From the School of Biological Sciences, Biosciences
Building, University of Liverpool, P. O. Box 147, Liverpool L69 7ZB,
United Kingdom, the ¶ Department of Molecular Biology and
Biotechnology, Firth Court, Sheffield S10 2TN, and the
** Department of Chemistry, Krebs Institute for Biomolecular
Research, University of Sheffield, Firth Court,
Sheffield S3 7HF, United Kingdom
Received for publication, November 25, 2002
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The contributions to substrate
binding and catalysis of 13 amino acid residues of the
Caenorhabditis elegans diadenosine tetraphosphate pyrophosphohydrolase (Ap4A hydrolase) predicted from the
crystal structure of an enzyme-inhibitor complex have been investigated by site-directed mutagenesis. Sixteen glutathione
S-transferase-Ap4A hydrolase fusion proteins
were expressed and their kcat and
Km values determined after removal of the
glutathione S-transferase domain. As expected for a Nudix
hydrolase, the wild type kcat of 23 s Ap4A1
hydrolases are enzymes that hydrolyze dinucleoside polyphosphates.
Structurally and mechanistically, they fall into two groups. The
symmetrically cleaving enzymes (EC 3.6.1.41), such as Escherichia
coli ApaH, generate 2-ADP from Ap4A, whereas the asymmetrically cleaving enzymes (EC 3.6.1.17) produce AMP and ATP (1,
2). The latter are members of the Nudix hydrolases, a family of
structurally and catalytically similar enzymes that act upon a wide
range of different nucleotide substrates. Some are highly specific
whereas others appear to have a broad substrate range in
vitro (3-5). The Nudix Ap4A hydrolases can be further subdivided into "plant" and "animal"-types, according to their primary structure (6). The plant-type includes enzymes from the
Proteobacteria that have in some cases been shown to be associated with
the invasion of mammalian cells, whereas the animal-type includes
putative Ap4A hydrolases from Archaea (6-10). Early
studies of both animal and plant Ap4A Nudix hydrolases
employing a combination of substrate analogues and labeling with heavy
isotopes of oxygen revealed the mechanism of hydrolysis to involve
in-line nucleophilic attack of a water molecule at the
P4 (P Detailed structural studies of E. coli MutT first showed the
importance of the highly conserved residues in the loop-helix-loop Nudix motif (Fig. 1). Glu53,
Glu56, Glu57, Glu98 (outside the
linear motif but structurally close), and the carbonyl of
Gly38 coordinate an enzyme-bound Mg2+ ion. A
water ligand of this ion is oriented or deprotonated for nucleophilic
attack by Glu53, which is itself oriented by
Arg52. A second metal ion is complexed to the substrate and
neutralizes the charge on the attacked phosphate while
Lys39 activates the NMP leaving group (18-21). The
importance of Glu57 was indicated by a 105-fold
reduction in kcat in a E57Q mutant (19). The
contributions of the other residues to catalysis were also confirmed by
site-directed mutagenesis: E53Q, E56Q, and E44Q led to
104.7-, 25-, and 14-fold decreases in
kcat, respectively (20), whereas K39Q and R52Q
produced 8-fold and >103-fold reductions, respectively
(22). The principle of this catalytic mechanism appears to be well
conserved among the Nudix hydrolases, including the lupin and
Bartonella bacilliformis Ap4A hydrolases (15-17, 23), human MTH1 (24), yeast Dcp2p (25), and human NUDT3
(DIPP1) (26).
1 was reduced by 105-, 103-, and
30-fold, respectively, by replacement of the conserved P4-phosphate-binding catalytic residues
Glu56, Glu52, and Glu103 by Gln.
