Identification of a catalytic residue of Clostridium paraputrificum N-acetyl-β-D-glucosaminidase Nag3A by site-directed mutagenesis

Article abstract of DOI:10.1271/bbb.70.1127

Clostridium paraputrificum M-21 β-N-acetylglucosaminidase 3A (Nag3A) is an enzyme classified in family 3 of the glycoside hydrolases. To identify catalytic residues of this enzyme, mutations were introduced into highly conserved Glu and Asp residues. Replacement of Asp175 with Ala abolished the catalytic activity without change in the circular dichroism spectrum, strongly suggesting that this residue is a catalytic residue, a nucleophile/base or a proton donor. Since the K_m values of mutant enzymes D119N, D229N, D229A and D274N increased 17 to 41 times as compared with that of wildtype enzyme, Asp119, Asp229, and Asp274 appear to be involved in substrate recognition and binding. Taking previous studies into consideration, we presume that Asp303 is the catalytic nucleophile and Asp175 is the proton donor of C. paraputrificum Nag3A.

Full text of DOI:10.1271/bbb.70.1127

Biosci. Biotechnol. Biochem., 70 (5), 1127–1133, 2006
Identification of a Catalytic Residue of Clostridium paraputrificum
N-Acetyl-ꢀ-D-glucosaminidase Nag3A by Site-Directed Mutagenesis
1
2
2
2
Huazhong LI, Guangshan ZHAO, Hideo MIYAKE, Hayato UMEKAWA,
2
2
2;y
Tetsuya KIMURA, Kunio OHMIYA, and Kazuo SAKKA
1
School of Medicine, Southern Yangtze University, 170, Huihe Road, Wuxi 214036, China
Faculty of Bioresources, Mie University, Kurima-Machiyacho 1577, Tsu 514-8507, Japan
2
Received August 26, 2005; Accepted January 7, 2006
3
)
Clostridium paraputrificum M-21 ꢀ-N-acetylglucosa-
minidase 3A (Nag3A) is an enzyme classified in family 3
of the glycoside hydrolases. To identify catalytic resi-
dues of this enzyme, mutations were introduced into
highly conserved Glu and Asp residues. Replacement of
Asp175 with Ala abolished the catalytic activity without
change in the circular dichroism spectrum, strongly
suggesting that this residue is a catalytic residue, a
nucleophile/base or a proton donor. Since the Km values
of mutant enzymes D119N, D229N, D229A and D274N
increased 17 to 41 times as compared with that of wild-
type enzyme, Asp119, Asp229, and Asp274 appear to be
involved in substrate recognition and binding. Taking
previous studies into consideration, we presume that
Asp303 is the catalytic nucleophile and Asp175 is the
proton donor of C. paraputrificum Nag3A.
hydrogenase gene. Therefore, the bacterium has appli-
cation possibilities in hydrogen production. Two chiti-
4
)
5)
nase genes, chiA and chiB, and two ꢀ-N-acetylglu-
6
)
7)
cosaminidase (Nag) genes, nag3A and nag84A of
C. paraputrificum M-21, were cloned from a gene
library and expressed in Escherichia coli, and their
translated products were characterized. Both chitinases
produced (GlcNAc)2 as the major end product from
chitin and chitooligosaccharides. Although Nag84A
hydrolyzed only chitooligosaccharides to produce
GlcNAc, Nag3A produced GlcNAc from either chitin
or chitooligosaccharides and showed particularly high
activity on chitobiose. It was also found that C. para-
putrificum M-21 secreted a 72-kD endo-chitinase
Chi18C other than ChiA and ChiB when cultivated on
ball-milled chitin as a carbon source (unpublished
result). Chi18C catalyzes the hydrolysis of chitin at
random sites along the polymer to produce chitooligo-
saccharides. The sequential hydrolysis conducted by
chi18C, chiA, chiB, nag84A, and nag3A gene products
must govern the breakdown of chitin and the growth of
C. paraputrificum M-21 when chitin was used as the
sole carbon source. Among these enzymes, Nag3A may
be a key enzyme for the final step of sequential
hydrolysis, since it prefers chitobiose to larger saccha-
rides as the substrate.
