Identification of a Catalytic Aspartyl Residue of
D-Ribulose 5-Phosphate 3-Epimerase by Site-directed
Mutagenesis*
Yuh-Ru
Chen
,
Frank W.
Larimer§¶,
Engin H.
Serpersu
, and
Fred C.
Hartman§¶
From the § Protein Engineering Program, Life Sciences
Division, Oak Ridge National Laboratory, and
University
of Tennessee-Oak Ridge Graduate School of Biomedical Sciences,
Oak Ridge, Tennessee 37831 and
Department of Biochemistry and
Cellular and Molecular Biology, University of Tennessee,
Knoxville, Tennessee 37996
 |
ABSTRACT |
Guided by comparative sequence considerations, we
have examined the possibility of a catalytic role of
Asp186 of D-ribulose 5-phosphate
epimerase by site-directed mutagenesis of the recombinant spinach
enzyme. Accordingly, D186A, D186N, and D186E mutants of the epimerase
were constructed, purified, and characterized; as judged by their
electrophoretic mobilities the mutants are properly assembled into
octamers like the wild-type enzyme. Based on the extent of internal
quenching of Trp fluorescence, the conformational integrity of the
wild-type enzyme is preserved in the mutants. The wild-type
kcat of 7.1 × 103
s
1 is lowered to 3.3 × 10
4
s
1 in D186A, 0.13 s
1 in D186N, and 1.1 s
1 in D186E; as gauged by D186A, altogether lacking a
functional side chain at position 186, the
-carboxyl of
Asp186 facilitates catalysis by >107-fold.
Relative to the wild-type enzyme, the Km for
D-ribulose 5-phosphate is essentially unaltered with D186N
and D186E but increased 10-fold with D186A. Apart from their
impairments in epimerase activity, the mutants are unable to catalyze
exchange between solvent protons and the C3 proton of substrates. This deficiency and the differential alterations of kinetic parameters among
the mutants are consistent with Asp186 serving as an
electrophile to facilitate
-proton abstraction. This study is the
first to identify a catalytic group of the epimerase.
 |
INTRODUCTION |
Despite its participation in the ubiquitous oxidative pentose
phosphate pathway and its concurrent role in the reductive pentose phosphate pathway (i.e. the Calvin cycle) of photosynthetic
organisms, D-ribulose 5-phosphate 3-epimerase (EC 5.1.3.1),
which catalyzes the interconversion of
Ru5P1 and Xu5P, has not been
well-characterized with respect to structure-function relationships.
This relative neglect is probably a consequence of low natural
abundance of the epimerase and, in the case of the plant enzyme,
pronounced instability. We have recently removed these barriers to
detailed structural and mechanistic studies by designing a heterologous
overexpression system for the gene that encodes spinach chloroplast
Ru5P epimerase and by devising a facile purification scheme for the
recombinant enzyme that is compatible with its lability (1).
The epimerase-catalyzed reaction probably entails a two-base mechanism
with an enediolate intermediate (Fig. 1).
This supposition is based on the observations that Ru5P formed from
[3-2H]Xu5P was completely free of deuterium, even though
deuterium was completely retained in the remaining Xu5P (2). Later
findings (3) that isomerization of D-erythrose 4-phosphate
to D-erythrulose 4-phosphate by Ru5P epimerase proceeded by
intramolecular proton transfer, whereas the C2 proton of
D-threose-4-phosphate formed concurrently by epimerization
of D-erythrose-4-phosphate was derived entirely from water,
lends additional credence to a two-base mechanism. In Fig. 1, we invoke
a general acid to facilitate
-proton abstraction and to stabilize
the enediolate intermediate based merely on the prevalence of such
groups among mechanistically characterized enzymes, which also abstract
-protons of carbon acids, inclusive of triosephosphate isomerase
(4-6), 3-ketosteroid isomerase (7-9), and mandelate racemase
(10, 11).

View larger version (8K):
[in this window]
[in a new window]
|
Fig. 1.
Likely reaction pathway for the Ru5P
epimerase catalyzed interconversion of Ru5P and Xu5P.
