From the Agrotechnological Research Institute
(ATO-DLO), P. O. Box 17, 6700 AA Wageningen, ¶ BIOSON Research
Institute and Laboratory of Biophysical Chemistry, Groningen
Biomolecular Sciences and Biotechnology Institute (GBB), University of
Groningen, Nijenborgh 4, 9747 AG Groningen, and
Department of
Microbiology, Groningen Biomolecular Sciences and Biotechnology
Institute (GBB), University of Groningen, Kerklaan 30, 9751 NN
Haren, The Netherlands
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ABSTRACT |
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The product specificity and pH optimum of the
thermostable cyclodextrin glycosyltransferase (CGTase) from
Thermoanaerobacterium thermosulfurigenes EM1 was engineered
using a combination of x-ray crystallography and site-directed
mutagenesis. Previously, a crystal soaking experiment with the
Bacillus circulans strain 251 -CGTase had revealed a
maltononaose inhibitor bound to the enzyme in an extended conformation.
An identical experiment with the CGTase from T. thermosulfurigenes EM1 resulted in a 2.6-Å resolution x-ray
structure of a complex with a maltohexaose inhibitor, bound in a
different conformation. We hypothesize that the new maltohexaose conformation is related to the enhanced
-cyclodextrin production of
the CGTase.
The detailed structural information subsequently allowed engineering of
the cyclodextrin product specificity of the CGTase from T. thermosulfurigenes EM1 by site-directed mutagenesis. Mutation D371R was aimed at hindering the maltohexaose conformation and resulted
in enhanced production of larger size cyclodextrins (- and
-CD).
Mutation D197H was aimed at stabilization of the new maltohexaose
conformation and resulted in increased production of
-CD.
Glu258 is involved in catalysis in CGTases as well as
-amylases, and is the proton donor in the first step of the
cyclization reaction. Amino acids close to Glu258 in the
CGTase from T. thermosulfurigenes EM1 were changed.
Phe284 was replaced by Lys and Asn327 by Asp.
The mutants showed changes in both the high and low pH slopes of the
optimum curve for cyclization and hydrolysis when compared with the
wild-type enzyme. This suggests that the pH optimum curve of CGTase is
determined only by residue Glu258.
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INTRODUCTION |
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Cyclodextrins (CDs)1 are
cyclic molecules composed of 6, 7, or 8 glucose units linked via
(1,4)-glycosidic bonds (
-,
-, and
-CD, respectively). The
ability of cyclodextrins to form inclusion complexes with small
hydrophobic molecules has provided a number of practical uses in the
food, pharmaceutical, and agrochemical industries (1-3). Cyclodextrins
are produced from starch by the action of the enzyme cyclodextrin
glycosyltransferase (CGTase; EC 2.4.1.19). However, apart from the
cyclization reaction, the enzyme can also catalyze disproportionation,
coupling, and hydrolysis reactions. All known CGTases produce a mixture
of
-,
-, and
-cyclodextrins. For the industrial production of
pure cyclodextrins,
-CD is selectively crystallized and
- and
-CD are complexed with organic solvents. The industrial production of cyclodextrins might be improved by the construction of mutant CGTases with improved product specificity (2, 3).
In the conventional commercial production of cyclodextrins, starch is
first liquefied by the action of a thermostable -amylase, whereafter
cyclodextrins are produced using the mesophilic CGTase from
Bacillus macerans. Two highly thermostable CGTases have been characterized that can directly be used for starch liquefaction, eliminating the need for
-amylase pretreatment (2, 3). These enzymes
are produced by thermophilic anaerobic bacteria belonging to the genus
Thermoanaerobacter (9, 10) and Thermoanaerobacterium thermosulfurigenes EM1 (11). The overall amino acid compositions of both enzymes show relatively minor differences, and also the biochemical characteristics of both enzymes are very similar (11). The
T. thermosulfurigenes EM1 CGTase displays maximum
cyclization activity at 80-85 °C and maximum starch hydrolyzing
activity at 90-95 °C. The pH optimum for cyclization is broad, in
the range of pH 4.5-7.0.
