From the
Departments of Food Engineering and
Biotechnology and
Chemistry and the
Institute of Catalysis Science and
Technology, Technion-Israel Institute of Technology, Haifa 32000, Israel, the
||Department of Inorganic Chemistry and The
Laboratory for Structural Chemistry and Biology, The Hebrew University of
Jerusalem, Jerusalem 91904, Israel, and the
**Architecture et Fonction des Macromolécules
Biologiques, UMR 6098, CNRS and Universités d'Aix-Marseille I and II,
31 Chemin Joseph Aiguier, 13402 Marseille cedex 20, France
Received for publication, April 21, 2003 , and in revised form, May 8, 2003.
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ABSTRACT |
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INTRODUCTION |
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The glycosidic bond is one of the most stable bonds in nature, with a half-life of over 5 million years (7). Glycoside hydrolases can accelerate the hydrolysis of the glycosidic bond by more than 1017-fold, making them the most efficient catalysts known. The enzymatic hydrolysis of the glycosidic bonds occurs via two major mechanisms, giving rise to either an overall retention or an inversion of the anomeric configuration. In both mechanisms, the hydrolysis usually requires two carboxylic acids, which are conserved within each glycoside hydrolase family, and proceeds through oxocarbenium ion-like transition states. Inverting glycosidases use a single displacement mechanism with the assistance of general acid and general base residues. Retaining glycosidases follow a two-step double displacement mechanism as shown in Fig. 1, involving two catalytic residues, one functioning as a nucleophile and the other functioning as an acid-base (8).
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Identification of the key active site residues is of great importance because it provides crucial information regarding the enzymatic catalytic mechanism and allows rational protein design for novel applications, such as for enzymatic synthesis (5). Candidates for the catalytic residues are primarily identified by searching for conserved carboxylates (Glu or Asp) throughout multiple amino acid sequence alignment. These conserved residues are replaced to a noncarboxylic residue, and the generated mutants are subjected to detailed kinetic analysis using substrates bearing different leaving groups, azide rescue analysis, and pH dependence activity profiles of the mutants and the wild type. In some cases, identification can be accomplished by labeling the catalytic residues using mechanism-based inactivators and affinity labels (9).
Based on amino acid sequence similarities, -D-xylosidases
are currently divided into families 3, 39, 43, 52, and 54 of glycoside
hydrolases (10,
11). These families together
with all other glycoside hydrolase families can be readily accessed at the
constantly updated web site
afmb.cnrs-mrs.fr/CAZY.
Although the identities of the catalytic residues for most of these
-D-xylosidase families are already known
(1216),
no such information is available for family 52.
Previously, we reported the cloning and purification of a -xylosidase
from Geobacillus stearothermophilus T-6 (XynB2) showing homology to
family 52 glycoside hydrolases. Its stereochemical course of hydrolysis showed
that the configuration of the anomeric carbon was retained, indicating that a
retaining mechanism prevails in family 52 glycoside hydrolases
(17). Because the
-xylosidase from G. stearothermophilus T-6 can be readily
overexpressed and purified, it can serve as an excellent representative of
family 52 glycoside hydrolases for the identification of the two key active
site residues. This paper describes a detailed kinetic analysis of the
putative acid-base and nucleophile mutants of XynB2, using substrates bearing
different leaving groups, chemical rescue, and pH dependence profiles. The
study provides for the first time unequivocal identification of the two
catalytic residues of family 52 glycoside hydrolases.
