukrishnan, 1997
; Shen and
Jacobs-Lorena, 1999
).
Individual chitinases possess different combinations of these three basic
domains. While the chitinases from Manduca, Bombyx and fall webworm
moth Hyphantria cunea exhibit the typical tripartite domain structure
(Kramer et al., 1993;
Kim et al., 1998
), some other
chitinases lack PEST-like or the typical chitin-binding regions
(Girard and Jouanin, 1999
;
Feix et al., 2000
;
Yan et al., 2002
). Other
insects, in turn, may express multi-modular enzymes. In Aedes, for
example, chitinases are encoded by two different genes. Nucleotide sequencing
has revealed that one of the genes contains tandemly arranged open reading
frames that encode three separate chitinases, each containing a catalytic- and
also a chitin-binding domain. The gene arrangement suggests co-regulated
transcription resembling bacterial operons
(Niehrs and Pollet, 1999
).
Post-transcriptional splicing, however, may also lead to a single,
multi-modular protein with three catalytic- and chitin-binding domains each
(de la Vega et al., 1998
;
Henrissat, 1999
).
TmChit5, the gene that encodes chitinase 5 of the beetle Tenebrio
molitor, also exhibits an unusual structure, since it contains five
chitinase units of approximately 480 amino acids that are separated by
putative PEST-like, chitin-binding and mucin-like domains
(Royer et al., 2002
). It is
speculated that multi-modular chitinases may be expressed as zymogens that are
subsequently cleaved by proteolysis to reveal multiple active enzymes.
The occurrence of conserved acidic residues seems to be a common
characteristic for the active site of glycohydrolases
(Bourne and Henrissat, 2001;
Henrissat, 1990
). Since the
tertiary structure of family 18 chitinases is similar to that of other
glycohydrolases, a common mechanism of hydrolysis involving conserved acidic
amino acids was postulated (Henrissat and
Bairoch, 1993
). The signature sequence FDxxDxDxE is found in the
active sites of family 18 chitinases, including a glutamate residue that is
essential for catalysis. The highly conserved sequence YDFDGLDLDWEYP found in
insect chitinases is consistent with the family 18 chitinase signature
(Terwisscha van Scheltinga et al.,
1994
; Choi et al.,
1997
; de la Vega et al.,
1998
). Consequently, site-directed mutagenesis of the essential
glutamate of the insect chitinase active site results in a loss of enzymatic
activity (Huang et al., 2000
;
Lu et al., 2002
;
Royer et al., 2002
). Based on
crystallographic data and theoretical models, the catalytic reaction of family
18 chitinases might take place through a substrate-assisted, double
displacement mechanism with a geometrically deformed oxocarbonium
intermediate, more conveniently referred to as a `retention mechanism'
(Sinnott, 1990
;
Hart et al., 1995
;
Robertus and Monzingo, 1999
).
It is postulated that the active site has a binding cleft for a hexamer of
N-acetylglucosamine. Following a convention developed for hen
egg-white lysozyme, the single sugar binding sites were termed A-F
(Blake et al., 1967
;
Kelly et al., 1979
). In a
first step, the sugar in binding site D is distorted to a boat conformation.
Subsequently, the catalytic glutamate breaks the glycosidic bond between the
sugars in sites D and E by protonation of the leaving group. This leads to a
positively charged oxocarbonium intermediate that is stabilized by a covalent
bond between the carboxyl oxygen of the N-acetyl-group and the C1
atom of the sugar. The cleaving group then leaves the active site and a water
molecule enters and attacks the C1 carbon from the ß-side and protonates
the glutamate. This reaction results in the retention of the stereochemistry
at the anomeric carbon of the product, in contrast to the inverting mechanism
of family 19 chitinases (Robertus and
Monzingo, 1999
). The soluble products of this catalytic mechanism
are chitotetraose, chitotriose and chitobiose, the latter
chito-oligosaccharide being predominant
(Terwisscha van Scheltinga et al.,
1995
; Kramer and
Muthukrishnan, 1997
; Brameld
et al., 1998
; Robertus and
Monzingo, 1999
; Fukamizo,
2000
; Abdel-Banat et al.,
2002
). The functional importance of active site residues has also
been demonstrated for an insect chitinase
(Lu et al., 2002
).
Site-directed mutagenesis of the Manduca chitinase revealed that E146
may function as an acid/base catalyst while D142 may influence the
pKa values of the catalytic residue E146 but also that of D144; the
latter residue may be an electrostatic stabilizer of the positively charged
transition state. Moreover, W145, which is also present in all family 18
chitinases, might extend the alkaline pH range in which the enzyme is active
and may increase affinity to the substrate
(Huang et al., 2000
).
![]() |
Regulation of chitin degradation |
---|
In the course of the molting process, the ecdysteroid titer increases
continuously and reaches its maximum shortly before apolysis
(Bollenbacher et al., 1981;
Riddiford, 1994
). Juvenile
hormone allows larval molting in response to ecdysteroids but prevents the
switching of gene expression necessary for metamorphosis
(Riddiford, 1996
). Therefore,
the delayed chitinase expression may be caused by differential sensing of
hormone titers during molting. Both secretion and activation of chitinolytic
enzymes are clearly controlled by ecdysteroids
(Reynolds and Samuels, 1996
).
Concordantly, Kimura (1973b
)
showed some 30 years ago that the activity of molting fluid enzymes can be
stimulated by ecdysteroid injections.
The chitinases of Bombyx and Manduca are induced at high
hemolymph levels of 20-hydroxyecdysone, while
ß-N-acetylglucosaminidase is already induced at low levels of
the steroid (Fukamizo and Kramer,
1987; Koga et al.,
1991
,
1992
;
Kramer et al., 1993
;
Zen et al., 1996
). As a
consequence, ß-N-acetylglucosaminidase expression was found to
start before that of the chitinase. Although it is obvious that the expression
of chitinolytic enzymes is hormonally coordinated, responsive elements that
would affect gene transcription have not been identified so far. Nevertheless,
the chitinase may be an early responsive gene and a direct target of the
ecdysone receptor because protein synthesis is not required for its induction
(Royer et al., 2002
).
Degradation of cuticles by chitinolytic enzymes certainly needs the
assistance of molting fluid proteases to degrade proteinaceous components
(Law et al., 1977). However,
these proteases may also function in the proteolytic activation of inactive
chitinase precursors, as was suggested for Manduca or
Tenebrio (Kramer et al.,
1993
; Samuels and Reynolds,
1993
). Proteolytic activation of chitinases may also occur in
insect gut systems, where, in the case of blood-sucking insects, the
activities of gut chitinase and ß-N-acetylglucosaminidase are
found to rise shortly after feeding (Filho
et al., 2002
). Indeed, the gut-specific chitinase of
Anopheles was shown to be secreted into the gut lumen as an inactive
pro-enzyme that needs to be trypsinized in order to develop chitinolytic
activity (Shen and Jacobs-Lorena,
1997
). Interestingly, the malaria parasite Plasmodium
also utilizes the protease-rich environment of the mosquito midgut to increase
the enzymatic activity of its own chitinase, which facilitates penetration of
the peritrophic matrix (Shahabuddin et
al., 1993
).
Proteolytic activation of chitinases has been extensively investigated in
Manduca and Bombyx. A probable zymogenic form of the
chitinase with a molecular mass of 215 kDa was observed during the spinning
period of Bombyx. Two to three days later, when enzyme activity is
detectable, three active fragments of 88 kDa, 65 kDa and 54 kDa appear, which
may be the result of successive, proteolytic processing (Koga et al.,
1989,
1992
,
1997
;
Abdel-Banat et al., 1999
). The
three active forms have a common N-terminal sequence, indicating that they
differ in length at the C-terminus. It was suggested that the 88 kDa enzyme
still contains a potent chitin-binding domain. This domain, however, may get
gradually lost by further proteolysis from the C-terminal side, resulting in
chitinase variants that are more active on shorter substrates. Consistent with
this, when the chitin-binding domain is attached to the catalytic domain, the
resulting recombinant fusion protein exhibits increased activity towards the
insoluble polymer but not towards the soluble chitin oligosaccharide
(Arakane et al., 2003
). Thus,
it seems that the domain structure of insect chitinases has evolved to
optimize degradation of insoluble polysaccharides to soluble oligosaccharides,
thereby accelerating the overall chitin degradation rate in addition to the
presence of ß-N-acetylglucosaminidases
(Koga et al., 1997
;
Abdel-Banat et al., 1999
).
Similar results were observed in the Manduca integument, where a
119 kDa protein might be interpreted as a zymogenic precursor and several
smaller proteins as proteolytically activated fragments
(Koga et al., 1992). In
general, studies on the zymogenic nature of insect chitinases are complicated
by the observed discrepancies between theoretical molecular masses deduced
from obviously complete cDNA sequences and apparent molecular masses estimated
from SDS-PAGE. Since these discrepancies cannot be explained by
post-translational processing, it has been concluded that at least some insect
chitinases may exhibit an anomalous electrophoretic migration behavior
(Kramer et al., 1993
).
The chitinase PEST sequences could possibly enhance the activation of
chitinase zymogens (Royer et al.,
2002). Consistent with this, activity of the fat body-specific
chitinase from the tsetse fly, which lacks a PEST-like region, is not
increased by trypsinization (Yan et al.,
2002
). In contrast to chitinases, the exo-cleaving
ß-N-acetylglucosaminidases of Manduca and
Bombyx do not seem to be zymogens (Koga et al.,
1991
,
1992
,
1997
).