The three-dimensional structures of several CGTases have been solved
(4, 5, 7). CGTase belongs to family 13 of the glycosyl hydrolases, a
group of homologous (/
)8-barrel proteins, to which
also the
-amylases belong (8). Recently, the three-dimensional structure of the CGTase from T. thermosulfurigenes EM1
(Tabium CGTase) was solved at 2.3-Å resolution (6). In the
present study we describe the three-dimensional structure of an
enzyme-substrate complex of Tabium CGTase. The x-ray
structure of a maltohexaose inhibitor complexed with Tabium
CGTase was solved at 2.6-Å resolution. The detailed information thus
obtained allowed rational engineering of the cyclodextrin product
specificity of Tabium CGTase.
Residues Glu258, Asp329, and Asp230
are directly involved in catalysis in CGTase (20). Glu258,
together with Asp230, is believed to cleave the
substrate's (1,4)-glycosidic bonds and to form the product's
(1,4)-glycosidic bonds by a double displacement mechanism (15). In
the first step of the reaction the general acid Glu258
protonates the oxygen of the glycosidic bond to be cleaved. After cleavage of this scissile bond, an oxocarbonium transition state is
formed, which is believed to collapse into a covalently linked intermediate by nucleophilic attack of Asp230 on the
anomeric C1. Subsequently, the reducing end diffuses out of the active
site and an acceptor comes in, which can be a water molecule (in case
of hydrolysis) or a carbohydrate C4-hydroxyl group (in case of
transglycosylation). In the second step of the reaction, the acceptor
hydroxyl group is activated through deprotonation by
Glu258, after which the acceptor performs a nucleophilic
attack on the covalent intermediate. Through another oxocarbonium
transition state, the product (
(1,4)-linked) is then formed (16,
17).
It appears from the double displacement mechanism that for optimal catalysis the nucleophile Asp230 must be deprotonated in the first step of the reaction, whereas the general acid Glu258 must be protonated in the first step, but deprotonated in the second step of the reaction. A third catalytic residue, Asp329, has been found to be hydrogen bonded to Glu258 in the unliganded CGTase, thereby elevating the pKa of Glu258 and assuring its protonation. After substrate binding, this hydrogen bond is lost, making deprotonation of Glu258 possible. It was suggested that this Glu258-Asp329 interaction is responsible for the broad pH optimum exhibited by CGTases (16). The importance of the pKa of Glu258 in the different reactions catalyzed by Tabium CGTase was further studied. By changing the electrostatic environment of Glu258 by site-directed mutagenesis, we could drastically shift the Tabium CGTase pH optimum.
The biochemical characteristics of the various mutant CGTases are presented with emphasis on the effects of these mutations on the CGTase cyclization and hydrolytic activities, pH optima, and product formation from starch.
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EXPERIMENTAL PROCEDURES |
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Structure Determination-- Crystals of the Tabium CGTase were grown from 21% saturated ammonium sulfate in 100 mM Tris buffer, pH 7.6. The crystals were stable in the presence of carbohydrates.
A double soaking experiment was started identical to that described earlier for the Bacillus circulans strain 251 CGTase (13). A Tabium CGTase crystal was soaked for 20 min in a solution of 0.25% w/v acarbose in 21% saturated ammonium sulfate and 100 mM CAPS buffer, pH 9.8, followed by 7 days of soaking in a solution of 0.5% maltohexaose in 21% saturated ammonium sulfate in 100 mM CAPS buffer, pH 9.8. Data were collected to a resolution of 2.6 Å on a MacScience Dip2000K Image Plate system and processed with XDS (23). Refinement of the structure was done with the TNT package (24), using the 2.3-Å structure of unliganded Tabium CGTase as a starting model (6). Rigid body refinement was followed by coordinate and all parameter (coordinates and individual atomic temperature factors) refinement. A test set for calculating a free R factor (25) comprised 8% (1956) of the unique reflections. Ideal protein bond lengths and angles were taken from Engh and Huber (26), ideal bond lengths and angles for glucose were taken from the crystal structure of maltose (27). Planarity, van der Waals contacts, and B factor correlations were restrained, whereas torsion angles were not. Chiral centers were watched. The model was manually adjusted using O (28), running on a Silicon Graphics workstation, in combination with the program OOPS (29). Electron density was displayed using
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Bacterial Strains, Plasmids, and Growth
Conditions--
Escherichia coli JM109 (35) was used for
recombinant DNA manipulations. E. coli PC1990 (36), known to
leak periplasmic proteins into the supernatant because of a mutation in
its tolB locus, was used for (extracellular) production of
CGTase (mutant) proteins. Plasmid pCT2, a derivative of pUC18
containing the amyA (cgt) gene of T. thermosulfurigenes EM1 (37), was used for site-directed mutagenesis, sequencing, and expression of the CGTase (mutant) proteins
(Fig. 1). Plasmid-carrying bacterial
strains were grown on LB medium with 100 µg/ml ampicillin. When
appropriate, isopropyl--D-thiogalactopyranoside was
added at a concentration of 0.1 mM for induction of protein expression.