Km values were not affected, indicating a lack of
importance for substrate binding. In contrast, mutating
His31 to Val or Ala and Lys83 to Met produced
10- and 16-fold increases in Km compared with the
wild type value of 8.8 µM. These residues stabilize the P1-phosphate. H31V and H31A had a normal
kcat but K83M showed a 37-fold reduction in
kcat. Lys36 also stabilizes the
P1-phosphate and a K36M mutant had a 10-fold
reduced kcat but a relatively normal
Km. Thus both Lys36 and
Lys83 may play a role in catalysis. The previously
suggested roles of Tyr27, His38,
Lys79, and Lys81 in stabilizing the
P2 and P3-phosphates
were not confirmed by mutagenesis, indicating the absence of
phosphate-specific binding contacts in this region. Also, mutating both
Tyr76 and Tyr121, which clamp one substrate
adenosine moiety between them in the crystal structure, to Ala only
increased Km 4-fold. It is concluded that
interactions with the P1- and
P4-phosphates are minimum and sufficient
requirements for substrate binding by this class of enzyme, indicating
that it may have a much wider substrate range then previously believed.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
) phosphate with subsequent breakage of
the P4-(O)P3 bond (8,
11-14). Recently, the catalytic residues of the lupin Ap4A
hydrolase involved in this process were identified by a combination of
structural analysis and site-directed mutagenesis (15-17). This study
supported the catalytic mechanism previously described in detail for
the prototypical Nudix hydrolase, the E. coli MutT 8-oxo-dGTPase.
View larger version (14K):
[in a new window]
Fig. 1.
The consensus Nudix motif and the actual
motifs of E. coli MutT protein, C. elegans (C.e.) Ap4A hydrolase, and
Lupinus angustifolius (L.a.)
Ap4A hydrolase. The numbers indicate the
positions in each primary structure.
Among the asymmetrically cleaving Ap4A hydrolases,
identification of residues responsible for substrate binding as well as catalysis is of interest for two reasons. First, it will help our
understanding of the evolution of substrate specificity among the Nudix
hydrolases. Second, if the plant-type Ap4A hydrolase of
invasive pathogenic bacteria is to be considered as a target for new
antibacterial agents, the design of such agents will require knowledge
of the subtle differences between the plant and animal types if
selectivity is to be achieved. Recently we reported the crystal
structure of an animal Ap4A hydrolase from the nematode Caenorhabditis elegans in both free form and after
crystallization in the presence of the substrate analogue,
AppCH2ppA (27, 28). The structure of the resulting binary
complex allowed some predictions to be made about the importance of
certain residues for substrate binding and catalysis and comparisons to
be drawn with the lupin enzyme. Here, we extend these studies to
include the effects of 19 site-specific mutations on Ap4A
binding and hydrolysis by the C. elegans enzyme.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Synthesis of C. elegans First Strand cDNA Library--
Total
RNA was isolated and purified from washed adult nematodes (C. elegans strain N2) using Trizol solution (Invitrogen) according to the manufacturer's instructions. Full-length C. elegans first strand cDNA was synthesized from this RNA using
a first strand cDNA synthesis kit (MPI Fermentas). RNA (2 µl of
2.5 µg/µl) was added to 10 µl of RNase-free ddH2O.
The solution was mixed gently, incubated at 70 °C for 5 min, and
chilled on ice for 3 min before adding to a mixture containing 4 µl
of 5× reaction buffer (250 mM Tris-HCl, pH 8.3, at
25 °C, 375 mM KCl, 15 mM MgCl2, 5 mM dithiothreitol), 1 µl of ribonuclease
inhibitor (20 units/µl), 2 µl of oligo(dT)18 (0.5 µg/µl), 2 µl of dNTPs (10 mM each), and 2 µl of
Moloney murine leukemia virus reverse transcriptase (20 units/µl,
Promega). The reaction was incubated at 42 °C for 1 h, and then
heated to 90 °C for 5 min. The library was stored at 20 °C.
Cloning of C. elegans Ap4A Hydrolase as a Glutathione S-Transferase (GST) Fusion Protein-- A cDNA corresponding to the C. elegans Y37H9A.6 Ap4A hydrolase gene (6) was amplified from the cDNA library by PCR using the forward and reverse primers d(CAGCGCCAGAATTCAATGGTCGTAAAAGCCGCGGG) and d(GAAATTACTCGAGAAAAATCGTTAAAATCCGGC), respectively. These primers provided an EcoRI restriction site at the start of amplified cDNA and a XhoI site at the end. After amplification with Taq DNA polymerase, the DNA was recovered, digested with EcoRI and XhoI, and the required restriction fragment ligated between the EcoRI and XhoI sites of the pGEX-6P-3 vector (Amersham Biosciences). The resulting construct, pGEX-Y37H9A, encoded the 137-amino acid Ap4A hydrolase fused to the C terminus of GST through a 6-amino acid linker.