![]() |
Inhibition of chitin metabolism |
---|
Inhibitors of chitin synthesis have been classified into three major
groups: peptidyl nucleosides, acyl ureas and substances interfering with
hormonal control. Peptidyl nucleosides isolated from diverse
Streptomyces species act as substrate analogues and include polyoxins
and nikkomycins (Zhang and Miller,
1999). They competitively inhibit both fungal and insect chitin
synthases. It is believed that inhibition occurs via binding to the
catalytic site (Ruiz-Herrera and
San-Blas, 2003
). Polyoxins have found some applications in the
control of phytopathogens, whereas the commercial application of nikkomycins
is pending, although they seem to be more potent inhibitors than polyoxins
(Cohen and Casida, 1980a
;
Zhang and Miller, 1999
;
Tellam et al., 2000
).
Generally, the application of peptidyl nucleosides is complicated by low
permeability, hydrolytic lability, varying susceptibility of fungal species
and the multitude of responses found in animals
(Zhang and Miller, 1999
;
Ruiz-Herrera and San-Blas,
2003
).
In contrast to the peptidyl nucleosides, acyl ureas play an important role
in integrated pest management. Although it is well established that acyl ureas
such as diflubenzuron and teflubenzuron affect chitin synthesis
(Post et al., 1974;
van Eck, 1979
), their mode of
action is still puzzling. However, several lines of experiment argue against a
direct interaction of these inhibitors with the chitin synthase. For instance,
in cell-free systems, acyl ureas do not inhibit chitin synthesis
(Cohen and Casida, 1980a
;
Mayer et al., 1981
;
Cohen, 1985
). Instead of
directly blocking chitin synthase activity, they may alter either vesicle
transport or fusion, inhibit the translocation of chitin fibrils across the
plasma membrane (Nakagawa and Matsumura,
1994
; Cohen, 2001
)
or interfere with the hormonal regulation of chitin synthesis by influencing
ecdysteroid production (Fournet et al.,
1995
).
The third group of inhibitors evidently affects hormonal regulation of
insect growth and development. One of the manifold effects of these substances
is certainly deregulation of chitin synthesis, probably by preventing the
expression of the chitin synthase or regulating factors. Hormonal regulation
can already be disturbed at the level of hormone biosynthesis. Some synthetic
imidazole and cholesterol derivates have been shown to prevent ecdysteroid
biosynthesis (Kadano-Okuda et al.,
1987; Roussel,
1994
; Lorenz et al.,
1995
). By contrast, the heterocyclic compound brevioxime and the
alkaloid arborine show significant blocking of juvenile hormone synthesis
(Moya et al., 1997
;
Muthukrishnan et al., 1999
).
The auxiliary application of isolated molting hormones or their synthetic
agonists and antagonists leads to abnormalities in insect development as well.
The ecdysteroid agonist tebufenozide manifests its effect by interacting with
the ecdysone receptor, the juvenile hormone agonists fenoxycarb, methoprene
and pyriproxyfen mimic the hormone action, and the juvenile hormone
antagonists precocene I and II act via their cytotoxicities on the
corpora allata (Schooneveld,
1979
; Mulla et al.,
1985
; Dhadialla et al.,
1998
; Hoffmann and Lorenz,
1998
; Kostyukovsky et al.,
2000
; Retnakaran et al.,
2001
).
Chitinase inhibitors can generally be grouped into two major classes: they
mimic either carbohydrate substrates or the oxocarbonium reaction intermediate
of family 18 chitinases. The most-studied chitinase inhibitor is allosamidin,
a pseudotrisaccharide. It was isolated from the mycelium of
Streptomyces sp. and exhibits a strong inhibitory activity against
family 18 chitinases of insects and fungi with a Ki in the
nano- to micromolar range (Sakuda et al.,
1987; Blattner et al.,
1996
; Berecibar et al.,
1999
). Most strikingly, it blocks malaria parasite transmission by
inhibiting the chitinase of Plasmodium that is essential to penetrate
the host's peritrophic matrix
(Shahabuddin et al., 1993
;
Tsai et al., 2001
;
Filho et al., 2002
). The
structural basis of interactions between the inhibitor and several family 18
chitinases has been solved by x-ray crystallography
(Terwisscha van Scheltinga et al.,
1995
; van Aalten et al.,
2001
). As a result of these studies, it was proposed that
allosamidin mimics the catalytic transition state. Allosamidin consists of two
N-acetylallosamine sugars linked to an allosamizoline that may
resemble the catalytic intermediate and cannot be hydrolyzed because it lacks
the pyranose oxygen (Bortone et al.,
2002
; Fusetti et al.,
2002
; Rao et al.,
2003
).
Allosamidin is a potent chitinase inhibitor; however, its production is
expensive because it is difficult to synthesize. A new, alternative class of
inhibitors includes the cyclopentapeptides argifin and argadin. These
molecules are as potent inhibitors as allosamidin but synthesis by peptide
chemistry is less expensive (Arai et al.,
2000; Omura et al.,
2000
; Houston et al.,
2002b
). While the cyclopentapeptides are carbohydrate mimics, the
small peptide CI-4, which was recently identified in the marine bacterium
Pseudomonas, functions like allosamidin as a mimic of the family 18
chitinases' catalytic transient state
(Izumida et al., 1996
;
Houston et al., 2002a
).
![]() |
Conclusion and outlook |
---|
So far, no cellular interaction partners for chitin synthases or chitinases are known in insects. Since protein-protein interactions are presumably essential for the regulation of enzyme biosynthesis, targeting and activity, identification of interacting proteins would provide new insights into cellular control mechanisms. To obtain first clues, the application of yeast two- or three-hybrid systems may yield putative binding partners, which have to be further analyzed regarding their binding capability by biochemical or cytological methods. Those proteins that interact either with chitin synthases or chitinases may again turn out to be suitable target sites for future biocides. In conclusion, understanding of the basic principles underlying insect chitin metabolism and its regulation will open up new vistas in pest management.
![]() |
Acknowledgments |
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References |
---|
Abdel-Banat, B. M., Kameyama, Y., Yoshioka, T. and Koga, D. (1999). Purification and characterization of a 54 kDa chitinase from Bombyx mori. Insect Biochem. Mol. Biol. 29,537 -547.[CrossRef]
Abdel-Banat, B. M., Zhou, W., Karasuda, S. and Koga, D. (2002). Analysis of hydrolytic activity of a 65-kDa chitinase from the silkworm, Bombyx mori. Biosci. Biotechnol. Biochem. 66,1119 -1122.[CrossRef][Medline]
Andersen, S. O. (1979). Biochemistry of the insect cuticle. Annu. Rev. Entomol. 24, 29-61.[CrossRef]
Andersen, S. O., Hojrup, P. and Roepstorff, P. (1995). Insect cuticular proteins. Insect Biochem. Mol. Biol. 25,153 -176.[CrossRef][Medline]
Andersen, S. O. and Weis-Fogh, T. (1964). Resilin. A rubberlike protein in arthropod cuticle. Adv. Insect Physiol. 2,1 -66.
Andres, A. J. and Cherbas, P. (1994). Tissue-specific regulation by ecdysone: distinct patterns of Eip28/29 expression are controlled by different ecdysone response elements. Dev. Genet. 15,320 -331.[Medline]
Antoniewski, C., Laval, M. and Lepesant, J. A. (1993). Structural features critical to the activity of an ecdysone receptor binding site. Insect Biochem. Mol. Biol. 23,105 -114.[CrossRef][Medline]
Apple, R. T. and Fristrom, J. W. (1991). 20-Hydroxyecdysone is required for, and negatively regulates, transcription of Drosophila pupal cuticle protein genes. Dev. Biol. 146,569 -582.[Medline]
Arai, N., Shiomi, K., Yamaguchi, Y., Masuma, R., Iwai, Y., Turberg, A., Kolbl, H. and Omura, S. (2000). Argadin, a new chitinase inhibitor, produced by Clonostachys sp. FO-7314. Chem. Pharm. Bull. 48,1442 -1446.[Medline]
Arakane, Y., Zhu, Q., Matsumiya, M., Muthukrishnan, S. and Kramer, K. J. (2003). Properties of catalytic, linker and chitin-binding domains of insect chitinase. Insect Biochem. Mol. Biol. 33,631 -648.[CrossRef][Medline]
Armand, S., Tomita, H., Heyraud, A., Gey, C., Watanabe, T. and Henrissat, B. (1994). Stereochemical course of the hydrolysis reaction catalyzed by chitinases A1 and D from Bacillus circulans WL-12. FEBS Lett. 343,177 -180.[CrossRef][Medline]
Aronson, N. N., Blanchard, C. J. and Madura, J. D. (1997). Homology modeling of glycosyl hydrolase family 18 enzymes and proteins. J. Chem. Inf. Comput. Sci. 37,999 -1005.[CrossRef][Medline]
Bade, M. L. (1983). Chitin biosynthesis: does it involve a lipid bound intermediate. J. Appl. Polym. Sci. 37,165 -178.
Bairoch, A., Bucher, P. and Hofmann, K. (1997).
The PROSITE database, its status in 1997. Nucleic Acids
Res. 25,217
-221.
