(Received for publication, September 18, 1995; and in revised form, December 6, 1995)
From the
The importance of eight nucleoside 2`-deoxyribosyltransferase
residues for catalysis was investigated by site-directed mutagenesis.
Each residue was selected because of its proximity to nucleophile
Glu-98 or on its potential contribution to intrinsic protein
fluorescence. Mutation of Asp-72, Asp-92, Tyr-7, Trp-12, and Met-125
resulted in over a 90% activity loss whereas mutation of Tyr-157,
Trp-64, and Trp-127 produced less than a 80% activity loss. The
magnitude of the perturbation on catalysis by mutation, however, was
dependent on donor substrate. The k values for
dIno hydrolysis by these mutants were greater than 25% of that for
native enzyme. Although mutant and native enzymes bound substrate
analogues with comparable affinities, K
values for dIno hydrolysis varied over a 1000-fold range.
The pH dependence of Glu-98 esterification by dCyd suggested that amino
acids with pK values of 4.2 and 7.5 were relevant for
catalysis. The intrinsic protein fluorescence was attributed primarily
to Trp-127 (
80%). Pre-steady-state kinetic parameters for
deoxyribosylation of mutant enzymes by dCyd, dThd, and dAdo were
determined by monitoring changes in enzyme fluorescence. Collectively,
results from mutagenesis suggest that, depending upon substrate, either
Asp-92 or Asp-72 functions as the general acid catalyst, and that this
enzyme undergoes a change in conformation upon Glu-98
deoxyribosylation.
Nucleoside 2`-deoxyribosyltransferase (EC 2.4.2.6, transferase) ()catalyzes cleavage of the glycosidic bonds of
2`-deoxyribonucleosides(1, 2, 3, 4, 5, 6, 7, 8) through
intervention of a covalent deoxyribosyl-enzyme (EX)
intermediate(9) . Glu-98 is the active site nucleophile
deoxyribosylated during catalysis (9) . Glycoside hydrolases
that catalyze analogous reactions with retention of stereochemical
configuration utilize the carboxyl group from either a
glutamyl(10, 11, 12, 13, 14, 15, 16) or an
aspartyl residue (17, 18, 19) as the active
site nucleophile, and, in general, additional acidic residues that
function as general
acids/bases(20, 21, 22, 23, 24) .
The distance between the carboxyl group of the active site nucleophile
and the carboxyl group of the general acid for retaining glycosidases
is 4.8 to 5.3 Å(23) . In addition to these carboxylates,
crystal structure data reveals that the Bacillus circulans xylanase has tyrosyl and tryptophanyl residues clustered about the
active site that are postulated to contribute to
catalysis(23) .
In nature, nucleoside
2`-deoxyribosyltransferase is found in various Lactobacilli species and
participates in nucleoside recycling in these
microorganisms(1, 2, 3, 4, 5) .
The ntd gene encoding this enzyme has been cloned, and the
recombinant protein expressed in Escherichia
coli(8, 9, 26) . Native transferase
purified from E. coli and Lactobacilli, is a hexamer composed
of identical 18,000-Da subunits(9, 26) . The
crystal structure of this enzyme has recently been solved and refined
to 2.5-Å resolution. (
)This study identified the
monomer fold as a single, doubly-wound
/
domain composed of a
central 5-stranded
-sheet flanked by 4
-helices. Although
there is one active site per subunit, each complete catalytic center
defined by the position of Glu-98 requires participation of side chains
from a neighboring subunit ( Fig. 1and Fig. 2). The
studies described herein were initiated to examine the effect that
mutagenesis of selected active site residues has on catalytic
efficiency. We present results that extend our understanding of this
process by comparing the pre-steady-state and steady-state kinetic
properties of native transferase with nine mutant transferases. Amino
acid residues were selected for mutagenesis because of their proximity
to the active site nucleophile Glu-98 or their fluorescence properties.
Mutation of Glu-98, Asp-72, Asp-92, Tyr-7, and Tyr-157 defined the
relative contribution of amino acid side chains to catalysis whereas
the W12A, W64A, W64F, and W127F proteins defined the relative
contribution of each tryptophanyl residue to the intrinsic protein
fluorescence. Collectively, results obtained with these two classes of
transferase mutants suggest a change in enzyme conformation attendant
upon deoxyribosylation of Glu-98.