Generation of Site-specific Mutants-- Site-directed mutagenesis was performed by PCR using the QuikChangeTM site-directed mutagenesis kit (Stratagene). PCR reactions contained pGEX-Y37H9A as template, Pfu Turbo DNA polymerase, and pairs of complementary oligonucleotide primers 37 to 43 nucleotides long containing the required mutations (Table I). Each reaction volume was 50 µl and contained the following: 50-100 ng of plasmid DNA, 125 ng of each mutagenic primer, 200 µM dNTPs, 10 mM KCl, 6 mM (NH4)2SO4, 20 mM Tris-HCl, pH 8.0, 2 mM MgCl2, 0.1% Triton X-100, 10 µg/ml bovine serum albumin, and 2.5 units of Pfu Turbo DNA polymerase. The PCR reaction protocol consisted of 2 min at 95 °C followed by 16 cycles of 95 °C for 1 min, 55 °C for 1 min, 68 °C for 14 min, followed by a final incubation at 72 °C for 15 min. Parental DNA was digested with 10 units of DpnI to degrade the methylated parental strands and the remaining plasmid DNA was used to transform E. coli XL1-Blue cells. For production of the Y76A/Y121A double mutant, the Y76A DNA construct was used as template in a PCR containing the Y121A mutagenic primers (Table I). The identities of all mutants were verified by complete sequencing of both DNA strands.
Expression and Purification of GST-Ap4A Hydrolase
Fusion Proteins--
E. coli strain BL21(DE3) was
transformed with pGEX-Y37H9A or its mutant derivatives. Cultures (250 ml) in LB medium containing 50 µg/ml ampicillin were grown to an
A600 of 0.7 at 37 °C.
Isopropyl-1-thio--D-galactopyranoside was added to 1 mM and incubation continued for 2 h. Induced cells (approximately 1.6 g) were harvested by centrifugation at
10,000 × g, washed, and resuspended in 10 ml of
ice-cold breakage buffer: 50 mM Tris-HCl, pH 8.0, 50 mM NaCl, 1 mM dithiothreitol, 1 mM phenylmethylsulfonyl fluoride, and 5 µM
trans-epoxysuccinyl-L-leucylamido-(4-guanidino)butane) (E-64, Sigma). Cell suspensions were sonicated and the resulting lysates cleared by centrifugation at 15,000 × g and
4 °C for 10 min. Supernatants were recovered and applied to columns
containing 2.5 ml of glutathione-Sepharose 4B (Amersham Biosciences).
Columns were washed with 25 ml of phosphate-buffered saline, followed by 25 ml of PreScission cleavage buffer (50 mM Tris-HCl, pH
7.0, 150 mM NaCl, 1 mM dithiothreitol, 1 mM EDTA, 5% (v/v) glycerol). Following complete elution of
the buffer, the outlets were closed and 100 units of PreScission
protease in 2.5 ml of cleavage buffer added to the resin and incubated
for 18-20 h with gentle rocking at 4 °C. Cleavage of the GST domain
from the Ap4A hydrolases was complete after 20 h.
Columns were remounted, the resin left to settle, and the free
Ap4A hydrolases containing the N-terminal extension
GPLGSPNS eluted.
Ap4A Hydrolase Assay-- Ap4A hydrolase activity was measured using a luciferase-based bioluminescence assay as previously described (6). One ng enzyme protein was used in each case, except for K83M (10 ng), K79M (20 ng), E52Q and E103Q (60 ng), and E56Q (600 ng). This sensitive, continuous assay permits direct evaluation of initial rates. The increase in luminescence was linear for several minutes for each enzyme.
Other Methods--
Protein concentrations were estimated by the
Coomassie Blue binding method (29) and protein molecular masses were
determined by electrospray mass spectrometry as previously described
(30).
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Expression and Purification of Wild Type and Mutant
GST-Ap4A Hydrolase Fusion Proteins--
A cDNA
corresponding to the C. elegans Y37H9A.6 Ap4A
hydrolase gene was amplified from a cDNA library by PCR and
inserted into the pGEX-6P-3 GST fusion vector to generate the
recombinant plasmid pGEX-Y37H9A. When E. coli BL21(DE3)
cells were transformed with this plasmid and induced with
isopropyl-1-thio--D-galactopyranoside, a major soluble
43-kDa band corresponding to the expected GST-Ap4A hydrolase fusion protein was detected (data not shown). The GST domain
of this protein was readily removed by on-column cleavage with
PreScission protease, resulting in the free Ap4A hydrolase with the N-terminal extension GPLGSPNS and mass 16.6 kDa.