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EXPERIMENTAL PROCEDURES |
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Mutagenesis, Protein Expression, and PurificationThe xynB2 gene (GenBankTM accession number AJ305327 [GenBank] ) from G. stearothermophilus T-6 was cloned in the pET9d vector, overexpressed in Escherichia coli BL21(DE3), and purified as previously reported (17). Site-directed mutagenesis was performed using the QuikChange site-directed mutagenesis kit (Stratagene, La Jolla, CA). The mutagenic primers for the mutations were as follows (the mutated nucleotides are in bold type): D495G, 5'-GGAAATCACAACGTACGGGAGTTTGGATGTTTCTCTTGG-3' and 5'-CCAAGAGAAACATCCAAACTCCCGTACGTTGTGATTTCC-3'; E335G, 5'-GCCGATTTGGGTCGTTAACGGCGGCGAGTACCGGATGATG-3' and 5'-CATCATCCGGTACTCGCCGCCGTTAACGACCCAAATCGGC-3'; E335A, 5'-GCCGATTTGGGTCGTTAACGCCGGCGAGTACCGGATGATG-3' and 5'-CATCATCCGGTACTCGCCGGCGTTAACGACCCAAATCGGC-3'; and E335Q, 5'-GCCGATTTGGGTCGTTAACCAGGGCGAGTACCGGATGATG-3' and 5'-CATCATCCGGTACTCGCCCTGGTTAACGACCCAAATCGGC-3'. The mutagenic primers were designed to include the mutation and when possible a restriction site to allow easy identification of the mutation. All of the mutations were created by a double base pair substitution to avoid translational misincorporation during protein synthesis by the host cell. The mutated genes were sequenced to confirm that only the desired mutations were inserted, and the proteins were overexpressed and purified as described for the wild type.
Kinetic StudiesSteady state kinetic studies were performed
by following the absorbance changes in the UV-visible range, using an
Ultrospec 2100 pro spectrophotometer (Amersham Biosciences) equipped
with a temperature-stabilized water circulating bath. Initial hydrolysis rates
were determined by incubating 500 µl of different substrate concentrations
(ranging from 0.1 to 7 Km where applicable) in
100 mM phosphate buffer (pH 7.0) containing 1 mg/ml bovine serum
albumin at 40 °C within the spectrophotometer until thermal equilibration
was achieved. The exact temperature inside the cuvette was verified using a
thermocouple. The reactions were initiated by the addition of 100 µl of
appropriately diluted enzyme, and the release of the phenol-derived product
was monitored at the appropriate wavelength. For very low
Km values, the initial rates were measured with
special care. For highly reactive substrates, blank mixtures containing all of
the reactants except the enzyme were used to correct for spontaneous
hydrolysis of the substrates. Sodium azide and formate were added to the
reaction mixtures where mentioned. The extinction coefficients used at pH 7.0
and 40 °C and the wavelength monitored for each substrate were as follows:
2,5-dinitrophenyl, 420 nm, = 3.68
mM1 cm1;
3,4-dinitrophenyl, 400 nm,
= 11.15
mM1 cm1;
2,4,6-trichlorophenyl, 312 nm,
= 3.97
mM1 cm1;
4-nitrophenyl, 420 nm,
= 7.61 mM1
cm1; 2-nitrophenyl, 420 nm,
= 1.91
mM1 cm1;
4-methylumbelliferyl, 355 nm,
= 2.87
mM1 cm1;
3-nitrophenyl, 380 nm,
= 0.455
mM1 cm1.
The values of Km and kcat
were determined by nonlinear regression analysis using the program GraFit 5.0
(19).
pH dependence studies were carried at 40 °C with pNPX1 as a substrate. Mixtures containing 600 µl of 1 mg/ml bovine serum albumin and different concentrations of substrate solutions in the appropriate buffer were prewarmed until the reaction was initiated by the addition of 200 µl of appropriately diluted enzyme. The buffers used were at a final concentration of 100 mM and were: citric acid-Na2HPO4 (pH 4.56.5), phosphate buffer (pH 6.08.0), and Tris-HCl buffer (pH 7.58.5). The pH range employed in this study included only pH values for which the enzyme was stable for at least 5 min. The reactions were monitored continuously at 40 °C, and upon completion the actual pH was measured to verify that the pH had not changed. The release of p-nitrophenol was monitored at 400 nm, and the mM extinction coefficients for p-nitrophenolate were determined at pH 4.63, 5.35, 6.02, 6.53, 6.93, 7.61, 8.0, 8.3, and 8.56 as follows: 1.43, 1.89, 4.45, 7.46, 11.2, 16.2, 17.0, 17.3, and 17.6 mM1 cm1, respectively. The pKa values assigned to the ionizable groups were determined using the program GraFit 5.0.