Figure 1:
The active site of nucleoside
2-deoxyribosyltransferase. The protein model was generated using
coordinates from the crystal structure refined at 2.5-Å
resolution. The residues examined by mutagenesis are shown.
The amber and blue tubes represent the peptide
backbones of the two subunits that form the active site. Tyr-157
(labeled Tyr357) and Met-125 (labeled Met325)
originate from the subunit adjacent to that containing
Glu-98.
Figure 2: Location of tryptophanyl residues relative to nucleophile Glu-98. Trp-12 and Trp-64 are from the same subunit as Glu-98, whereas Trp-127 (labeled Trp327) at the interface between the two subunits is from the neighboring subunit.
The activity of transferase mutants was determined with cytosine
as the acceptor substrate and three donor acceptor substrate pairs in
which the concentration of each substrate was equal to 1 mM.
The reaction of dIno and Cyt was monitored at 292 nm as described
above. The reaction of dAdo with Cyt was monitored at 285 nm
( = 2.7 mM
cm
). The reaction of dUrd with Cyt was
monitored at 286 nm. (
= 2.2
mM
cm
).
If the time course of the reaction was first-order, F was set equal to 0. These pseudo-first-order
rate constants (k
) were determined as a function
of substrate concentration (S).
When the reaction of mutant enzymes
with different donor substrates was examined, these results were
interpreted in terms of the mechanisms of , ,
and . The reaction of dCyd with transferase is a
monophasic process(27) . Initial complex formation (K) occurs within the dead time of the
stopped-flow spectrophotometer (1.6 ms) followed by a first-order
reaction to yield deoxyribosylated enzyme ()
The observed first-order rate constant for this process is given by .
The reaction of dThd with transferase is a biphasic process (27) . The early phase
represents binding of dThd to give EdThd. In
contrast to dCyd binding, the binding of dThd to transferase is
observable on the stopped-flow spectrophotometer. The second phase of
the reaction is the result of enzyme deoxyribosylation. The two
observed rate constants for this mechanism are given by and .
The reaction of dAdo with transferase is monophasic in the presence of Ade(27) . Because of the fluorescent properties of intermediates in the reaction, it was necessary to include Ade in the reaction to observe a significant change in protein fluorescence during the course of the reaction. This reaction is described by .
Under conditions in which the concentration of Ade is
sufficiently large such that EX is present mostly as EXAde, the observed rate constant for this mechanism is
given by .
where F([L]) is the corrected fractional
fluorescence of the transferase solution at a ligand concentration
equal to [L]. Because the dissociation constants of the
substrate analogues (L) used to titrate transferase were much larger
than the enzyme concentration, a correction for depletion of the
substrate analogue resulting from binding to transferase was not
necessary.
The pH dependence of the dissociation constant (K) of transferase for a substrate analogue was
fitted to , which was analogous to .
The following buffer systems were used for the pH titration: 0.05 M sodium acetate (pH 3.95 to 5.03), 0.05 M sodium MES (pH 5.5 to 6.56), 0.05 M sodium HEPES (pH 7.70 to 8.09), and 0.05 sodium TAPS (pH 8.26 to 9.01).
Native enzyme catalyzes
hydrolysis of 2`-deoxyribosylnucleosides in addition to its transferase
activity(6, 7, 8, 9) . The
steady-state kinetic parameters for dIno hydrolysis by mutant
transferases are summarized in Table 2. The k values for the hydrolysis reaction, except for E98A, showed less
than a 4-fold variation (Table 2). In contrast, the K
values varied between 8 µM and 7100
µM. The steady-state kinetic parameters for the transfer
reaction were determined for those mutant transferases having catalytic
activities with 1 mM dIno and 1 mM Cyt (Table 1) that were at least 5-fold larger than k
for the hydrolytic reaction (Table 2).
The equation for a ping-pong kinetic mechanism () was
fitted to the initial velocity data to give values for k
, K
, and K
. The k
values for the
transferase reaction with these mutants varied between 1 s
and 38 s
. The K
values
varied from 2300 µM to 8000 µM, whereas the K
values varied between 52 µM and
700 µM.