Baker, F. C., Tsai, L. W., Reuter, C. C. and Schooley, D. A. (1987). In vivo fluctuation of JH, JH acid, and ecdysteroid titer, and JH esterase activity during development of fifth stadium Manduca sexta. Insect Biochem. 17,989 -996.[CrossRef]
Bakkers, J., Kijne, J. W. and Spaink, H. P. (1999). Function of chitin oligosaccharides in plant and animal development. EXS 87,71 -83.[Medline]
Berecibar, A., Grandjean, C. and Siriwardena, A. (1999). Synthesis and biological activity of natural aminocyclopentitol glycosidase inhibitors: mannostatins, trehazolin, allosamidins, and their analogues. Chem. Rev. 99,779 -844.[CrossRef][Medline]
Berger, B., Wilson, D. B., Wolf, E., Tonchev, T., Milla, M. and Kim, P. S. (1995). Predicting coiled coils by use of pairwise residue correlations. Proc. Natl. Acad. Sci. USA 92,8259 -8263.[Abstract]
Binnington, K. C. (1985). Ultrastructural changes in the cuticle of the sheep blowfly, Lucilia, induced by certain insecticides and biological inhibitors. Tissue Cell 17,131 -140.[Medline]
Blake, C. C., Johnson, L. N., Mair, G. A., North, A. C., Phillips, D. C. and Sarma, V. R. (1967). Crystallographic studies of the activity of hen egg-white lysozyme. Proc. R. Soc. Lond. B Biol. Sci. 167,378 -388.[Medline]
Blattner, R., Furneaux, R. H. and Lynch, G. P. (1996). Synthesis of allosamidin analogues. Carbohydr Res. 294,29 -39.[CrossRef][Medline]
Bollenbacher, W. E., Smith, S. L., Goodman, W. and Gilbert, L. I. (1981). Ecdysteroid titer during larval-pupal-adult development of the tobacco hornworm, Manduca sexta. Gen. Comp. Endocrinol. 44,302 -306.[Medline]
Boot, R. G., Renkema, G. H., Strijland, A., van Zonneveld, A. J.
and Aerts, J. M. (1995). Cloning of a cDNA encoding
chitotriosidase, a human chitinase produced by macrophages. J.
Biol. Chem. 270,26252
-26256.
Boot, R. G., Renkema, G. H., Verhoek, M., Strijland, A., Bliek,
J., de Meulemeester, T. M., Mannens, M. M. and Aerts, J. M.
(1998). The human chitotriosidase gene. Nature of inherited
enzyme deficiency. J. Biol. Chem.
273,25680
-25685.
Bortone, K., Monzingo, A. F., Ernst, S. and Robertus, J. D. (2002). The structure of an allosamidin complex with the Coccidioides immitis chitinase defines a role for a second acid residue in substrate-assisted mechanism. J. Mol. Biol. 320,293 -302.[CrossRef][Medline]
Bouligand, Y. (1972). Twisted fibrous arrangements in biological materials and cholesteric mesophases. Tissue Cell 4,189 -217.[Medline]
Bourne, Y. and Henrissat, B. (2001). Glycoside hydrolases and glycosyltransferases: families and functional modules. Curr. Opin. Struct. Biol. 11,593 -600.[CrossRef][Medline]
Bownes, M., Ronaldson, E. and Mauchline, D. (1996). 20-Hydroxyecdysone, but not juvenile hormone, regulation of yolk protein gene expression can be mapped to cis-acting DNA sequences. Dev. Biol. 173,475 -489.[CrossRef][Medline]
Bracker, C. E., Ruiz-Herrera, J. and Bartnicki-Garcia, S. (1976). Structure and transformation of chitin synthetase particles (chitosomes) during microfibril synthesis in vitro. Proc. Natl. Acad. Sci. USA 73,4570 -4574.[Abstract]
Brameld, K. A., Shrader, W. D., Imperiali, B. and Goddard, W. A., III (1998). Substrate assistance in the mechanism of family 18 chitinases: theoretical studies of potential intermediates and inhibitors. J. Mol. Biol. 280,913 -923.[CrossRef][Medline]
Breton, C., Mucha, J. and Jeanneau, C. (2001). Structural and functional features of glycosyltransferases. Biochimie 83,713 -718.[CrossRef][Medline]
Burkhard, P., Stetefeld, J. and Strelkov, S. V. (2001). Coiled coils: a highly versatile protein folding motif. Trends Cell Biol. 11,82 -88.[CrossRef][Medline]
Candy, D. J. and Kilby, B. A. (1962). Studies on chitin synthesis in the desert locust. J. Exp. Biol. 39,129 -140.
Carlini, C. R. and Grossi-de-Sa, M. F. (2002). Plant toxic proteins with insecticidal properties. A review on their potentialities as bioinsecticides. Toxicon 40,1515 -1539.[CrossRef][Medline]
Carlson, J. R. and Bentley, D. (1977). Ecdysis: neural orchestration of a complex behavioral performance. Science 195,1006 -1008.[Medline]
Cecchelli, R., Cacan, R. and Verbert, A. (1986). Mechanism of UDP-sugar transport into intracellular vesicles. Occurrence of UDP-GlcNAc/UDP and UDP-Gal/UDP antiports. FEBS Lett. 208,407 -412.[CrossRef][Medline]
Chang, R., Yeager, A. R. and Finney, N. S. (2003). Probing the mechanism of a fungal glycosyltransferase essential for cell wall biosynthesis. UDP-chitobiose is not a substrate for chitin synthase. Org. Biomol. Chem. 1, 39-41.[CrossRef][Medline]
Charpentier, M. and Percheron, F. (1983). The chitin-degrading enzyme system of a Streptomyces species. Int. J. Biochem. 15,289 -292.[CrossRef][Medline]
Choi, H. K., Choi, K. H., Kramer, K. J. and Muthukrishnan, S. (1997). Isolation and characterization of a genomic clone for the gene of an insect molting enzyme, chitinase. Insect Biochem. Mol. Biol. 27,37 -47.[CrossRef][Medline]
Cioffi, M. (1979). The morphology and fine structure of the larval midgut of a moth (Manduca sexta) in relation to active ion transport. Tissue Cell 11,467 -479.[CrossRef][Medline]
Cohen, E. (1982). In vitro chitin synthesis in an insect: formation and structure of microfibrils. Eur. J. Cell Biol. 26,289 -294.[Medline]
Cohen, E. (1985). Chitin synthetase activity and inhibition in different insect microsomal preparations. EXS 41,470 -472.
Cohen, E. (2001). Chitin synthesis and inhibition: a revisit. Pest Manag. Sci. 57,946 -950.[CrossRef][Medline]
Cohen, E. and Casida, J. E. (1980a). Inhibition of Tribolium gut chitin synthetase. Pestic. Biochem. Physiol. 13,129 -136.
Cohen, E. and Casida, J. E. (1980b). Properties of Tribolium gut chitin synthetase. Pestic. Biochem. Physiol. 13,121 -128.
Cohen, E. and Casida, J. E. (1982). Properties and inhibition of insect integumental chitin synthetase. Pestic. Biochem. Physiol. 17,301 -306.
Coutinho, P. M., Deleury, E., Davies, G. J. and Henrissat, B. (2003). An evolving hierarchical family classification for glycosyltransferases. J. Mol. Biol. 328,307 -317.[CrossRef][Medline]
Dahn, U., Hagenmaier, H., Hohne, H., Konig, W. A., Wolf, G. and Zahner, H. (1976). Stoffwechselprodukte von Mikroorganismen. 154. Mitteilung. Nikkomycin, ein neuer Hemmstoff der Chitinsynthese bei Pilzen. Arch. Microbiol. 107,143 -160.[Medline]
de la Vega, H., Specht, C. A., Liu, Y. and Robbins, P. W. (1998). Chitinases are a multi-gene family in Aedes, Anopheles and Drosophila. Insect Mol. Biol. 7, 233-239.[Medline]
DeAngelis, P. L., Yang, N. and Weigel, P. H. (1994). The Streptococcus pyogenes hyaluronan synthase: sequence comparison and conservation among various group A strains. Biochem. Biophys. Res. Commun. 199, 1-10.[CrossRef][Medline]
Dhadialla, T. S., Carlson, G. R. and Le, D. P. (1998). New insecticides with ecdysteroidal and juvenile hormone activity. Annu. Rev. Entomol. 43,545 -569.[CrossRef][Medline]
Ding, X., Gopalakrishnan, B., Johnson, L. B., White, F. F., Wang, X., Morgan, T. D., Kramer, K. J. and Muthukrishnan, S. (1998). Insect resistance of transgenic tobacco expressing an insect chitinase gene. Transgenic Res. 7, 77-84.[CrossRef][Medline]
Doblin, M. S., Kurek, I., Jacob-Wilk, D. and Delmer, D. P.
(2002). Cellulose biosynthesis in plants: from genes to rosettes.
Plant Cell Physiol. 43,1407
-1420.
Dorstyn, L., Colussi, P. A., Quinn, L. M., Richardson, H. and
Kumar, S. (1999). DRONC, an ecdysone-inducible
Drosophila caspase. Proc. Natl. Acad. Sci.
USA 96,4307
-4312.