Isolation and Analysis of Reaction Products in the Presence of Sodium
AzideThe enzymatic reactions included 0.4 mg/ml of either
XynB2-E335G or XynB2-D495G, 10 mM of 2,5-DNPX, and 1 M
sodium azide in a final volume of 10 ml of 100 mM phosphate buffer,
pH 7.0. The mixtures were incubated at 40 °C, and the reaction was
monitored by TLC. TLC analysis was performed using precoated plates (Silica
Gel 60 F254, 0.25 mm; Merck), and MeOH/CHCl3 1:4 as the
running solvents. The spots were visualized by charring with a yellow solution
containing 120 g of
(NH4)Mo7O24·4H2Oand5gof(NH4)2Ce(NO3)6
in 800 ml of 10% H2SO4. After complete hydrolysis of the
substrate (5 h), the mixtures were lyophilized, and the resulting solid
was extracted with methanol (4 x 5 ml). The extracts were combined and
evaporated to dryness. The crude material was purified by flash chromatography
(MeOH:CHCl3, 1:9) on a silica gel (Merck; 63200 mesh) column
to yield the pure product as a white solid (40 mg). 1H NMR and
13C NMR spectra were recorded at an ambient temperature on a Bruker
Avance 500 MHz spectrometer. The mass spectrum was obtained on a TSQ-70B mass
spectrometer (Finnigan Mat) by negative chemical ionization in isobutane or on
a Bruker Daltomics Apex-III (ICR-MS) by the method of electrospray ionization.
Fourier transform infrared spectroscopy (FTIR) was recorded on a Bruker vector
22 spectrometer.
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RESULTS |
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Catalytic Properties of the Glu335 and Asp495 Mutants Glu335 was replaced with Gly, Ala, or Gln, and the catalytic properties of the E335G, E335A, and E335Q mutants using pNPX and 2,5-DNPX as substrates were determined and summarized in Table I. The kcat values measured for the E335G, E335A, and E335Q mutants were all significantly reduced and are about 106-fold of wild type activity with both pNPX and 2,5-DNPX as substrates. The very low activity precluded reliable Km determination.
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Similarly, Asp495 was replaced with Gly, and the kinetic
constants of the D495G mutant were determined using different aryl
-D-xylopyranosides with different leaving groups
(Table II and
Fig. 2). The
kcat values of the D495G mutant were 103-fold
lower than for the wild type. For both the wild type enzyme and the D495G
mutant, kcat values were invariant for hydrolysis of all
of the substrates. Although the Km values of the
wild type enzyme were roughly similar with all substrates, with the D495G
mutant these values increased as the substrate reactivity decreased
(increasing pKa). Consequently, the decrease in
the specificity constant
(kcat/Km) values for the
D495G mutant was more pronounced, because the substrates are less
reactive.
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Chemical Rescue of the Catalytic MutantsRate acceleration
by exogenous nucleophilic anions is the most definitive tool for the
identification of the catalytic residues
(21). In the presence of 1.4
M azide, kcat values increased by 4, 33,
and 400 times for E335Q, E335A, and E335G mutants, respectively
(Table I), indicating that the
effect of azide decreases as the side chain is longer. The addition of 2.3
M formate resulted in a 103-fold increase of
kcat for the E335G mutants
(Table I). Both azide and
formate accelerated the reaction in a concentration-dependent manner.
The kinetic constants of hydrolysis of 2,5-DNPX by the D495G mutant in the
presence of different concentrations of azide were measured and are plotted in
Fig. 3. Both
kcat and Km increased with
increasing azide concentrations until leveling off at about 0.5 M
azide. Consequently, the
kcat/Km values remained
unchanged. The effect of azide was also tested for the hydrolysis of
substrates with different leaving groups at substrate saturating conditions
(Fig. 4). All of the
kcat values increased with increasing concentrations of
azide until reaching a plateau. Finally, the kinetic constants for hydrolysis
of various aryl -D-xylopyranosides were determined in the
presence of 0.63 M azide (Table
II and Fig. 5).
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Characterization of Reaction Products in the Presence of Sodium
AzideThe formation of a glycosyl-azide product is a powerful
diagnostic tool for identifying the catalytic residues, and determining its
anomeric configuration is useful for distinguishing between the acid-base and
the nucleophile. TLC analysis of the reaction mixture containing E335G,
2,5-DNPX, and azide revealed the formation of a new product
(Rf = 0.5) distinct from xylopyranoside
(Rf = 0.18) and 2,5-DNPX (Rf = 0.51).