The activity of mutants E98A, E98D, D72A, D92A, and Y7A was less than 0.2% of the activity of native enzyme. This residual activity could result from trace amounts of native enzyme arising from translational misincorporation. If this were the case, then the substrate activity ratios for mutant and native transferases should be similar. The data of Table 1suggest that this is not the case. The dAdo/dIno and dUrd/dIno substrate activity ratios for native transferase were 5.7 and 2.7, respectively, whereas these ratios for E98A were 4.3 and 18, respectively. These ratios for other transferase mutants were also different from those of the native enzyme (Table 1).
An alternative approach to address the identity of
residual activity is to treat the mutant enzyme with an inactivator of
native transferase. Previously, dFDAP was shown to inhibit native
transferase by covalently modifying Glu-98. When 200 µM enzyme was incubated with 1000 µM dFDAP for 50 min,
the activity of D72A, D92A, and Y7A transferases with dAdo and Cyt as
substrates did not markedly decrease. In contrast, native enzyme lost
over 95% of its activity under these conditions. Incubation of E98A
with dFDAP decreased the ratio of E98A activity to that of native
enzyme from 4 10
to 5
10
.
Under certain circumstances, the activity of
a mutant enzyme can be increased by providing the missing functional
group in the form of a small, soluble acid or base. When catalytic
rescue was attempted with a mutant -glucosidase from Agrobacterium faecalis lacking an essential glutamyl residue,
the addition of a high concentration of formate increased k
(10
-fold), although product
stereochemistry was opposite that formed with the wild-type
enzyme(35) . When native transferase was assayed in 4 M formate at pH 6.0 with 1 mM dAdo and 1 mM Cyt as
substrates, its activity was reduced 48%. Assay of mutant transferases
in buffer containing 4 M formate reduced the activities of the
D72A and E98A enzymes by 38% and 78%, respectively, but enhanced the
activity of the D92A mutant 26-fold. Nonetheless, the activity of D92A
transferase in 4 M formate was less than 1% that of native
transferase in this buffer.
Trp-12, Trp-64, and Trp-127 were changed individually to an alanyl or a phenylalanyl residue by site-directed mutagenesis. Substitution of an alanyl residue for Trp-12 or Trp-64 did not markedly affect the fluorescence emission intensity of the protein (Table 3). Similarly, substitution of a phenylalanyl residue for Trp-64 had little effect on protein fluorescence (Table 3). In contrast, substitution of a phenylalanyl residue for Trp-127 resulted in a dramatic decrease in fluorescence emission intensity. The fluorescence of W127F was 20% of that measured for native transferase ( Table 3and Fig. 3).
Figure 3:
Fluorescence emission spectra of selected
mutants of transferase. The fluorescence emission spectra (uncorrected)
of W64F, W12A, W127F, and native transferase were recorded with
= 280 nm. The enzymes (1 µM)
were in 0.1 M potassium phosphate at pH 6.0 and 25 °C.
Corrections for the slightly different absorbance values of the
different enzymes at 280 nm were not made (<3%). The order of the
spectra is given by the inset.
Native enzyme, W12A, and W64F had
emission maximum at 322 nm in 0.1 M potassium phosphate at pH
6.0 ( = 280 nm), but that for W127F was 313
nm. Denaturation of native and mutant enzymes in 6 M guanidinium chloride at pH 6.0 shifted the emission maximum
wavelength to 360 nm, similar to that of N-acetyltryptophan in
this solvent. The ratio of the fluorescence emission intensities for
native enzyme in 0.1 M potassium phosphate at pH 6.0 in the
absence (
= 322 nm) and presence (
= 360 nm) of
6 M guanidinium chloride was 1.05. The fluorescence emission
intensity for each Trp substitution mutant in 6 M guanidinium
chloride was reduced approximately 40% relative to that of native
enzyme (Table 3), which is the expected fluorescence of denatured
native enzyme after loss of one tryptophanyl residue.
Titrations of the fluorescence of native, W12A, and W64A
transferases by N-methyladenine are presented in Fig. 4. For native enzyme, dissociation constants (K
) with dTAdo and dFThd were 30 ± 1
µM and 263 ± 9 µM, respectively (Table 4). These value were similar to the K
values of 46 ± 3 µM and 490 ± 20
µM, respectively, determined for these substrate analogues
in a transfer reaction between dCyd and Ade. Thus, the dissociation
constants represent substrate analogue binding to the active site.