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DNA Manipulations--
DNA manipulations and transformation of
E. coli were essentially as described by Sambrook et
al. (38). Electrotransformation of E. coli was
performed using the Bio-Rad gene pulser apparatus (Bio-Rad, Veenendaal,
The Netherlands). The selected conditions were 2.5 kV, 25 µF, and 200 .
Site-directed Mutagenesis-- Mutant CGTase genes were constructed via a double PCR method using Pfu DNA polymerase (Stratagene, Westburg, Leusden, The Netherlands). A first PCR reaction was carried out with the mutagenesis primer for the coding strand plus a primer 195-715 base pairs downstream on the template strand. The reaction product was subsequently used as primer in a second PCR reaction together with a primer 295-815 base pairs upstream on the coding strand. The product of the last reaction was cut with NcoI and MunI and exchanged with the corresponding fragment (900 base pairs) from the vector pCT2 (Fig. 1). The resulting (mutant) plasmid was transformed to E. coli JM109 for sequencing and to E. coli PC1990 for production of the (mutant) proteins. The following oligonucleotides were used to produce the mutations: D197H, 5'-CGTAACTTATTTCATTTAGCAGATCTAAATCAACAG-3'; F284K, 5'-GTCTTTTGGACAAGAGGTTTTCTC-3'; N327D, 5'-GGTTACTTTTATTGATGATCATGATATGG-3'; D371R, 5'-GACAGGCAATGGACGTCCTTATAATAGAGC-3'. The bold codons indicate the changed amino acids. Successful mutagenesis resulted in appearance of the underlined restriction sites (BglII for D197H, BclI for N327D and AatII for D371R), which allowed rapid screening of potential mutants. It was not possible to find a convenient restriction site for mutant F284K. Mutations were verified by DNA sequencing (39). All 900 base pairs on the MunI-NcoI fragment obtained by PCR were checked by DNA sequencing.
Production and Purification of CGTase Proteins--
For
production of CGTase proteins, E. coli PC1990 (pCT2) was
grown in a 2-liter fermentor at pH 7.0 and 30 °C. The medium contained 2% (w/w) trypton (Oxoid, Boom BV, Meppel, The Netherlands), 1% (w/w) yeast extract (Oxoid), 1% (w/w) sodium chloride, 1% (w/w) casein hydrolysate (Merck, Darmstadt, Germany), 100 µg/liter
ampicillin, and 0.1 mM
isopropyl--D-thiogalactopyranoside. Growth was monitored by measuring the absorbance (A) at 450 nm. At an
A450 nm of 2-3, an extra amount of 50 g
of trypton was added to the fermentor. Cells were harvested after
20-24 h of growth (8000 × g, 30 min, 4 °C), at
absorbance values of 8-12. The supernatant was directly applied to an
-CD-Sepharose-6FF affinity column (40) for further purification of
the CGTase proteins. After washing the column with 10 mM
sodium acetate, pH 5.5, the CGTase was eluted with the same buffer
supplemented with 1% (w/w)
-CD. Purity and molecular weight of the
CGTase (mutant) proteins were checked on SDS-polyacrylamide gel
electrophoresis (11). Protein concentrations were determined by the
method of Bradford (42), using the Coomassie protein assay reagent of
Pierce (Pierce Europe bv, Oud-Beijerland, The Netherlands).
