Structural and kinetic insights into the mechanism of 5-hydroxyisourate hydrolase from Klebsiella pneumoniae.
ABSTRACT The stereospecific oxidative degradation of uric acid to (S)-allantoin has recently been demonstrated to proceed via two unstable intermediates and requires three separate enzymatic reactions. The second step of this reaction, the conversion of 5-hydroxyisourate (HIU) to 2-oxo-4-hydroxy-4-carboxy-5-ureidoimidazoline, is catalyzed by HIU hydrolase (HIUH). The high-resolution crystal structure of HIUH from the opportunistic pathogen Klebsiella pneumoniae (KpHIUH) has been determined. KpHIUH is a homotetrameric protein that, based on sequence and structural similarity, belongs to the transthyretin-related protein family. In addition, the steady-state kinetic parameters for this enzyme and four active-site mutants have been measured. These data provide valuable insight into the functional roles of the active-site residues. Based upon the structural and kinetic data, a mechanism is proposed for the KpHIUH-catalyzed reaction.
- SourceAvailable from: Lokesh Gakhar[Show abstract] [Hide abstract]
ABSTRACT: The molecular basis of the ability of bacteria to live on caffeine via the C-8 oxidation pathway is unknown. The first step of this pathway, caffeine to trimethyluric acid (TMU), has been attributed to poorly characterized caffeine oxidases and a novel quinone-dependent caffeine dehydrogenase. Here, we report the detailed characterization of the second enzyme, a novel NADH-dependent trimethyluric acid monooxygenase (TmuM), a flavoprotein that catalyzes the conversion of TMU to 1,3,7-trimethyl-5-hydroxyisourate (TM-HIU). This product spontaneously decomposes to racemic 3,6,8-trimethylallantoin (TMA). TmuM prefers trimethyluric acids and, to a lesser extent, dimethyluric acids as substrates, but it exhibits no activity on uric acid. Homology models of TmuM against uric acid oxidase HpxO (which catalyzes uric acid to 5-hydroxyisourate) reveal a much bigger and hydrophobic cavity to accommodate the larger substrates. Genes involved in the caffeine C-8 oxidation pathway are located in a 25.2-kb genomic DNA fragment of CBB1, including cdhABC (coding for caffeine dehydrogenase) and tmuM (coding for TmuM). Comparison of this gene cluster to the uric acid-metabolizing gene cluster and pathway of Klebsiella pneumoniae revealed two major open reading frames coding for the conversion of TM-HIU to S-(+)-trimethylallantoin [S-(+)-TMA]. The first one, designated tmuH, codes for a putative TM-HIU hydrolase, which catalyzes the conversion of TM-HIU to 3,6,8-trimethyl-2-oxo-4-hydroxy-4-carboxy-5-ureidoimidazoline (TM-OHCU). The second one, designated tmuD, codes for a putative TM-OHCU decarboxylase which catalyzes the conversion of TM-OHCU to S-(+)-TMA. Based on a combination of enzymology and gene-analysis, a new degradative pathway for caffeine has been proposed via TMU, TM-HIU, TM-OHCU to S-(+)-TMA.Journal of bacteriology 05/2012; 194(15):3872-82. · 3.94 Impact Factor
- [Show abstract] [Hide abstract]
ABSTRACT: Lip is a membrane-bound lipoprotein and a core component of the type VI secretion system found in Gram-negative bacteria. The structure of a Lip construct (residues 29-176) from Serratia marcescens (SmLip) has been determined at 1.92 Å resolution. Experimental phases were derived using a single-wavelength anomalous dispersion approach on a sample cocrystallized with iodide. The membrane localization of the native protein was confirmed. The structure is that of the globular domain lacking only the lipoprotein signal peptide and the lipidated N-terminus of the mature protein. The protein fold is dominated by an eight-stranded β-sandwich and identifies SmLip as a new member of the transthyretin family of proteins. Transthyretin and the only other member of the family fold, 5-hydroxyisourate hydrolase, form homotetramers important for their function. The