Except for a few instances, binding of nucleobase or nucleoside
analogue by native and mutant proteins were similar; the ratio of K
values for mutant to native enzymes was, in
general, less than 10 (Table 4).
Figure 4:
Titration of transferase proteins by N-methyladenine. Titrations of native, W64A, and
W12A enzymes with N
-methyladenine were monitored
by the quenching of each protein's intrinsic fluorescence upon
complex formation (
= 300 nm;
= 340 nm). The data were corrected for inner filter
effects as described under ``Experimental Procedures.'' Other
conditions were as described in Fig. 3. Fluorescence values were
normalized to the fluorescence of enzyme in the absence of ligand. The solid lines were calculated with , and the values
for the parameters are tabulated in Table 4.
Reactions of native transferase with
dThd and dAdo, analyzed by the mechanism of and , have been presented previously(27) . The time
course for reaction of native transferase with dCyd (Fig. 5, A and B) was analyzed by the mechanism of . Binding of dCyd to transferase is a rapid equilibrium
process that yields an initial complex (EdCyd) from
which deoxyribosylated enzyme (EX) is formed. For dCyd
concentrations much greater that the K
for initial
complex formation, a maximal fluorescence change is observed and
represents the transformation of E
dCyd to EX
and Cyt (Fig. 5A). The fluorescence change for EX formation from E and dCyd decreases as the dCyd
concentration is decreased (Fig. 5B). Because the
concentration of E
dCyd accumulating transiently at low
dCyd concentrations and the net fluorescence change approaches 0 under
these conditions, E and EX have similar fluorescence
but that for E
dCyd is quenched. Equivalent results were
observed with W127F transferase at high dCyd concentrations (Fig. 5A). At low dCyd, however, the sign of the
fluorescence change reversed (Fig. 5B), which suggested
that, for this mutant enzyme, fluorescence of E was greater
than that of EX. The concentration dependences of the
pseudo-first-order rate constant describing the half-reaction for
native and W127F transferases with dCyd are described by (Fig. 6). The kinetic parameters for native enzyme
were k
= 280 ± 10 s
and K
= 240 ± 30
µM. The analogous values for W127F were 260 ± 10
s
and 250 ± 40 µM.
Figure 5:
Comparison of the time course for the
reaction of dCyd with native and W127F transferases. The reaction of
dCyd with native transferase (10 µM) and W127F transferase
(9 µM) was monitored by the changes of intrinsic protein
fluorescence upon enzyme deoxyribosylation ( =
280 nm;
> 305 nm). A, the reaction with
1 mM dCyd. B, the reaction with 0.1 mM dCyd.
Fluorescence values are normalized to the fluorescence of each enzyme
at the end of the reaction. Because the fluorescence of native
transferase is 5-fold larger than that of W127F, the fluorescence
values for W127F need to be divided by 5 if absolute fluorescence
changes of the two enzymes are to be
compared.
Figure 6: Concentration dependences of the pseudo-first-order rate constants for the reaction of dCyd with native transferase or W127F transferase. The pseudo-first-order rate constants were calculated from data such as that presented in Fig. 3. The solid lines were calculated with , and the parameters are listed in Table 5.
Kinetic parameters for the half-reaction of each mutant transferase with dCyd, dThd, and dAdo are summarized in Table 5and Table 6. Substitution of Glu-98 with either an alanyl or aspartyl residue profoundly affected the rate of EX formation from both dCyd and dThd. This effect on catalysis was similar to that observed for these mutant enzymes in their transfer reaction rates (Table 1) and in their rate of dIno hydrolysis (Table 2). It should be noted that, although the fluorescence of W127F was reduced by 80%, the pre-steady-state parameters determined for this mutant enzyme were equivalent to those of native transferase ( Table 5and Table 6).