This new product was isolated and identified as
-D-xylopyranosyl azide, as determined by 1H NMR
(Fig. 6a),
13C NMR, mass spectrometry, and FTIR: 1H NMR (500 MHz,
CD3OD)
3.48 (m, 3H, H-2, H-3, H-5), 3.57 (t, 1H, J
= 11.0 Hz, H-4), 3.70 (dd, 1H, J = 5.0, 12.0 Hz, H-5'), 5.24
(d, 1H, J = 2.5 Hz, H-1); 13C NMR (125.8 MHz,
CD3OD)
65.4, 71.0, 73.3, 74.6, 91.6 (C-1); negative CIMS
m/z 173.9 (M-H,
C5H9O4N3 requires 175.1); FTIR
(mineral oil)
2116 cm1 (N3).
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Likewise, TLC analysis of the reaction mixture containing D495G, 2,5-DNPX,
and azide revealed the formation of a new product (Rf =
0.42) distinct from xylopyranoside (Rf = 0.18) and
2,5-DNPX (Rf = 0.51). This new product was isolated and
identified as -D-xylopyranosyl azide, as determined by
1H NMR (Fig.
6b), 13C NMR, mass spectrometry, and FTIR:
1H NMR (500.1 MHz, CD3OD)
3.03 (t, 1H,
J2,3 = 9.0 Hz, H-2), 3.21 (t, 1H,
J5a,5b = 11.5 Hz, H-5a), 3.23 (t, 1H,
J3,4 = 9.0 Hz, H-3), 3.39 (ddd, 1H,
J4,5a = 9.5 Hz, J4,5b = 5.5 Hz, H-4),
3.84 (dd, 1H, H-5b), 4.33 (d, 1H, J1,2 = 8.1 Hz, H-1).
13C NMR (125.8 MHz, CD3OD)
69.0 (C-5), 70.8
(C-4), 74.7 (C-2), 78.1 (C-3), 92.7 (C-1); electrospray ionization
m/z: 198.1 (M+ + Na,
C5H9O4N3, requires 175.1); FTIR
(mineral oil)
2116 cm1 (N3).
pH DependenceThe kcat values of the wild type enzyme and the D495G mutant for hydrolysis of pNPX were determined at different pH values in the range of 4.58.5 (Fig. 7). The pH activity profile of the wild type enzyme showed strong dependence upon pH changes with pKa values of <4 and 7.3. Conversely, no such dependence is observed for the mutant enzyme within these pH values. The activity of the mutant at pH levels lower than 4 could not be determined because the enzyme was insoluble.
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DISCUSSION |
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The kcat values measured for the E335G, E335A, and E335Q mutants were all drastically reduced with activities of about 106-fold lower than the wild type for hydrolysis of both pNPX and 2,5-DNPX. This magnitude of decrease in activity is typical for nucleophile mutants and was also observed in other retaining glycoside hydrolases (21). However, the substantial decrease in the catalytic activity is insufficient for the unequivocal assignment of Glu335 as the catalytic nucleophile. There are several examples where single mutations in proposed catalytic residues of glycoside hydrolases resulted in reduced or undetectable activity (23, 24), whereas crystallographic and biochemical analyses showed that these residues are not involved directly in catalysis (25, 26). To unambiguously identify Glu335 as the catalytic nucleophilic residue of XynB2, the rescue methodology was applied. In this procedure (Fig. 8a), the catalytic activities of the putative catalytic mutants are monitored in the presence of small nucleophilic anions, such as azide or formate. The small anion can enter the vacant place created by the elimination of the nucleophilic residue and attack the anomeric carbon of the sugar substrate to form a glycosyl-azide product (when azide is added) with inverted anomeric configuration. Rate acceleration of the mutant in the presence of external anions is a strong indication that the mutation is indeed in the catalytic residue.
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Although no rate enhancement was observed for the wild type enzyme, the
hydrolysis rates of 2,5-DNPX by the E335G mutant significantly accelerated
with increasing concentrations of azide and formate. The presence of 2.3
M formate increased kcat up to
103-fold, only 100 times lower from the wild type activity.