Specific mutations were introduced into the Ap4A hydrolase
coding region of pGEX-Y37H9A by PCR (Table
I). The mutations were confirmed by DNA
sequencing. A total of 16 single mutants and one double mutant
involving 13 different residues was generated in this way. Their
positions in the primary structure of the Ap4A hydrolase
are shown in Fig. 2A and the
locations of their -carbon atoms in the three-dimensional structure
of the binary complex are shown in Fig. 2B. Each was
expressed as a GST fusion protein and purified after on-column cleavage
and elution as described for the wild type. The predicted masses of the
cleaved proteins were confirmed by mass spectrometry. With the
exception of five (K36M, Y76A, Y76A/Y121A, K79M, and W32G) mutant
proteins were substantially expressed in the soluble fraction (at least
40% of the total expressed recombinant protein) and the yields of purified proteins were nearly the same as for the wild type. The first
four exceptions yielded about 5% of the recombinant protein in a
soluble form, whereas W32G was completely insoluble when expressed. All
kinetic data were determined using enzymes purified from the soluble
fractions, which were all judged to be more than 95% pure by
SDS-PAGE.
|
|
Effects of Mutations on the Catalytic Properties of
Ap4A Hydrolase--
The values of Km
(8.8 µM) and kcat (23 s1) estimated for the N-terminal extended wild type
Ap4A hydrolase after cleavage from the GST domain were
close to those previously reported for the native recombinant enzyme
purified by conventional procedures (7.0 µM and 27 s
1, respectively) (6). From this we conclude that the
8-amino acid N-terminal extension does not interfere significantly with the binding of the substrate or with catalysis. The linearity of light
output from the luminometric assay used was the same for all mutants as
for the wild type, indicating the stability of the mutants under assay
conditions. Km and kcat
values were then determined for each mutant enzyme to determine the
effects of each mutation on substrate binding and catalysis (Table
II). For this enzyme,
Km can be taken to approximate the dissociation constant of the ES complex(es), and hence as an inverse measure of
affinity, based on the lack of effect of active site
(kcat) mutants on the value of
Km. Previous mutational studies with Nudix
hydrolases have highlighted the importance of the Glu residues within
the Nudix motif for catalysis (Fig. 1) (16, 20, 25, 26, 31, 32). Not
surprisingly, therefore, the E56Q mutation was found to result in a
105-fold reduction in
kcat and virtual abolition of
detectable enzyme activity, exactly as was found for the equivalent
residue (Glu59) in the lupin Ap4A hydrolase; in
contrast, the Km was unaffected, indicating that the
mutation has no effect on substrate binding. Similarly, neutralization
of the charge on Glu52, the second of the three highly
conserved Glu residues within the Nudix motif, by conversion to Gln
(E52Q) reduced kcat by a factor of
103 but again had little effect on Km
(Table II). On the basis of the 105-fold reduction in
kcat, we previously proposed that
Glu56 was most likely to be the catalytic base that
deprotonates the attacking water molecule. However, the structural
equivalents of Glu52 in the lupin Ap4A
hydrolase (Glu55) and in the E. coli MutT
protein (Glu53) have been proposed as the deprotonating
base (11, 16, 20). Glu103, although not in the Nudix motif,
is positioned close to it in the three-dimensional structure and
coordinates two of the four Mg2+ ions located in the
catalytic site (27). E103Q has a 30-fold lower
kcat than the wild type and a similar
Km. The equivalent mutations in E. coli
MutT (E98Q) and the lupin Ap4A hydrolase (E125Q) produced
6.3- and 140-fold reductions in kcat,
respectively (16, 20). Whereas these values suggest that
Glu103 and its equivalents are unlikely to be the catalytic
base in these enzymes, a detailed structural analysis of E. coli ADP-ribose pyrophosphatase has led to the conclusion that the
equivalent Glu162 has this role in that enzyme (33, 34).