6
)
Key words: ꢀ-N-acetylglucosaminidase; Clostridium
paraputrificum; catalytic residue; site-di-
rected mutagenesis
Chitin is the second most abundant polysaccharide
found in nature. It consists of variable-length linear
chains of ꢀ-1,4-linked polymers of N-acetylglucosamine
hydrogen bonded into an ordered insoluble crystalline
structure. The enormous amounts of chitin produced
annually in the biosphere are degraded by chitinases.
Chitinases are ubiquitous in nature, being found in
eukaryotes, prokaryotes, archaea, and viruses. They
consist of a group of hydrolytic enzymes that are able to
break down polymeric chitin to chitin oligosaccharides,
Nag3A is an enzyme classified in family 3 of the
glycoside hydrolases (http://afmb.cnrs-mrs.fr/CAZY/
index.html). Twelve Nags are found in this family,
and four of them have been characterized as to their
8
)
6
,9–11)
enzyme properties.
From the alignment of the
diacetylchitobiose [(GlcNAc) ], and N-acetylglucosa-
2
catalytic domains of the 12 Nags (Fig. 1), it can be seen
that two Glu residues and six Asp residues are highly
conserved among them and are candidates for catalytic
amino acid residues. Barley ꢀ-glucan glucanohydrolase
ExoI in family 3 is composed of 605 amino acid
residues and is the only enzyme of which crystalline
mine (GlcNAc).
Clostridium paraputrificum M-21 was isolated and
characterized as a chitin-degrading hydrogen-producing
1
,2)
anaerobe in this laboratory. It produces a significant
amount of hydrogen gas, approximately 2 mol H2/mol
GlcNAc. Its hydrogen gas production was enhanced to
1
2,13)
structure has been reported. Structural analysis of
ExoI indicated that this enzyme is a globular protein
1
.7-fold of that of the wild type by overexpression of a
y
To whom correspondence should be addressed. Tel: +81-59-231-9621; Fax: +81-59-231-9684; E-mail: sakka@bio.mie-u.ac.jp
Abbreviations: Nag, ꢀ-N-acetylglucosaminidase; GlcNAc, N-acetylglucosamine; 4-MU-(GlcNAc), 4-methylumbelliferyl ꢀ-D-N-acetylgluco-
samine; PAGE, polyacrylamide gel electrophoresis; PCR, polymerase chain reaction; Tris, Tris(hydroxymethyl)aminomethane

1128
H. LI et al.
Fig. 1. Multiple Alignment of the Catalytic Domains of Family-3 Nags.
Amino acids conserved in at least 9 of 11 sequences are highlighted; - gap left to improve alignment. Asterisks indicate the amino acids
selected for site-directed mutagenesis. Numbers at the end of each line refer to amino acid residue positions. All sequences except ExoI are
numbered from Met-1 of the peptide and ExoI sequence is numbered from the Asp residue of the mature form without signal peptide. C-p,
C. paraputrificum Nag3A; A-s, Alteromonas sp. Cht60; B-s, Bacillus subtilis YbbD; C-f, Cellulomonas fimi Nag3A; E-c1, E. coli CFT073
NagZm; E-c2, E. coli K-12 NagZ; S-t, Streptomyces thermoviolaceus NagII; T-n, Thermotoga neapolitana CbsA; V-c, Vibrio cholerae NagZ;
V-f, Vibrio furnissii ExoII; Y-p, Yersinia pestis NagZ; ExoI, Barley ꢀ-D-glucan exohydrolase isoenzyme ExoI.
composed of two domains: the first 357 residues
constitute an ðꢁ=ꢀÞ TIM-barrel domain, and the second
Asp285 in the first domain and Glu491 in the second
domain were identified as the catalytic nucleophile and
proton donor respectively. On the other hand,
C. paraputrificum Nag3A consists of 413 amino acid
8
1
2–14)
domain is arranged in a six-stranded ꢀ sandwich with
three ꢁ helices on either side of the sheet. Furthermore,

Clostridium paraputrificum N-Acetyl-ꢀ-D-glucosaminidase
1129
Table 1. Mutants and Synthetic Oligonucleotides Used for Amplifi-
cation and Mutagenesis of the nag3A Gene in This Study
residues, suggesting that it is a single-domain protein
devoid of a second domain. Therefore, although a
possible nucleophile identified in ExoI is conserved as
Asp303 in C. paraputrificum Nag3A, no possible proton
donor residue is found in Nag3A. To identify catalytic
residues of C. paraputrificum Nag3A, we constructed a
number of mutant Nag3A proteins with substitutions at
possible catalytic residues by site-directed mutagenesis
and characterized their enzyme properties.