Figure 7:
pH dependences of the transferase
reaction. A, the pH dependence of k/K
and k
for the reaction of dCyd with native transferase. Experimental
values for these parameters were calculated from data similar to those
presented in Fig. 4. The solid line was calculated by , and the values for the parameters are given under
``Results.'' B, the pH dependences for the binding
of dFThd to native and E98A transferases. K
values were calculated from fluorescence quenching data as
described in Fig. 4. The solid line for the binding
data with native transferase was calculated with an equation analogous
to , and the parameters are given under
``Results.''
The pH
dependence of the affinity of native transferase for dFThd, a
competitive inhibitor of the transferase reaction, was determined to
complement results obtained with dCyd. The ionizations associated with
this binding process should be related only to pK values for
free enzyme as dFThd does not have a pK value between pH 5 and
8.5. The dependence of 1/K of native enzyme for
dFThd (Fig. 7B) was fitted to to give a pH
independent K
= 290 ± 20
µM, K
= (2.0 ± 0.6)
10
M (pK
= 4.7 ± 0.1), and K
=
(1 ± 0.3)
10
M (pK
= 8.0 ± 0.1). When the
ionizable carboxymethyl group of Glu-98 of native enzyme was replaced
by a methyl group in mutant E98A, the pH dependence of dFThd binding to
this mutant enzyme was eliminated (Fig. 7B).
The intrinsic protein fluorescence of transferase is markedly
perturbed by substrates and substrate analogues. Recently, these
fluorescence changes were used to monitor transfer of the deoxyribosyl
sugar of a donor substrate to the enzyme(27) . Native
transferase is a hexamer composed of identical subunits, each of which
contains three tryptophanyl residues(9) . Crystal structure
data places Trp-12 and Trp-64 in the vicinity of the active
site defined by the position of Glu-98, whereas Trp-127 is located at
the subunit interface distal from the active site nucleophile (Fig. 2). The effect of substitution on each of these
tryptophanyl residues with an alanyl or phenylalanyl residue
individually established that 80% of the intrinsic protein fluorescence
is due to Trp-127. Substitution of an alanyl residue for Trp-12 or
Trp-64 caused a 5% decrease or a 13% increase in the intrinsic protein
fluorescence, respectively ( Fig. 2and Table 3). The
wavelength for maximal fluorescence emission of native transferase was
322 nm and is characteristic for a tryptophanyl residue buried in an
hydrophobic environment(36) . Furthermore, the relatively
larger quantum yield of Trp-127 relative to either Trp-12 or Trp-64 is
consistent with their location in hydrophilic environments.
The
magnitude of the fluorescence changes (20%) during catalysis and
the relative contributions of the fluorescence of Trp-12, Trp-64, and
Trp-127 to the intrinsic fluorescence of the enzyme suggested that most
of the fluorescence change was due to perturbation of the Trp-127
environment. This was confirmed by comparison of the fluorescence
changes associated with reaction of dCyd with native and W127F
transferases (Fig. 5A). The reaction of 1 mM dCyd with native enzyme (K
= 240
µM) and W127F (K
= 250
µM) resulted in fractional fluorescence changes of 0.28
and 0.09, respectively. For comparison of the absolute fluorescence
changes associated with each protein, the fractional fluorescence
change associated with W127F was multiplied by the ratio of the
relative fluorescence of W127F to that of native enzyme. Thus, the
normalized fractional change in fluorescence of native enzyme during
the reaction with dCyd was 0.28 and that with W127F was 0.018. Clearly,
over 90% of the fluorescence change associated with the half-reaction
of dCyd with native transferase is due to perturbation in the
fluorescence of Trp-127. Because Trp-127 is located 16 Å from the
active site nucleophile, (
)these fluorescence changes result
from a conformation change at the subunit-subunit interface and not
from a direct interaction of dCyd with Trp-127 (Fig. 2).
The
nucleoside 2`-deoxyribosyltransferase-catalyzed reaction results in
retention of product configuration(6, 7) . This result
plus the fact that initial velocity data for the transfer reaction is
described by a ping-pong kinetic mechanism suggests formation of a
covalently deoxyribosylated enzyme intermediate(8) . Recently,
this intermediate was trapped by reaction of transferase with
2,6-diamino-9-(2`-deoxy-2`-fluoro--D-arabinofuranosyl)-9H-purine,
and the residue modified was identified as Glu-98(9) . Many
glycosidases catalyze reactions that are mechanistically similar to the
transferase reaction and which result in retention of product
stereochemical configuration. For these enzymes, catalysis proceeds by
the double displacement mechanism originally described by Koshland (37) that has been confirmed by Withers and co-workers (10, 11, 38, 39, 40) who
pioneered the use of 2-deoxy-2-fluoroglycoside analogues for
identification of the active site nucleophile. In addition to this
carboxylic acid, retaining glycosidases have one or more aspartyl or
glutamyl residues that participate in transition state stabilization or
function as a general acid/base(23, 24, 41) .