asymmetric unit of SmLip is a tetramer with 222 symmetry, but the assembly is distinct from that previously noted for the transthyretin protein family. However, structural comparisons and bacterial two-hybrid data suggest that the SmLip tetramer is not relevant to its role as a core component of the type VI secretion system, but rather reflects a propensity for SmLip to participate in protein-protein interactions. A relatively low level of sequence conservation amongst Lip homologues is noted and is restricted to parts of the structure that might be involved in interactions with physiological partners.Acta Crystallographica Section D Biological Crystallography 12/2011; 67(Pt 12):1065-72. · 12.67 Impact Factor
- [Show abstract] [Hide abstract]
ABSTRACT: Transthyretin-like proteins are a family of proteins that share remarkable structural similarities to transthyretin, that have been identified in a variety of taxa such as bacteria, fungi, plants, invertebrates and vertebrates. Despite the enormous progress in the study of transthyretin-like protein, little is known about it in amphioxus, a model organism for insights into the origin and evolution of vertebrates. Here we identified a transthyretin-like protein gene in Branchiostoma japonicum, named Bjtlp, which possessed a TLP-HIUase (an enzyme hydrolyzing 5-hydroxyisourate) domain and a consensus C-terminal tetrapeptide Tyr-Arg-Gly-Ser that are both characteristics of all known transthyretin-like proteins. Phylogenetic and intron-exon structure analyses support that TTR likely arose from a vertebrate specific duplication after vertebrates diverged from invertebrate chordates. Quantitative real-time PCR analysis revealed that Bjtlp was expressed in a tissue-specific fashion, with the transcript levels being most abundant in the hepatic caecum and hind gut. Enzymatic activity assays demonstrated that recombinant BjTLP had the capacity to hydrolyze 5-hydroxyisourate. Site-directed mutagenesis showed that both Y156 and R93 residues were critical for 5-hydroxyisourate hydrolase activity of recombinant BjTLP. Moreover, the single mutation, Y156T, at the active site of BjTLP caused approximately 97% loss of its enzymatic activity, and meanwhile gained the thyroxine binding activity. All these data together suggest that the single mutation Y156T is critical for converting BjTLP to a new transport protein capable of distributing thyroxine.Comparative biochemistry and physiology. Part B, Biochemistry & molecular biology 12/2012; · 1.61 Impact Factor
Acta Cryst. (2011). D67, 671–677doi:10.1107/S090744491101746X
Acta Crystallographica Section D
Structural and kinetic insights into the mechanism
of 5-hydroxyisourate hydrolase from Klebsiella
Jarrod B. French and Steven E.
Department of Chemistry and Chemical Biology,
Cornell University, Ithaca, NY 14853-1301,
Correspondence e-mail: firstname.lastname@example.org
# 2011 International Union of Crystallography
Printed in Singapore – all rights reserved
The stereospecific oxidative degradation of uric acid to
(S)-allantoin has recently been demonstrated to proceed
via two unstable intermediates and requires three separate
enzymatic reactions. The second step of this reaction, the
conversion of 5-hydroxyisourate (HIU) to 2-oxo-4-hydroxy-
4-carboxy-5-ureidoimidazoline, is catalyzed by HIU hydrolase
(HIUH). The high-resolution crystal structure of HIUH from
the opportunistic pathogen Klebsiella pneumoniae (KpHIUH)
has been determined. KpHIUH is a homotetrameric protein
that, based on sequence and structural similarity, belongs
to the transthyretin-related protein family. In addition, the
steady-state kinetic parameters for this enzyme and four
active-site mutants have been measured. These data provide
valuable insight into the functional roles of the active-site
residues. Based upon the structural and kinetic data, a
mechanism is proposed for the KpHIUH-catalyzed reaction.
