From the Dipartimento di Scienze Biochimiche "A. Rossi Fanelli" and Centro di Biologia Molecolare del Consiglio Nazionale delle Ricerche, Università "La Sapienza", Roma, Italy and the § School of Molecular and Medical Biosciences, University of Wales, Cardiff CF1 1XL, United Kingdom
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ABSTRACT |
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Two histidine residues in glutamate decarboxylase
from Escherichia coli, potential participants in catalysis
because they are conserved among amino acid decarboxylases and because
they are at the active site in the homologous enzyme ornithine
decarboxylase, were mutated. His-275 is shown to bind the cofactor
pyridoxal 5-phosphate but not to contribute directly to catalysis. The H275N enzyme was unable to bind the cofactor whereas the H275Q mutant
contained 50% of the normal complement of cofactor and its specific
activity (expressed per mole of cofactor) was 70% of that of the
wild-type enzyme. The H167N mutant bound the cofactor tightly, its
specific activity was approximately half that of the wild-type enzyme
and experiments in D2O showed that it catalyzed replacement
of the carboxyl group with retention of configuration as does the
wild-type enzyme. Comparison of reaction profiles by observing changes
in the absorbance of the cofactor after stopped-flow mixing, revealed
that a slow reaction, in which approximately one-third of the wild-type
enzyme is converted to an unreactive complex during catalysis, does not
occur with the H167N mutant enzyme. This reaction is attributed to a
substrate-induced conformational change, a proposal that is supported
by differential scanning calorimetry.
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INTRODUCTION |
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Glutamate decarboxylase is a member of a large family of
pyridoxal-phosphate
(PLP)1-dependent
enzymes which catalyze a wide variety of different reactions on their
amino acid substrates (1). Although the enzyme from Escherichia
coli is the most studied of the amino acid decarboxylases, its
three-dimensional structure has not been solved. However, sequence
comparisons show that it is homologous to ornithine decarboxylase from
Lactobacillus 30a, the three-dimensional structure of which
(2, 3) shows two histidine residues, His-223 and His-354, close to and
on either side of the coenzyme (2, 3). These histidines are conserved
in the sequences of most, if not all, of the PLP-dependent
amino acid decarboxylases (2). There is no doubt that His-275 of
E. coli glutamate decarboxylase aligns with His-354 of
ornithine decarboxylase since, in both enzymes as in other amino acid
decarboxylases, it immediately precedes the lysine residue that forms
an imine with the coenzyme (Fig. ins;1821f1}1). In
ornithine decarboxylase, the imidazole ring of His-354 contributes to
binding the 5-phosphate of the cofactor and is considered unlikely to
participate in catalysis unless it is displaced by a substrate-induced
conformational change (2, 3). Alignment of His-167 of glutamate
decarboxylase is less clear because sequence similarity between the two
enzymes is weak in this region (Fig. 1). However, the presence, in
ornithine decarboxylase, of the imidazole ring of His-223 just below
the imine of the cofactor on the re face, suggests that this
arrangement may be a common feature of the amino acid
decarboxylases. His-167 in glutamate decarboxylase is the only
histidine residue that can reasonably occupy this position unless there
are major differences between the folds of the two proteins.
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Several stages in the catalytic mechanism of amino acid decarboxylation
involve proton transfers for which histidine residues might be
responsible (Scheme ins;1821s1}1). The formation of
an external aldimine (III) requires multiple proton transfers and, after decarboxylation to give the quinonoid intermediate (IV), the
carboxyl group is replaced by a proton. Further proton transfers, analogous to those occurring in external aldimine formation, are required for liberation of the product 4-aminobutyrate. An additional protonation occurs in a side reaction where a proton is added to C4 of
the cofactor rather than C
of the substrate to give an external
aldimine of pyridoxamine phosphate with succinic semialdehyde. This
reaction, which occurs only once in 3 × 105 turnovers
in E. coli glutamate decarboxylase (4), is a feature of most
amino acid decarboxylases (5-9). In some of these enzymes this
abortive transamination is kinetically more prominent and is almost
certainly metabolically important because it inactivates the enzyme by
leaving the cofactor as pyridoxamine phosphate.