The data herein, and that reported
previously(8, 9, 28) , indicate an analogous
mechanism for transferase.
The assignment of Glu-98 as the active
site nucleophile (9) and its position within the crystal
structure of native transferase define those amino acids
that form the active site pocket and which could participate in
catalysis. Identification of the transferase general acid/base is of
particular interest. The combination of structural data and the
mechanism-based requirement of a second carboxylate residue guided our
selection of amino acids to be replaced by site-directed mutagenesis.
For retaining glycosidases, the average distance between the pair of
catalytic carboxylates is 4.8 to 5.3 Å(23) . Only Asp-72
and Asp-92 are candidates for the role of the general acid/base in
transferase. The distance between Glu-98 (O
2) and Asp-72 (O
2)
is 6.0 Å and that between Glu-98 (O
2) and Asp-92 (O
2)
is 5.6 Å (Fig. 1). Thus, based on geometry, either Asp-72
or Asp-92, or possibly both, could function as the general acid/base.
The role of each in catalysis was assessed by the effects that
replacement with alanyl or asparaginyl residues had on substrate
binding and catalysis. The ratio of catalytic activity of native enzyme
to D92A was approximately 10
for all substrates examined
whereas this ratio for D72A was as small as 30 for dIno and Cyt as
substrates (Table 1). In accord with these findings, only the
activity of D92A was rescued by 4 M formate. These results
suggest that Asp-92 functions as the general acid/base. However, the
ratio of catalytic activity of native transferase to that of D92N was
less than 10 with dUrd and Cyt as substrates (Table 1). Since
asparagine would not be expected to function efficiently as a general
acid/base, this result brings into question the exclusivity of Asp-92
as the acid catalyst. Alternatively, this observation may reflect the
relationship between acid/base-assisted catalysis and the nature of the
leaving group. For example, the nucleophile and general acid/base of
the exoglucanase/xylanase from Cellulomonas fimi (Cex), which
has a mechanism similar to that of transferase, are Glu-233 and
Glu-127, respectively(20) . When the ratio of catalytic
efficiency (k
/K
) of native
Cex to E127A enzyme was systematically studied, this value varied
between 6000 and 1 depending on the leaving group ability of the
substrate(20) . If a similar circumstance exists for
transferase, these studies implicate Asp-92 as the general acid/base.
The role of Asp-72 is less clear; perhaps this residue participates in
stabilizing the oxycarbonium ion-like transition state.
Replacement
of the active site nucleophile Glu-98 by an alanyl or an aspartyl
residue had a more profound effect on catalysis than did substitution
of other amino acid residues (Table 1, Table 2, and Table 4). However, the ratio of the catalytic activity of E98A to
that of native enzyme was greater than expected from previous work with
a comparable mutant of the A. faecalis -glycosidase(42) . The source of this difference is
unknown but is too large to be accounted for by translational
misincorporation. It is noteworthy that the carboxylates of Asp-92 and
Asp-72 are separated by 9 to 11 Å which is similar to the
distance between the catalytic carboxylate pair of the inverting
glycosidases(23, 35) . However, when Glu-98 was
replaced by an aspartyl residue, the ratio of catalytic activity of the
mutant transferase relative to that of the native enzyme was roughly
comparable to the analogous ratios for the A. faecalis
-glycosidase and the B. circulans xylanase(35, 42) . Because substrates bind to
mutant transferases with similar affinity (Table 4), the effects
of substitution at Glu-98 reside in the chemistry of the reaction.