0
0
Oligonucleotide
Sequence (5 !3 )
N- and C-terminal flanking oligonucleotides
NAG-For ggatccactattttaacagggaaaaatg
NAG-Rev gtcgacctatttatacattgcttcaactac
Mutation-introducing oligonucleotides
D119N
E121Q
E121A
D175N
D175A
D229N
D229A
D274N
E293Q
D303N
D339N
fwd
rev
attgggtttAaCgaagaagga
tccttcttcGtTaaacccaat
tttgatgaaCaaggaggaagt
acttcctccttGttcatcaaa
tttgatgaagCaggaggaagt
acttcctcctGcttcatcaaa
ggtactgttttaAaCattaatact
agtattaatGtTtaaaacagtacc
ggtactgttttagCtattaatact
agtattaataGctaaaacagtacc
acggaggtgAaCtctcattt
aaatgagaGtTcacctccgt
acggaggtggCttctcattt
aaatgagaaGccacctccgt
ccacaaattAaCaaagagtac
gtactctttGtTaatttgtgg
ttacgtgatCaaatgggttata
tataacccatttGatcacgtaa
attattactAaCgatcttgaaa
tttcaagatcGtTagtaataat
gcaggtgcaAaCattttatta
taataaaatGtTtgcacctgc
fwd
rev
Materials and Methods
fwd
rev
fwd
rev
Bacterial strains, plasmids, and culture conditions.
pQE-30T, a derivative of pQE-30 (Qiagen, Hilden,
Germany), which contains a short stretch of nucleotides
encoding a thrombin recognition sequence (LVPRGS)
between the six-His affinity tag–coding region and the
multiple cloning sites, was used as the expression
vector.15) E. coli strain XL1-Blue was used as the host
for construction and isolation of recombinant plasmids.
E. coli M15 (pREP4) (Qiagen) was used as the host for
expression of the recombinant proteins from plasmids
fwd
rev
fwd
rev
fwd
rev
fwd
rev
fwd
rev
fwd
rev
derived from pQE-30T. All E. coli strains were grown at
ꢀ
37 C on LB medium supplemented with appropriate
antibiotics.
fwd
rev
The positions underlined are the restriction recognition sites, the mutated
codons are shown in bold, and the capital letters indicate the displaced bases
from the wild type.
Construction of site-directed mutants and heterolo-
gous gene expression in E. coli. Heterologous expres-
sion of the nag3A gene (DDBJ accession no. AB078776)
using the pQE-30T vector and an N-terminal fusion to
a six-His-tag (plasmid pNAG3A) were described else-
where.6 The site-directed mutants of several codons
)
did not bind to the column were collected and desalted
by dialysis as described above, and further purified using
a MonoQ HR5/5 column (0:5 ꢁ 5 cm; Amersham
Pharmacia Biotech) equilibrated with 50 mM Tris–HCl
buffer (pH 7.5).
1
6)
were constructed using the overlap extension method
with pNAG3A as a template. The mutants constructed
and the oligonucleotides used for site-directed muta-
genesis are listed in Table 1. The PCR products were
digested with BamHI and SalI and ligated to the pQE-
3
0T vector digested with the same enzymes. After
Enzyme and protein assays. 4-Methylumbelliferyl ꢀ-
D-N-acetylglucosamine [4-MU-(GlcNAc); Sigma] was
transformation of E. coli XL1-Blue cells, the resulting
plasmid constructs were prepared and sequenced to
verify the presence of the desired mutations. E. coli
M15 cells were transformed with the original pNAG3A
and the mutant plasmids for gene expression. Expression
of the nag3A gene and its derivatives was induced by the
addition of isopropyl-ꢀ-D-thiogalactopyranoside (final
concentration, 1 mM) to the cultures growing in LB
0
dissolved in N,N -dimethylformamide at a concentration
ꢀ
of 10 mM and stored at ꢂ20 C. Nag activity was
measured in 10 mM potassium phosphate buffer (pH 7.0)
containing 0.1 mM 4-MU-(GlcNAc) as a substrate
according to a modification of the procedure of
1
7)
O’Brien. Reaction mixtures (1 ml) containing 0.5 unit
ꢀ
of the respective enzymes were incubated at 37 C for
ꢀ
medium at 37 C at an OD600 ¼ 0:6{0:8, and incubated
10 min, and the amount of 4-methylumbelliferone
released from the substrate was spectrofluorometrically
measured with an excitation wavelength at 355 nm and
an emission wavelength at 465 nm on a Shimadzu
spectrofluorometer RF-1500 (Shimadzu, Tokyo). One
unit of Nag activity was defined as the amount of
enzyme that produces 1 mmol of 4-MU per min. Protein
concentration was measured according to the method of
with vigorous shaking for 4 h after induction.