Interestingly, in the presence of 1.4 M azide, the rate of
hydrolysis by the E335G mutant was accelerated by about 4 x
102-fold, whereas hydrolysis by the E335A and E335Q mutants with
the same azide concentration resulted in only 32- and 4-fold rate enhancement,
respectively. Hence, as the length of the side chain increases, the vacant
place is smaller, preventing azide facile penetration. Interestingly, formate
had a greater effect on rate acceleration, although it exhibits lower
nucleophilicity. This was also observed with the nucleophile-less mutants of
the -glucosidase from Agrobacterium faecalis
(27) and the
1,3-1,4-
-glucanase from Geobacillus licheniformis
(28). It is possible that the
strong resemblance between formate and the missing carboxylate nucleophile
allows better accommodation of formate in the created cavity. Similar chemical
rescue of activity in the presence of azide and formate was observed for many
other retaining glycoside hydrolases
(20,
2730).
Azide can rescue the activity of both the nucleophile and acid-base
mutants. However, the anomeric configuration of the glycosyl-azide product is
inverted or retained for the nucleophile and acid-base mutant, respectively
(Fig. 8). To verify the role of
Glu335 as the catalytic nucleophile, azide was added to the
reaction mixture containing the E335G mutant and 2,5-DNPX. The isolated
product revealed the formation of
1-azido-1-deoxy--D-xylopyranoside
(Fig. 6a). The
inverted anomeric configuration of the xylopyranosyl-azide product is exactly
as would be expected if Glu335 is the catalytic nucleophile.
Together, these results provide unambiguous confirmation for the assignment of
the conserved Glu335 as the catalytic nucleophile of XynB2 and, by
extension, of the other members of family 52 glycoside hydrolases.
Catalytic Properties of the D495G MutantCatalysis by
retaining glycosidases proceeds via a double displacement mechanism involving
two catalytic residues (nucleophile and acid-base) as shown in
Fig. 1. According to this
mechanistic pathway, substitution of the acid-base catalytic residue by
mutation with a nonacidic residue should affect the rates of both chemical
steps, although the effect on each step will be different. The effect on the
glycosylation step will depend strongly on the leaving group ability of the
aglycone. The rates of hydrolysis for substrates with a poor leaving group
should be more affected than those with a good leaving group. Therefore, if
this step is rate-limiting, kcat values will vary with
substrate reactivity. The deglycosylation step, however, will be affected
equally for all substrates carrying different leaving groups because the same
glycosyl-enzyme intermediate is hydrolyzed during this step. Thus, if this
step is rate-limiting, the kcat values will be invariant
with substrate reactivity. The kcat values of the D495G
mutant were reduced by 103-fold, consistent with Asp495
acting as a catalytic residue (Table
II). The Brønsted plot of log(kcat)
versus pKa shown in
Fig. 2a reveals that
for the wild type enzyme, the deglycosylation is the rate-limiting step
because kcat values are invariant for all substrates.
Likewise, this step is also rate-limiting for the D495G mutant for hydrolysis
of substrates with pKa < 8. Thus, for these
substrates the deglycosylation step was more affected upon the removal of
Asp495. The 103-fold reduction of this step, as evident
by the relative kcat values of the wild type and the
mutant (Table II), is
consistent with Asp495 acting as a general base catalyst. Such a
large rate reduction in the deglycosylation step was also observed for other
acid-base mutants from the -L-arabinofuranosidase of G.
stearothermophillus (31)
and the family 1
-glucosidases of A. faecalis
(32) and Streptomyces
sp. (33). Furthermore,
hydrolysis of these substrates by the mutant D495G resulted in very low
Km values
(Table II), suggesting that a
glycosyl enzyme intermediate accumulates, as would be expected if the second
step is rate-limiting. Interestingly, hydrolysis of 3-nitrophenyl
-D-xylopyranoside (pKa = 8.39)
by the mutant resulted in kcat that is approximately three
times lower than the kcat values for hydrolysis of
substrates with pKa < 8, suggesting that in
this case the glycosylation step is, at least partially, rate-limiting. This
claim will gain more support upon applying chemical rescue with exogenous
anions, as will be described in the following section.