Drouillard, S., Armand, S., Davies, G. J., Vorgias, C. E. and Henrissat, B. (1997). Serratia marcescens chitobiase is a retaining glycosidase utilizing substrate acetamido group participation. Biochem. J. 328,945 -949.[Medline]
Duran, A., Bowers, B. and Cabib, E. (1975). Chitin synthetase zymogen is attached to the yeast plasma membrane. Proc. Natl. Acad. Sci. USA 72,3952 -3955.[Abstract]
Duran, A. and Cabib, E. (1978). Solubilization and partial purification of yeast chitin synthetase. Confirmation of the zymogenic nature of the enzyme. J. Biol. Chem. 253,4419 -4425.[Abstract]
Dziadik-Turner, C., Koga, D., Mai, M. S. and Kramer, K. J. (1981). Purification and characterization of two ß-N-acetylhexosaminidases from the tobacco hornworm, Manduca sexta (L.) (Lepidoptera:Sphingidae). Arch. Biochem. Biophys. 212,546 -560.[Medline]
Easterby, J. S. (1973). Coupled enzyme assays: a general expression for the transient. Biochim. Biophys. Acta 293,552 -558.[Medline]
Elango, N., Correa, J. U. and Cabib, E. (1982).
Secretory character of yeast chitinase. J. Biol. Chem.
257,1398
-1400.
Feix, M., Gloggler, S., Londershausen, M., Weidemann, W., Spindler, K. D. and Spindler-Barth, M. (2000). A cDNA encoding a chitinase from the epithelial cell line of Chironomus tentans (Insecta, diptera) and its functional expression. Arch. Insect Biochem. Physiol. 45, 24-36.[CrossRef][Medline]
Filho, B. P., Lemos, F. J., Secundino, N. F., Pascoa, V., Pereira, S. T. and Pimenta, P. F. (2002). Presence of chitinase and beta-N-acetylglucosaminidase in the Aedes aegypti: a chitinolytic system involving peritrophic matrix formation and degradation. Insect Biochem. Mol. Biol. 32,1723 -1729.[CrossRef][Medline]
Florez-Martinez, A., Lopez-Romero, E., Martinez, J. P., Bracker, C. E., Ruiz-Herrera, J. and Bartnicki-Garcia, S. (1990). Protein composition of purified chitosomes of Mucor rouxii. Exp. Mycol. 14,160 -168.
Fournet, F., Sannier, C., Moriniere, M., Porcheron, P. and Monteny, N. (1995). Effects of two insect growth regulators on ecdysteroid production in Aedes aegypti (Diptera: Culicidae). J. Med. Entomol. 32,588 -593.[Medline]
Fristrom, J. W., Doctor, J., Fristrom, D. K., Logan, W. R. and Silvert, D. J. (1982). The formation of the pupal cuticle by Drosophila imaginal discs in vitro. Dev. Biol. 91,337 -350.[Medline]
Fukamizo, T. (2000). Chitinolytic enzymes: catalysis, substrate binding, and their application. Curr. Protein Pept. Sci. 1,105 -124.[Medline]
Fukamizo, T. and Kramer, K. J. (1985a). Mechanism of chitin hydrolysis by the binary chitinase system in insect moulting fluid. Insect Biochem. 15,141 -145.[CrossRef]
Fukamizo, T. and Kramer, K. J. (1985b). Mechanism of chitin oligosaccharide hydrolysis by the binary enzyme chitinase system in insect moulting fluid. Insect Biochem. 15, 1-7.[CrossRef]
Fukamizo, T. and Kramer, K. J. (1987). Effect of 20-hydroxyecdysone on chitinase and ß-N-acetylglucosaminidase during the larval-pupal transformation of Manduca sexta (L.). Insect Biochem. 17,547 -550.[CrossRef]
Fusetti, F., von Moeller, H., Houston, D., Rozeboom, H. J.,
Dijkstra, B. W., Boot, R. G., Aerts, J. M. and van Aalten, D. M.
(2002). Structure of human chitotriosidase. Implications for
specific inhibitor design and function of mammalian chitinase-like lectins.
J. Biol. Chem. 277,25537
-25544.
Gagou, M. E., Kapsetaki, M., Turberg, A. and Kafetzopoulos, D. (2002). Stage-specific expression of the chitin synthase DmeChSA and DmeChSB genes during the onset of Drosophila metamorphosis. Insect Biochem. Mol. Biol. 32,141 -146.[CrossRef][Medline]
Geremia, R. A., Mergaert, P., Geelen, D., Van Montagu, M. and Holsters, M. (1994). The NodC protein of Azorhizobium caulinodans is an N-acetylglucosaminyltransferase. Proc. Natl. Acad. Sci. USA 91,2669 -2673.[Abstract]
Girard, C. and Jouanin, L. (1999). Molecular cloning of a gut-specific chitinase cDNA from the beetle Phaedon cochleariae. Insect Biochem. Mol. Biol. 29,549 -556.[CrossRef][Medline]
Giraud-Guille, M. M. and Bouligand, Y. (1986). Chitin-protein molecular organization in arthropods. In Chitin in Nature and Technology (ed. R. Muzzarelli, C. Jeuniaux and G. W. Gooday), pp. 29-35. New York: Plenum Press.
Glaser, L. and Brown, D. H. (1957). The
synthesis of chitin in cell-free extracts of Neurospora crassa. J.
Biol. Chem. 228,729
-742.
Gooday, G. W. (1972). The effect of polyoxin D on morphogenesis in Coprinus cinereus. Biochem. J. 129,17P -18P.
Gooday, G. W. (1999). Agressive and defensive roles for chitinases. In Chitin and Chitinases (ed. R. A. A. Muzzarelli and P. Jolles), pp. 157-169. Basel: Birkhäuserverlag.
Gopalakrishnan, B., Muthukrishnan, S. and Kramer, K. J. (1995). Baculovirus-mediated expression of a Manduca sexta chitinase gene: properties of the recombinant protein. Insect Biochem. Mol. Biol. 25,255 -265.[CrossRef]
Hansen, J. E., Lund, O., Rapacki, K. and Brunak, S.
(1997). O GLYCBASE version 2.0: a revised database of
O-glycosylated proteins. Nucleic Acids Res.
25,278
-282.
Hardy, J. C. and Gooday, G. W. (1983). Stability and zymogenic nature of chitin synthase from Candida albicans.Curr. Microbiol. 9,51 -54.
Hart, P. J., Ready, M. P. and Robertus, J. D. (1992). Crystallization of an endochitinase from Hordeum vulgare L. seeds. J. Mol. Biol. 225,565 -567.[Medline]
Hart, P. J., Pfluger, H. D., Monzingo, A. F., Hollis, T. and Robertus, J. D. (1995). The refined crystal structure of an endochitinase from Hordeum vulgare L. seeds at 1.8 Å resolution. J. Mol. Biol. 248,402 -413.[CrossRef][Medline]
Hawkins, C. J., Yoo, S. J., Peterson, E. P., Wang, S. L.,
Vernooy, S. Y. and Hay, B. A. (2000). The Drosophila
caspase DRONC cleaves following glutamate or aspartate and is regulated by
DIAP1, HID, and GRIM. J. Biol. Chem.
275,27084
-27093.
Hawtin, R. E., Arnold, K., Ayres, M. D., Zanotto, P. M., Howard, S. C., Gooday, G. W., Chappell, L. H., Kitts, P. A., King, L. A. and Possee, R. D. (1995). Identification and preliminary characterization of a chitinase gene in the Autographa californica nuclear polyhedrosis virus genome. Virology 212,673 -685.[CrossRef][Medline]
Heifetz, A., Keenan, R. W. and Elbein, A. D. (1979). Mechanism of action of tunicamycin on the UDP-GlcNAc:dolichyl-phosphate Glc-NAc-1-phosphate transferase. Biochemistry 18,2186 -2192.[Medline]
Henrissat, B. (1990). Weak sequence homologies among chitinases detected by clustering analysis. Protein Seq. Data Anal. 3,523 -526.[Medline]
Henrissat, B. (1991). A classification of glycosyl hydrolases based on amino acid sequence similarities. Biochem. J. 280,309 -316.[Medline]
Henrissat, B. (1999). Classification of chitinases modules. EXS 87,137 -156.[Medline]
Henrissat, B. and Bairoch, A. (1993). New families in the classification of glycosyl hydrolases based on amino acid sequence similarities. Biochem. J. 293,781 -788.[Medline]
Hernandez, J., Lopez-Romero, E., Cerbon, J. and Ruiz-Herrera, J. (1981). Lipid analysis of chitosomes, chitin synthesizing microvesicles from Mucor rouxii. Exp. Mycol. 5, 349-356.
Herrera-Estrella, A. and Chet, I. (1999). Chitinases in biological control. EXS 87,171 -184.[Medline]
Hiruma, K., Carter, M. S. and Riddiford, L. M. (1995). Characterization of the dopa decarboxylase gene of Manduca sexta and its suppression by 20-hydroxyecdysone. Dev. Biol. 169,195 -209.[CrossRef][Medline]
Hiruma, K., Hardie, J. and Riddiford, L. M. (1991). Hormonal regulation of epidermal metamorphosis in vitro: control of expression of a larval-specific cuticle gene. Dev. Biol. 144,369 -378.[Medline]
Hoffmann, K. H. and Lorenz, M. W. (1998). Recent advances on hormones in insect pest control. Phytoparasitica 26,323 -330.