Purification of recombinant Nags. The wild-type and
mutant Nags produced by the recombinant E. coli M15
were purified with a HiTrap chelating HP column (1 ml;
Amersham Pharmacia Biotech, Tokyo) according to the
supplier’s protocol. Active fractions were collected and
desalted by dialysis against 20 mM Tris–HCl buffer
1
8)
Bradford with bovine serum albumin (Sigma) as a
standard.
(
pH 7.5). The enzyme was treated with thrombin
protease (10 units per mg of each Nag) and subjected
to purification on a Ni-NTA spin column. Proteins that
Circular dichroism. Circular dichroism measurements

1130
H. LI et al.
were performed using a Jasco automatic recording
spectropolarimeter Model J-720M (Jasco, Tokyo). The
measurements were made in 20 mM Tris–HCl buffer
D175N
D229N
Nag3A
M
M
kDa
CE E1 E2 CE E1 E2 PP E2
97
(
pH 7.5) in a cylindrical cell with a light path of 1 mm
66
and constant nitrogen flushing. The scan wavelength was
from 200 to 250 nm. The protein concentration was
about 200 mg/ml. Each spectrum was scanned once at
45
5
nm/min. Calculations were made according to ½ꢂꢃ ¼
3
2
0
0
ꢀ
ꢂ M=100 lc, where M is the mean residue molecular
weight (taking 110.68 for the wild type, D119N, D175N,
D229N, D274N, D303N, and D329N, 110.57 for D175A
and D229A, and 110.53 for E121A), ꢂ is the measured
ellipticity in degree, l is light pass of the cell in
decimeters, and c is the protein concentration in grams
ꢀ
Fig. 2. SDS–PAGE of D175N, D229N, and Wild-Type Nag3A.
CE, crude enzyme; E1, eluate from HiTrap column; E2, eluate
from MonoQ column; PP, precipitated protein; M, molecular mass
markers (molecular masses shown at the right).
1
9)
per cubic centimeter.
Results
Heterologous gene expression and purification of
recombinant wild-type and mutant Nags
Enzyme activity of wild-type and mutant Nags
The wild-type and mutant Nags purified from E. coli
were analyzed for their specific activities and Km and
Vmax values. Specific activity was measured in 10 mM
phosphate buffer (pH 7.0) containing 100 mM 4-MU-
(GlcNAc) as substrate, and the Km and Vmax values were
determined with the same substrate at a concentration
from 1/2 Km to 2 Km by Lineweaver-Burk plots. As
shown in Table 2, neither D175N nor D175A showed
any detectable activity, strongly suggesting that Asp175
residue has an important role, probably as one of the
catalytic residues in the catalysis of Nag3A. Mutant
enzymes E121Q, E121A, D229N, D229A, D119N, and
D303N lost almost all activity (less than 5% left), and
E293Q kept about 80% of the activity of the wild-type
enzyme. On the other hand, D329N showed a little
higher activity than the original activity. When the Km
values of these enzymes were compared, mutants
D119N, D229A, D229N, and D274N showed extremely
high Km values, 17–41 times as high as that of the wild
type, two mutants in Glu121 showed 50% increases, and
the other three mutants kept the Km value of the wild
type.
Eleven different mutant genes derived from nag3A
were successfully constructed using the strategy describ-
ed in ‘‘Materials and Methods.’’ Sequencing of the
constructs showed that all of them carried the planned
mutation. The wild-type and mutant Nag proteins were
purified from recombinant E. coli M15 cells in a one-
step procedure over the HiTrap chelating HP column,
and all of the purified preparations appeared to be more
than 95% homogeneous as judged by SDS–GAGE.