R1 = -CH2OPO32 ; R2 = -HC(OH)-CH2OPO32 .
|
|
The catalytically functional groups depicted in Fig. 1 have not been
identified, nor have any residues of Ru5P epimerase even been assigned
to the active site. We wish to rectify this situation by applying
site-directed mutagenesis to the cloned spinach gene. Because
three-dimensional structural information and chemical modification data
of Ru5P epimerase are unavailable to guide selections of site-directed
mutants, we have assessed comparative sequences and predictive
secondary structural analyses for candidate catalytic residues. These
assessments prompted scrutiny of Asp186. As reported
herein, properties of the D186E, D186N, and D186A mutants of Ru5P
epimerase are entirely compatible with a crucial catalytic role for
Asp186.
 |
EXPERIMENTAL PROCEDURES |
Materials--
Materials and vendors were as follows:
transketolase, triosephosphate isomerase, glycerophosphate
dehydrogenase, thiamine pyrophosphate,
DL-
-glycerophosphate, D-ribose-5-phosphate,
Ru5P, leupeptin, phenylmethylsulfonyl fluoride,
N-acetyl-L-tryptophan, D2O (99.9 atom % D), lactate dehydrogenase, phospho(enol)pyruvate, pyruvate
kinase, phosphoriboisomerase, ATP, and NADH, Sigma; Pfu DNA
polymerase, Stratagene; DpnI restriction endonuclease, New England Biolabs; oligonucleotide primers for polymerase chain reaction
mutagenesis, Life Technologies; 4-(2-aminoethyl) benzenesulfonyl fluoride, Calbiochem; 3,3'-diaminobenzidine tetrahydrochloride dihydrate and Bio-Spin 6 columns, Bio-Rad; ultrapure urea, ICN; and
molecular weight calibration kits, Amersham Pharmacia Biotech. Common
laboratory reagents used in conjunction with mutagenesis, expression,
and enzyme purification were procured at the highest level of purity
readily available. Spinach phosphoribulokinase was prepared as
described earlier (12, 13).
Site-directed Mutagenesis and Expression of Ru5P Epimerase
Mutants--
The Ru5P epimerase mutants were constructed by use of
linear polymerase chain reaction in conjunction with Pfu DNA
polymerase (14). Plasmid pFL506 (GenBank accession number AF070943), which encodes mature wild-type spinach Ru5P epimerase ligated as an
NcoI-BamHI fragment adjacent to the tac promoter
of expression vector pFL260 (1, 15), was used as the template for
mutagenesis. The mutagenic oligonucleotide primers (for forward and
reverse replication) and the encoded amino acid substitutions are shown in Table I. After replication, the
polymerase chain reaction reaction mixture was treated with
DpnI restriction endonuclease to cleave the methylated
template DNA; unmethylated product DNA is not a substrate for
DpnI. Product DNA was then ethanol-precipitated, redissolved
in 10 mM Tris and 1 mM EDTA (pH 8.0), and
electroporated into Escherichia coli XL 1-Blue (16). Plasmid
template was isolated from the resulting ampR transformants
and sequenced across the region of interest to confirm the desired
mutation.
An overnight culture (25 ml) of the mutant expression vector in host
strain MV1190 or XL 1-Blue was grown in 2X-YT medium (17) containing
1% (v/v) glycerol and 50 µg/ml ampicillin at 37 °C. The culture
was diluted 1:100 into the same medium and incubated for 4 h with
shaking (250 rpm). Isopropyl
-D-thiogalactopyranoside was added to 0.1 mM, and the incubation was continued for
an additional 3 h at 37 °C, at which time the cells were
harvested by centrifugation.
Purification of Recombinant Ru5P Epimerase Mutants--
Mutants
were purified by a protocol designed for wild-type Ru5P epimerase,
which entails successive chromatography on DEAE-cellulose, hydroxyapatite, and MonoQ (1). Processing of ~10 g of cell paste,
harvested from 1-liter cultures of transformed E. coli, typically gave 5-6 mg of the purified mutant. Final preparations (>5
mg/ml) in 10 mM DL-
-glycerophosphate, 50 mM NaCl, and 1 mM EDTA (pH 8.0) were stored at
80 °C in the absence of cryoprotectant; DL-
-glycerophosphate was present at all stages of
purification to mitigate spontaneous loss of activity (1).
Progression of purifications was routinely followed by SDS-PAGE of
fractions generated at each step. Because of the inherently low
epimerase activity of the mutants, which was overshadowed by the far
greater levels from indigenous E. coli Ru5P epimerase, assays of the initial extracts were uninformative. However, because E. coli and recombinant spinach Ru5P epimerase are widely
separated by DEAE-cellulose in the first chromatographic step of the
purification (1), assays of epimerase activity in fractions collected
from this column and at all subsequent steps could be used to monitor D186N and D186E. The paucity of activity associated with D186A precluded reliable activity measurement before concentration of pooled
fractions from the final chromatographic step. Similarly to the
observed behavior of the wild-type epimerase (1), each of the mutants
was resolved into two peaks by chromatography on MonoQ. The catalytic
parameters of the two peaks were indistinguishable.