The effect of the mutation on the glycosylation step can be obtained by
comparing the specificities
(kcat/Km) of the wild type
and the D495G mutant (Table
II). Because
kcat/Km =
k1
k2/(k1+
k2), and k1 and
k1 are the rate constants for the
Michaelis substrate-enzyme complex association and deassociation,
respectively, it is likely that
kcat/Km reflects the first
glycosylation step. For hydrolysis of substrates with good leaving groups,
kcat/Km are similar for both
the wild type and the mutant. However, for substrates with poor leaving
groups, the kcat/Km values of
the D495G mutant were two orders of magnitude smaller than those of the wild
type. Thus, the replacement of Asp495 had little effect on the
hydrolysis rates of the first step for substrates with good leaving groups
that require little or no acid assistance. Conversely, the first step was
severely affected with poor leaving groups substrates that require strong acid
assistance. This behavior is consistent with Asp495 acting as a
general acid catalyst. More information regarding proton assistance during the
glycosylation step can be obtained from the Brønsted plot of
log(kcat/Km) versus
the pKa of the leaving group. The Brønsted
plot shown in Fig. 2b
reveals good correlation between the pKa and
log(kcat/Km) with a slope of
1g = 1 for XynB2-D495G. The corresponding correlation
for XynB2 was derived only for poor substrates with
pKa of >8 because a biphasic relationship was
observed, probably reflecting a change in the enzyme-substrate association
rate constant, which becomes rate-limiting for the most highly reactive
substrates (33,
34). Thus, for the wild type
enzyme a slope of
1g = 0.82 was observed (data not
shown), and regardless of the reason for this biphasic nature, it is clear
that the reaction catalyzed by the mutant is much more dependent on the
aglycone leaving group ability. The larger value of
1g for
the mutant as compared with that of the wild type suggests a higher amount of
negative charge on the glycosidic oxygen of the leaving group in the
glycosylation transition state. This difference probably results from little
proton donation by the mutant, exactly as would be expected in the absence of
acid catalysis.
Chemical Rescue of the D495G MutantThe function of
Asp495 as the acid-base catalyst can also be verified by the use of
the azide rescue methodology (Fig.
8b). Small anions can accelerate the second step,
therefore increasing kcat values if the second step is
rate-limiting, and in this case a -glycosyl azide product is expected.
Indeed, in the presence of increasing concentrations of azide, the
kcat values significantly increased for hydrolysis of
2,5-DNPX (a substrate for which deglycosylation is rate-limiting) by
XynB2-D495G (Fig. 3).
Interestingly, the rates of hydrolysis increased until reaching a plateau,
which probably reflects a change in the rate-limiting step from
deglycosylation to glycosylation step. Thus, the value of
kcat at the plateau reflects the rate of the glycosylation
step, which is not affected by azide. This increase in activity is markedly
high, with kcat values increasing by almost
103-fold, approaching the rates of the wild type enzyme. Rate
enhancements by azide were also observed for other acid-base mutants, although
in these cases only up to a 3 x 102-fold increase was
achieved (32,
35). The presence of azide
also affected the Km values, and these increased
dramatically until leveling off (Fig.
3). For retaining glycoside hydrolases
Km = (k1 +
k2)k3/(k2 +
k3)k1. This
Km value is small for hydrolysis of a good
substrate, such as 2,5-DNPX (pKa = 5.15), by
XynB2-D495G, because the deglycosylation step is severely affected
(k3 is low), and the glycosylation step is virtually
unaffected (k2 is high relative to
k3). However, because k3 increases
relative to k2 in the presence of azide, the
Km values also increase to a point at which
k2 << k3 and
Km = (k1 +
k2)/k1 reaching the highest possible
Km value that can be obtained for hydrolysis of
any given substrate. Although Km and
kcat increase in the presence of azide,
kcat/Km, which reflects the
glycosylation step, is unaffected (Fig.
3), confirming that azide affects primarily the deglycosylation
step.
As mentioned above, the presence of azide changed the rate-limiting step
from deglycosylation to glycosylation, with kcat values
probably reflecting the rate of hydrolysis of the first glycosylation step at
the plateau. Hydrolysis by XynB2-D495G of additional substrates with different
leaving groups in the presence of azide also resulted in rate enhancement
until reaching a similar plateau (Fig.
4). However, the plateau level of kcat values
drops as the pKa of the leaving group elevates,
and higher concentrations of azide are required for reaching this plateau,
because the substrate is more reactive. The correlation between the plateau
level and the pKa values provides additional
confirmation that indeed glycosylation becomes the rate-limiting step upon the
addition of azide. The Brønsted plot relating
log(kcat) values obtained in the absence and presence of
0.63 M azide (already at the plateau) and the
pKa of the leaving group shows again the change
in the rate-limiting step from deglycosylation to glycosylation because the
kcat values vary with substrate reactivity upon the
addition of azide (Fig.