Typical examples of SDS–PAGE profiles are shown in
Fig. 2. All the proteins were produced in soluble state in
E. coli cell lysates, i.e., there were no inclusion bodies
formed in any of the mutants. The yield was 30 to 40 mg
of each soluble Nag protein per liter of E. coli culture,
but partial protein precipitation of mutant proteins
D229A and D229N occurred during dialysis against
2
0 mM Tris–HCl buffer (pH 7.5) after purification on the
HiTrap chelating HP column. The precipitated proteins
showed migration in SDS–PAGE identical to that of the
same proteins in soluble form (Fig. 2).
Table 2. Enzyme Activities, Km and Vmax of Wild-Type and Mutant Nags
Vmax
Enzyme activity
Specific activity (U/mg)
Nag
Km (mM)
Relative activity (%)
ꢂ1
ꢂ1
)
(
mmol min mg
Wild type
D119N
E121Q
E121A
D175A
D175N
D229N
D229A
D274N
E293Q
D303N
D329N
20.3
349.2
30.3
30.7
ND
41.3
2.9
41.3
2.0
100
4.8
1.1
0.45
0.42
ND
ND
0.81
0.80
7.5
0.45
0.42
ND
ND
0.52
0.51
5.4
1.0
ND
ND
1.3
ND
833.5
823.6
474.5
24.5
20.8
21.5
1.2
13.1
78.9
4.1
32.6
1.7
32.6
1.7
45.1
45.1
109.2
ND, not detectable

Clostridium paraputrificum N-Acetyl-ꢀ-D-glucosaminidase
1131
A
A
1
00
1
00
WT
Wild type
D119N
E121Q
D229N
E293Q
8
6
4
2
0
0
0
0
0
D119N
E121Q
D229N
E293Q
8
6
4
0
0
0
20
0
3
4
5
6
7
pH
8
9
10 11 12
2
0
30
40
50
60
70
80
Temperature (°C)
B
1
00
E121A
D229A
D274N
D303N
D329N
8
6
4
2
0
0
0
B
E121A
D229A
D274N
D303N
D329N
1
00
8
6
4
2
0
0
0
0
0
0
0
3
4
5
6
7
8
9
10 11 12
pH
2
0
30
40
50
60
70
80
Fig. 3. Optimum pH of Wild-Type and Mutant Nags.
Temperature (°C)
The enzyme activity was measured in Brittion-Robinson’s
universal buffer solution (40 mM phosphoric acid-40 mM boric
acid-40 mM acetic acid; pH was adjusted with NaOH) at various
pHs containing 0.1 mM 4-MU-(GlcNAc) as a substrate. Reaction
Fig. 4. Optimum Temperature of Wild-Type and Mutant Nags.
The enzyme activity was measured in 10 mM potassium phosphate
buffer (pH 7.0) containing 0.1 mM 4-MU-(GlcNAc) as a substrate.
ꢀ
mixtures were incubated at 37 C for 10 min. The maximum activity
was expressed as 100%. Figures are arbitrarily divided into A and B
for easier identification of graphic symbols.
ꢀ
Reaction mixtures were incubated at 37 C for 10 min. The
maximum activity was expressed as 100%. Figures were arbitrarily
divided into A and B for easier identification of graphic symbols.
Optimum pH and optimum temperature of wild-type
and mutant Nags
10
The optimum pH was measured in Brittion-Robin-
son’s universal buffer solution (40 mM phosphoric acid-
4
0 mM boric acid-40 mM acetic acid; pH was adjusted
0
with NaOH) at various pHs. As shown in Fig. 3, it was
found that the optimum pH of mutant D229N and
D229A shifted from pH 7.0 to pH 6.0, but there was no
significant difference in optimum pHs of the other
mutant Nags as compared with the wild-type enzyme.
On the other hand, (Fig. 4), changes in their optimum
temperatures were remarkable. Only mutants D329N
and E293Q kept the optimum temperature of the wild-
type enzyme. The optimum temperature for all the other
-10
-
20
200
210
220
230
240
250
ꢀ
ꢀ
Wavelength (nm)
mutant enzymes shifted from 50 C to 40 C.
Wild type
E121Q
D175N
D175A
Circular dichroism of wild-type and mutant Nags
Circular dichroism provides a direct method for
determining the gross conformational properties of
protein. Although there was a small difference in
circular dichroism profiles among D175N, E121Q, and
wild-type Nag3A, the other mutant proteins showed
almost the same profiles as the wild type, as shown in
Fig. 5. This means that most of the mutant proteins
retain the native conformation of the wild-type enzyme
Nag3A.