Hojrup, P., Andersen, S. O. and Roepstorff, P. (1986). Isolation, characterization, and N-terminal sequence studies of cuticular proteins from the migratory locust, Locusta migratoria. Eur. J. Biochem. 154,153 -159.[Abstract]
Hopkins, T. L. and Harper, M. S. (2001). Lepidopteran peritrophic membranes and effects of dietary wheat germ agglutinin on their formation and structure. Arch. Insect Biochem. Physiol. 47,100 -109.[CrossRef][Medline]
Horsch, M., Mayer, C. and Rast, D. M. (1996). Stereochemical requirements of chitin synthase for ligand binding at the allosteric site for N-acetylglucosamine. Eur. J. Biochem. 237,476 -482.[Abstract]
Horst, M. N. (1983). The biosynthesis of crustacean chitin. Isolation and characterization of polyprenol-linked intermediates from brine shrimp microsomes. Arch. Biochem. Biophys. 223,254 -263.[Medline]
Horst, M. N. and Walker, A. N. (1993). Crustacean chitin synthesis and the role of the Golgi apparatus: in vivo and in vitro studies. In Chitin Enzymology, vol. 1 (ed. R. A. A. Muzzarelli), pp. 109-118. Atec: Grottammare.
Houston, D. R., Eggleston, I., Synstad, B., Eijsink, V. G. and van Aalten, D. M. (2002a). The cyclic dipeptide CI-4 [cyclo-(l-Arg-d-Pro)] inhibits family 18 chitinases by structural mimicry of a reaction intermediate. Biochem. J. 368, 23-27.[CrossRef][Medline]
Houston, D. R., Shiomi, K., Arai, N., Omura, S., Peter, M. G.,
Turberg, A., Synstad, B., Eijsink, V. G. and van Aalten, D. M.
(2002b). High-resolution structures of a chitinase complexed with
natural product cyclopentapeptide inhibitors: mimicry of carbohydrate
substrate. Proc. Natl. Acad. Sci. USA
99,9127
-9132.
Huang, X., Zhang, H., Zen, K. C., Muthukrishnan, S. and Kramer, K. J. (2000). Homology modeling of the insect chitinase catalytic domain-oligosaccharide complex and the role of a putative active site tryptophan in catalysis. Insect Biochem. Mol. Biol. 30,107 -117.[CrossRef][Medline]
Ibrahim, G. H., Smartt, C. T., Kiley, L. M. and Christensen, B. M. (2000). Cloning and characterization of a chitin synthase cDNA from the mosquito Aedes aegypti. Insect Biochem. Mol. Biol. 30,1213 -1222.[CrossRef][Medline]
Izumida, H., Imamura, N. and Sano, H. (1996). A novel chitinase inhibitor from a marine bacterium, Pseudomonas sp. J. Antibiot. 49,76 -80.[Medline]
Jaworski, E., Wang, L. and Margo, G. (1963). Synthesis of chitin in cell-free extracts of Prodenia eridania.Nature 198,790 .[Medline]
Jindra, M., Malone, F., Hiruma, K. and Riddiford, L. M. (1996). Developmental profiles and ecdysteroid regulation of the mRNAs for two ecdysone receptor isoforms in the epidermis and wings of the tobacco hornworm, Manduca sexta. Dev. Biol. 180,258 -272.[CrossRef][Medline]
Jones, D., Wache, S. and Chhokar, V. (1996). Toxins produced by arthropod parasites: salivary gland proteins of human body lice and venom proteins of chelonine wasps. Toxicon 34,1421 -1429.[CrossRef][Medline]
Kadano-Okuda, K., Kuwano, E., Eto, M. and Yamashita, O. (1987). Inhibitory action of an imidazole compound on ecdysone synthesis in prothoracic glands of the silkworm, Bombyx mori.Growth Diff. 29,527 -533.
Kang, M. S., Elango, N., Mattia, E., Au-Young, J., Robbins, P.
W. and Cabib, E. (1984). Isolation of chitin synthetase from
Saccharomyces cerevisiae. Purification of an enzyme by entrapment in
the reaction product. J. Biol. Chem.
259,14966
-14972.
Kaznowski, C., Schneiderman, H. A. and Bryant, P. J. (1986). The incorporation of precursors into Drosophila larval cuticle. J. Insect Physiol. 32,133 -142.[CrossRef]
Keller, F. A. and Cabib, E. (1971). Chitin and
yeast budding. Properties of chitin synthetase from Saccharomyces
carlsbergensis. J. Biol. Chem.
246,160
-166.
Kelly, J. A., Sielecki, A. R., Sykes, B. D., James, M. N. and Phillips, D. C. (1979). X-ray crystallography of the binding of the bacterial cell wall trisaccharide NAM-NAG-NAM to lysozyme. Nature 282,875 -578.[Medline]
Kenchington, W. (1976). Adaption of insect peritrophic membranes to form cocoon fabrics. In The Insect Integument (ed. H. R. Hepburn), pp.497 -513. Amsterdam: Elsevier Science.
Kim, M. G., Shin, S. W., Bae, K. S., Kim, S. C. and Park, H. Y. (1998). Molecular cloning of chitinase cDNAs from the silkworm, Bombyx mori and the fall webworm, Hyphantria cunea.Insect Biochem. Mol. Biol. 28,163 -171.[CrossRef][Medline]
Kimura, S. (1973a). Chitinolytic enzymes in the larval development of the silkworm, Bombyx mori. Appl. Ent. Zool. 8,234 -236.
Kimura, S. (1973b). The control of chitinase activity by ecdysterone in larvae of Bombyx mori. J. Insect Physiol. 19,115 -123.[CrossRef]
Kimura, S. (1977). Exo-ß-N-acetylglucosaminidase and chitobiase in Bombyx mori.Insect Biochem. 7,237 -245.[CrossRef]
Kingan, T. G. and Adams, M. E. (2000).
Ecdysteroids regulate secretory competence in Inka cells. J. Exp.
Biol. 203,3011
-3018.
Klesov, A. A. and Grigorash, S. Y. (1982). Kinetic mechanisms of hydrolysis of insoluble cellulose by multienzyme cellulase systems under nonsteady-state reaction conditions. Biokhimiia 42,198 -213.
Koga, D., Fujimoto, H., Funakoshi, T., Utsumi, T. and Ide, A. (1989). Appearance of chitinolytic enzymes in integument of Bombyx mori during the larval-pupal transformation. Evidence for zymogenic forms. Insect Biochem. 19,123 -128.[CrossRef]
Koga, D., Funakoshi, T., Fujimoto, H., Kuwano, E., Eto, M. and Ide, A. (1991). Effects of 20-hydroxyecdysone and KK-42 on chitinase and ß-N-acetylglucosaminidase during the larval-pupal transformation of Bombyx mori. Insect Biochem. 21,277 -284.[CrossRef]
Koga, D., Funakoshi, T., Mizuki, K., Ide, A., Kramer, K. J., Zen, K. C., Choi, H. and Muthukrishnan, S. (1992). Immunoblot analysis of chitinolytic enzymes in integument and molting fluid of the silkworm, Bombyx mori, and the tobacco hornworm, Manduca sexta.Insect Biochem. Mol. Biol. 22,305 -311.[CrossRef]
Koga, D., Jilka, J. and Kramer, K. J. (1983). Insect endochitinases: glycoproteins from moulting fluid, integument and pupal haemolymph of Manduca sexta L. Insect Biochem. 13,295 -305.[CrossRef]
Koga, D., Mai, M. S., Dziadik-Turner, C. and Kramer, K. J. (1982). Kinetics and Mechanism of exochitinase and ß-N-acetylhexosaminidase from the tobacco hornworm, Manduca sexta L. (Lepidoptera: Sphingidae). Insect Biochem. 12,493 -499.[CrossRef]
Koga, D., Sasaki, Y., Uchiumi, Y., Hirai, N., Arakane, Y. and Nagamatsu, Y. (1997). Purification and characterization of Bombyx mori chitinases. Insect Biochem. Mol. Biol. 27,757 -767.[CrossRef][Medline]
Kondo, K., Matsumoto, M., Kojo, A. and Mauda, R. (2002). Purification and characterization of chitinase from pupae Pieris rapae crucivora (Boiduval). J. Chem. Eng. Jap. 35,241 -246.[CrossRef]
Kostyukovsky, M., Chen, B., Atsmi, S. and Shaaya, E. (2000). Biological activity of two juvenoids and two ecdysteroids against three stored product insects. Insect Biochem. Mol. Biol. 30,891 -897.[CrossRef][Medline]
Kramer, K. J., Corpuz, L., Choi, H. K. and Muthukrishnan, S. (1993). Sequence of a cDNA and expression of the gene encoding epidermal and gut chitinases of Manduca sexta. Insect Biochem. Mol. Biol. 23,691 -701.[CrossRef][Medline]
Kramer, K. J., Hopkins, T. L. and Schaefer, J. (1995). Applications of solids NMR to the analysis of insect sclerotized structures. Insect Biochem. Mol. Biol. 25,1067 -1080.[CrossRef]
Kramer, K. J. and Koga, D. (1986). Insect chitin: physical state, synthesis, degradation and metabolic regulation. Insect Biochem. 16,851 -877.[CrossRef]
Kramer, K. J. and Muthukrishnan, S. (1997). Insect chitinases: molecular biology and potential use as biopesticides. Insect Biochem. Mol. Biol. 27,887 -900.[CrossRef][Medline]
Krishnan, A., Nair, P. N. and Jones, D. (1994).