5a). The plot reveals good correlation between the
pKa and log(kcat) in the
presence of azide with a slope of 1g = 1, which is
exactly the same as derived from the Brønsted plot of
log(kcat/Km) in the previous
section (Fig. 2b). The
Brønsted plot of
log(kcat/Km) shown in
(Fig. 5b), which
reflects the glycosylation step, reveals similar relationships in the absence
or presence of azide, verifying again that azide has no effect on the
glycosylation step. As speculated in the previous section, the predominantly
rate-limiting step for hydrolysis of 3-nitrophenyl
-D-xylopyranoside (pKa = 8.39)
by the D495G mutant is glycosylation because azide did not affect the reaction
rate and the Km value
(Table II).
Finally, the reaction mixture containing the D495G mutant, 2,5-DNPX, and
azide revealed the formation of -D-xylopyranosyl azide
(Fig. 6b). The fact
that the anomeric configuration of the xylopyranosyl-azide product is retained
provides definite evidence that Asp495 is indeed the catalytic
acid-base. Together, these results are consistent with the assignment of
Asp495 as the catalytic acid-base. pH Dependence Profiles of
XynB2 and XynB2-D495GThe assignment of Asp495 as the
acid-base catalyst can also be done by testing the pH dependence profiles for
the wild type and the D495G mutant. The pH dependence profile of the wild type
presented in Fig. 7 is a
typical bell-shaped curve, suggesting that hydrolysis requires one group in
its protonated form and the other in its deprotonated form, as observed for
many other glycoside hydrolases. Thus, enzymatic catalysis depends upon two
ionizable amino acid residues with pKa values of
<4 and 7.3, which are ascribed to the catalytic pair, with the higher
pKa attributed to the acid-base catalyst and the
lower pKa assigned to the nucleophile
(36). However, with the D495G
mutant no dependence in activity was observed at high pH values, suggesting
that the protonated group had been removed exactly as would be expected from a
glycoside hydrolases lacking its acid catalyst. Unfortunately, determination
of activity at pH values lower than 4.5 was precluded because the enzyme
precipitated rapidly. Similar behavior was also observed for many other
acid-base mutants from different glycoside hydrolases families
(12,
16,
31,
32,
37,
38). Collectively, these
results provide unequivocal confirmation for the assignment of the conserved
Asp495 as the catalytic acid-base of the family 52 XynB2.
In conclusion, Glu335 and Asp495 are the catalytic nucleophile and acid-base, respectively, of the family 52 XynB2. The large rate reduction for the E335G mutant, together with chemical rescue of activity, and the formation of a xylosyl-azide product with inverted configuration provide unequivocal evidence for the assignment of Glu335 as the catalytic nucleophile. With the D495G mutant, the large decrease in the deglycosylation step together with the large decrease in the glycosylation step for hydrolysis of poor substrates indicates that both general acid and general base catalysis were severely affected. Further, rate enhancement by chemical rescue accompanied with the generation of a xylosyl-azide product with retained configuration, together with the absence of acid catalysis as indicated by pH dependence profiles provide unambiguous confirmation for the assignment of Asp495 as the catalytic acid-base. Thus, these results allow for the first time direct identification of the catalytic residues of XynB2, and by extension, those of all members of family 52 glycoside hydrolases.
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FOOTNOTES |
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¶ Supported by the Center of Absorption in Science, the Ministry of
Immigration Absorption, and the Ministry of Science and Arts, Israel (Kamea
Program).
To whom correspondence should be addressed. Tel.: 972-4-8292590; Fax:
972-4-8233735; E-mail:
chtimor{at}tx.technion.ac.il.
¶¶ To whom correspondence should be addressed. Tel.: 972-4-8293072; Fax: 972-4-8293399; E-mail: yshoham{at}tx.technion.ac.il.
1 The abbreviations used are: pNPX, p-nitrophenyl
-D-xylopyranoside; 2,5-DNPX, 2,5-dinitrophenyl
-D-xylopyranoside; FTIR, Fourier transform infrared
spectroscopy.
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REFERENCES |
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