Fig. 5. Far-UV CD Spectra of Wild-Type and Mutant D175N,
D175A and E121Q Nags.
Far-UV CD spectra were measured in 20 mM Tris–HCl, pH 7.5.
Start wavelength 250 nm, end wavelength 200 nm, response 8
second, step resolution 0.1 nm, scan speed 5 nm/min, number of
scan 1, sensitivity 50 mdeg. Mutants other than D175N, D175A and
E121Q Nags gave profiles similar to the wild type.
cnrs-mrs.fr/CAZY/index.html) contains ꢀ-glucosidase
(EC 3.2.1.21); xylan 1,4-ꢀ-xylosidase (EC 3.2.1.37); ꢀ-
Discussion
N-acetylhexosaminidase (EC 3.2.1.52); glucan 1,3-ꢀ-
glucosidase (EC 3.2.1.58); glucan 1,4-ꢀ-glucosidase
(EC 3.2.1.74); exo-1,3-1,4-glucanase (EC 3.2.1.-); and
Family 3 of the glycoside hydrolases (http://afmb.

1132
H. LI et al.
ꢁ
-L-arabinofuranosidase (EC 3.2.1.55). In general, the
that of ExoI. Although some mutations affected opti-
mum pH and temperature of the parental enzyme, as
shown in Figs. 3 and 4, these effects do not appear to
have had a substantial influence on the Km and Vmax
values of the mutant enzymes shown in Table 2, because
the enzyme assay condition, viz., 10 min incubation at
active sites of glycoside hydrolases are composed of
aspartic acid or/and glutamic acid residues: one serves
as catalytic nucleophile and another as catalytic proton
donor. Barley ꢀ-D-glucan exohydrolase isoenzyme ExoI
was the first reported structure of a family-3 glycoside
hydrolase, which was determined by X-ray crystallog-
raphy to be a globular protein composed of two
domains. In this enzyme, Asp285 in the N-terminal
domain and Glu491 in the C-terminal domain are
considered to be involved in catalysis as nucleophile
and proton donor, respectively.12–14) Based on this
finding, a putative active site of Nags in this family
was also deduced to be Asp242 of Vibrio furnissii
ꢀ
pH 7.0 and 37 C, was not far from optimal conditions
for all the enzymes. All the mutant enzymes were
ꢀ
optimally active at 40 C or higher, and D229N and
D229A at pH 7.0 retained about 80% of the highest
activity at pH 6.0.
Asp119, Asp229, and Asp274 residues must be
involved in substrate recognition and binding because
mutations in these positions reduced the affinity of
Nag3A for the substrate at least 17 times as judged from
their Km values. Asp229 is different from the other
highly conserved Asp and Glu residues in that mutation
Asp229 into Asn and Ala strongly affected various
enzyme properties such as kinetic parameters, optimum
temperature, and optimum pH. Furthermore, the mutant
proteins D229A and D229N became unstable; i.e.,
partial precipitation of the enzyme proteins occurred
while the elutates from the HiTrap column were
dialyzed against 20 mM Tris–HCl buffer (pH 7.5).
Mutations in Asp119, Glu121, Asp229, Asp274, and
Asp303 not only affected the strength of catalytic
activity but also decreased their optimum temperature
ExoII1 and Asp303 of C. paraputrificum Nag3A,
0)
6)
1
2)
corresponding to Asp285 of ExoI. However, another
active site, a putative proton donor, was not expected in
single-domain enzymes, including C. paraputrificum
Nag3A, from the information about barley ExoI, since
Nag3A and other single-domain enzymes of family 3
did not contain a second domain. On the other hand, in
V. furnissii ExoII, a single-domain enzyme composed of
3
29 amino acid residues, Asp242 was identified as the
catalytic nucleophile by use of the mechanism-based
reagent 2-acetamido-2-deoxy-5-fluoro-ꢁ-L-idopyranosyl
fluoride. As described above, Asp242 of V. furnissii
ExoII corresponds to Asp285 of barley ExoI, identified
2
0)
ꢀ
ꢀ
as the nucleophile.
We expected that a second
from 50 C to 40 C, i.e., mutations in these positions
strongly affected the structural stability of the enzyme.