Thus, although the architecture of the catalytic sites are broadly
similar among the Nudix hydrolases, the mechanism of proton abstraction
from the attacking water/hydroxyl appears to be subtly different in different family members.
|
Whereas several mutational studies of Nudix hydrolases have highlighted
the importance to catalysis of particular residues in and adjacent to
the catalytic motif, less attention has been focused on residues
elsewhere that may be involved in substrate recognition and binding as
well as catalysis. The crystal structure of the C. elegans
Ap4A hydrolase binary complex showed that the adenine ring
in the "AMP-binding pocket" distal to the
P4-phosphate (the site of nucleophilic attack)
was sandwiched between the phenolic rings of Tyr76 and
Tyr121 and formed extensive -
stacking interactions
with these residues. To achieve this, a 90° rotation of the phenolic
ring of Tyr121 about the
1 dihedral angle and an
associated shift in Tyr76 occurred upon substrate binding
(27). Both of these residues are highly conserved in structure-based
sequence alignments of animal and plant Ap4A hydrolases.
This, coupled with further interactions between the side chain hydroxyl
group of Tyr121 and the 2'-OH of the attached ribose and
between the ring of Tyr76 and the ribose O4
oxygen suggested that both Tyr residues should be important for substrate binding, therefore the effects of replacing each with Ala
were investigated. As expected, both Y76A and Y121A showed an increased
Km, but only by a factor of about 8 (Table II).
Surprisingly, the combination of mutations in the double mutant
Y76A/Y121A appeared to reduce the Km, again such that it was only 4-fold higher than the wild type. This suggests that
the substrate may be able to bind effectively in a way that is
independent of the Tyr residues (see "Discussion"). This
alternative binding does lead to a lower catalytic rate, as evidenced
by the reduced kcat values (20-fold less in the
double mutant).
The crystal structure of the binary complex also showed that the
P1-phosphate attached to the above adenosine
moiety is stabilized on the enzyme via a series of hydrogen bonds/salt
bridges between the phosphate oxygens and the side chain NZ nitrogens
of Lys36 and Lys83, the side chain hydroxyl
group of Tyr76, and the side chain imidazole ring
N2 of His31 (27). Replacement of
His31 with Ala (H31A) or Val (H31V) had a significant and
specific impact on the binding of Ap4A, increasing the
Km 8-12-fold while only marginally reducing
kcat. Loss of the NZ nitrogen of Lys83 (K83M) led to an even greater increase in
Km (16-fold) but in this case
kcat was also substantially reduced (37-fold). The K36M mutant also had a reduced kcat
(10-fold) but a relatively normal Km. These results
confirm the predictions of the crystal structure and indicate the
importance of Lys36 and Lys83, which is
positioned such that it could also stabilize the
P2-phosphate, to catalysis (27).
As there was no interpretable electron density for either the
P2- or P3-phosphates in
the binary complex, it was suggested that the side chains of
His38 (within the Nudix motif) plus Lys79,
Lys81, and Tyr27 (outside the Nudix motif)
might be in appropriate positions to participate in
P2- and P3-phosphate
stabilization either by direct interaction or by metal coordination.
The main chain amide of His38 is also the only direct
protein contact with P4 via a hydrogen bond to
one of the oxygen atoms (27). Potentially, His38
(structurally equivalent to Lys39 in the E. coli
MutT protein) and/or Lys79 and/or Lys81 could
neutralize the developing negative charge on the ATP leaving group, in
much the same way as has been proposed for MutT Lys39 (18,
20). Therefore, appropriate mutants were generated to test these
suggestions. Surprisingly, of the mutants analyzed (Y27A, Y27D, H38G,
H38K, K79M, and K81M), only K79M showed a significant change in any
kinetic constant, a substantial 140-fold reduction in
kcat. However, Lys79, like
Tyr27, is not well conserved among the Ap4A
hydrolases, so this reduced activity may reflect a slight structural
alteration in the protein rather than an important catalytic role. In
contrast, Lys81 is well conserved as a basic residue in
animal and plant Ap4A hydrolases. However, its mutation to
Met resulted in a slight increase in kcat to 30 s1, so it seems unlikely to be involved in stabilizing
the leaving group. H38G and H38K also showed slight increases in
kcat such that the
kcat/Km ratio was 2.5-fold
higher than the wild type. The equivalent residue in plant
Ap4A hydrolases is a Gly, so, unlike Lys39 in
MutT, His38 does not appear to be important for catalysis
either. As all other residues in the region are too small to make
contact, the conclusion is that there are few, if any, structurally or
mechanistically important binding contacts for the
P2- and P3-phosphates.
This interesting point is discussed further below.
Finally, Trp32 is a completely conserved residue among
Ap4A hydrolases of the Nudix family and is commonly found
in other family members. This residue does not form interactions with
the substrate but appears to stabilize the protein fold through
interactions with Leu22 and Gln24 in the B
strand and with Ile118 in the
II helix (27). Consistent
with this essential structural role is the fact that the W32G mutant
was completely insoluble and inactive when expressed.