Electrophoresis--
SDS-PAGE was carried out at 15 °C and pH
8.1 on 12.5% (w/v) PhastGels, and nondenaturing PAGE was carried out
at 15 °C and pH 8.8 on 8-25% (w/v) gradient PhastGels, in
conjunction with a Phast System apparatus (Amersham Pharmacia Biotech).
IEF under denaturing conditions was performed at 15 °C on PhastGel
IEF 5-8 as described earlier (1). Gels were stained with Coomassie Blue
according to the supplier's protocol.
Protein and Enzyme Assays--
Protein concentration was
determined by a dye binding method (18) with reagent from Bio-Rad;
bovine serum albumin served as the standard for comparison. Ru5P
epimerase activity was assayed spectrophotometrically at 340 nm by
coupling Xu5P formation to NADH oxidation via reactions catalyzed by
transketolase, triosephosphate isomerase, and glycerophosphate
dehydrogenase (19). Stock preparations of mutants for use in
determination of catalytic parameters were centrifuged at 4 °C
through a Bio-Spin column (8 × 20 mm) equilibrated with 50 mM Bicine (pH 8.0) to remove
DL-
-glycerophosphate. If dilutions were required before
initiation of enzyme assays, the 50 mM Bicine buffer
containing bovine serum albumin at 1 mg/ml was used. Final
concentrations of the mutant proteins in the 120-µl assay solutions
were 0.01, 0.1, and 5 mg/ml for D186E, D186N, and D186A, respectively;
these amounts contrast to 1.2-1.8 ng/ml as were ample to assess
wild-type enzyme.
Fluorescence Measurements--
Steady-state fluorescence
emission spectra were recorded with an SLM-AB2 fluorimeter; excitation
and emission bandwidths were set at 4 nm. All measurements were carried
out at 25 °C with excitation at 295 nm to selectively excite the
indole side chains. Protein samples (0.1 mg/ml with
A295 of ~0.01) were prepared in 50 mM Bicine (pH 8.0); 8 M urea was included for
examination of denatured wild-type epimerase.
N-Acetyl-L-tryptophan (5 µM,
A295 of 0.01) in 50 mM Bicine (pH
8.0) was used as a standard for fluorescence emission. Controls lacking
protein were used to subtract out background signals. Recorded emission
spectra were automatically corrected, because the instrument accounts
in real time for wavelength-dependent efficiencies of the
light source.
NMR Spectroscopy--
NMR spectra were collected at 300K with a
wide-bore Bruker AMX 400 MHz spectrometer. One-dimensional
1H NMR spectra of samples in D2O (pH 8.0) were
collected over a spectral width of 4807 Hz with 32K data points.
Proton chemical shifts were referenced to external disodium
2,2-dimethyl-2-silapentane-5-sulfonate.
 |
RESULTS |
Comparative Sequences and Predictive Secondary Structure of Ru5P
Epimerase--
Comparisons of DNA-deduced amino acid sequences of Ru5P
epimerases from 16 species,2
including representatives from prokaryotes, eukaryotes, and archaea, identify only 25 invariant residues (Fig.
2). Among the eight with ionizable side
chains (Asp39, Asp44, Asp47,
Asp186, His42, His75,
His99, and Glu101), Asp186 occurs
in the only segment of contiguous invariant residues. When assessed by
neural networks for predictions of secondary structure (20, 21), the
epimerase sequence describes a series of alternating
-strands and
-helices, compatible with an eight-stranded
/
barrel folding
motif.

View larger version (24K):
[in this window]
[in a new window]
|
Fig. 2.
Primary structure of Ru5P epimerase.
Species invariant residues are denoted by boldface.