Received 25 March 2011
Accepted 9 May 2011
The catabolism of uric acid, a key intermediate in the degra-
dation of purines, varies among different organisms. Birds,
reptiles, humans and some bacteria lack the necessary
enzymes to degrade uric acid (Vogels & Van der Drift, 1976;
Zrenner et al., 2006). In humans, the inability to metabolize
uric acid can lead to the crystallization of urate in the joints,
causing the painful condition known as gout. In many plants
and bacteria, however, the high nitrogen content of this
molecule makes it an attractive nitrogen source. In all known
organisms that catabolize uric acid, a conserved pathway
converts this molecule stereospecifically to (S)-allantoin
(Vogels & Van der Drift, 1976). Until recently, it was believed
that a single enzyme, urate oxidase, catalyzed the slow
conversion of urate to allantoin. Over the last few years,
however, conclusive experiments have been conducted that
demonstrate the involvement of two additional enzymes:
5-hydroxyisourate hydrolase (HIUH) and 2-oxo-4-hydroxy-
(Kahn & Tipton, 1997, 1998; Ramazzina et al., 2006). This
enzymatic pathway from uric acid to allantoin is shown in
Klebsiella pneumoniae is an opportunistic human pathogen
affecting patients with chronic pulmonary disease and ranks
second to Escherichia coli as the cause of urinary-tract infec-
tions in the elderly (Podschun & Ullmann, 1998). In addition,
K. pneumoniae is known to cause liver abscesses with meta-
static complications (Lederman & Crum, 2005). Two recent
genetic studies of Klebsiella sp. revealed a cluster of genes that
are responsible for the catabolism of uric acid (de la Riva et al.,
2008; Pope et al., 2009). Structural and biochemical studies
of the proteins encoded by this gene cluster revealed several
enzymes with novel functions, including an FAD-dependent
urate oxidase and a ureidoglycine aminotransferase (French &
Ealick, 2010a; O’Leary et al., 2009).
In this work, we report the crystal structure of K. pneu-
moniae HIUH (KpHIUH) at 1.8 A˚resolution. In this struc-
ture, a phosphate ion occupies the active site and provides
clues as to the nature of several important protein–substrate
interactions. Guided by the KpHIUH structure, several active-
site mutants were constructed and kinetically characterized.
These results were used to construct a model of substrate
binding and to propose a mechanism of catalysis that is
consistent with both the structural and the kinetic data.
2. Materials and methods
2.1. Protein expression and purification
The KpHIUH gene (hpxT) was cloned from genomic DNA
from K. pneumoniae subsp. pneumoniae (Schroeter) Trevisan
MGH78578 (ATCC 700721) and inserted into a pET-28-based
plasmid with an N-terminal His6 tag. The protein was
expressed in BL21 (DE3) cells at 288 K after induction with
0.5 mM isopropyl ?-d-1-thiogalactopyranoside. KpHIUH was
first purified by Ni–NTA (Qiagen) affinity chromatography,
which was followed by gel-filtration chromatography on an
A¨KTAexplorer FPLC with a HiLoad 26/60 Superdex prep-
grade G200 column running 10 mM Tris buffer pH 7.6 with
30 mM NaCl. After purification, the protein was concentrated
to 20 mg ml?1using a centrifugal concentrator and aliquots
were flash-frozen and stored at 193 K.
The H7N, R41K, H92N and S108A mutants were made at
the Cornell Protein Production and Purification Facility by
using site-directed mutagenesis of the native gene. Briefly, site-
directed mutagenesis was performed on the KpHIUH gene by
a standard PCR protocol using PfuTurbo DNA polymerase
(Invitrogen) and DpnI (New England Biolabs) to digest
the methylated parental DNA prior to
transformation. The presence of the
enzymes were expressed and purified in
the same manner as the native protein.
