(Received for publication, September 17, 1996, and in revised form, January 30, 1997)
From the Laboratory of Biochemistry, Institut
Químic de Sarrià, Universitat Ramon Llull, 08017 Barcelona, Spain and the § Institut de Biologia Fonamental
V. Villar Palasí and the Department de Bioquímica i
Biologia Molecular, Universitat Autonoma de Barcelona,
08193 Bellaterra, Barcelona, Spain
The carbohydrate-binding cleft of Bacillus
licheniformis 1,3-1,4--D-glucan 4-glucanohydrolase
is partially covered by the surface loop between residues 51 and 67, which is linked to
-strand-(87-95) of the minor
-sheet III of
the protein core by a single disulfide bond at
Cys61-Cys90. An alanine scanning mutagenesis
approach has been applied to analyze the role of loop residues from
Asp51 to Arg64 in substrate binding and
stability by means of equilibrium urea denaturation, enzyme
thermotolerance, and kinetics. The
GU
between oxidized and reduced forms is approximately constant for all
mutants, with a contribution of 5.3 ± 0.2 kcal·mol
1 for the disulfide bridge to protein
stability. A good correlation is observed between
GU values by reversible unfolding and enzyme thermotolerance. The N57A mutant, however, is more thermotolerant than
the wild-type enzyme, whereas it is slightly less stable to reversible
urea denaturation. Mutants with a <2-fold increase in
Km correspond to mutations at residues not involved in substrate binding, for which the reduction in catalytic efficiency (kcat/Km) is proportional
to the loss of stability relative to the wild-type enzyme. Y53A, N55A,
F59A, and W63A, on the other hand, show a pronounced effect on
catalytic efficiency, with Km > 2-fold and
kcat < 5% of the wild-type values. These
mutated residues are directly involved in substrate binding or in
hydrophobic packing of the loop. Interestingly, the mutation M58A
yields an enzyme that is more active than the wild-type enzyme (7-fold
increase in kcat), but it is slightly less
stable.
1,3-1,4--D-Glucan 4-glucanohydrolase
(1,3-1,4-
-glucanase1; EC 3.2.1.73) is an
endo-glycosidase that hydrolyzes
-glucans containing
mixed
-1,3- and
-1,4-linkages as lichenin and cereal
-glucans.
The enzyme has a strict cleavage specificity for
-1,4-glycosidic bonds in 3-O-substituted glucopyranose units (1, 2). Genes encoding bacterial 1,3-1,4-
-glucanases have been cloned and
sequenced from different Bacillus species (3-9),
Fibrobacter succinogenes (10), Ruminococcus
flavofaciens (11), and Clostridium thermocellum (12).
Together with 1,3-
-glucanases ("laminarinases"), all bacterial
1,3-1,4-
-glucanases share a high degree of sequence similarity and
have been classified as members of family 16 of glycosylhydrolases (13,
14).
Bacillus licheniformis 1,3-1,4--glucanase is a retaining
glycosidase (2), acting by general acid/base catalysis in a double displacement mechanism (15). Glu138 has been proposed as
the proton donor residue and Glu134 as the catalytic
nucleophile (16, 17). The three-dimensional structure, recently refined
at 0.18-nm resolution by x-ray crystallography (18), is almost
identical to that of the hybrid H(A16M) between Bacillus
amyloliquefaciens and Bacillus macerans (19) and the B. macerans (20) enzymes. It has a jelly-roll
-sandwich
fold, with the carbohydrate-binding cleft located on the concave face of a
-sheet formed by seven antiparallel
-strands (see Fig. 1).
The Bacillus enzymes are unrelated to the plant
1,3-1,4-
-glucanases in both sequence similarity (family 17 of
glycosylhydrolases) and three-dimensional structure (
/
-barrel
structure), clearly indicating that the identical substrate
specificities have arisen by convergent evolution (21). On the
other hand, the Bacillus enzymes show structural
similarities to plant legume lectins and family 7 cellulases.
Cellobiohydrolase I from Trichoderma reesei (22) has a very
similar fold, with most of the
-sandwich residues in the protein
core being superimposable, but it has long loops shaping the
substrate-binding tunnel that are missing in the 1,3-1,4-
-glucanase structure.