Assignment of residues to -strands (E) or -helices
(H) is based on predictive secondary structure analysis (20,
21). The N-terminal Ala is a consequence of the expression construct
used to produce the recombinant enzyme (1); the authentic enzyme
isolated from spinach chloroplast begins with Thr (28), which is
therefore designated residue 1.
|
|
Purity and Structural Integrity of Ru5P Epimerase
Mutants--
Electrophoretic analyses indicate a high degree of purity
of the epimerase mutants, which comigrate with the wild-type enzyme during denaturing (Fig. 3A)
and nondenaturing (Fig. 3B) PAGE. A pI of 6.6 for the D186A
and D186N subunits in contrast to a pI of 6.3 for wild-type and D186E
subunits, as determined by denaturing IEF (Fig. 3C), agrees
with the calculated impact of changing the net charge from
4 to
3.
The uncharacterized microheterogeneity of the wild-type enzyme, which
is observed by nondenaturing IEF (1), persists with each of the mutants
(data not shown).

View larger version (40K):
[in this window]
[in a new window]
|
Fig. 3.
Electrophoretic comparisons of wild-type and
mutant Ru5P epimerases by SDS-PAGE (A), nondenaturing PAGE
(B), and denaturing IEF (C). The molecular
mass markers (from top to bottom) in A
are phosphorylase b, 94 kDa; albumin, 67 kDa; ovalbumin, 43 kDa; carbonic anhydrase, 30 kDa; trypsin inhibitor, 20.1 kDa; and
lactalbumin, 14.4 kDa. The molecular mass markers (from top
to bottom) in B are thyroglobulin, 669 kDa;
ferritin, 440 kDa; catalase 232 kDa; lactate dehydrogenase, 140 kDa;
and bovine serum albumin, 67 kDa. The pH gradient (8-5) in
C decreases from top to bottom.
|
|
Trp fluorescence has been examined as a diagnostic of conformational
integrity of the mutants (Fig. 4).
Compared with a typical emission spectrum for Trp that is observed for
the wild-type epimerase in the presence of 8 M urea,
fluorescence is almost totally quenched under nondenaturing conditions.
Likewise, little Trp fluorescence can be detected with the nondenatured
mutants.

View larger version (27K):
[in this window]
[in a new window]
|
Fig. 4.
Corrected fluorescence emission spectra of
wild-type and mutant Ru5P epimerases. Spectra were collected at 2 nm/s; additional details are provided under "Experimental
Procedures." w.t., wild type.
|
|
Catalytic Properties of Ru5P Epimerase Mutants--
Although each
mutant is severely impaired catalytically, sufficient activity remains
to quantify kcat and Km of
Ru5P (Table II). Only slight increases in
Km associated with substitution of Asp by Asn or Glu
implicate Asp186 directly in catalysis.
View this table:
[in this window]
[in a new window]
|
Table II
Kinetic parameters of spinach recombinant Ru5P epimerase
Experimental details are provided under "Experimental Procedures."
Based on replicate assays, Km and
Vmax values vary by ±10%.
|
|
The proton resonances of Ru5P and Xu5P, including the doublets
reflecting their respective C3 protons, are sufficiently different (Fig. 5, A and B)
to allow the epimerase-catalyzed conversion of either compound in
D2O to be monitored by 1H NMR. After
equilibrium has been reached (Fig. 5C), the spectra lack the
doublet given by the C3 proton of Ru5P or Xu5P, because the C3 proton
of substrate is replaced in the product by deuterium from solvent. In
the experiments depicted, spectra of 5 mM Ru5P or Xu5P were
recorded before and 5 min after the addition of wild-type epimerase to
the respective solutions at a final concentration of 2.2 µg/ml. By
contrast, the spectra were unaltered by incubation of either substrate
for 30 min with D186N at 28 µg/ml. Thus, evidence of preferential
retention of proton exchange activity by the mutant, independent of
epimerase activity, is lacking.

View larger version (17K):
[in this window]
[in a new window]
|
Fig. 5.
Proton NMR spectra of Ru5P (A),
Xu5P (B), and Ru5P or Xu5P after incubation with wild-type
Ru5P epimerase (C). Only one spectrum is shown for the
two incubations, because the spectra generated were superimposable.
Assignment of protons is designated by the numbered carbon atoms
(C1-C5) to which they are bonded. Additional details are
provided under "Experimental Procedures."
|
|
 |
DISCUSSION |
Because of the numerous enzymes that abstract protons
to
carbonyl or carboxyl groups, including D-ribose 5-phosphate
isomerase that shares a common substrate with Ru5P epimerase and
several others that transform substrates structurally similar to Ru5P, we expected to uncover some enzymes with regions of sequence similarity with Ru5P epimerase through data base searches. Somewhat surprisingly, none were found, thereby precluding prospects of extrapolating known
active site features from functionally similar enzymes to Ru5P
epimerase. However, predictive secondary structural analyses did
suggest that the Ru5P epimerase subunit folds into an eight-stranded
/
barrel, as commonly found among isomerases, racemases, and epimerases (22, 24, 25).