Enzyme Assays--
Specific assays were used to determine the
activities (initial rates) of the four different reactions catalyzed by
CGTases (14). In the cyclization reaction the reducing end of a sugar is transferred to another sugar residue in the same oligosaccharide chain, resulting in the formation of cyclic compounds. Coupling is the
reverse reaction in which a cyclodextrin molecule is linked to a linear
oligosaccharide chain, producing a longer oligosaccharide chain. In the
disproportionation reaction, part of a linear donor-oligosaccharide is
transferred to a linear acceptor chain. The saccharifying activity is
the hydrolysis of starch into linear oligosaccharides. All assays were
standardly performed at pH 6.0 and 60 °C. Cyclization and
saccharifying assays were performed as described by Penninga et
al. (14). Coupling activity was measured essentially as described by Nakamura et al. (41). -CD (2.5 mM) was
used as donor substrate and methyl
-D-glucopyranoside
(100 mM) as acceptor substrate. The linear oligosaccharide
formed in the reaction was converted to single glucose units by the
action of amyloglucosidase (Sigma, Darmstadt, Germany). Glucose was
detected with the glucose/GOD-Perid method of Boehringer Mannheim
(Almere, The Netherlands). Disproportionation activity was measured as
described by Nakamura et al. (18). EPS,
4-nitrophenyl-
-D-maltoheptaoside-4-6-O-ethylidene
(3 mM, Boehringer Mannheim), was used as donor substrate
and maltose (10 mM) as acceptor substrate. The reaction
product containing the nitrophenyl group was cleaved by the action of
-glucosidase (Boehringer Mannheim). For each reaction units were
defined as the amount of enzyme producing/converting 1 µmol of
product/substrate at pH 6.0 and 60 °C.
HPLC Product Analysis--
Formation of cyclodextrins was
measured under industrial process conditions by incubation of 0.1 unit/ml CGTase (-CD forming activity) with 10% Paselli WA4
(pregelatinized drum-dried starch with a high degree of polymerization;
AVEBE) in 10 mM sodium citrate buffer, pH 6.0, at 60 °C
for 45 h. Samples were taken at regular time intervals and boiled
for 10 min. Products formed were analyzed by HPLC, using a 25-cm
Econosil-NH2 10-µm column (Alltech Nederland bv, Breda,
The Netherlands) eluted with acetonitrile/water (65:45) at 1 ml/min.
Products were detected by a refractive index detector (Waters 410, Waters Chromatography Division, Milford, MA). The temperature of the
flow cell and column was set at 50 °C, to avoid possible
precipitation of starch. Formation of linear products was directly
analyzed. Formation of CDs was analyzed after incubation of the samples
with an appropriate amount of
-amylase (type I-B from sweet potato,
Sigma, Boom BV, Meppel, The Netherlands), degrading linear sugars (but
not CDs) to glucose. The retention times for
-,
-, and
-CD
were the same as those for G4, G5, and G6 linear oligosaccharides,
respectively.
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RESULTS AND DISCUSSION |
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Binding of the Maltohexaose Inhibitor
The maltohexaose inhibitor complexed with the CGTase from T. thermosulfurigenes EM1 was bound at subsites 3 to +3 (Fig.
2). In complexes with BC251
CGTase (13) or other CGTases, binding at subsite
3 has never been
observed. The present study thus reveals the nature of subsite
3 for
the first time. The glucose at subsite
3 (overall B
factor: 58 Å2) has long distance interactions with
Glu264 O-
1 and Thr262 N, both at 3.6 Å. A
better contact is formed at 3.4 Å with the Asn591 O
1
from a symmetry related molecule. This stabilization by a crystal
contact may explain why in BC251 CGTase crystals a glucose at subsite
3 has never been observed. The interactions that do not
result from a crystal contact are very weak, indicating that subsite
3 is not of large relevance.
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In the vicinity of the scissile bond the active site architecture is
identical in Tabium and BC251 CGTase. It is
therefore not surprising that at subsites 2 to +2, the maltohexaose
inhibitor is bound in the same fashion to Tabium CGTase as
the maltononaose inhibitor to BC251 CGTase (13). Even though
the acarbose binding mode has been modeled differently, the same amino
acids are providing similar interactions from subsites
2 to +2,
suggesting that acarbose can adapt its conformation easily to that
required by the active site. It is clear that differences in
characteristics between Tabium and BC251 CGTase
must originate from interactions at more distant subsites.