Except for the Bacillus brevis isozyme, all
Bacillus 1,3-1,4--glucanases possess a single disulfide
bond at Cys61-Cys90 (B. licheniformis numbering) that connects a
-strand (residues 87-95) with a loop from residues 51 to 67 (see Fig. 1). This major loop is located on the concave side of the molecule, is
solvent-exposed, and partially covers the active-site cleft. Even
though no three-dimensional structure of an enzyme-inhibitor complex
with a carbohydrate inhibitor filling the entire binding cleft is yet
available, the three-dimensional structure of a covalent complex
between the hybrid H(A16M) and epoxybutyl
-cellobioside (19) and the
molecular model of an enzyme-substrate complex made by computational
methods (23) indicate that some loop residues might interact with a
substrate occupying distant subsites on the nonreducing end of the
binding site cleft.
Here we use the technique of alanine scanning mutagenesis (24) to
analyze the role of loop residues (from Asp51 to
Arg64) in B. licheniformis 1,3-1,4--glucanase
in substrate binding and stability by means of equilibrium urea
denaturation, enzyme thermotolerance, and kinetics. Previous studies of
the disulfide bond at Cys61-Cys90 have shown
the deleterious effect of cysteine-to-alanine mutations on protein
stability and activity, but no effect of disulfide bond reduction on
activity (25). These results suggested that the loop has little
flexibility and that the disulfide bond is not required to keep the
structural integrity of the loop. Other hydrophobic interactions may
position the loop to shape the active-site cleft.
Escherichia
coli TG1 (supE hsd5 thi
(lac-proAB)
F
[traD36 proAB+
lacIq lacZ
M15]) was used for plasmid
propagation, transformation with the mutagenic polymerase chain
reaction (PCR), and protein expression. For plasmid isolation, bacteria
were grown in 2YT medium (26), and 2SB medium (27) was used for protein
expression. Ampicillin at 100 µg/ml was added when appropriate.
Urea (molecular biology-grade) was
purchased from Sigma; dithiothreitol, 3,5-dinitrosalicilic acid, and
5,5-dithiobis(2-nitrobenzoic acid) were from Fluka. Restriction
endonucleases and T4 DNA ligase were from Boehringer Mannheim, and
DeepVent® polymerase was from New England Biolabs Inc.
-35S-ATP was purchased from Amersham Corp. DNA
sequencing was performed with the T7 sequencing kit from Pharmacia
Biotech Inc. Oligonucleotides were synthesized by Boehringer Mannheim.
Barley
-glucan was from Megazyme (Sydney, Australia). All buffers
and solutions for kinetic and urea denaturation experiments were
degassed prior to use.
The gene coding for
B. licheniformis -glucanase previously cloned from the
genomic DNA (8) and subcloned in pUC119 as a 1.21-kilobase
SacI/SphI fragment (16) was used as the template for mutagenic PCR following the method previously reported for other
1,3-1,4-
-glucanase mutants (17). The first PCR used the mutagenic
primers and the reverse universal primer flanking the 5
-end of the
1,3-1,4-
-glucanase gene. The primers were as follows (mismatches are
in boldface): D51A,
5
-CCATTCGAGTACCCAGCTGCTTTTTGCCAT-3
; G52A,
5
-GTTTCCATTCGAGTACGCATCTGCTTTTTGCC-3
; Y53A,
5
-AAACATGTTTCCATTCGAGGCCCCATCTGCTTTTTGCC-3
; S54A,
5
-AAACATGTTTCCATTCGCGTACCCATCTGCTTTTTGCC-3
; N55A, 5
-TAAACATGTTTCCAGCCGAGTACCCATCTGCTTTTTGCC-3
; G56A,
5
-ACAGTTAAACATGTTTGCATTCGAGTACCCATC-3
; N57A,
5
-ACAGTTAAACATGGCTCCATTCGAGTACCC-3
; M58A,
5
-CGCCACGTACAGTTAAACGCGTTTCCATTCGAGT-3
; F59A,
5
-GCACGCCACGTACAGTTAGCCATGTTTCCATTCG-3
; N60A,
5
-GCACGCCACGTACAGGCAAACATGTTTCCATTCG-3
; T62A,
5
-TGTTTGCACGCCACGCACAGTTAAACATGTTTCC-3
; W63A,
5
-GGAGACATTGTTTGCACGCGCCGTACAGTTAAACATGTTTCC-3
; and R64A,
5
-GGAGACATTGTTTGCAGCCCACGTACAGTTAAAC-3
. The second
PCR used the product of the first PCR as a primer and the forward
universal primer to yield the whole 1,3-1,4-
-glucanase gene with the
desired mutation. The mutated gene was cut with EcoRI/HindIII and ligated again to a pUC119
vector. After transformation of E. coli TG1 cells,
transformants were screened by DNA sequencing using appropriate primers
located ~100 bases from the mutation point. Positive clones were
confirmed by complete sequencing of the entire gene.