In contrast to data base searches for proteins with sequence
similarities to Ru5P epimerase, consideration of sequences of the
epimerase from evolutionarily diverse sources proved instructive to
selecting targets for mutagenesis. Only 8 of the 25 invariant residues
have ionizable side chains. One of these, Asp186, is
located in the only stretch of 3 contiguous invariant residues that
occurs throughout the entire 235-residue polypeptide chain. Furthermore, this location represents the C-terminal end of
-strand 8 of the predicted
/
barrel; most active site residues of
proteins having this motif are positioned at the C-terminal ends of
-strands or in interconnecting loops between C termini of
-strands and the N termini of the following helices (22-24).
Envisioning Asp186 as one of the two prospective general
bases for the abstraction of the C3 proton from either Ru5P or Xu5P or
the prospective general acid to assist deprotonation in both the
forward and reverse directions of catalysis, we wished to ascertain the
consequences of deletion of the aspartyl
-carboxyl group and of
substitution with groups that might partially fulfill its normal
function. Thus, we characterized the D186A, D186N, and D186E mutants of
Ru5P epimerase.
Each mutant, which could be purified by the regimen that proved
successful for wild-type enzymes, displays the same mobility as the
wild-type enzyme during both denaturing and nondenaturing PAGE; thus,
the mutant subunits are full-length translation products, which fold
and assemble properly into octameric structures akin to wild type.
Despite the presence of two tryptophanyl residues (Trp40
and Trp182) in each subunit of Ru5P epimerase,
fluorescence emission elicited by excitation at 295 nm is barely
discernible. Although we have not determined the side chain
interactions responsible for the virtually complete quenching of Trp
fluorescence, retention of this striking feature by the mutants
indicates that their conformations are not perturbed substantially
relative to wild type.
As judged by the severe impairment of kcat
values of the mutants (2 × 107-fold for D186A,
5.5 × 104-fold for D186N, and 6.5 × 103-fold for D186E), in conjunction with essentially
unaltered Km values for Ru5P with D186N and D186E,
there can be little doubt that Asp186 of Ru5P epimerase
plays a critical role in catalysis. The 10-fold increase in the
Km for Ru5P with D186A, in contrast to the wild-type
values with the other two mutants, which retain hydrogen binding
potential at position 186, is consistent with an interaction between
the
-carboxyl of Asp186 and the carbonyl group of bound
substrate, as would occur if this residue served as the electrophilic
catalyst. The much higher levels of activity associated with D186N and
D186E, compared with D186A, are also compatible with Asp186
functioning as an electrophile to facilitate abstraction of the
-proton from substrate. The 10-fold greater
kcat of D186E relative to that of D186N might
reflect a mechanism whereby the enediolate intermediate is normally
stabilized by partial transfer of the active site carboxyl proton to
the carbonyl of the intermediate (i.e. formation of a short,
strong hydrogen bond), as demonstrated in the case of mandelate
racemases (10, 11, 26, 27). Whereas the side chain of Asn could engage
the substrate and intermediate through hydrogen bonding, partial proton
transfer is unlikely because of the much higher pKa
of an amide. If Asp186 rather serves as one of the two
bases responsible for
-proton abstraction from substrates, D186N
should be as debilitated as D186A; however, it is 400 times more active
than the latter and only 10 times less active than D186E.
In attempts to further distinguish possible roles of Asp186
in electrophilic catalysis versus proton abstraction, we
examined the ability of D186N to catalyze exchange of solvent protons
with the C3 proton of Ru5P or Xu5P by 1H NMR. In the case
of mandelate racemase, replacement of His297, the base that
abstracts the
-proton from (R)-mandelate, by Asn
eliminates both racemase activity and exchange activity between D2O and (R)-mandelate; however, the H297N mutant
catalyzed the exchange reaction between D2O and the
-proton of (S)-mandelate at a rate only 3.3-fold less
than observed with wild-type enzyme (11). This observation of
stereospecific exchange activity was indeed instrumental in assigning
the role of His297. At concentrations of the D186N mutant
in D2O that would have readily allowed detection of
exchange activity at 0.1% of wild type, the results were totally
negative with both Ru5P and Xu5P as substrate. Lack of differential
exchange activity, relative to overall epimerase activity, associated
with D186N is in accord with Asp186 serving as the
electrophile, rather than the base, in facilitation of the initial
substrate deprotonation step.