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The results of this analysis confirm the importance of residues previously implicated by structural analysis in binding and catalysis at the P4-phosphate and binding of the P1-phosphate of Ap4A by the C. elegans Ap4A hydrolase. It also provides important information on two further aspects of substrate binding by this enzyme that have implications for substrate recognition in the Nudix hydrolase family as a whole. First, there appear to be few, if any, important binding contacts for the P2- and P3-phosphates. Our previous structural analysis was unable to provide this information. It is possible that a nucleotide-bound metal ion is more important for stabilization of the negative charge during catalysis than any individual amino acid side chain. Such an ion appears to be required by the B. bacilliformis Ap4A hydrolase (23) and the E. coli ADP-ribose pyrophosphatase (33). However, the apparent absence of specific interactions in this region is entirely consistent with our recent discovery that this and related dinucleoside polyphosphate hydrolases can bind and hydrolyze 5-phosphoribosyl 1-pyrophosphate (35). In this case the ribose ring would occupy the P2,P3 site, with binding dependent solely on interactions in the P1 and P4 sites, in agreement with the data above.
Second, the mutational data relating to Tyr76 and
Tyr121 were somewhat unexpected. The 90° rotation of the
phenolic ring of Tyr121 in the binary complex compared with
the apoenzyme and the -
stacking interactions between both rings
and the adenine ring of the adenosine moiety attached to the
P1-phosphate originally suggested an essential
role for these residues. However, substitution of both Tyr residues by
Ala did not have the expected dramatic effect on substrate binding and
yielded only a 4-fold increase in Km. Again, this is
consistent with the finding that 5-phosphoribosyl 1-pyrophosphate,
which lacks a base altogether, is a substrate. Thus, binding of one adenine ring between the Tyr residues is not an essential requirement for catalysis, although it undoubtedly contributes to the higher specificity constant for Ap4A compared with
5-phosphoribosyl 1-pyrophosphate (35). Interestingly, in the NMR
structure of the lupin Ap4A hydrolase complexed with the
substrate analogue ATP-MgFx, the adenine ring is not
located between the structurally equivalent residues Tyr77
and Phe144, and instead Tyr77 is suggested to
be important for the structural integrity of the enzyme-substrate
complex rather than for direct substrate binding (17). Although
structurally very similar, plant and animal Ap4A hydrolases
typically show only 25-30% sequence similarity outside the Nudix
motif and appear to form two distinct evolutionary groups within the
family (6). Thus, either the animal and plant enzymes differ
substantially in the way they bind substrate, or the possibility exists
that Ap4A can bind to both Ap4A hydrolases in
two different ways. Conceivably, one site represents the true substrate
binding site before hydrolysis whereas the other is a transitional site
for the ATP leaving group. Indeed, in view of the lack of electron
density for the P2- and
P3-phosphates and the second adenosine moiety in
the crystal structure of the C. elegans binary complex, we
have suggested that the analogue AppCH2ppA was probably
hydrolyzed during crystallization (27). Thus, the visible AMP may be
the AppCH2p product with its mobile P2- and P3-phosphates
disordered in the crystal lattice and therefore invisible, or even the
AMP product that has re-bound between the Tyr residues after departure
of the ATP. If binding of the adenine ring between Tyr76
and Tyr121 is not essential, this could also explain our
observations that phosphonate analogues with isopolar halomethylene
groups bridging P1 and P2
and P3 and P4 such as
ApCHFppCHFpA, and
P1,P4-thiophosphates such
as ApspppsA can be cleaved symmetrically by
Nudix Ap4A hydrolases (14, 36). This requires attack at P2 or P3. Assuming that
the loop-helix-loop structural motif containing the catalytic residues
cannot move significantly, symmetrical hydrolysis implies that the
substrate is bound in such a way as to present
P3 rather than the usual
P4 to the attacking nucleophile. Thus,
substrates are able to bind in more than one location. Taken
together, the ability of some Nudix hydrolases to use non-nucleotide
substrates such as 5-phosphoribosyl 1-pyrophosphate and
diphosphoinositol polyphosphates and the flexibility of substrate
binding noted above suggest that the substrate range and function of
Nudix hydrolases may be much wider than previously believed.