Proteins were purified from the supernatant of E. coli TG1 cultures harboring the mutagenized plasmids basically as described before (28) with an additional purification step of fast protein liquid chromatography on an ion-exchange TSK CM-3SW column in 5 mM acetate buffer, pH 5.6, and elution with a linear gradient of 0-0.4 M NaCl in the same buffer. The proteins were analyzed by SDS-polyacrylamide gel electrophoresis as described (29) and by fast protein liquid chromatography on a TSK CM-3SW column at pH 5.6. Enzyme concentrations were determined by absorbance at 280 nm using A1 mg/ml = 14.5 absorbance units for the wild-type enzyme and by the Bradford protein assay (30) for the mutants using the wild-type enzyme as a standard. Spectrophotometric and kinetic measurements were performed on a Varian Cary 4 spectrophotometer with a Peltier temperature control system.
Enzyme Assay and Kinetics1,3-1,4--Glucanase activity on
plates was detected by the Congo red assay after growing the E. coli TG1 cells containing the mutagenized plasmids on LB plates
supplemented with 0.04% (w/v) barley
-glucan (31). Activity in the
supernatant from liquid cultures and the enzyme activity of purified
enzymes were determined as previously reported (28) by measuring the
net release of reducing sugars from barley
-glucan in
citrate/phosphate buffer (6.5 mM citric acid, 87 mM Na2HPO4), pH 7.2, 0.1 mM CaCl2 using the 3,5-dinitrosalicilic acid
reagent (32). Kinetic measurements were performed using the synthetic
substrate 4-methylumbelliferyl 3-O-
-cellobiosyl-
-D-glucopyranoside in
citrate/phosphate buffer, pH 7.2, 0.1 mM CaCl2
by measuring the release of 4-methylumbelliferone at 365 nm (33, 34).
Kinetic parameters were derived by fitting the data to a substrate
inhibition model (v = kcat·[E]0/([S] + Km + [I]2/KI)) by
means of nonlinear regression analysis (50).
Unfolding was monitored by
fluorescence spectroscopy in a Perkin-Elmer LS50 spectrofluorometer,
with excitation at 282 nm (3-nm slit) and the emission spectra being
recorded from 270 to 440 nm (8-nm slit) and measured at 340 nm, in
thermostatted cuvette holders at 37 °C. For each data point
collected, wild-type or mutant 1,3-1,4--glucanases in
citrate/phosphate buffer, pH 7.2, were diluted to 1 µg/ml in degassed
urea solution in the same buffer and incubated overnight at 37 °C.
To obtain reduced enzymes, the protein stock solutions (25 µg/ml)
were made 200 mM dithiothreitol and incubated for 30 min at
25 °C before being added to the denaturant solution. The final
dithiothreitol concentration was 10 mM, enough to avoid
reoxidation for at least 24 h as shown by free sulfhydryl titration with 5,5
-dithiobis(2-nitrobenzoic acid) (after removal of
the excess dithiothreitol by means of a pD10 G25 desalting column from
Pharmacia).
For
determination of enzyme thermotolerance, samples of 50 µg/ml enzyme
in 50 mM sodium acetate buffer, pH 6.0, 20 mM
CaCl2 were incubated at 65 or 70 °C. Aliquots of 80 µl
were withdrawn at various time intervals (until complete inactivation)
and immediately diluted 5-fold in ice-cold water. The residual activity
was determined at 45 °C using 4-methylumbelliferyl
3-O--cellobiosyl-
-D-glucopyranoside (3 mM assay concentration) and barley
-glucan (5 mg/ml
assay concentration) in citrate/phosphate buffer, pH 7.2, 0.1 mM CaCl2. The enzymatic half-life
(t50) was calculated by fitting the first phase
of the plot residual activity versus incubation time to a
single exponential decay.