Isolation, cloning, and characterization of new chitinase stored in active
form in chitin-lined venom reservoir. J. Biol. Chem.
269,20971
-20976.
Krogh, A., Larsson, B., von Heijne, G. and Sonnhammer, E. L. L. (2001). Predicting transmembrane protein topology with a hidden Markov model: application to complete genomes. J. Mol. Biol. 305,567 -580.[CrossRef][Medline]
Kuranda, M. J. and Robbins, P. W. (1991).
Chitinase is required for cell separation during growth of Saccharomyces
cerevisiae. J. Biol. Chem.
266,19758
-19767.
Kurek, I., Kawagoe, Y., Jacob-Wilk, D., Doblin, M. and Delmer,
D. (2002). Dimerization of cotton fiber cellulose synthase
catalytic subunits occurs via oxidation of the zinc-binding domains.
Proc. Natl. Acad. Sc.i USA
99,11109
-11114.
Lasters, I., Wodak, S. J., Alard, P. and van Cutsem, E. (1988). Structural principles of parallel beta-barrels in proteins. Proc. Natl. Acad. Sci. USA 85,3338 -3342.[Abstract]
Law, J. H., Dunn, P. E. and Kramer, K. J. (1977). Insect proteases and peptidases. Adv. Enzymol. Relat. Areas Mol. Biol. 45,389 -425.[Medline]
Leah, R., Tommerup, H., Svendsen, I. and Mundy, J.
(1991). Biochemical and molecular characterization of three
barley seed proteins with antifungal properties. J. Biol.
Chem. 266,1564
-1573.
Lehane, M. J. (1997). Peritrophic matrix structure and function. Annu. Rev. Entomol. 42,525 -550.[CrossRef]
Locke, M. (1991). Insect epidermal cells. In Physiology of the Insect Epidermis (ed. K. Binnington and A. Retnakaran), pp. 1-22. Melbourne: CRISCO Publ.
Locke, M. and Huie, P. (1979). Apolysis and the turnover of plasma membrane plaques during cuticle formation in an insect. Tissue Cell 11,277 -291.[CrossRef][Medline]
Londershausen, M., Kammann, V., Spindler-Barth, M., Spindler, K. D. and Thomas, H. (1988). Chitin synthesis in insect cell lines. Insect Biochem. 18,631 -636.[CrossRef]
Lorenz, J., Lenz, M. and Hoffmann, K. H. (1995). Effects of pharmacological agents on ecdysteroid synthesis in vitro in ovaries and abdominal integument from female adult crickets, Gryllus bimaculatus de Geer (Ensifera, Gryllidae). Z. Naturforsch. 50,286 -293.
Lu, Y. M., Zen, K. C., Muthukrishnan, S. and Kramer, K. J. (2002). Site-directed mutagenesis and functional analysis of active site acidic amino acid residues D142, D144 and E146 in Manduca sexta (tobacco hornworm) chitinase. Insect Biochem. Mol. Biol. 32,1369 -1382.[CrossRef][Medline]
Luo, Y., Amin, J. and Voellmy, R. (1991). Ecdysterone receptor is a sequence-specific transcription factor involved in the developmental regulation of heat shock genes. Mol. Cell. Biol. 11,3660 -3675.[Medline]
Machida, S. and Saito, M. (1993). Purification
and characterization of membrane-bound chitin synthase. J. Biol.
Chem. 268,1702
-1707.
Marks, E. P. (1972). Effects of ecdysterone on the deposition of cockroach cuticle in vitro. Biol. Bull. 142,293 -301.[Medline]
Marks, E. P., Balke, J. and Klosterman, H. (1984). Evidence for chitin synthesis in an insect cell line. Arch. Insect Biochem. Physiol. 1, 225-230.
Marks, E. P. and Leopold, R. A. (1971). Deposition of cuticular substances in vitro by leg regenerates from the cockroach, Leucophaea maderae. Biol. Bull. 140, 73-83.[Medline]
Mayer, R. T., Chen, A. C. and DeLoach, J. R. (1980). Characterization of a chitin synthase from the stable fly, Stomoxys calcitrans (L.). Insect Biochem. 10,549 -556.[CrossRef]
Mayer, R. T., Meola, S. M. and DeLoach, J. R. (1981). Chitin synthesis inhibiting insect growth regulators do not inhibit chitin synthetase. EXS 37,337 -338.
McMurrough, I. and Bartnicki-Garcia, S. (1971).
Properties of a particulate chitin synthetase from Mucor rouxii. J.
Biol. Chem. 246,4008
-4016.
Merz, R. A., Horsch, M., Nyhlen, L. E. and Rast, D. M. (1999a). Biochemistry of chitin synthase. EXS 87,9 -37.[Medline]
Merz, R. A., Horsch, M., Ruffner, H. P. and Rast, D. M. (1999b). Interaction between chitosomes and concanavalin A. Phytochemistry 52,213 -226.
Meszaros, M. and Morton, D. B. (1997). Up- and downregulation of esr20, an ecdysteroid-regulated gene expressed in the tracheae of Manduca sexta. Arch. Insect Biochem. Physiol. 34,159 -174.[CrossRef][Medline]
Mikitani, K., Sugasaki, T., Shimada, T., Kobayashi, M. and
Gustafsson, J. A. (2000). The chitinase gene of the silkworm,
Bombyx mori, contains a novel Tc-like transposable element.
J. Biol. Chem.
275,37725
-37732.
Mommsen, T. P. (1980). Chitinase and beta-N-acetylglucosaminidase from the digestive fluid of the spider, Cupiennius salei. Biochim. Biophys. Acta 612,361 -372.[Medline]
Moya, P., Castillo, M., Primo-Yufera, E., Couillaud, F., Martinez-Manez, R., Garcera, M. D., Miranda, M. A., Primo, J. and Martinez-Pardo, R. (1997). Brevioxime: a new juvenile hormone biosynthesis inhibitor isolated from Penicillium brevicompactum. J. Org. Chem. 62,8544 -8545.[CrossRef][Medline]
Mulla, M. S., Darwazeh, H. A., Ede, L. and Kennedy, B. (1985). Laboratory and field evaluation of the IGR fenoxycarb against mosquitoes. J. Am. Mosq. Control Assoc. 1, 442-448.[Medline]
Müller, M. (1992). Proteolysis in protein import and export: signal peptide processing in eu- and prokaryotes. EXS 48,118 -129.
Munro, C. A. and Gow, N. A. R. (2001). Chitin synthesis in human pathogenic fungi. Med. Myc. 39, 41-53.
Muthukrishnan, J., Seifert, K., Hoffmann, K. H. and Lorenz, M. W. (1999). Inhibition of juvenile hormone biosynthesis in Gryllus bimaculatus by Glycosmis pentaphylla leaf compounds. Phytochemistry 50,249 -254.[CrossRef]
Nagahashi, S., Sudoh, M., Ono, N., Sawada, R., Yamaguchi, E.,
Uchida, Y., Mio, T., Takagi, M., Arisawa, M. and Yamada-Okabe, H.
(1995). Characterization of chitin synthase 2 of
Saccharomyces cerevisiae. Implication of two highly conserved domains
as possible catalytic sites. J. Biol. Chem.
270,13961
-13967.
Nakagawa, Y. and Matsumura, F. (1994). Diflubenzuron affects gammathio GTP stimulated Ca2+ transport in vitro in intracellular vesicles from the integument of the newly molted American cockroach, Periplaneta americana L. Insect Biochem. Mol. Biol. 24,1009 -1015.[CrossRef][Medline]
Niehrs, C. and Pollet, N. (1999). Synexpression groups in eukaryotes. Nature 402,483 -487.[CrossRef][Medline]
Nielsen, H., Engelbrecht, J., Brunak, S. and von Heijne, G. (1997). Identification of prokaryotic and eukaryotic signal peptides and prediction of their cleavage sites. Protein Eng. 10,1 -6.[Abstract]
Oberlander, H. (1976). Hormonal control of growth and differentiation of insect tissues cultured in vitro. In Vitro 12,225 -235.[Medline]
Oberlander, H., Lynn, D. A. and Leach, C. A. (1983). Inhibition of cuticle production in imaginal disks Plodia interpunctella (cultured in vitro): Effects of colcemid and vinblastine. J. Insect Physiol. 29, 47-53.[CrossRef]
Omura, S., Arai, N., Yamaguchi, Y., Masuma, R., Iwai, Y., Namikoshi, M., Turberg, A., Kolbl, H. and Shiomi, K. (2000). Argifin, a new chitinase inhibitor, produced by Gliocladium sp. FTD-0668. I. Taxonomy, fermentation, and biological activities. J. Antibiot. 53,603 -608.[Medline]
Ono, N., Yabe, T., Sudoh, M., Nakajima, T., Yamada-Okabe, T.,
Arisawa, M. and Yamada-Okabe, H. (2000). The yeast Chs4
protein stimulates the trypsin-sensitive activity of chitin synthase 3 through
an apparent protein-protein interaction. Microbiology
146,385
-391.
Orlean, P. (1987). Two chitin synthases in
Saccharomyces cerevisiae. J. Biol. Chem.
262,5732
-5739.