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Considerable attention has been given to the stereochemistry of amino
acid decarboxylation and subsequent protonation. Decarboxylation requires the carboxyl group to be orthogonal to the plane comprising the cofactor pyridinium ring and the imine double bond (10). The proton
which replaces the carboxyl group at C arrives from the same
direction in which the carboxyl group has left (11). Retention of
configuration has also been observed in all other amino acid
decarboxylases in which the question has been investigated (12-14).
Kinetic evidence (15) indicates that, in methionine decarboxylase, a
histidine residue protonates at C
, whereas a lysine residue
protonates at C4
and it has been argued that lysine and histidine
perform the corresponding protonations in glutamate decarboxylase (16).
In the present work, we have examined the effect of mutations at
His-167 and at His-275 on the kinetic and structural properties of the
enzyme.
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EXPERIMENTAL PROCEDURES |
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Analytical Reagents--
4-Aminobutyrate aminotransferase was
prepared as described (17). D2O was from Sigma, Vent
polymerase was from New England Biolabs. Restriction enzymes, T4 DNA
ligase, the agarose gel DNA extraction kit, and Gabase were from
Boehringer. The T7 sequencing kit, DEAE-Sepharose, and Sephadex G-25
were from Pharmacia. [-35S]dATP (1000 Ci/mmol) was
from NEN Life Science Products. Ingredients for bacterial growth were
from Difco. Oligonucleotides were from Genenco. Other chemicals were
from BDH.
Site-directed Mutagenesis--
Site-directed mutagenesis was
performed by overlap extension polymerase chain reactions (18).
External primers annealing over the N- and C-terminal sequences were
those used in the construction of the expression plasmid containing the
gadB gene (19). Mutagenic primers for H167N were
5-GCTGGAATAAATTCGCCC-3
and its complementary sequence.
Those for H275N and H275Q were 5
-GCTTCAGGCCAGAAATTCG-3
and 5
-GCTTCAGGCAATAAATTCG-3
and their complementary
sequences, respectively. Plasmid pQgadB was used as template. The
products of polymerase chain reactions (25 cycles), carried out with
2.5 units of Vent polymerase with denaturation at 95 °C for 1 min, annealing at 45 °C (H167N) or 48 °C (H275N and H275Q) for 1 min, and extension at 74 °C for 2 min, were used as templates with the
external primers to generate the complete coding sequence of glutamate
decarboxylase. Fragments (NcoI/EcoRV for H167N
and EcoRV/HindIII for H275N and H275Q) were
subcloned into pQgadB. The newly inserted parts of the
expression constructs, pQgadH167N, pQgadH275N,
and pQgadH275Q, were sequenced and the plasmids were used to
transform E. coli JM109 carrying the plasmid pREP4.
Expression, purification, and assay of mutant forms of glutamate
decarboxylase were as described for the wild-type enzyme (19) except
where stated.
Reconstitution of Apo-enzyme with
N-(5-Phosphopyridoxyl)glutamate and Calorimetric
Analysis--
N-(5
-Phosphopyridoxyl)glutamate was prepared
by treating 0.5 M sodium glutamate and 0.5 mM
PLP (pH 4.5) with sodium cyanoborohydride (10 mM).
Apo-forms of wild-type and mutant forms of glutamate decarboxylase (10 mg/ml in 0.1 M piperazine-HCl, pH 4.5, containing 0.1 M dithiothreitol) were reconstituted with a 5-fold molar
excess of N-(5
-phosphopyridoxyl)glutamate for 1 h
(25 °C). Samples were concentrated, separated on Sephadex G-25, and
the proteins dialyzed against 20 mM sodium acetate (pH 3.6)
containing 0.1 mM dithiothreitol. Thermal unfolding of
degassed samples (1.5-2.0 mg/ml) was analyzed under nitrogen pressure
on a MicroCal MC-2D differential scanning calorimeter (MicroCal, Inc.,
Northampton, MA). Results were corrected for instrumental baseline and
normalized for protein concentration. No reversibility was observed in
a second heating cycle.