2.2. Crystallization, data collection,
data processing and structure
using the hanging-drop vapor-diffusion
method at 291 K. Crystal Screen and
Crystal Screen 2 (Hampton Research)
and Wizard I and II (Emerald Bio-
Systems) sparse-matrix screens were
used to identify initial crystallization
leads. The protein crystallized well,
forming rod-shaped crystals in several
of the initial screening conditions. Two of these were opti-
mized and yielded diffraction-quality crystals of similar size
and morphology. The optimized conditions included (i) 6–12%
PEG 3000, 0.2–0.4 M MgCl2or NaCl and 0.1 M cacodylate pH
6.0–6.5 or (ii) 0.5–1.0 M ammonium phosphate and 0.1 M
imidazole pH 7.5–8.5. The rod-shaped crystals were robust and
grew over 1–2 weeks. Microseeding of the drops prior to
sealing the trays led to crystals of equivalent quality that grew
in 2–5 d. The crystals tolerated several different cryoprotec-
tion strategies, but were typically cryoprotected in either 30%
PEG 3000, 0.2 M MgCl2and 0.1 M cacodylate pH 6.0 or 15%
glycerol, 15% ethylene glycol, 1.0 M ammonium phosphate
and 0.1 M imidazole pH 7.9. After a brief soak in the cryo-
protectant, the crystals were frozen by plunging them into
liquid nitrogen and were stored in liquid nitrogen prior to data
Data sets were collected at 100 K on the Northeastern
Collaborative Access Team (NE-CAT) beamline 24-ID-E at
Argonne National Laboratory (wavelength of 0.979 A˚)usinga
Quantum 315 CCD detector (Area Detector Systems) with 1 s
exposure times and 1?oscillations. The data were indexed,
integrated and scaled using HKL-2000 (Otwinowski & Minor,
1997). The structure was solved by molecular replacement
using MOLREP (Vagin & Teplyakov, 2010) with an all-alanine
version of the structure of Bacillus subtilis PucM (PDB entry
2h0e; Jung et al., 2006) as a search model. The model was
refined through successive rounds of manual model building
using Coot (Emsley & Cowtan, 2004) and restrained refine-
ment using REFMAC5 (Murshudov et al., 2011). Water
molecules were added only after the model converged and this
was followed by two additional rounds of refinement. Data-
collection and refinement statistics are provided in Table 1.
All kinetic measurements were carried out in 10 mM Tris
buffer pH 7.6 containing 30 mM NaCl. The substrate for the
reaction, 5-hydroxyisourate (HIU), is relatively unstable in
French & Ealick
? 5-Hydroxyisourate hydrolase
Acta Cryst. (2011). D67, 671–677
Enzymatic pathway for the conversion of uric acid to allantoin in K. pneumoniae.
aqueous solutions and must therefore be generated in situ just
prior to kinetic analysis. Briefly, an appropriate amount of
recombinant uricase from Candida sp. (Sigma–Aldrich) was
added to a solution of uric acid, resulting in complete
conversion to HIU in less than 1 min. The conversion could be
monitored by the change in the absorbance spectrum from
uric acid to HIU to OHCU (French & Ealick, 2010b). Owing
to the high rate of turnover of the HIUH enzyme, a stopped-
flow instrument (SF-2004, KinTek) was used to measure
progress curves for the reaction. Conversion of HIU to OHCU
was measured by monitoring the change in absorbance at
293 nm. To calculate kinetic parameters, instantaneous rates at
each time, tx, were approximated by calculating the rate at
points along the progress curve using the slope of the line from
tx?2 to tx+2. These approximate instantaneous rates were
plotted against the substrate concentrations and fitted using
the Michaelis–Menten equation in order to derive steady-state
kinetic parameters (French et al., 2010; Yun & Suelter, 1977).
The calculated rate of non-enzymatic hydrolysis of HIU
(2.7 ? 10?3) was consistent with a previously reported rate
(Kahn & Tipton, 1998) and was taken into account in all rate
2.4. Modeling of HIU into the KpHIUH active site
The modeling of the ligand in the active site of KpHIUH
was performed using v.9.7.211 of MacroModel (Mohamadi
et al., 1990; http://www.schrodinger.com/products/14/11/). The
protein was truncated to a shell containing all atoms within
20 A˚of the ligand. Water molecules were removed and the
protein-preparation utility was used to add H atoms and to
ensure proper ionic states of amino-acid side chains. In order
to initially place the substrate, HIU was positioned in the
active site by superposition with 8-azaxanthine from the
homologous PucM structure (Jung et al., 2006). For the
modeling runs, both the active-site residues and the ligand
were allowed the freedom to move. The calculations were
completed by energy minimization using the AMBER* force
field (Weiner et al., 1984) with a distance-dependent dielectric
that was further attenuated by a factor of 4. The energy
minimization relied upon the TNCG technique (Ponder &
Richards, 1987) and was considered
to have converged when the energy
gradient was less than 0.01 kJ mol?1.
All of the figures presented here were
prepared using PyMOL (DeLano, 2002)
and ChemBioDraw (CambridgeSoft).
The sequence alignment was generated
by ESPript (Gouet et al., 2003).
3. Results and discussion
3.1. Overall structure of KpHIUH
KpHIUH crystallized in an ortho-
rhombic space group and contained
four protomers in the asymmetric unit.