![]() |
FOOTNOTES |
---|
* This work was supported in part by the Biotechnology and Biological Sciences Research Council and the Wellcome Trust (to J. B. R. and A. G. McL.).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.
§ Recipient of a postgraduate scholarship from the Egyptian government.
To whom correspondence should be addressed: School of
Biological Sciences, Biosciences Building, University of Liverpool, P. O. Box 147, Liverpool L69 7ZB, United Kingdom. Tel.: 151-795-4426; Fax: 151-795-4404; E-mail: agmclen@liv.ac.uk.
Royal Society Olga Kennard Fellow.
§§ Present address: Dept. of Molecular Biophysics and Biochemistry, Yale University, Bass Center, 266 Whitney Ave., New Haven, CT 06520-8114.
Published, JBC Papers in Press, December 9, 2002, DOI 10.1074/jbc.M211983200
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
Ap4A, diadenosine
5',5-P1,P4-tetraphosphate;
AppCH2ppA, diadenosine
5',5
-(P2,P3-methylene)-P1,P4-tetraphosphate;
Nudix, nucleoside diphosphate linked to
X;
GST, glutathione S-transferase.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Guranowski, A., and Sillero, A. (1992) in Ap4A and Other Dinucleoside Polyphosphates (McLennan, A. G., ed) , pp. 81-133, CRC Press Inc., Boca Raton, FL |
2. | Guranowski, A. (2000) Pharmacol. Ther. 87, 117-139[CrossRef][Medline] [Order article via Infotrieve] |
3. |
Bessman, M. J.,
Frick, D. N.,
and O'Handley, S. F.
(1996)
J. Biol. Chem.
271,
25059-25062 |
4. | McLennan, A. G. (1999) Int. J. Mol. Med. 4, 79-89[Medline] [Order article via Infotrieve] |
5. | McLennan, A. G., Cartwright, J. L., and Gasmi, L. (2000) in Purine and Pyrimidine Metabolism in Man X (Zoref-Shani, E. , and Sperling, O., eds), Vol. 486 , pp. 115-118, Kluwer Academic/Plenum Publishers, New York |
6. | Abdelghany, H. M., Gasmi, L., Cartwright, J. L., Bailey, S., Rafferty, J. B., and McLennan, A. G. (2001) Biochim. Biophys. Acta 1550, 27-36[Medline] [Order article via Infotrieve] |
7. |
Bessman, M. J.,
Walsh, J. D.,
Dunn, C. A.,
Swaminathan, J.,
Weldon, J. E.,
and Shen, J. Y.
(2001)
J. Biol. Chem.
276,
37834-37838 |
8. | Cartwright, J. L., Britton, P., Minnick, M. F., and McLennan, A. G. (1999) Biochem. Biophys. Res. Commun. 256, 474-479[CrossRef][Medline] [Order article via Infotrieve] |
9. |
Conyers, G. B.,
and Bessman, M. J.
(1999)
J. Biol. Chem.
274,
1203-1206 |
10. | Badger, J. L., Wass, C. A., and Kim, K. S. (2000) Mol. Microbiol. 36, 174-182[CrossRef][Medline] [Order article via Infotrieve] |
11. | Guranowski, A., Brown, P., Ashton, P. A., and Blackburn, G. M. (1994) Biochemistry 33, 235-240[Medline] [Order article via Infotrieve] |
12. |
Dixon, R. M.,
and Lowe, G.
(1989)