Deuterium Exchange Reactions-- Enzyme samples were brought into 99.5% D2O by repeated concentration and dilution. The enzyme (0.8 µM) was mixed with glutamate (14.25 mM). DCl was added to maintain constant pD of 4.6 (reading on pH meter = 4.2). At the end of the reaction (2 h) the solution was neutralized by adding solid Tris, lyophilized, and redissolved in 0.5 ml of D2O. NMR spectra were determined using a Bruker AMX 360 spectrometer. The stereochemistry of the deuterated 4-aminobutyrate was determined by repeating the NMR analysis after an overnight incubation in the presence of 4-aminobutyrate aminotransferase (0.36 mg/ml).
Stopped and Quenched Flow Measurements, Spectrophotometry-- Stopped-flow experiments were performed on a SF-1 stopped-flow spectophotometer (Hi-Tech, Salisbury, United Kingdom). Product formation during the period from 0.2 to 7 s was measured using a quenched flow apparatus (8, 20). Reactions lasting longer than 7 s were stopped manually. Curve fitting and statistical analyses were performed using the data manipulation software Scientist (Micromath, Salt Lake City, UT). Absorption spectra were measured with a Hewlett-Packard model 8452 diode-array spectophotometer. CD spectra were recorded as the average of 3 scans on a Jasco 710 spectropolarimeter equipped with a DP 520 processor at 25 °C using a 2-mm quartz cell.
Quantitation of 4-Aminobutyrate-- 4-Aminobutyrate was measured by high performance liquid chromatography (8) or using a commercial preparation containing 4-aminobutyrate aminotransferase and succinic semialdehyde dehydrogenase (19).
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RESULTS |
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Stability, Specific Activity, Cofactor Content, and pH Dependence
of Absorption Spectra--
The far UV CD spectra of the H167N and
H275Q mutants were identical with that of the wild-type enzyme (Fig.
ins;1821f2}2) indicating that these mutations had
not introduced major changes in the global structure of the enzyme. The
H275N mutant precipitated too rapidly at low concentrations to permit
Far UV CD analysis. Yields of the H167N mutant enzyme after the
standard purification, which does not include added PLP, were as high
as those of the wild-type enzyme. This form, like the wild-type enzyme,
was stable for many months at 4 °C and for several hours at room
temperature. Its absorption and CD spectra (Fig. 2) were almost
identical with those of the wild-type enzyme, indicating that one
molecule of cofactor was bound per monomer. The specific activity (126 µmol min1 mg
1) was approximately half
that of the wild-type enzyme. Differential scanning calorimetry of the
wild-type enzyme (Fig. ins;1821f3}3) showed that the
transition temperature of the apoenzyme reconstituted with the covalent
substrate-cofactor adduct,
N-(5
-phosphopyridoxyl)glutamate, was 8 °C higher
than that of the native holo-enzyme (51 °C) suggesting that binding
of glutamate at the active site stabilizes the structure considerably.
When the same experiment was carried out on the H167N enzyme, the
holoenzyme was found to be 4 °C more stable than the wild-type
enzyme but the mutant was not further stabilized when the cofactor was
replaced by N-(5
-phosphopyridoxyl)glutamate (Fig. 3).
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Deuterium Exchange Reactions--
In ornithine decarboxylase,
His-223 which we hypothesize to be equivalent to His-167 in glutamate
decarboxylase, occupies a position on the re face of the
cofactor. Lys-355 in ornithine decarboxylase is on the si
face and the equivalent residue in glutamate decarboxylase is Lys-276.
In the aminotransferases from the same family, the proton transfers
occur on the si face and are mediated by the equivalent of
Lys-276 (23). It is known that decarboxylation of glutamate occurs with
retention of configuration (11) but it is not known from which side the
carboxyl group leaves. Quantum mechanical calculations (4) confirm that
the C-COO
bond has two positions of maximal lability,
each perpendicular to the coenzyme ring as predicted by Dunathan (10),
but pointing in opposite directions. It is possible therefore that the
carboxyl group leaves from the re face and that protonation
of C
is mediated from this face by His-167. To test this
possibility, the 4-aminobutyrate produced by wild-type and H167N
glutamate decarboxylase in D2O was analyzed by NMR. In both
cases the signal from protons at C4 was halved, indicating that the
4-aminobutyrate produced by each form of the enzyme was monodeuterated
at C4 (equivalent to C
of glutamate). When the products were treated
in D2O with 4-aminobutyrate aminotransferase, which
labilizes the pro-S proton at C4 of 4-aminobutyrate exclusively (24), the signal from C4 protons was lost in each case.
