(Received for publication, July 18, 1995; and in revised form, August 22, 1995)
From the
Chitin deacetylase (EC 3.5.1.41), the enzyme that catalyzes the
hydrolysis of acetamido groups of N-acetyl-D-glucosamine in chitin, has been purified
to homogeneity from the culture filtrate of the fungus Colletotrichum lindemuthianum and further characterized. The
enzyme is a glycoprotein, and its apparent molecular mass was
determined to be 150 kDa. The glycosylation pattern of the enzyme
is consistent with a mixture of N-linked glycans including
oligomannosidic hybrid and/or complex type, and its carbohydrate
content is approximately 67% by weight. Chitin deacetylase is active on
several chitinous substrates and chitin derivatives, is not
considerably inhibited by carboxylic acids, especially acetic acid, and
exhibits a remarkable thermostability. The enzyme requires at least two N-acetyl-D-glucosamine residues (chitobiose) for
catalysis. When glycol chitin (a water-soluble chitin derivative) was
used as substrate, the optimum temperature for enzyme activity was
determined to be 50 °C, and the optimum pH was
8.5.
Chitin, a homopolymer of -(1-4)-linked N-acetyl-D-glucosamine, is one of the most abundant,
easily obtained, and renewable natural polymers, second only to
cellulose. It is commonly found in the exoskeletons or cuticles of many
invertebrates (1) and in the cell walls of most fungi and algae (2) .
Chitin is an extremely insoluble material and has yet to find an important industrial use, whereas chitosan (the deacetylated form of chitin) is water-soluble and a much more tractable material with a large number and a broad variety of reported applications(3, 4, 5) . At present, chitosan is produced by the thermochemical deacetylation of chitin. An alternative or complementary procedure exploiting the enzymatic deacetylation of chitin could potentially be employed, especially when a controlled, non-degradative, and well defined process is required.
Chitin deacetylase (CDA), ()the enzyme that catalyzes the
conversion of chitin to chitosan by the deacetylation of N-acetyl-D-glucosamine residues, was first identified
and partially purified from extracts of the fungus Mucor
rouxii(6) . Since then, the presence of this enzyme
activity has been reported in several other fungi (7, 8, 9) and in some insect
species(10) . The purification of the first chitin deacetylase
to homogeneity and further characterization of the enzyme from the
fungus M. rouxii has been recently reported. The enzyme is an
acidic glycoprotein of
75 kDa with a carbohydrate content of
30% by weight. It exhibits a very stringent specificity for
-(1,4)-linked N-acetyl-D-glucosamine
homopolymers, requires at least four residues
(tetra-N-acetylchitotetraose) for catalysis, and is inhibited
by carboxylic acids, particularly acetic acid. The optimum temperature
for enzyme activity was determined to be
50 °C and the optimum
pH
4.5. A cDNA of the M. rouxii encoding CDA was
isolated, sequenced, and further characterized(13) . Protein
sequence comparisons revealed significant similarities between the
fungal chitin deacetylase and the rhizobial nodB proteins, suggesting
functional homology of these evolutionarily distant proteins. The
functional assignment of the nodB protein in Nod factor biosynthesis,
as deduced from its sequence similarity, was subsequently verified
biochemically(14) . The purification and partial
characterization of chitin deacetylases from Absidia coerulea(15) and Aspergillus nidulans(16) has
also been recently reported. CDA from A. coerulea as compared
to the M. rouxii enzyme exhibited similar molecular weight,
amino-terminal sequence, pH and temperature optimum, and substrate
specificity. However, CDA from A. nidulans as compared to the
above enzymes exhibited different molecular weight, pH optimum, and
substrate specificity.
Initiating a study to elucidate the potential biological role of CDA activity and further evaluate the potential use of an enzymatic process for deacetylation of chitin substrates, we now report the isolation and characterization of CDA from the fungus Colletotrichum lindemuthianum.