J. Biol. Chem.
264,
2069-2074 |
13. | McLennan, A. G., Prescott, M., and Evershed, R. P. (1989) Biomed. Environ. Mass Spectrom. 18, 450-452[Medline] [Order article via Infotrieve] |
14. | McLennan, A. G., Taylor, G. E., Prescott, M., and Blackburn, G. M. (1989) Biochemistry 28, 3868-3875[Medline] [Order article via Infotrieve] |
15. | Swarbrick, J. D., Bashtannyk, T., Maksel, D., Zhang, X. R., Blackburn, G. M., Gayler, K. R., and Gooley, P. R. (2000) J. Mol. Biol. 302, 1165-1177[CrossRef][Medline] [Order article via Infotrieve] |
16. | Maksel, D., Gooley, P. R., Swarbrick, J. D., Guranowski, A., Gange, C., Blackburn, G. M., and Gayler, K. R. (2001) Biochem. J. 357, 399-405[CrossRef][Medline] [Order article via Infotrieve] |
17. | Fletcher, J. I., Swarbrick, J. D., Maksel, D., Gayler, K. R., and Gooley, P. R. (2002) Structure 10, 205-213[CrossRef][Medline] [Order article via Infotrieve] |
18. | Lin, J., Abeygunawardana, C., Frick, D. N., Bessman, M. J., and Mildvan, A. S. (1997) Biochemistry 36, 1199-1211[CrossRef][Medline] [Order article via Infotrieve] |
19. | Lin, J., Abeygunawardana, C., Frick, D. N., Bessman, M. J., and Mildvan, A. S. (1996) Biochemistry 35, 6715-6726[CrossRef][Medline] [Order article via Infotrieve] |
20. | Harris, T. K., Wu, G., Massiah, M. A., and Mildvan, A. S. (2000) Biochemistry 39, 1655-1674[CrossRef][Medline] [Order article via Infotrieve] |
21. | Mildvan, A. S., Weber, D. J., and Abeygunawardana, C. (1999) Adv. Enzymol. Relat. Areas Mol. Biol. 73, 183-209[Medline] [Order article via Infotrieve] |
22. | Frick, D. N., Weber, D. J., Abeygunawardana, C., Gittis, A. G., Bessman, M. J., and Mildvan, A. S. (1995) Biochemistry 34, 5577-5586[Medline] [Order article via Infotrieve] |
23. | Conyers, G. B., Wu, G., Bessman, M. J., and Mildvan, A. S. (2000) Biochemistry 39, 2347-2354[CrossRef][Medline] [Order article via Infotrieve] |
24. |
Cai, J. P.,
Kawate, H.,
Ihara, K.,
Yakushiji, H.,
Nakabeppu, Y.,
Tsuzuki, T.,
and Sekiguchi, M.
(1997)
Nucleic Acids Res.
25,
1170-1176 |
25. |
Dunckley, T.,
and Parker, R.
(1999)
EMBO J.
18,
5411-5422 |
26. |
Yang, X. N.,
Safrany, S. T.,
and Shears, S. B.
(1999)
J. Biol. Chem.
274,
35434-35440 |
27. | Bailey, S., Sedelnikova, S. E., Blackburn, G. M., Abdelghany, H. M., Baker, P. J., McLennan, A. G., and Rafferty, J. B. (2002) Structure 10, 589-600[CrossRef][Medline] [Order article via Infotrieve] |
28. | Bailey, S., Sedelnikova, S. E., Blackburn, G. M., Abdelghany, H. M., McLennan, A. G., and Rafferty, J. B. (2002) Acta Crystallogr. Sect. D 58, 526-528[CrossRef][Medline] [Order article via Infotrieve] |
29. | Peterson, G. L. (1983) Methods Enzymol. 91, 95-119[Medline] [Order article via Infotrieve] |
30. | Thorne, N. M. H., Hankin, S., Wilkinson, M. C., Nuñez, C., Barraclough, R., and McLennan, A. G. (1995) Biochem. J. 311, 717-721[Medline] [Order article via Infotrieve] |
31. |
Fujii, Y.,
Shimokawa, H.,
Sekiguchi, M.,
and Nakabeppu, Y.
(1999)
J. Biol. Chem.
274,
38251-38259 |
32. |
Shimokawa, H.,
Fujii, Y.,
Furuichi, M.,
Sekiguchi, M.,
and Nakabeppu, Y.
(2000)
Nucleic Acids Res.
28,
3240-3249 |
33. | Gabelli, S. B., Bianchet, M. A., Ohnishi, Y., Ichikawa, Y., Bessman, M. J., and Amzel, L. M. (2002) Biochemistry 41, 9279-9285[CrossRef][Medline] [Order article via Infotrieve] |
34. | Gabelli, S. B., Bianchet, M. A., Bessman, M. J., and Amzel, L. M. (2001) Nat. Struct. Biol. 8, 467-472[CrossRef][Medline] [Order article via Infotrieve] |
35. |
Fisher, D. I.,
Safrany, S. T.,
Strike, P.,
McLennan, A. G.,
and Cartwright, J. L.
(2002)
J. Biol. Chem.
277,
47313-47317 |
36. | Blackburn, G. M., Taylor, G. E., Thatcher, G. R., Prescott, M., and McLennan, A. G. (1987) Nucleic Acids Res. 15, 6991-7004[Abstract] |