This indicated incorporation of a second deuterium at C4 .
Rapid Mixing and Quenching Experiments-- Significant differences between the H167N mutant and the wild-type enzyme were observed when changes in the absorption of the cofactor were measured after stopped-flow mixing with glutamate.
The reaction with the most readily interpreted kinetic behavior was that of the H167N mutant. Changes in absorbance were largest at 322 nm. A large increase, complete within the mixing time (2 ms), was followed by a smaller increase which followed a single exponential (Fig. ins;1821f4}4a, k = 30 ± 1 s
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Solvent Isotope Effects--
During the investigations aimed at
measuring the incorporation of deuterium into the product
4-aminobutyrate (see "Deuterium Exchange Reactions"), we noticed
large differences in the rates at which acid needed to be added to
maintain constant pH in the reactions catalyzed by the wild-type
enzyme. An earlier study of the effects of D2O on the
reaction catalyzed by glutamate decarboxylase has already shown large
solvent isotope effects (25) and these have been cited in support of a
proposal that, after the decarboxylation step, protonation at C is
mediated by a histidine residue (16). To investigate these effects
further, we conducted rapid mixing experiments in D2O with
wild-type enzyme and with the H167N mutant. We measured the steady
state constants by analyzing the complete reaction profile after
omitting the first 5 s containing the transient. Our results for
the wild-type enzyme (Fig. 5b) are in broad agreement with
those published earlier (25). We found kcat
(26.3 ± 0.8 s
1) to be 2.7 times smaller and
kcat/Km (2.3 ± 0.1 mM
1 s
1) to be 3.5 times smaller
than in H2O. We also observed a pronounced solvent isotope
effect on the slow transient, the amplitude of which was greatly
increased both at 420 and 322 nm. The whole process, including the slow
transient, fitted well to Scheme 2 and gave constants of best fit
k1 = 0.84 ± 0.37 s
1,
k2 = 0.4 ± 0.1 s
1,
kc = 74 ± 38 s
1, and
Ks = 32 ± 12 mM. The fit assigned
extinction coefficients to the species ES and EX
of 1982 ± 130 M
1 cm
1 and
4320 ± 610 M
1 cm
1,
respectively. The increased size of the transient suggested that, if it
is due to slow formation of a species off the reaction pathway, the
course of product formation should be characterized by a more
pronounced burst when the reaction is carried out in D2O.
Fig. 5b (inset) shows the course of
4-aminobutyrate production measured by quenched-flow over the first
10 s of reaction and confirms that a pronounced burst is present.
The constants of best fit for these data were k1 = 2.3 ± 1.7 s
1, k2 = 0.65 ± 0.15 s
1, kc = 42 ± 19 s
1, Ks = 32 ± 12 mM. Although the constants derived from the different types
of experiment are not as closely similar as those from the
corresponding experiments in H2O, we consider that, in view
of the large standard deviations, Scheme 2 also provides a satisfactory
explanation.
Decarboxylation-dependent Transamination--
The
345-nm chromophore formed in the slow side reaction has been proposed
(4) to arise by tautomerization of the quinonoid structure formed after
decarboxylation (Scheme 1; IV). In this proposal, the proton on N of
the cofactor-substrate imine has transferred to O3 of the cofactor to
give an intermediate which was considered to be more likely to
protonate at C4
than at C
and thereby to lead to the abortive
transamination reaction. The H167N mutant provides a test of this
proposal because it clearly does not undergo the proposed
tautomerization and should therefore not lose activity progressively by
transamination. The transamination reaction was measured experimentally
(4) by following the course of product formation over a long period
(100 min). Despite the fact that substrate concentration remained high,
the reaction slowed and stopped but was restarted by addition of PLP.