Figure 1:
Purification of chitin deacetylase on Q
Sepharose fast flow column. A sample (500 ml, 635 mg of protein) of
concentrated and dialyzed against 20 mM Bis-Tris-HCl, pH 5.8
(buffer A), culture filtrate, was applied onto a Q Sepharose fast flow
adsorbent (44 160 mm). The column was washed with buffer A and
then developed with a linear gradient of NaCl (2000 ml; 0-0.5 M) at a flow rate of 300 ml/h. Chitin deacetylase activity
appeared in fractions 145-180, corresponding to
0.12 M NaCl. The protein content was followed by a UV monitor at 280
nm.
Fractions containing enzyme activity were pooled,
concentrated (3 ml) by ultrafiltration, and subsequently applied onto a
Sephacryl S300 HR column (26 600 mm) previously equilibrated in
20 mM Tris-HCl, pH 7.4, 0.5 mM NaCl (Fig. 2).
Figure 2:
Purification of chitin deacetylase on
Sephacryl S300 HR column. A sample (3 ml, 59.5 mg of protein) of
partially purified chitin deacetylase on a Q Sepharose column was
applied onto a Sephacryl S300 HR column (26 600 mm). Active
fractions were collected. The protein content was followed by a UV
monitor at 280 nm.
Active fractions were pooled, dialyzed against 50 mM sodium
formate buffer, pH 4.0 (buffer B), and loaded onto a MonoS column (5
50 mm) equilibrated in buffer B. The column was washed with
buffer B and subsequently developed with a linear gradient of NaCl (30
ml; 0-1 M) in buffer B at a flow rate of 30 ml/h.
Fractions containing enzyme activity were pooled and stored at
-20 °C (Fig. 3).
Figure 3:
Purification of chitin deacetylase on
MonoS column. A sample (5 ml, 9.24 mg of protein) of partially purified
chitin deacetylase on a Sephacryl S300 HR column, was concentrated,
dialyzed against 50 mM sodium formate buffer pH 4.0 (buffer
B), and applied onto a MonoS adsorbent (5 50 mm). The column
was washed with buffer B and then developed with a linear gradient of
NaCl (30 ml; 0-1 M) at a flow rate of 30 ml/h. Fractions
containing enzyme activity were collected. The protein content was
followed by a UV monitor at 280 nm.
Figure 4:
Electrophoretic pattern of chitin
deacetylase. Purified enzyme preparation was electrophoresed on a 10%
polyacrylamide gel under denaturing and reducing conditions. Protein
bands were visualized by staining with Coomassie Brilliant Blue R.
Identities and amount of protein loaded were: Lane 1, culture
filtrate (50 µg); lane 2, Q Sepharose eluate (
50
µg); lane 3, Sephacryl 300 eluate (
15 µg); lane 3, MonoS eluate (
5 µg); lane 5, N-deglycosylated enzyme (treatment with N-glycosidase
F (
5 µg)).
Figure 5: Fluorophore assisted carbohydrate electrophoresis of chitin deacetylase N-linked glycans. N-Linked glycans were cleaved and labeled as described under ``Experimental Procedures'' before loading on polyacrylamide gel. Identities of samples loaded were: lane 1, N-glycosidase F digest of chitin deacetylase; lane 2, wheat starch-labeled polymers of glucose; lane 3, 50 pmol of maltotetraose.
Figure 6:
Thermal stability of chitin deacetylase.
Thermostability of CDA was tested after preincubation of (0.2
milliunit) of the enzyme for different time periods at 37 °C
(), 50 °C (
), and 60 °C (
). After cooling on
ice, each sample was assayed under standard conditions to determine
remaining activity.
In this report, we describe the purification of the enzyme
chitin deacetylase from the culture filtrate of the fungus C.
lindemuthianum to apparent electrophoretic homogeneity. The enzyme
is a glycoprotein. From the carbohydrate and amino acid analysis in
combination with the apparent molecular mass of the deglycosylated
enzyme (50 kDa), its apparent molecular mass was determined to be
150 kDa, while its carbohydrate content
67% by weight.