The amino acids in the native enzyme were chosen for mutagenesis
based on their proximity to Glu-98. Three of these, Asp-72, Asp-92, and
Tyr-7 originate from the subunit containing Glu-98 while Met-125,
Trp-127, and Tyr-157 belong to the neighboring subunit (see Fig. 1and Fig. 2). Mutant enzymes with a phenylalanyl
residue in place of Trp-64, Trp-127, or Tyr-157 suffered less than an
85% decrease in catalytic activity and substrate affinity ( Table 1and Table 4). In contrast, replacement of Tyr-7,
Trp-12, or Met-125 with an alanyl residue and deletion of Tyr-157, the
C-terminal residue, caused a significantly larger decrease in catalytic
activity; but, as noted above, the magnitude of these effects was
substrate-dependent. Comparison of the of Y157F and Y157 suggest
that the C-terminal carboxylate and not the phenolic oxygen is
important for catalysis. For Y7A and M125A, the reduction in enzymatic
activity is consistent with these mutations affecting reaction
chemistry since the effects on substrate binding were minor (Table 4). The effects of these substitutions on catalysis may be
due to subtle changes in orientation of either the substrates within
the active site or the groups involved in catalysis. The phenolic
oxygen of Tyr-7 is 3.0 Å from the catalytic carboxylate of Glu-98
and could serve to orient this group for catalysis or hydrogen bond to
the 3`-OH of the deoxynucleoside substrate.
A similar
function for a tyrosyl residue has been suggested following mutagenesis
of the B. circulans xylanase(25) . The role of Met-125
remains to be defined, but the consequence of its replacement and
proximity to Trp-127, the source of 90% of the fluorescent change
observed during catalysis, is most interesting. Further insight into
the roles of these residues in catalysis may come from characterization
of revertant enzymes derived from genetic selections that require
expression of a functional transferase.
Except for mutation of
Glu-98, the mutations examined here had little effect on k for transferase hydrolase activity, but had a
large effect on K
. This result is consistent with
hydrolysis of deoxyribosylated enzyme being the rate-limiting step that
is not subject to significant catalysis by any of the amino acid side
chains mutated for this study.
The pH dependence of k/K
for the deoxyribosylation
of native transferase by dCyd indicated two groups with pK values of 4.2 and 7.5 (Fig. 7A). Because substrate
ionization was corrected for, these represent pK values for
free enzyme. When the pH dependence for binding of dFThd to native
enzyme was measured, the pK values obtained were 4.7 and 8.0 (Fig. 7B). The pH dependence for binding of dFThd to
E98A suggests that the lower pK value measured with native
enzyme is the property of active site nucleophile Glu-98, although a
definitive assignment is not possible due to inaccuracy inherent in
signal strength as the assay pH became more alkaline (Fig. 7B). The bell-shaped profile noted for the pH
dependence of k
/K
of the
native enzyme suggests that the monoprotonated form of the free enzyme
binds substrate as described by . For native enzyme, k
was independent of pH. This suggests that, upon ES formation, other ionizations are unimportant for catalysis
or that substrate binding results in their pK values being
shifted outside the pH range of these experiments. The pH dependence of
the transferase reaction is similar to that for the hydrolysis of
2",4"-dinitrophenyl-
-cellobioside by the xylanase from C.
fimi(13) . For this enzyme, the active site nucleophile
was identified as Glu-233 and the general acid as Glu-127. The pH
dependence data for transferase and C. fimi xylanase (13) indicate that two ionizable groups contribute to catalysis
and that only one of these groups is protonated in the active form of
the enzyme. These data do not, however, indicate the protonation state
of the specific transferase residues in the active form of the enzyme;
but, it seems likely that one of these two ionizable residues is
Glu-98.
In summary, we have established that Asp-72, Asp-92, Trp-12, Met-125, Tyr-7, and the C-terminal carboxylate of Tyr-157 contribute significantly to the catalytic efficiency of nucleoside 2-deoxyribosyltransferase. The detailed role of each residue in catalysis will have to await the crystal structures of the mutant enzymes complexed with appropriate substrates in their active sites. We had hoped that these mutagenesis studies would identify the general acid for the transferase reaction, but Asp-92 and Asp-72 both remain as possible candidates. Perhaps the ambiguity in assigning the role of general acid exclusively to one of these residues is founded in the relaxed substrate specificity of transferase that may allow both residues to function as the general acid in a substrate-dependent manner.