We have confirmed these observations and additionally found that the
H167N mutant shows the same behavior (Fig.
ins;1821f6}6). It is clear that the rate of
inactivation through transamination is the same when His-167 is absent
even though the 345-nm chromophore is not formed. Therefore the
formation of the 345-nm chromophore is not required as a precursor to
transamination and its identification as the tautomer of the quinonoid
(4) is probably incorrect. It was noticeable that, using the conditions
described in Ref. 4 (pyridine HCl as buffer), the rate of the reactions
catalyzed by the H167N mutant was only about 30% less than that of the
wild-type enzyme compared with 50% in acetate buffer.
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DISCUSSION |
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The experiments with glutamate decarboxylase mutated at His-275
show that this residue contributes significantly to binding the
cofactor and that glutamine is an adequate replacement whereas asparagine is not. Because the amide nitrogens of glutamine and asparagine superimpose on the and
nitrogens of histidine, respectively, it is proposed (26) that substitution of histidine by
these two residues allows identification of the imidazole nitrogen involved in a functional hydrogen bond. Our observations indicate that
the
nitrogen of His-275 makes a hydrogen bond with the cofactor.
The corresponding histidine (His-231) in PLP-dependent histidine decarboxylase has also been replaced either by asparagine or
glutamine (27). Estimates of the activity of these mutants in crude
extracts showed that the glutamine mutant had 12% of the specific
activity of the wild-type enzyme whereas the asparagine mutant was
barely active (0.16%). The authors considered inefficient coenzyme
binding to be the cause of the lower activity and the
nitrogen was
identified as the atom responsible for hydrogen bonding (27). This also
appears to be the arrangement in the crystallographic structure of
ornithine decarboxylase where His-354 makes a hydrogen bond with an
oxygen of the cofactor 5
-phosphate (3). The evidence thus suggests
that a hydrogen bond between the cofactor phosphate and the histidine
that almost invariably precedes the cofactor-binding lysine in the
sequence, is a common feature in the family of amino acid
decarboxylases.
The similarity of the values obtained from analyzing the complete
reaction profile of the H275Q mutant to those obtained for the
wild-type enzyme agrees well with the observation that this form of the
enzyme has a specific activity 70% that of the wild-type enzyme. We
conclude that His-275 is not essential for catalyzing any of the proton
transfers in the reaction. This conclusion is to be expected if, as
argued above, His-275 occupies a position equivalent to that of the
corresponding histidine in ornithine decarboxylase (3) where the
nitrogens of the imidazole ring are too far from C of the substrate
to act as proton donor.
The fact that His-275 can be replaced by glutamine without serious loss
of the ability to bind the coenzyme or to catalyze the reaction, raises
the question of why it is so strongly conserved within the amino acid
decarboxylases. In the evolutionarily-related B6 enzyme, serine
hydroxymethyltransferase, it can be replaced by asparagine without
substantial loss of activity or ability to bind the cofactor (28). It
was suggested that in this enzyme it interacts with the substrate to
induce a closure of the enzyme necessary for specificity (28). The
coenzyme-binding lysine in tryptophan synthase, which belongs to a
completely different family, is also preceded by a histidine which can
be mutated without major loss of enzyme activity or ability to bind the
cofactor (29). In this case the structure shows the imidazole to be
near the cofactor phosphate but not as near as would be expected for bonding. This histidine is not present in any of the 51 sequences of 14 different aminotransferases even though some of these are undoubtedly
evolutionarily related to the group of amino acid decarboxylases of
which E. coli glutamate decarboxylase is a member (1). In
many of the aminotransferases, the corresponding residue is serine and
in E. coli aspartate aminotransferase, for which a structure
is available, this serine makes a hydrogen bond with the 5-phosphate
of the cofactor just as we propose for the histidine of glutamate
decarboxylase. The fact that preparations of the H275Q glutamate
decarboxylase mutant have less than the full complement of cofactor
shows that glutamine is not a perfect replacement so that substitution
of the histidine incurs a penalty which may have been sufficient to
ensure that this residue was conserved.