In contrast to subsites 2 to +2, at subsite +3 the binding mode of
the maltohexaose inhibitor is radically different from the maltononaose
binding mode, the former being more bent. In Fig.
3, an overlay of the two inhibitor
conformations can be seen. All enzyme-substrate interactions are given
in Table II. The glucose at subsite 3 of
the maltohexaose inhibitor occupies a position more bent toward
Phe196. This conformation at subsite 3 is stabilized by
Lys47. In BC251 CGTase this residue is
Arg47, which so far has never been found to be involved in
substrate binding. Furthermore, in the BC251 enzyme
Tyr89 has strong interactions at subsite 3, but in the
Tabium CGTase this residue is an Asp. The conformation of
Asp89 does not allow any interactions with substrate. Apart
from these two differences at subsite 3, residues in Tabium
CGTase at subsites 4-7 might be unfavorable to the maltononaose
binding mode (straight), although we could not find evidence for that
from the structure of Tabium CGTase. The maltohexaose
conformation observed is only stabilized by the protein through the
contact with Lys47. The maltohexaose conformation, however,
might have less internal strain because it allows the O2-O3
interglucoside hydrogen bond between subsites 2 and 3 to be formed at
2.7 Å, whereas in the straight conformation the O2-O3 distance is 4.0 Å (13) prohibiting hydrogen bond formation. In addition, the lack of
interactions with the enzyme might allow for more flexibility of the
maltohexaose chain, which would thus be entropically stabilized. On the
basis of these data, we thought it possible to favor one binding mode over the other by site-directed mutagenesis, thereby investigating whether it could be related to one of the Tabium CGTase
characteristics.
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The Tabium and BC251 CGTases are different in
many respects. The molecular basis for thermostability of the
Tabium CGTase has been extensively discussed (6).
Furthermore, Tabium CGTase displays a relatively high
hydrolytic activity (24 units/mg), compared with BC251
CGTase (3.5 units/mg), which results in formation of substantial
amounts of linear sugars besides cyclodextrins from starch (11).
Tabium CGTase produces a mixture of -,
-, and
-CD
at a ratio of 28:58:14, respectively, whereas BC251 CGTase has a product ratio of 13:64:23 (14). A mutant BC251 CGTase with a substantially higher
-cyclodextrin production also showed preference for a bent maltohexaose inhibitor over a straightly bound
maltononaose inhibitor (43). This suggests a relation between the bent
conformation and
-cyclodextrin production. The maltohexaose and
maltononaose binding modes thus may reflect (part of) specific
intermediates for
-CD and
-CD production by Tabium CGTase and BC251 CGTase, respectively. Further experimental
evidence for this was sought by modifying relevant residues in
Tabium CGTase, using site-directed mutagenesis. Since the
maltohexaose and maltononaose inhibitors are synthesized by CGTase
in situ in the crystal, no sufficient quantities were
available to determine their binding or inhibitor constants. Therefore
we designed our mutants only by qualitative arguments.
Our first approach was to design a mutation that would hinder the
maltononaose binding mode and possibly bind a substrate in the
maltohexaose conformation. We found that the replacement of
Asp197 by His fulfilled these requirements, since modeling
of the mutation D197H (Fig. 4) shows that
the His ring cannot assume a conformation in which all the atoms are
more than 2.0 Å away from the atoms in the maltononaose inhibitor. So
His197 is likely to block the straight conformation.
Moreover, the His197 N-2 atom could potentially
stabilize the bent conformation by forming a hydrogen bond with the
glucoside O6 at subsite +3. The D197H mutation changes the
electrostatic field of the active site, which could have long distance
effects by changing charge-charge interactions. However, since the
substrate is uncharged, effects on the substrate binding mode will have
to be indirect, in contrast to the van der Waals and hydrogen bonding
interactions.
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Our second approach was to construct a mutant that would stimulate the maltononaose conformation over the maltohexaose conformation, which is easiest achieved, not by constructing a specific interaction with the straight conformation, but by destabilizing the bent conformation. With the mutation Asp371 to Arg we aimed at introducing a bulky residue that would clash with the maltohexaose conformation, but be less hindering to the straight maltononaose conformation (Fig. 4). The long and flexible side chain of Arg371 could probably extend its effect to subsite +3, where the bent and straight conformations differ most. The mutation D371R changes again the electrostatics of the active site, with possible indirect consequences for substrate binding. However, the mutation conserves the polarity of residue 371.