Point mutations to alanine in the loop residues from Asp51 to Arg64 (Fig. 1) were prepared by site-directed mutagenesis by PCR. The mutant proteins were purified up to 95% as judged by SDS-polyacrylamide gel electrophoresis following the procedure described for the wild-type enzyme (28). Expression and purification yields were similar for all the mutant and wild-type enzymes. Proteins were stored in their oxidized form. Reduction of the disulfide bond at Cys61-Cys90 (reduced enzymes) was done just before their use as described under "Materials and Methods."
Analysis of Stability by Equilibrium Urea DenaturationThe
stability of the enzymes reported in this study was examined by urea
denaturation assuming a two-state transition using the model of Clarke
and Fersht (35). Unfolding was monitored by measuring the dependence of
fluorescence intensity on urea concentration. The data were analyzed as
described previously for other -glucanase mutants (25) using
Equation 1 (for its derivation, see Ref. 35),
![]() |
(Eq. 1) |
![]() |
(Eq. 2) |
The calculated values for m and
[D]50% are given in Table I,
and Fig. 2 plots the transition curves of the wild-type enzyme and one mutant as an example. Inspection of the m
values for the oxidized and reduced enzymes shows that they are grouped in two clusters, one for the oxidized proteins with an average m value (mav) of 2.3 ± 0.4 kcal·mol1·M
1 and the other
for the reduced forms with an mav value of
1.2 ± 0.2 kcal·mol
1·M
1. This
difference has proved to be significant by a Student's t
test (
= 0.05), indicating that the presence or absence of the
disulfide bond has a significant and constant effect on the unfolding
behavior.
|
Since individual m values for each mutant are subjected to large standard errors, we used the corresponding mav value to calculate the free energies of unfolding in the absence of denaturant for the oxidized and reduced enzymes, respectively. Then, Equation 2 becomes Equation 3.
![]() |
(Eq. 3) |
![]() |
(Eq. 4) |
|
The energetic contribution of the disulfide bridge to stability
(GUS-S) was calculated using
Equation 4 in which a and b are the reduced and
oxidized forms of the same protein, respectively. The calculated values
in Table II display, within the experimental error, a constant stabilizing effect of 5.3 ± 0.2 kcal·mol
1 for the
disulfide bridge in all wild-type and mutant enzymes.
kcat and Km
values for wild-type and mutant enzymes were determined with a specific
substrate for 1,3-1,4--glucanases recently developed by our group
(33, 34). 4-Methylumbelliferyl 3-O-
-cellobiosyl-
-D-glucopyranoside
undergoes a single glycosidic bond cleavage upon enzymatic hydrolysis
with release of the 4-methylumbelliferone chromophore, which can be
continuously monitored at 365 nm. Reactions were done in
citrate/phosphate buffer, pH 7.3, 0.1 mM CaCl2.
While the optimal temperature for the wild-type enzyme is 55 °C
(36), some of the alanine mutants are more thermolabile. 45 °C was
found to be the highest temperature for which all the proteins studied showed a linear progress curve during the initial 15 min of reaction. Substrate inhibition was observed at high concentrations, so the data
were fitted to an uncompetitive substrate inhibition model by nonlinear
regression (36). Calculated values for kcat and Km are summarized in Table III.
|
A measure of the enzyme
thermotolerance can be obtained by deducing t50
at a specified temperature. Residual activity of the enzymes was
measured after various periods of incubation at a given temperature by
steady-state kinetics with 4-methylumbelliferyl 3-O--cellobiosyl-
-D-glucopyranoside
substrate at 45 °C. Preliminary experiments with the wild-type
enzyme at 50 and 500 µg·ml
1 at 65 and 70 °C in
sodium acetate buffer, pH 6.0, showed that extensive protein
aggregation took place at high enzyme concentration and that a very low
t50 (<10 min) was obtained at 70 °C.