Strong electron density was present for
all 108 residues, with the exception of a
disordered loop region comprised of
residues 84–88. All of the modelled
residues have torsion angles consistent
Acta Cryst. (2011). D67, 671–677 French & Ealick
? 5-Hydroxyisourate hydrolase
Structure of K. pneumoniae 5-hydroxyisourate hydrolase. (a) KpHIUH tetramer, (b) KpHIUH
protomer. Helices are colored blue,?-strands are colored green and loop regions arecolored yellow.
The N- and C-termini are labeled N and C, respectively.
Data-collection and refinement statistics.
Values in parentheses are for the highest resolution shell.
No. of reflections
No. of protein atoms
No. of ligand atoms
No. of water atoms
R.m.s.d. bonds (A˚)
R.m.s.d. angles (?)
Mean B factor (A˚2)
Most favored (%)
Additionally allowed (%)
Generously allowed (%)
† Rmerge =
intensity of i reflections with intensities Ii(hkl).
where Fobsand Fcalcare observed and calculated structure factors, respectively.
Rfreethe sum extends over a subset of reflections (5%) that were excluded from all stages
ijIiðhklÞ ? hIðhklÞij=P
iIiðhklÞ, where hI(hkl)i is the mean
‡ R =P
??jFobsj ? jFcalcj??=P
with the most favored region of the Ramachandran plot. The
four chains of the enzyme come together to form a homo-
tetramer with 222 symmetry (Fig. 2a). Size-exclusion chro-
matography results are also consistent with a tetrameric
species (data not shown). The KpHIUH protomer consists of
nine ?-strands and one ?-helix organized into a ?-sandwich
structure (Fig. 2b). The tetramer is a dimer of dimers, with two
protomers arranged to create an extended ?-sheet that makes
up the dimer–dimer interface. While other HIUH enzymes
showed some slight differences in conformation between
protomers, there is little structural difference between the
chains of KpHIUH [the average root-mean-square deviation
(r.m.s.d.) for superposition is 0.28 A˚].
3.2. Comparison with other structures
A BLAST search (Altschul et al., 1990) using the KpHIUH
sequence places this enzyme in the transthyretin-like protein
superfamily. This family includes transthyretin, a protein that
distributes thyroid hormones throughout the body, and the
structurally similar 5-hydroxyisourate hydrolase. All of these
proteins are homotetramers that contain an eight-stranded
?-sandwich and a small ?-helix. The HIUH enzymes are
distinguished from transthyretin primarily by the YRGS
C-terminal motif. A sequence alignment of KpHIUH, trans-
thyretin and several other transthyretin-like protein family
members is shown in Fig. 3(a).
A search for structurally related proteins using the DALI
server (Holm et al., 2008) identified several structural homo-
logs. All of the highly similar structures (Z > 10) were
members of the transthyretin-like protein family and included
Salmonella dublin HIUH (PDB entry 2gpz, r.m.s.d. of 1.1 A˚;
Hennebry et al., 2006), E. coli HIUH (PDB entry 2igl, r.m.s.d.
of 1.0 A˚; Y. Zuo, J. Blanco, J. Shah, Y. Wang, S. Rivera, T. J.
Ragan, G. Hernandez, C. M. Nelersa, R. Mitchell, K. E. Rudd
& A. Malhotra, unpublished work), zebrafish HIUH (PDB
entry 3iwv, r.m.s.d. of 1.0 A˚; Cendron et al., 2011), B. subtilis
HIUH (PDB entry 2h0e, r.m.s.d. of 1.4 A˚;Jung et al., 2006) and
several transthyretin structures (rat, PDB entry 1kgj, r.m.s.d.