Ostrowski, S., Dierick, H. A. and Bejsovec, A.
(2002). Genetic control of cuticle formation during embryonic
development of Drosophila melanogaster. Genetics
161,171
-182.
Passonneau, J. V. and Williams, C. M. (1953). The moulting fluid of the Cecropia silkmoth. J. Exp. Biol. 246,124 -131.
Perez, M. and Hirschberg, C. B. (1985). Translocation of UDP-N-acetylglucosamine into vesicles derived from rat liver rough endoplasmic reticulum and Golgi apparatus. J. Biol. Chem. 260,4671 -4678.[Abstract]
Perrakis, A., Ouzounis, C., Wilson, K. S. and Vorgias, C. (1996). Implications of the tertiary structure determination of chitinase A. Similarities with Fibronectin III domains, oviductal proteins and narbonin. In Advances in Chitin Science, vol.1 (ed. A. Domard C. Jeuniaux, R. A. A. Muzzarelli and G. Roberts), pp. 34-41. Lyon: Jacques André.
Peters, W. (1992). Peritrophic membranes. In Zoophysiology, vol. 30 (ed. S. D. Bradshaw, W. Burggren, H. C. Heller, S. Ishii, H. Langer, G. Neuweiler and D. J. Randall), pp. 1-238. Berlin: Springer.
Peters, W., Heitmann, S. and D'Haese, J. (1979). Formation and fine structure of peritrophic membranes in the earwig, Forficula auricularia. Entomol. Gen. 5, 241-254.
Post, L. V., LJong, B. J. and Vincent, W. R. (1974). 1-(2,6-Disubsituted benzoyl)-3-phenylurea insecticides: inhibitors of chitin synthesis. Pestic. Biochem. Physiol. 4,473 -483.
Pummill, P. E., Achyuthan, A. M. and DeAngelis, P. L.
(1998). Enzymological characterization of recombinant Xenopus
DG42, a vertebrate hyaluronan synthase. J. Biol. Chem.
273,4976
-4981.
Quesada-Allue, L. A. (1982). The inhibition of insect chitin synthesis by tunicamycin. Biochem. Biophys. Res. Commun. 105,312 -319.[Medline]
Quesada-Allue, L. A., Marechal, L. R. and Belocopitow, E. (1976). Chitin synthesis in Triatoma infestans and other insects. Acta Physiol. Lat. Am. 26,349 -363.[Medline]
Rao, F. V., Houston, D. R., Boot, R. G., Aerts, J. M., Sakuda, S. and Van Aalten, D. M. (2003). Crystal structures of allosamidin derivatives in complex with human macrophage chitinase. J. Biol. Chem. 14,20110 -20116.[CrossRef]
Rechsteiner, M. and Rogers, S. W. (1996). PEST sequences and regulation by proteolysis. Trends Biochem. Sci. 21,267 -271.[CrossRef][Medline]
Retnakaran, A., Gelbic, I., Sundaram, M., Tomkins, W., Ladd, T., Primavera, M., Feng, Q., Arif, B., Palli, R. and Krell, P. (2001). Mode of action of the ecdysone agonist tebufenozide (RH-5992), and an exclusion mechanism to explain resistance to it. Pest Manag. Sci. 57,951 -957.[CrossRef][Medline]
Reynolds, S. E. and Samuels, R. I. (1996). Physiology and biochemistry of insect moulting fluid. Adv. Insect Physiol. 26,157 -232.
Riddiford, L. M. (1994). Cellular and molecular actions of juvenile hormone I. General considerations and premetamorphic actions. Adv. Insect Physiol. 24,211 -274.
Riddiford, L. M. (1996). Juvenile hormone: the status of its "status quo" action. Arch. Insect Biochem. Physiol. 32,271 -286.[CrossRef][Medline]
Robertus, J. D. and Monzingo, A. F. (1999). The structure and action of chitinases. EXS 87,125 -135.[Medline]
Rogers, S., Wells, R. and Rechsteiner, M. (1986). Amino acid sequences common to rapidly degraded proteins: the PEST hypothesis. Science 234,364 -368.[Medline]
Rosa, F., Sargent, T. D., Rebbert, M. L., Michaels, G. S., Jamrich, M., Grunz, H., Jonas, E., Winkles, J. A. and Dawid, I. B. (1988). Accumulation and decay of DG42 gene products follow a gradient pattern during Xenopus embryogenesis. Dev. Biol. 129,114 -123.[Medline]
Roussel, J.-P. (1994). Synthetic molecules designed as potential inhibitors in ecdysone biosynthesis. Entomol. Exp. Appl. 71,193 -199.
Royer, V., Fraichard, S. and Bouhin, H. (2002). A novel putative insect chitinase with multiple catalytic domains: hormonal regulation during metamorphosis. Biochem. J. 366,921 -928.[Medline]
Rudall, K. M. and Kenchington, W. (1973). The chitin system. Biol. Rev. 48,597 -636.
Ruiz-Herrera, J. and Martinez-Espinoza, A. D. (1999). Chitin biosynthesis and structural organization in vivo. EXS 87,39 -53.[Medline]
Ruiz-Herrera, J. and San-Blas, G. (2003). Chitin synthesis as target for antifungal drugs. Curr. Drug Targets Infect. Disord. 3,77 -91.[Medline]
Sakuda, S., Isogai, A., Matsumoto, S. and Suzuki, A. (1987). Search for microbial insect growth regulators. II. Allosamidin, a novel insect chitinase inhibitor. J. Antibiot. 40,296 -300.[Medline]
Samuels, R. and Reynolds, S. E. (1993). Moulting fluid enzymes of the tobacco hornworm, Manduca sexta: timing of proteolytic and chitinolytic activity in relation to pre-ecdysial development. Arch Insect Biochem. Physiol. 24, 33-44.
Samuels, R. I. and Paterson, I. C. (1995). Cuticle degrading proteases from insect moulting fluid and culture filtrates of entomopathogenic fungi. Comp. Biochem. Physiol. B 110,661 -669.[CrossRef][Medline]
Santos, B. and Snyder, M. (1997). Targeting of
chitin synthase 3 to polarized growth sites in yeast requires Chs5p and Myo2p.
J. Cell Biol. 136,95
-110.
Sanz, M., Trilla, J. A., Duran, A. and Roncero, C. (2002). Control of chitin synthesis through Shc1p, a functional homologue of Chs4p specifically induced during sporulation. Mol. Microbiol. 43,1183 -1195.[CrossRef][Medline]
Saville, G. P., Thomas, C. J., Possee, R. D. and King, L. A.
(2002). Partial redistribution of the Autographa
californica nucleopolyhedrovirus chitinase in virus-infected cells
accompanies mutation of the carboxy-terminal KDEL ER-retention motif.
J. Gen. Virol. 83,685
-694.
Saxena, I. M., Brown, R. M., Jr and Dandekar, T. (2001). Structure-function characterization of cellulose synthase: relationship to other glycosyltransferases. Phytochemistry 57,1135 -1148.[CrossRef][Medline]
Schooneveld, H. (1979). Precocene-induced necrosis and haemocyte-mediated breakdown of corpora allata in nymps of the locust Locusta migratoria. Cell Tissue Res. 203, 25-33.[Medline]
Schrempf, H. (2001). Recognition and degradation of chitin by streptomycetes. Antonie Van Leeuwenhoek 79,285 -289.[CrossRef][Medline]
Segawa, H., Kawakita, M. and Ishida, N. (2002).
Human and Drosophila UDP-galactose transporters transport
UDP-N-acetylgalactosamine in addition to UDP-galactose.
Eur. J. Biochem. 269,128
-138.
Sentandreu, R., Martinez-Ramon, A. and Ruiz-Herrera, J. (1984). Localization of chitin synthase in Mucor rouxii by an autoradiographic method. J. Gen. Microbiol. 130,1193 -1199.[Medline]
Shahabuddin, M., Toyoshima, T., Aikawa, M. and Kaslow, D. C. (1993). Transmission-blocking activity of a chitinase inhibitor and activation of malarial parasite chitinase by mosquito protease. Proc. Natl. Acad. Sci. USA 90,4266 -4270.[Abstract]
Shen, Z. and Jacobs-Lorena, M. (1999). Evolution of chitin-binding proteins in invertebrates. J. Mol. Evol. 48,341 -347.[Medline]
Shen, Z. C. and Jacobs-Lorena, M. (1997).
Characterization of a novel gut-specific chitinase gene from the human malaria
vector Anopheles gambiae. J. Biol. Chem.
272,28895
-28900.
Sinnott, M. L. (1990). Catalytic mechanisms of enzymatic glycosyl transfer. Chem. Rev. 90,1171 -1202.
Skehel, J. J. and Wiley, D. C. (1998). Coiled coils in both intracellular vesicle and viral membrane fusion. Cell 95,871 -874.[Medline]
Spindler, K.-D., Przibilla, S. and Spindler-Barth, M. (2001). Moulting hormones of arthropods: molecular mechanisms. Zoology 103,189 -201.
Surholt, B. (1975). Studies in vitro and in vivo on chitin synthesis during larval-adult moulting cycle of the migratory locust, Locusta migratoria L. J. Comp. Physiol. 102,135 -142.
Tellam, R. L. (1996). The peritrophic matrix. In Biology of the Insect Midgut (ed. M. J. Lehane and P. F. Billingsley), pp. 86-114. Cambridge: Chapman and Hall.