Further incubation of the N-deglycosylated enzyme with O-glycosidase, did not result in any further reduction in the
molecular mass of the enzyme, suggesting that oligosaccharide chains
are N-linked. The fluorophore-labeled N-glycan
migration range was between glucose oligomers, having a degree of
polymerization between 4 and 12. Most N-linked
oligosaccharides migrate in this region of the gel and usually consist
of between 10 and 20 monosaccharide units. The glycosylation pattern of
the enzyme is consistent with a mixture of N-linked glycans
including the oligomannosidic hybrid and/or complex type.
Although several other purification schemes were also employed in our laboratory resulting in a final enzyme preparation of comparable purity, the one we describe here was chosen for several reasons. It is simple, since it can be completed within 3 days; it employs standard protein purification media and equipment; it is economic, as expensive adsorbents are avoided; and it is easy to scale up. Polyclonal antibodies raised against the M. rouxii CDA do not react with the C. lindemuthianum enzyme, even though we have observed immunological homology of CDAs within the Zygomycetes class (data not shown). Thus, immunoaffinity chromatography based on antibodies against the M. rouxii enzyme, which has been successfully employed for the purification of CDA from M. rouxii to homogeneity in a one-step procedure(12) , cannot be effective for the isolation of the enzyme from C. lindemuthianum.
We have used two different assays for the determination of CDA activity. A radiometric assay, using radiolabeled glycol chitin as substrate, proved to be a rapid and sensitive way of screening chromatographic fractions, whereas the estimation of acetate released by an enzymatic method was used for monitoring the deacetylation process of nonradiolabeled substrates(24) .
Glycol chitin has been used as a model substrate for the determination of CDA activity(6) . Since it is not easy to evaluate, (i) the extent and distribution of derivatization (O-hydroxyethyl groups) in glycol chitin commercially available and (ii) the effect of derivatization on enzyme activity, we have used hexa-N-acetylchitohexaose as a model substrate for the determination of the enzyme activity as described previously(11) .
When glycol chitin was used as substrate, the optimum temperature for enzyme activity was determined to be 50 °C, similar to all CDAs examined so far, while the optimum pH was estimated to be 8.5, which is the highest reported. Furthermore, the enzyme is not inhibited by acetate and exhibits a remarkable thermostability.
CDA from C. lindemuthianum appears to
exhibit a very narrow specificity, acting only on N-acetyl-D-glucosamine homopolymers, similarly to the
enzyme from M. rouxii and A. coerulea, while the
corresponding enzyme from A. nidulans was found to exhibit a
wider specificity. Overincubation of the enzyme with N-acetylchitooligosaccharides revealed that the enzyme
requires at least two N-acetyl-D-glucosamine residues
for catalysis. However, shorter incubation times (10 min) indicated
that the rate of deacetylation was considerably higher with chitin
oligomers having more that three N-acetyl-D-glucosamine residues and that the enzyme
can deacetylate (GlcNAc), (GlcNAc)
, and
(GlcNAc)
with approximately the same efficiency. In the
case of M. rouxii and A. coerulea enzymes, the rate
of deacetylation was higher the longer the chitooligosaccharide was.
Chitin deacetylase from A. nidulans showed maximum activity
with (GlcNAc)
while it was less active on
(GlcNAc)
. In order to test CDA effectiveness in
deacetylating chitin and chitosan substrates, two crystalline and two
amorphous chitin samples as well as a chitosan substrate (chitin 50)
were incubated with the enzyme under the standard assay conditions.
When the enzyme was incubated with the 40% deacetylated chitin and
various N-acetylchitooligosaccharides for 10 min, the degrees
of deacetylation obtained were comparable (Table 4). When
crystalline chitin and its chemically modified form, amorphous chitin,
were incubated with the enzyme for 24 h, approximately 0.5 and 5%
deacetylation was achieved, respectively (Table 5). This
indicates that the enzyme is not very effective in deacetylating
insoluble chitin substrates and that pretreatment of crystalline chitin
substrates prior to enzyme addition is necessary, in order to improve
the accessibility of the acetyl groups to the enzyme and therefore
enhance the yield and rate of the deacetylation reaction. Similar
results employing the same substrates and CDA from M. rouxii have been reported(24) . CDA from A. coerulea was
also not active toward crystalline chitin while it was effective in
deacetylating colloidal chitin, an amorphous chitin substrate.