of 1.2 A˚; sea bream, PDB entry 1sn0, r.m.s.d. of 1.3 A˚; human,
PDB entry 1bzd, r.m.s.d. of 1.2 A˚; chicken, PDB entry 1tfp,
r.m.s.d. of 1.6 A˚; Muziol et al., 2001; Eneqvist et al., 2004;
Schormann et al., 1998; Sunde et al., 1996). Despite the high
degree of similarity to transthyretin (Fig. 3b), several recent
studies have shown that HIUH enzymes are unable to bind
hormones but catalyze the conversion of HIU to OHCU
(Hennebry et al., 2006; Kahn & Tipton, 1997, 1998). In addi-
tion, the genes for HIUH and OHCUD have been shown to
French & Ealick
? 5-Hydroxyisourate hydrolase
Acta Cryst. (2011). D67, 671–677
Alignment of KpHIUH with other transthyretin-related proteins (TRPs). (a) Sequence alignment of KpHIUH and four other TRPs. The sequences used
are those of K. pneumoniae HIUH, E. coli transthyretin-related protein (EcTTP; PDB entry 2g2n; Lundberg et al., 2006), E. coli YedX (EcYEDX; PDB
entry 2igl), B. subtilis PucM (BsPUCM; PDB entry 2h0e), zebrafish HIUH (DrHIUH; PDB entry 2h1x; Zanotti et al., 2006) and human transthyretin
(HsTTR; PDB entry 1f41; Hornberg et al., 2000). In the alignment, residues with red lettering and a blue box are highly conserved, while those with white
lettering on a red background are completely conserved. The residues that line the active site of KpHIUH are indicated by blue triangles. The secondary-
structure elements shown above the alignment correspond to those observed in the structure of KpHIUH. This figure was generated with ESPript
(Gouet et al., 2003). (b) Structural alignment of KpHIUH (blue) and human transthyretin (green).
appear within clusters of known purine-degradative enzymes
(de la Riva et al., 2008; Pope et al., 2009) and are expressed as a
single fused gene in several cases (Ramazzina et al., 2006).
3.3. KpHIUH active site
Transthyretin-like proteins are known to possess two active
sites per tetramer that are situated at a dimer interface near
the surface of the protein. Residues believed to be important
for substrate binding and catalysis include the C-terminal
tetrapeptide, two histidine residues and an arginine residue
from each chain (Fig. 3, blue arrows). These residues and
the active-site architecture are also conserved in KpHIUH
(Fig. 4a). In this structure, a phosphate is seen to be coordi-
nated by the active-site arginine residue (Arg41) and both
active-site histidine residues (His7 and His92). This phosphate
occupies a position on a plane of noncrystallographic
symmetry and appears to bind equally well to both active sites
at the dimer interface. To investigate the likely interactions of
the active-site residues with the substrate, 5-hydroxyisourate
was modeled into the active site. A schematic of the contacts
made between the modeled ligand and the active-site residues
is shown in Fig. 4(b). The positions of the active-site side
chains in this model are consistent with those observed in the
structure of the B. subtilis HIUH complex with 8-azaxanthine
and can be used to infer putative roles for the active-site
residues. Of the four amino acids of the conserved tetra-
peptide sequence, only Tyr105 appears to make significant
interactions for binding or catalysis. This residue interacts with
the O atom on C8 of the purine ring, most likely to stabilize
the charge at this atom and help to orient the HIU molecule
for catalysis. Ser108 may also participate in catalysis indirectly
by helping to orient and inductively activate His7. The two
histidine residues located in the active site, His7 and His92, are
situated near the reactive center of the substrate, although
only His92 appears to have the correct orientation and
distance to form a hydrogen bond to the substrate. The posi-
tively charged Arg41 is situated at a hydrogen-bonding
distance to the carbonyl at C6 of the purine ring, placing it in
an ideal position to stabilize a negative charge at this atom
(see mechanistic discussion below).