On the other hand, Asp329 and Glu293, which are less
conserved among family-3 Nags, are not important in
the catalytic mechanism of Nag3A, because both D329N
and E293Q mutants maintained enzymatic properties
catalytic residue existed in highly conserved Asp or
Glu residues of the 430 amino acids of C. paraputrii-
ficum Nag3A, and we introduced mutations into con-
served Asp and Glu residues.
Mutation of Asp303 into Asn did not abolish but
strongly affected the catalytic activity of Nag3A, i.e., the
remaining activity accounted for 4.1% of the original
activity of the wild-type enzyme (Table 2). This result is
similar to the observation that displacement of Asp52,
the catalytic nucleophile, of chicken egg white lysozyme
did not result in complete loss of the original activ-
such as K , Vmax, optimum pH, and optimum temper-
m
ature quite similar to those of the wild-type enzyme.
In conclusion, it is quite possible that Asp303 is the
catalytic nucleophile and Asp175 the proton donor of
C. paraputrificum Nag3A. Tertiary structure analysis of
Nag3A should be done to identify the catalytic residues
correctly.
21,22)
ity.
Taking previous studies about barley ExoI and
V. furnissii ExoII into consideration, it is most plausible
that Asp303 is the catalytic nucleophile of Nag3A.
When mutations were separately introduced into
Glu121, Asp175, and Asp229 that were completely
conserved among all Nags except barley ExoI, the
mutations in all these residues drastically lowered Vmax,
suggesting that one of them was the proton donor.
Neither D175N nor D175A showed any catalytic
activity, although the mutations in Glu121 did not
completely deaden the activity. Therefore, we assume
that Asp175 is the first candidate for a proton donor and
Glu121 the second candidate. Only a slight difference
in circular dichroism profiles was observed between
D175N and the wild type, and no difference was
detected between the circular dichroism profiles of
D175A and the wild type (Fig. 5). This Asp residue is
not conserved in barley ExoI (Fig. 1), suggesting that
the proton donor of Nag3A is completely different from
Acknowledgments
This work was partly supported by the New Energy
and Industrial Technology Development Organization
(NEDO) of Japan.
References
1
)
Evvyernie, D., Yamazaki, S., Morimoto, K., Karita, S.,
Kimura, T., Sakka, K., and Ohmiya, K., Identification
and characterization of Clostridium paraputrificum M-
2
1, a chitinolytic, mesophilic and hydrogen-producing
bacterium. J. Biosci. Bioeng., 89, 596–601 (2000).
Evvyernie, D., Morimoto, K., Karita, S., Kimura, T.,
Sakka, K., and Ohmiya, K., Conversion of chitinous
wastes to hydrogen gas by Clostridium paraputrificum
M-21. J. Biosci. Bioeng., 91, 339–343 (2001).
2
)
3) Morimoto, K., Kimura, T., Sakka, K., and Ohmiya, K.,

Clostridium paraputrificum N-Acetyl-ꢀ-D-glucosaminidase
1133
Overexpression of a hydrogenase gene in Clostridium
paraputrificum to enhance hydrogen gas production.
FEMS Microbiol. Lett., 246, 229–234 (2005).
Morimoto, K., Karita, S., Kimura, T., Sakka, K., and
Ohmiya, K., Cloning, sequencing, and expression of the
gene encoding Clostridium paraputrificum chitinase
ChiB and analysis of the functions of novel cadherin-
like domains and a chitin-binding domain. J. Bacteriol.,
190 (1999).
13) Hrmova, M., Gori, R. D., Smith, B. J., Vasella, A.,
Varghese, J. N., and Fincher, G. B., Three-dimensional
structure of the barley ꢀ-D-glucan glucohydrolase in
complex with a transition state mimic. J. Biol. Chem.,
279, 4970–4980 (2004).
14) Hrmova, M., Varghese, J. N., Gori, R. D., Smith, B. J.,
Driguez, H., and Fincher, G. B., Catalytic mechanism
and reaction intermediates along the hydrolytic pathway
of a plant ꢀ-D-glucan glucohydrolase. Structure, 9,
1005–1016 (2001).
15) Mursheda, K. A., Hayashi, H., Karita, S., Goto, M.,
Kimura, T., Sakka, K., and Ohmiya, K., Importance of
the carbohydrate-binding module of Clostridium ster-
corarium Xyn10B to xylan hydrolysis. Biosci. Biotech-
nol. Biochem., 65, 41–47 (2001).