The requirements for substrate recognition and the mechanism of enzyme action on N-acetylchitooligosaccharides as well as chitin and chitosan polymers, need to be further studied.
In summary, CDA from C. lindemuthianum, as compared to all other corresponding enzymes, exhibits different properties, e.g. increased thermostability and different pH optimum, while it is not inhibited by acetate. These properties could potentially be exploited for the effective deacetylation of chitinous substrates.
Even though chitin
biosynthesis, enzymology, and cytology in fungi have been extensively
studied, there is limited information on chitosan biosynthesis. It has
been reasonably suggested that CDA from M. rouxii is a
secreted enzyme and that its function is localized in the periplasmic
space(25) . Preliminary immunolocalization experiments in this
fungus reinforce this suggestion. ()It has been also
recently demonstrated, by immunoelectron microscopy, that CDA from A. coerulea is localized near the inner face of the cell wall
(periplasmic space). In contrast, CDA from C. lindemuthianum exhibits a 10-fold higher specific activity in the culture
filtrate than in mycelial extracts, suggesting that the enzyme may have
another (or additional) role than in chitosan biosynthesis in this
fungus.
In considering other possible biological roles for CDA the following hypotheses can be envisaged, taking into account the fact that C. lindemuthianum is a plant pathogen. First, it is known that chitin oligomers (tetramer to hexamer), elicit lignification in wounded wheat leaves(26) , formation of callose and coumarin derivatives in parsley cells(27) , as well as in protoplasts and cells of Catharanthus roseus(28) . The deacetylated forms of these oligomers do not possess any significant elicitor activity in any of the above systems. On the other hand, it has been reported that chitosan oligomers induce the synthesis of pisatin in pea pods (29) and of proteinase inhibitors in tomato leaves(30) . These observations, in combination with our finding that CDA from C. lindemuthianum is active on chitin oligomers, suggest that the enzyme might play a role in plant pathogen interactions. A possible scenario is that endochitinases of plant and/or fungal origin may cause the release of chitin oligomers arising from the cell walls of C. lindemuthianum. These oligomers could act as elicitors for the synthesis of callose, lignin, and phytoalexins, substances which potentially contribute to the plant's defense mechanism. CDA being extracellular in this fungus could convert these oligomers to their deacetylated forms, thereby diminishing their elicitor activity. Second, plants produce chitinases which are generally thought to contribute in defense against fungal pathogens(31, 32) . Deacetylation of cell wall chitin chains in C. lindemuthianum might make the polymer resistant to degradation by the plant chitinases. However, it has been reported that cell walls of this fungus do not contain chitosan even though they do contain chitin(33) . The above hypotheses could be tested once the structural gene has been cloned (work in progress in this laboratory) by carrying out gene disruption experiments in this fungus.
Chitin deacetylases isolated so far have been reported to be involved either in the formation of the cell wall (M. rouxii, A. coerulea) in combination with chitin synthetases operating in tandem or in deacetylating chitin oligosaccharides during autolysis after the action of endochitinase on cell walls (A. nidulans). In this report, a new role for C. lindemuthianum CDA is proposed, its involvement in plant-pathogen interactions; since the enzyme is extracellular, its cell wall does not contain chitosan, while it is active on chitin oligomers.
The similarities and differences between nodB proteins and CDAs from M. rouxii and C. lindemuthianum make these enzymes an attractive system for studying structure and function relationships in substrate recognition and catalysis. Comparative analysis of sequence similarities between these proteins can provide the basis for developing a protein engineering strategy in order to modify the specificity of CDAs and/or nodB protein. The ultimate goal could be the design of an enzyme with improved efficiency for the synthesis of new Nod factors or oligomeric bioactive chitooligosaccharides and polymeric chitosan substrates.