Of the two mutants D197H, designed to relatively stabilize the
maltohexaose conformation, is expected to produce more
-cyclodextrin. Mutant D371R, designed to prefer the maltononaose
conformation, is expected to produce less
-cyclodextrin. The
experimental data show that these mutants display altered cyclodextrin
product specificity according to expectation (Table
III).
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Production and Purification of CGTase (Mutant) Proteins
Mutants of Tabium CGTase were successfully constructed by site-directed mutagenesis via PCR; all mutations were verified by restriction site analysis (except for mutant F284K) and DNA sequencing. Amounts of 0.4-1.2 mg of pure protein were obtained in a single fermentor run depending on the construct used (Table IV). Purification yields varied between 11% for mutant D371R and 58% for mutant D197H. Purity and molecular weight of the (mutant) CGTases were checked on SDS-polyacrylamide gel electrophoresis. All proteins were purified to apparent homogeneity and displayed a molecular mass of 68 kDa.
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Characterization of CGTase Mutant Proteins
Cyclodextrin Product Specificity--
Mutant D197H has a
cyclodextrin production profile different from wild-type CGTase (Fig.
5). At the initial stages of the reaction, the mutant has an increased preference for -cyclodextrin production, mostly due to a collapse of the production of
-cyclodextrin. At later stages of the reaction, the production of
-cyclodextrin increases, but the product ratio is then a result of a
subtle equilibrium between cyclization and cyclodextrin breakdown
specificities, as well as the solubilities of the diverse
cyclodextrins. However, the cyclodextrin production ratio after 45 h still shows a small preference for
-cyclodextrin (Table III),
proving that the total product ratio can be modified by changing the
initial reaction rates. The initial reaction's preference for
-cyclodextrin is according to the expectations we had upon designing
the mutant (see above). An additional effect of the mutation D197H is
that coupling activity is reduced by a factor 4 when compared with the
activity of the wild-type enzyme (Table
V), possibly because coupling activity
was measured with
-cyclodextrin whereby a maltononaose was formed as
an intermediate. The mutation was, however, designed to make binding of
maltononaose less favorable.
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Site-directed Mutations Close to the Proton Donor Glu258: Implications for pH Optima of CGTases-- To further investigate the role of the environment of Glu258 on the pKa of Glu258 in the different reactions catalyzed by CGTases we replaced Phe284 by Lys (F284K) in Tabium CGTase. Residue Phe284 is located in a hydrophobic cavity immediately above the active site, close to Glu258 (Fig. 6). At most physiological pH values Lys can be expected to bear a positive charge. Positioning of a positive charge near Glu258 stabilizes its deprotonated form, so decreasing its pKa. This would predict a shift of the pH optimum for this mutant toward acidity. Fig. 7, A and B, show that this indeed happens for both the cyclization and hydrolysis activity. Apart from this effect, the optimum curves also become more narrow, an observation that was also made for a F184L mutant in Bacillus sp. 1011 CGTase (18). The mutation F184L does not introduce any charges; therefore, the narrowing effect can be best explained by a reduction of the hydrophobicity of the environment of Glu258 (see below).