Beyond the inherent limitations to interpretation of negative results,
a particular caveat pertains to the NMR-based exchange assays.
Incorporation of deuterium from D2O into substrate can only
occur if the substrate-derived proton, transferred to an enzyme
monoprotic base, can exchange with solvent while the deprotonated substrate remains enzyme bound. This precondition may not be met. During the turnover of [3-2H]Xu5P in H2O by
wild-type Ru5P epimerase, deuterium is not lost from the remaining pool
of unprocessed substrate (2). Thus, either the substrate-derived proton
cannot exchange with solvent while the enediolate intermediate resides
at the active site or the protonation of the intermediate by the second
base to form product is far more rapid than exchange of protons between
the first base and solvent.
In conclusion, we have offered rather compelling evidence that
Asp186 of Ru5P epimerase contributes directly and
profoundly to catalysis. Circumstantial evidence favors
Asp186 as the electrophile that facilitates
-proton
abstraction by either of the general base catalysts, but its identity
as one of the two bases cannot be rigorously excluded. Future studies of solvent and substrate isotope effects may clarify this issue.
 |
ACKNOWLEDGEMENTS |
We gratefully acknowledge the expert
technical assistance of Tse-Yuan S. Lu and Alice A. Hardigree.
 |
FOOTNOTES |
*
This work was supported by the Office of Biological and
Environmental Research, United States Department of Energy, under Contract DE-AC05-96OR22464 with Lockheed Martin Energy Research Corp.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.
¶
Contributed equally to this research. To whom correspondence
should be addressed: Life Sciences Division, Oak Ridge National Laboratory, P. O. Box 2009, Oak Ridge, TN 37831-8080. Tel.:
423-574-0959; Fax: 423-574-0793; E-mail: ffh{at}ornl.gov (F. C. H.) or fwl{at}ornl.gov (F. W. L.).
The abbreviations used are:
Ru5P, ribulose
5-phosphate; IEF, isoelectric focusing; PAGE, polyacrylamide gel
electrophoresis; Xu5P, xylulose 5-phosphate.
2
Species of Ru5P epimerases that were compared
and GenBank accession numbers for their sequences are as follows:
Caenorhabditis elegans, U28991; Saccharomyces
cerevisiae, P46969; Schizosaccharomyces pombe, Z98979;
E. coli, P32661; Serratia marcescens, P45455; Hemophilus influenzae, P44756; Alcaligenes
eutrophus, P40117; Spinacia oleracea, AF070943;
Solanum tuberosum, Q43157; Synechocystis sp,
P74061; Bacillus subtilis, Y13937; Rhodobacter capsulatus, U23145; Mycobacterium tuberculosis, P71676; Helicobacter pylori, P56188; Rhodospirillum
rubrum, P51013; and Methanococcus jannashii,
Q58093.
 |
REFERENCES |
-
Chen, Y.-R.,
Hartman, F. C.,
Lu, T.-Y. S.,
and Larimer, F. W.
(1998)
Plant Physiol.
118,
199-207[Abstract/Free Full Text]
-
Davis, L.,
Lee, N.,
and Glaser, L.
(1972)
J. Biol. Chem.
247,
5862-5866[Abstract/Free Full Text]
-
Hosomi, S.,
Nakai, N.,
Kogita, J.,
Terada, T.,
and Mizoguchi, T.
(1986)
Biochem. J.
239,
739-743[Medline]
[Order article via Infotrieve]
-
Bash, P. A.,
Field, M. J.,
Davenport, R. C.,
Petsko, G. A.,
Ringe, D.,
and Karplus, M.
(1991)
Biochemistry
30,
5826-5832[Medline]
[Order article via Infotrieve]
-
Lodi, P. J.,
and Knowles, J. R.
(1991)
Biochemistry
30,
6948-6956[Medline]
[Order article via Infotrieve]
-
Alagona, G.,
Ghio, C.,
and Kollman, P. A.
(1995)
J. Am. Chem. Soc.
117,
9855-9862
-
Wu, Z. R.,
Ebrahimian, S.,
Zawrotny, M. E.,
Thornburg, L. D.,
Perez-Alvarado, G. C.,
Brothers, P.,
Pollack, R. M.,
and Summers, M. F.