Therefore, we chose a protein concentration of 50 µg·ml
1 and an incubation temperature of 65 °C as
standard assay conditions. The plot of residual activity
versus incubation time follows a double exponential curve,
with the value of t50 being in the first phase
of the inactivation decay. Mutants Y53A, N55A, F59A, and W63A could not
be analyzed under these conditions due to their low activity, which
required enzyme concentrations above 500 µg·ml
1.
Values of t50 are summarized in Table I.
The carbohydrate-binding cleft of the 1,3-1,4--glucanase of
B. licheniformis is partially covered by the surface loop
between residues 51 and 67, which is linked to
-strand-(87-95) of
the minor
-sheet III (18) by the single disulfide bridge at
Cys61-Cys90. The technique of alanine scanning
mutagenesis has been applied to analyze the role of loop residues
(Asp51-Arg64) in substrate binding and
stability as well as the contribution of the disulfide bridge to
stability.
Unfolding transition curves are
described by two parameters in a two-state model:
[D]50% is a measure of the midpoint of the
transition region, and m is a measure of the steepness of
the transition region and reflects the cooperativity of the unfolding
process. A clear distinction is observed between reduced and oxidized
forms in terms of m values, with the oxidized enzymes having
a steeper transition. Common to all models that have been proposed to
describe the dependence of the free energy of unfolding on denaturant
concentration (37, 38) is the premise that denaturants alter the
equilibrium N U through a preferential interaction with the
denatured state. Schellman (37) proposed that the parameter that can
describe the differential interaction of the native and denatured state
to denaturants is the different solvent-accessible area between both
states, AN and AU, respectively (Equation 5),
![]() |
(Eq. 5) |
The spatial distribution of the mutated residues in the
crystallographic structure of the wild-type enzyme (18) suggests that
the destabilizing effect is larger near the N- and C-terminal ends of
the loop (Table IV). This observation is in agreement with the idea that the loop edges are rigid, with a central part being
more flexible and the C-terminal end being more tightly packed as
judged by the side chain solvent accessibility and van der Waals
interaction data shown in Table IV. No correlation was found between
the experimental GU values and the free
energy of transfer of amino acid side chains from water to octanol
(corrected or not for solvent-exposed area) or the number of atoms
inside a sphere around C-
of the mutated amino acid residue (Table
IV). Such correlations have been shown to work properly in a number of
proteins (41-44) for series of mutants in hydrophobic regions of the
protein structure. This is not the case for the 1,3-1,4-
-glucanase mutants probably because the loop is partially solvent-exposed and some
of the residues are hydrophilic.
|
The
calculated values for GUS-S
in Table III indicate a constant stabilizing effect of 5.3 ± 0.2 kcal·mol
1 for the disulfide bridge in all mutant and
wild-type enzymes. This value is larger than that previously estimated
for the wild-type enzyme (0.7 kcal·mol
1 (25)). The
m value for the oxidized wild-type enzyme deviates from the
general trend observed for the mutants, and it is much closer to the
m value for the reduced wild-type enzyme. In the absence of
mutant data, our first estimation was performed using an
mav value of 1.31 kcal·mol
1·M
1 for both forms
of the wild-type enzyme. However, the large number of mutants studied
here clearly shows a significant difference between oxidized and
reduced forms. Even though the behavior of the wild-type enzyme might
be different, the general trend observed here allows us to conclude
that the disulfide bond has a larger contribution to protein
stability.