Our hypothesis that His-167 functions as the general acid that
protonates at C was constructed on the basis that this residue is
also highly conserved in amino acid decarboxylases. The hypothesis was
clearly wrong. Comparison of specific activities and of
kcat and
kcat/Km values showed that
the H167N mutant was only 2-3-fold less active than the wild-type
enzyme. Thus His-167 does not contribute greatly to any rate-limiting
protonation in the mechanism. However, on this evidence alone, it
remained possible that His-167 ensured very rapid protonation from the
re side. In this case, substitution of His-167 could yield a
form of the enzyme which still catalyzes efficient decarboxylation but
without the stereochemical fidelity of the wild-type enzyme. The
deuterium exchange experiments demonstrate that the direction in which
the carboxyl group is replaced is unaffected by mutation of His-167, showing that this residue is not responsible for protonating at C
of
the substrate. We conclude that His-167 is not directly involved in
catalysis.
However, His-167 does play a significant part in events that occur
after glutamate has bound. The slow conversion of the wild-type enzyme
to an unreactive complex during the first few seconds of reaction, is
not detectable in the H167N mutant. The absorbance changes that the
cofactor undergoes in the first few seconds after mixing with glutamate
have been reported earlier (21, 4) and it has been shown that the
chromophore formed absorbs maximally at 345 nm (4). However, the
kinetic constants characterizing the different phases were not
determined and it was not shown that the process leads to a less active
form of the enzyme. In Scheme 2, the complex EX lies off the
path leading to product. Its formation is analogous to the slow
appearance of uncompetitive inhibition. Km and
kcat are decreased in the same proportion by its
existence and kcat/Km is
therefore unchanged. Thus, when [S] Km, the
formation of this unreactive complex will not result in significantly
lower steady-state rates. If, as is common for most enzymes, glutamate
decarboxylase normally operates far from saturation, the side reaction
will not be metabolically disadvantageous.
Some deductions can be made about the nature of the side reaction
promoted by His-167. Immediately after mixing, the system has high
absorbance at 420 nm. At the substrate concentrations used, the enzyme
is almost saturated so that the intermediate at highest concentration
must be one of the two external aldimines, either that with glutamate
(III in Scheme 2) or that with the product 4-aminobutyrate. The most
reasonable explanation for the large difference in coenzyme spectrum
from a 420-nm absorbing chromophore to one that absorbs at 345 nm is
that a proton has been lost from the external aldimine, either
dissociated altogether or transferred to O3 to give a less resonant
tautomer. However, the process is far too slow to be due directly to
proton transfer and it seems likely that it results from a slow
conformational change, induced by the binding of glutamate, to a form
with a lower pK for the external aldimine or a lower
polarity favoring the less polar tautomer. The calorimetric evidence
(Fig. 3) supports this proposal because it shows that the binding of
glutamate stabilizes the wild-type enzyme but not the H167N mutant. The
fact that the transition temperature is already 4 °C higher in the
H167N holoenzyme indicates that this mutant enzyme has probably already
adopted at least part of the conformational change normally induced by substrate. Effects of this kind are seen with other
PLP-dependent enzymes which are known to undergo
substrate-induced conformational changes (30, 31).
The absence of the side reaction in decarboxylation catalyzed by the H167N mutant shows that most of the previously observed and unexpectedly large solvent isotope effect on kcat (25) is due to the increase in the extent to which the unproductive side reaction occurs and not to an effect on a protonation that is directly involved in the chemical transformations. This observation illustrates the difficulties associated with interpreting solvent isotope effects in enzyme-catalyzed reactions.
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FOOTNOTES |
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* This work was supported by European Community Human Capital Mobility Program Contract CHRX-CT93-0179 and the Biotechnology and Biological Sciences Research Council.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.
Part of this work to be submitted in partial fulfillment of the
requirements for the Ph.D degree at the Università "La
Sapienza", Rome.
¶ To whom correspondence should be addressed: MOMED, University of Wales, P. O. Box 911, Cardiff CF1 3US, United Kingdom. Tel.: 44-1222-874114; Fax: 44-1222-874116.
1
The abbreviation used is: PLP, pyridoxal
5-phosphate.
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REFERENCES |
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