(1997)
Science
276,
415-418[Abstract/Free Full Text]
-
Cho, H.-S.,
Choi, G.,
Choi, K. Y.,
and Oh, B.-H.
(1998)
Biochemistry
37,
8325-8330[CrossRef][Medline]
[Order article via Infotrieve]
-
Thornburg, L. D.,
Hénot, F.,
Bash, D. P.,
Hawkinson, D. C.,
Bartel, S. D.,
and Pollack, R. M.
(1998)
Biochemistry
37,
10499-10506[CrossRef][Medline]
[Order article via Infotrieve]
-
Landro, J. A.,
Gerlt, J. A.,
Kozarich, J. W.,
Koo, C. W.,
Shah, V. J.,
Kenyon, G. L.,
Neidhart, D. J.,
Fujita, S.,
and Petsko, G. A.
(1994)
Biochemistry
33,
635-643[Medline]
[Order article via Infotrieve]
-
Mitra, B.,
Kallarakal, A. T.,
Kozarich, J. W.,
Gerlt, J. A.,
Clifton, J. G.,
Petsko, G. A.,
and Kenyon, G. L.
(1995)
Biochemistry
34,
2777-2787[Medline]
[Order article via Infotrieve]
-
Porter, M. A.,
Milanez, S.,
Stringer, C. D.,
and Hartman, F. C.
(1986)
Arch. Biochem. Biophys.
245,
14-23[Medline]
[Order article via Infotrieve]
-
Porter, M. A.,
Stringer, C. D.,
and Hartman, F. C.
(1988)
J. Biol. Chem.
263,
123-129[Abstract/Free Full Text]
-
Weiner, M. P.,
Costa, G. L.,
Schoettlin, W.,
Cline, J.,
Mathur, E.,
and Bauer, J. C.
(1994)
Gene (Amst.)
151,
119-123[CrossRef][Medline]
[Order article via Infotrieve]
-
Larimer, F. W.,
Mural, R. J.,
and Soper, T. S.
(1990)
Protein Eng.
3,
227-231[Medline]
[Order article via Infotrieve]
-
Dower, W. J.,
Miller, J. F.,
and Ragsdale, C. W.
(1988)
Nucleic Acids Res.
16,
6127-6145[Abstract]
-
Sambrook, J.,
Fritsch, E. F.,
and Manniatis, T.
(1989)
Molecular Cloning: A Laboratory Manual, Vol. 3, p. A.3, Cold Spring Harbor Laboratory Press, New York
-
Bradford, M. M.
(1976)
Anal. Biochem.
72,
248-254[CrossRef][Medline]
[Order article via Infotrieve]
-
Kiely, M. E.,
Stuart, A. L.,
and Wood, T.
(1973)
Biochim. Biophys. Acta
293,
534-541[Medline]
[Order article via Infotrieve]
-
Rost, B.,
and Sander, C.
(1994)
Proteins
19,
55-72[Medline]
[Order article via Infotrieve]
-
Rost, B.,
Sander, C.,
and Scheider, R.
(1994)
Comput. Appl. Biosci.
10,
53-60[Abstract]
-
Farber, G. K.,
and Petsko, G. A.
(1990)
Trends Biochem. Sci.
15,
228-234[CrossRef][Medline]
[Order article via Infotrieve]
-
Brändén, C.
(1987)
in
Crystallography in Molecular Biology (Moras, D., Drenth, J., Strandberg, B., Suck, K., and Wilson, K., eds), pp. 359-364, Plenum Press, New York
-
Brändén, C.,
and Tooze, J.
(1991)
Introduction to Protein Structure, pp. 43-57, Garland Publishing, Inc., New York
-
Babbitt, P. C.,
and Gerlt, J. A.
(1997)
J. Biol. Chem.
272,
30591-30594[Free Full Text]
-
Gerlt, J. A.,
and Gassman, P. G.
(1993)
J. Am. Chem. Soc.
115,
11552-11568
-
Gerlt, J. A.,
and Gassman, P. G.
(1993)
Biochemistry
32,
11943-11952[Medline]
[Order article via Infotrieve]
-
Teige, M.,
Melzer, M.,
and Süss, K.-H.
(1998)
Eur. J. Biochem.
252,
237-244[Abstract]
Copyright © 1999 by The American Society for Biochemistry and Molecular Biology, Inc.