16) Ho, S. N., Hunt, H. D., Horton, R. M., Pullen, J. K., and
Pease, L. R., Site-directed mutagenesis by overlap
extension using the polymerase chain reaction. Gene,
77, 51–59 (1989).
17) O’Brien, M., and Colwell, R. R., A rapid test of chitinase
activity that uses 4-methylumbelliferyl-N-acetyl-ꢀ-D-
glucosaminide. Appl. Environ. Microbiol., 53, 1718–
1720 (1987).
18) Bradford, M. M., A rapid and sensitive method for the
quantitation of protein utilizing the principle of protein-
dye binding. Anal. Biochem., 72, 248–254 (1976).
19) Umekawa, H., Endo, T., and Hidaka, H., A rapid
separation of bovine brain S-100a and S-100b protein
and related conformation studies. Arch. Biochem. Bio-
phys., 227, 147–153 (1983).
20) Vocadlo, D. J., Mayer, C., He, S., and Withers, S. G.,
Mechanism of action and identification of Asp242 as the
catalytic nucleophile of Vibrio furnisii N-acetyl-ꢀ-D-
glucosaminidase using 2-acetamido-2-deoxy-5-fluoro-ꢁ-
L-idopyranosyl fluoride. Biochemistry, 39, 117–126
(2000).
21) Malcolm, B. A., Rosenberg, S., Corey, M. J., Allen, J. S.,
Baetselier, A. D., and Kirsch, J. F., Site-directed muta-
genesis of the catalytic residues Asp-52 and Glu-35 of
chicken egg white lysozyme. Proc. Natl. Acad. Sci. USA,
86, 133–137 (1989).
4
)
179, 7306–7314 (1997).
5
)
)
Morimoto, K., Karita, S., Kimura, T., Sakka, K., and
Ohmiya, K., Sequencing, expression, and transcription
analysis of the Clostridium paraputrificum chiA gene
encoding chitinase ChiA. Appl. Microbiol. Biotechnol.,
51, 340–347(1999).
6
Li, H., Morimoto, K., Katagiri, N., Kimura, T., Sakka,
K., and Ohmiya, K., A novel ꢀ-N-acetylglucosaminidase
of Clostridium paraputrificum M-21 with high activity
on chitobiose. Appl. Microbiol. Biotechnol., 60, 420–427
(
2002).
7
)
)
Li, H., Morimoto, K., Kimura, T., Sakka, K., and
Ohmiya, K., A new type of ꢀ-N-acetylglucosaminidase
from hydrogen-producing Clostridium paraputrificum
M-21. J. Biosci. Bioeng., 96, 268–274 (2003).
Coutinho, P. M., and Henrissat, B., The modular
structure of cellulases and other carbohydrate-active
enzymes: an integrated database approach. In ‘‘Genetics,
Biochemistry and Ecology of Cellulose Degradation,’’
eds. Ohmiya, K., Hayashi, K., Sakka, K., Kobayashi, Y.,
Karita, S., and Kimura, T., Uni Publishers, Tokyo,
pp. 15–23 (1999).
Tsujibo, H., Fujimoto, K., Tanno, H., Miyamoto, K.,
Imada, C., Okami, Y., and Inamori, Y., Gene sequence,
purification and characterization of N-acetyl-beta-gluco-
saminidase from a marine bacterium, Alteromonas sp.
strain O-7. Gene, 146, 111–115 (1994).
8
9)
1
1
1
0) Chitlaru, E., and Roseman, S., Molecular cloning and
characterization of a novel beta-N-acetyl-D-glucosami-
nidase from Vibrio furnissii. J. Biol. Chem., 271, 33433–
33439 (1996).
1) Cheng, Q., Li, H., Merdek, K., and Park, J. T., Molecular
characterization of the ꢀ-N-acetylglucosaminidase of
Escherichia coli and its role in cell wall recycling. J.
Bacteriol., 182, 4836–4840 (2000).
2) Varghese, J. N., Hrmova, M., and Fincher, G. B., Three-
dimensional structure of a barley ꢀ-D-glucan exohydro-
lase, a family 3 glycosyl hydrolase. Structure, 7, 179–
22) Hashimoto, Y., Yamada, K., Motoshima, H., Omura, T.,
Yamada, H., Yasukochi, T., Miki, T., Ueda, T., and
Imoto, T., A mutation study of catalytic residue Asp 52
in hen egg lysozyme. J. Biochem., 119, 145–150 (1996).