Enzyme thermotolerance was determined at
65 °C as the incubation time required to irreversibly inactivate the
enzyme to 50% of its initial activity (t50). A
good correlation was observed between GU
(from equilibrium urea denaturation) and t50 for
the mutants in their oxidized form (Fig. 3), except for
N57A, which is surprisingly more thermotolerant than the wild-type
enzyme. Even though direct comparison of thermal stability and urea
denaturation is not possible in general (kinetic versus
equilibrium experiments), the results may be rationalized considering a
fast irreversible process from the denatured state at high temperature
(Reaction 1),
![]() |
![]() |
Effects on Enzyme Kinetics
The kcat
and Km values of the 1,3-1,4--glucanase mutants
(Table II) show that most of the mutations have an effect on enzyme
activity. It could be a direct effect of removing an amino acid side
chain involved in substrate binding (or interacting with an essential
catalytic residue) or an indirect effect of local rearrangements
produced by the mutation (which are also reflected in a decrease in
protein stability). kcat/Km values are plotted against stability data (as
GUH2O values) for the
oxidized enzyme forms in Fig. 4. Inspection of this plot
suggests that the mutants can be classified in four groups. Group A
(wild-type, D51A, G52A, G56A, N60A, T62A, and R64A) is formed by those
enzymes showing a good correlation between catalytic efficiency and
enzyme stability. For these mutants, the decrease in catalytic
efficiency is mainly due to kcat since Km values are <2-fold larger than the wild-type
Km value. Moreover, the mutated residues in this
group have no specific role in substrate binding as proposed from the
structure of the modeled enzyme-substrate complex. Therefore, the
reduction in kcat/Km is
interpreted as the result of local rearrangements in the protein
structure induced by the mutations, which also have a proportional
effect on protein stability. Group B mutants (S54A and N57A) slightly
deviate from the correlation. They show <2-fold reduction in
kcat and almost no effect on
Km as compared with the wild-type enzyme. Group C
(Y53A, N55A, F59A, and W63A) is composed of mutants that have a
pronounced deleterious effect on enzyme activity, with
Km > 2-fold and kcat < 5%
of the wild-type kinetic parameters. These mutated residues are
directly involved in substrate binding or in hydrophobic packing of the
loop.2 Tyr53 forms a hydrogen
bond with the 3-OH of the glucopyranose unit of the substrate in
subsite
III, whereas the amide nitrogen of Asn55
hydrogen-bonds with the 6-OH of the glucopyranose unit in subsite
II
according to the modeled enzyme-substrate complex for the B. licheniformis enzyme2 or the modeled complex for the
hybrid H(A16M)
-glucanase (23). On the other hand, Phe59
has the aromatic side chain pointing toward the core of the protein, and it has a strong hydrophobic (stacking) interaction with
Trp213, which belongs to a
-chain of the major
-sheet
on the concave face of the molecule. This interaction might be
important to position the loop and to create a hydrophobic environment
in this portion of the cleft when the substrate binds.
Trp63 also has the aromatic side chain interacting with the
main core and very close to Phe59, contributing to the
structural integrity of the loop. Finally, group D comprises the single
mutant M58A, which is more active, with a kcat
value 7-fold higher than that of the wild-type enzyme. This surprising
result was unpredictable from a simple structural analysis since the
side chain of Met58 does not interact with the substrate or
with any essential catalytic residue in the three-dimensional structure
of the free enzyme. However, replacement of the side chain by a smaller
methyl group (M58A) might allow some readjustments of the loop in the
enzyme-substrate complex that may have a favorable effect on transition
state stabilization. Up to date, only the structure of a covalent
complex between the hybrid H(A16M) 1,3-1,4-
-glucanase and epoxybutyl
-cellobioside has been solved by x-ray crystallography (19).
Inhibitor binding has no significant effect on the active-site geometry
as observed by comparing the structures of the covalent complex and the
free H(A16M) enzyme. However, the cellobiose unit of this suicide
inhibitor only fills subsites
II and
III, with subsite
I being
occupied by an alkyl chain instead of a glucopyranose ring. Since ring distortion in subsite
I is expected, the structure of this covalent complex is not a good model to analyze the structural effects that the
reported mutations may have on substrate binding and on transition
state stabilization. New three-dimensional structures of
enzyme-inhibitor complexes (or inactive mutant-substrate complexes) are
required to evaluate small but significant structural changes that
might occur upon ligand binding.
When applying an alanine scanning mutagenesis strategy, comparison between catalytic efficiency and enzyme stability provides a useful method to identify those residues that have an important role in ligand binding or in structural packing of the protein. Taking as a reference the mutants for which the reduction in catalytic efficiency is proportional to the loss of protein stability, mutations that deviate from this correlation indicate that these residues are involved in substrate binding or in maintaining the active-site structure. It is remarkable that two mutants, N57A with increased thermostability and M58A with higher catalytic efficiency, have been obtained. The effects of these mutations were unpredictable with the current knowledge of protein structure/function relationships, supporting the fact that scanning and random mutagenesis strategies are useful approaches to obtain proteins with improved properties for biotechnological applications.
We are indebted to Teresa Dot for the molecular modeling analysis.