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pH Optimum Curve for Cyclization and Hydrolysis-- In Tabium CGTase residue Glu258 is the proton donor in the first step of catalysis, so a protonated state of Glu258 is essential. In the second step of catalysis, Glu258 is suggested to activate the acceptor by deprotonation, in this case a deprotonated state of Glu258 would be essential. Furthermore, for optimal activity the catalytic nucleophile, Asp230, must remain deprotonated in the first step of catalysis. The Tabium CGTase displays a broad pH optimum for cyclization in the range of pH 4-7 (Fig. 7A). Other CGTases have an efficient cyclization reaction from pH 5 to 7 (16, 18, 21, 22). It might be expected that the drop in enzyme activity at high pH is caused by deprotonation of Glu258 in the first step of the reaction and that the activity drop at low pH is caused by protonation of Asp230. The combination of these two effects would result in a pH optimum curve. In Fig. 7A, however, it can be seen that the mutations near Glu258 shift both the slopes at high and low pH, while we had expected only to see effects on the high pH slope of the optimum curve, which is affected by Glu258. Similar observations have been described before in literature. The mutation F284L in Bacillus sp. 1011 CGTase shifts the pH optimum both over acidic and alkaline pH ranges (18). This mutation is close to the proton donor and is unlikely to have long distance effects. Mutation of Asp329 in BC251 CGTase to Asn had a most pronounced effect on the low pH slope of the optimum curve (20). Furthermore, mutation of Asp230 to Asn in the BC251 CGTase did not result in any shift of the pH optimum, whereas a shift was observed with the Glu258 to Gln mutation (20). Therefore, it is likely that the protonation state of Glu258 determines both slopes of the pH optimum curve. At high pH, the first step of the double displacement mechanism is hindered by lack of a proton donor, at low pH the second step is inhibited by incomplete substrate activation. If the protonation state of Glu258 determines both slopes of the pH profile, it is unexpected that the pH optimum is so broad. This broad pH range of activity must be explained by a different environment for Glu258 in the first and second step of catalysis. Indeed, in Tabium CGTase (6) the third catalytic residue Asp329 probably forms a hydrogen bond with Glu258 initially, increasing the pKa of Glu258 and ensuring its protonation. This bond is broken after substrate binding, facilitating again deprotonation of Glu258 in the ensuing second catalytic step. The impact of changes near Glu258 during catalysis is enhanced by its hydrophobic environment, in which electrostatic interactions are much stronger. Changes in the environment of Glu258 before and after substrate binding have also been observed in the structure of BC251 CGTase (13, 16).
The pH optimum for hydrolysis is much sharper and lies about 1 pH unit lower than the pH optimum for cyclization in the Tabium CGTase (Fig. 7B). The only difference between these two reactions is the acceptor molecule, water in the hydrolysis reaction, and a C4-hydroxyl group in the cyclization reaction. The fact that the pH optimum for hydrolysis is less broad could result from a decreased hydrophobicity of the Glu258 environment with water as an acceptor. The shift toward lower pH might be explained from the fact that water as an acceptor is much easier activated, making the requirement for an unprotonated Glu258 in the second reaction step less stringent, thus increasing activity at low pH. ![]() |
CONCLUSIONS |
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At subsite 3 the binding mode of the maltohexaose inhibitor in
Tabium CGTase is radically different from the maltononaose binding mode in the BC251 CGTase. The bent conformation of
the maltohexaose inhibitor, in contrast to the straight conformation of
the maltononaose inhibitor, was found to be correlated to enhanced -cyclodextrin production. Mutations stabilizing the bent
conformation but hindering the straight conformation resulted in
enhanced production of
-CD, whereas mutations hindering the bent
conformation but stabilizing the straight conformation resulted in
decreased production of
-CD. The Tabium CGTase can hence
be changed from an
/
-cyclodextrin producer to a
/
-cyclodextrin producer by a single mutation, illustrating the
feasibility of CGTase protein engineering.
Mutations near the proton donor Glu258 suggest that the pH optimum curve of CGTase may be determined only by the protonation state of residue Glu258. Both the high and low slopes of the pH optimum curve could be manipulated by site-directed mutations close to Glu258. Changes in the environment of Glu258 before and after substrate binding can account for its broad pH optimum.
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ACKNOWLEDGEMENTS |
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The assistance of Dirk Penninga, Jan Springer, and Gerard Rouwendaal with construction of the mutants and the assistance of Gert-Jan van Alebeek with the enzyme assays is gratefully acknowledged.
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FOOTNOTES |
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* This work was supported by European Community Grants AIR-CT-93-1023 (to R. D. W. and R. M. B.) and ERBIO2-CT-94-3071 (to J. U., B. W. D., and L. D.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
§ To whom correspondence should be addressed: ATO-DLO, P. O. Box 17, 6700 AA Wageningen, The Netherlands. Tel.: 31-317-475321; Fax: 31-317-475347; E-mail: r.d.wind{at}ato.dlo.nl.
1 The abbreviations used are: CD(s), cyclodextrin(s); CGTase, cyclodextrin glycosyltransferase; CAPS, 3-(cyclohexylamino)propanesulfonic acid; MBS, maltose binding site; PCR, polymerase chain reaction; HPLC, high performance liquid chromatography.
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REFERENCES |
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