Tellam, R. L., Smith, D. H. and Willadsen, P. (1992). Vaccination against ticks. In Animal Parasite Control Utilizing Biotechnology (ed. W. K. Yong), pp.303 -331. Boca Raton: CRC Press.
Tellam, R. L., Vuocolo, T., Johnson, S. E., Jarmey, J. and
Pearson, R. D. (2000). Insect chitin synthase cDNA sequence,
gene organization and expression. Eur. J. Biochem.
267,6025
-6043.
Terra, W. R., Ferreira, C., Jordao, B. P. and Dillon, R. J. (1996). Digestive enzymes. In Biology of the insect midgut (ed. M. J. Lehane and P. F. Billingsley), pp.153 -194. Cambridge: Chapman and Hall.
Terwisscha van Scheltinga, A. C., Armand, S., Kalk, K. H., Isogai, A., Henrissat, B. and Dijkstra, B. W. (1995). Stereochemistry of chitin hydrolysis by a plant chitinase/lysozyme and X-ray structure of a complex with allosamidin: evidence for substrate assisted catalysis. Biochemistry 34,15619 -15623.[Medline]
Terwisscha van Scheltinga, A. C., Kalk, K. H., Beintema, J. J. and Dijkstra, B. W. (1994). Crystal structures of hevamine, a plant defence protein with chitinase and lysozyme activity, and its complex with an inhibitor. Structure 2,1181 -1189.[Medline]
Thomas, C. J., Gooday, G. W., King, L. A. and Possee, R. D.
(2000). Mutagenesis of the active site coding region of the
Autographa californica nucleopolyhedrovirus chiA gene.
J. Gen. Virol. 81,1403
-1411.
Trilla, J. A., Duran, A. and Roncero, C.
(1999). Chs7p, a new protein involved in the control of protein
export from the endoplasmic reticulum that is specifically engaged in the
regulation of chitin synthesis in Saccharomyces cerevisiae. J. Cell
Biol. 145,1153
-1163.
Truman, J. W. and Riddiford, L. M. (1970). Neuroendocrine control of ecdysis in silkmoths. Science 167,1624 -1626.
Tsai, Y. L., Hayward, R. E., Langer, R. C., Fidock, D. A. and
Vinetz, J. M. (2001). Disruption of Plasmodium
falciparum chitinase markedly impairs parasite invasion of mosquito
midgut. Infect. Immun.
69,4048
-4054.
Turnbull, I. F. and Howells, A. J. (1983). Integumental chitin synthase activity in cell-free extracts of larvae of the Australian sheep blowfly, Lucilia cuprina, and two other species of diptera. Aust. J. Biol. Sci. 36,251 -262.[Medline]
Uchida, Y., Shimmi, O., Sudoh, M., Arisawa, M. and Yamada-Okabe, H. (1996). Characterization of chitin synthase 2 of Saccharomyces cerevisiae. II. Both full size and processed enzymes are active for chitin synthesis. J. Biochem. 119,659 -666.[Abstract]
Usui, T., Hayashi, Y., Nanjo, F., Sakai, K. and Ishido, Y. (1987). Transglycosylation reaction of a chitinase purified from Nocardia orientalis. Biochim. Biophys. Acta 923,302 -309.[Medline]
van Aalten, D. M., Komander, D., Synstad, B., Gaseidnes, S.,
Peter, M. G. and Eijsink, V. G. (2001). Structural insights
into the catalytic mechanism of a family 18 exochitinase. Proc.
Natl. Acad. Sci. USA 98,8979
-8984.
van Eck, W. H. (1979). Mode of action of two benzoylphenyl ureas as inhibitors of chitin synthesis in insects. Insect Biochem. 9,295 -300.[CrossRef]
Vardanis, A. (1976). An in vitro assay system for chitin synthesis in insect tissue. Life Sci. 19,1949 -1956.[CrossRef][Medline]
Vardanis, A. (1979). Characteristics of the chitin-synthesizing system of insect tissue. Biochim Biophys Acta 588,142 -147.[Medline]
Venegas, A., Goldstein, J. C., Beauregard, K., Oles, A., Abdulhayoglu, N. and Fuhrman, J. A. (1996). Expression of recombinant microfilarial chitinase and analysis of domain function. Mol. Biochem. Parasitol. 78,149 -159.[CrossRef][Medline]
Vermeulen, C. A. and Wessels, J. G. (1986). Chitin biosynthesis by a fungal membrane preparation. Evidence for a transient non-crystalline state of chitin. Eur. J. Biochem. 158,411 -415.[Abstract]
von Heijne, G. (1990). The signal peptide. J. Membr. Biol. 115,195 -201.[Medline]
Vorgias, C. E., Kingswell, A. J., Dauter, Z. and Oppenheim, A. B. (1992). Crystallization of recombinant chitinase from the cloned chiA gene of Serratia marcescens. J. Mol. Biol. 226,897 -898.[Medline]
Wang, X., Ding, X., Gopalakrishnan, B., Morgan, T. D., Johnson, L., White, F., Muthukrishnan, S. and Kramer, K. J. (1996). Characterization of a 46 kDa insect chitinase from transgenic tobacco. Insect Biochem. Mol. Biol. 26,1055 -1064.[CrossRef]
Wigglesworth, V. B. (1930). The formation of the peritrophic membrane in insects, with special reference to the larvae of mosquitoes. Q. J. Microsc. Sci. 73,593 -616.
Wingender, E., Kel, A. E., Kel, O. V., Karas, H., Heinemeyer,
T., Dietze, P., Knuppel, R., Romaschenko, A. G. and Kolchanov, N. A.
(1997). TRANSFAC, TRRD and COMPEL: towards a federated database
system on transcriptional regulation. Nucleic Acids
Res. 25,265
-268.
Yabe, T., Yamada-Okabe, T., Nakajima, T., Sudoh, M., Arisawa, M. and Yamada-Okabe, H. (1998). Mutational analysis of chitin synthase 2 of Saccharomyces cerevisiae. Identification of additional amino acid residues involved in its catalytic activity. Eur. J. Biochem. 258,941 -947.[Abstract]
Yan, J., Cheng, Q., Narashimhan, S., Li, C. B. and Aksoy, S. (2002). Cloning and functional expression of a fat body-specific chitinase cDNA from the tsetse fly, Glossina morsitans. Insect Biochem. Mol. Biol. 32,979 -989.[CrossRef][Medline]
Yao, T. P., Forman, B. M., Jiang, Z., Cherbas, L., Chen, J. D., McKeown, M., Cherbas, P. and Evans, R. M. (1993). Functional ecdysone receptor is the product of EcR and Ultraspiracle genes. Nature 366,476 -479.[CrossRef][Medline]
Yarema, C., McLean, H. and Caveney, S. (2000). L-Glutamate retrieved with the moulting fluid is processed by a glutamine synthetase in the pupal midgut of Calpodes ethlius. J. Insect Physiol. 46,1497 -1507.[CrossRef][Medline]
Zen, K. C., Choi, H. K., Krishnamachary, N., Muthukrishnan, S. and Kramer, K. J. (1996). Cloning, expression, and hormonal regulation of an insect beta-N-acetylglucosaminidase gene. Insect Biochem. Mol. Biol. 26,435 -444.[CrossRef][Medline]
Zhang, D. and Miller, M. J. (1999). Polyoxins and nikkomycins: progress in synthetic and biological studies. Curr. Pharm. Des. 5,73 -99.[Medline]
Zhang, H., Huang, X., Fukamizo, T., Muthukrishnan, S. and Kramer, K. J. (2002). Site-directed mutagenesis and functional analysis of an active site tryptophan of insect chitinase. Insect Biochem. Mol. Biol. 32,1477 -1488.[CrossRef][Medline]
Zhu, X., Zhang, H., Fukamizo, T., Muthukrishnan, S. and Kramer, K. J. (2001). Properties of Manduca sexta chitinase and its C-terminal deletions. Insect Biochem. Mol. Biol. 31,1221 -1230.[CrossRef][Medline]
Zhu, Y. C., Specht, C. A., Dittmer, N. T., Muthukrishnan, S., Kanost, M. R. and Kramer, K. J. (2002). Sequence of a cDNA and expression of the gene encoding a putative epidermal chitin synthase of Manduca sexta. Insect Biochem. Mol. Biol. 32,1497 -1506.[CrossRef][Medline]
Zielkowski, R. and Spindler, K. (1978). Chitinase and chitobiase from the integument of Locusta migratoria: characteristics and titer during the fith instar. Insect Biochem. 8,67 -71.[CrossRef]
Ziman, M., Chuang, J. S., Tsung, M., Hamamoto, S. and Schekman,
R. (1998). Chs6p-dependent anterograde transport of Chs3p
from the chitosome to the plasma membrane in Saccharomyces cerevisiae.Mol. Biol. Cell 9,1565
-1576.
Zimoch, L. and Merzendorfer, H. (2002). Immunolocalization of chitin synthase in the tobacco hornworm. Cell Tissue Res. 308,287 -297.[CrossRef][Medline]
Zitnan, D., Ross, L. S., Zitnanova, I., Hermesman, J. L., Gill, S. S. and Adams, M. E. (1999). Steroid induction of a peptide hormone gene leads to orchestration of a defined behavioral sequence. Neuron 23,523 -535.[Medline]