3.4. Kinetics of native enzyme and active-site mutants
To better understand the different functions that the active-
site residues play in catalysis, the kinetic parameters of native
KpHIUH and several mutants were characterized. The results
(Table 2) are consistent with the structural data and provide
additional details about the roles of these residues. Both of
the histidine-to-asparagine mutations effectively inactivate
the enzyme. Despite the low kcatvalue, however, the H92N
mutant has a Kmthat is only fourfold higher than that of the
native enzyme. The major difference in kcatvalues between the
native and H92N mutant suggests that His92 plays a direct role
in catalysis. The H7N mutant, however, not only experiences
a diminished kcatbut also has a significantly higher Km. This
indicates that this residue may play a more complicated role in
the enzyme mechanism and perhaps participates in substrate
binding. An apparent interaction between His7 and the
hydroxyl group of HIU (Fig. 4b) would explain a possible role
in orienting the substrate for catalysis. The S108A mutation
does not substantially alter the Kmand leads to only a twofold
reduction in rate, implying that Ser108 is not likely to
Acta Cryst. (2011). D67, 671–677 French & Ealick
? 5-Hydroxyisourate hydrolase
KpHIUH active site. (a) A stereodiagram of the active site of the KpHIUH–phosphate complex is shown. Residue names followed by an asterisk (*) are
from the adjoining chain. In this figure C atoms are colored green, O atoms are colored red, N atoms are colored blue, P atoms are colored orange and
water atoms are shown as red spheres and labeled with a ‘w’. The electron density shown is from an Fo? Fomap contoured at 3? that was calculated
before adding the phosphate. Note that as the active site coincides with a noncrystallographic symmetry plane, the phosphate molecule is modeled in two
alternative orientations. (b) A schematic of 5-hydroxyisourate modeled into the active site of KpHIUH. Residues from the two different chains are
colored red and blue, while the ligand is shown in black.
Kinetic parameters of KpHIUH.
172 ? 2
0.8 ? 0.09
34.7 ? 0.4
0.35 ? 0.01
81 ? 0.7
103 ? 2
4400 ? 500
195 ? 3
430 ? 10
93 ? 2
1.7 ? 106
1.8 ? 102
1.8 ? 105
8.2 ? 102
8.7 ? 105
participate in binding or catalysis. Similarly, the R41K mutant
retains much of its activity, but the fivefold decrease in rate
and twofold increase in Kmsuggest some minor role in the
reaction mechanism. As suggested by the structure, it is likely
that Arg41 primarily interacts with the C6 carbonyl O atom,
hydrogen bonding to the substrate and possibly helping to
stabilize a negative charge at this atom during the reaction.
3.5. Proposed mechanism of KpHIUH
A mechanism for the HIUH reaction was first proposed by
Tipton and coworkers for the soybean (Glycine max) enzyme
and was based primarily upon structural alignments with
proteins of known function and kinetic data from site-directed
mutants (Raychaudhuri & Tipton, 2003; Sarma et al., 1999). In
this mechanism an active-site glutamate is implicated as the
nucleophile that initiates the reaction. This protein, however,
belongs to the type I glycosidase family and is not structurally
homologous to the transthyretin-like HIUH enzymes. The
transthyretin-like HIUH enzymes not only lack such a gluta-
mate residue but do not possess any strong nucleophile in
their active sites.
Based upon the structural and kinetic data for KpHIUH,
a mechanistic proposal for the transthyretin-like HIUH is
presented in Fig. 5. Owing to the lack of a strong nucleophile
in the active site, the reaction is likely to be initiated by a water
molecule that is first activated by deprotonation. Considering
the geometry of the two histidine residues in the active site as
well as the kinetic data, it is likely that both of these residues
take part in this deprotonation step. The hydrogen-bonding
interactions between the water molecule and the two histidine
residues, His7 and His92, would serve to orient the water
ideally for attack at C6 of the purine ring. The highly
conserved C-terminal serine residue, Ser108 in KpHIUH, is in
position to form a hydrogen bond to His7 and may indirectly
participate in catalysis by inductively activating this residue.
Deprotonation of the water by His7 creates a hydroxide
nucleophile that attacks C6 of the purine ring, leading to a
tetrahedral oxyanion intermediate. The charge on the
resulting oxyanion would be stabilized by the positively
charged guanidinium group of Arg41. It is clear from the
direction of the attack and the geometry of the resulting
tetrahedral center that Arg41 from the neighboring chain
helps to stabilize the charge on the oxyanion intermediate.
Collapse of the oxyanion would
then lead to ring opening, with the
final proton coming from the
proton abstracted from a water
molecule by His7 would then be
transferred to Arg41 to complete
the catalytic cycle.
presented here provide important
details about the mechanism of
this enzyme and the roles that the
active-site residues play during substrate binding and catalysis.
These results move us a step closer to complete character-
ization of the enzymes involved in the purine catabolic
pathway in the opportunistic human pathogen K. pneumoniae.
In addition, this structure, together with others involved in the
pathway, helps to suggest potential targets for antimicrobial
therapies that target purine metabolism in this organism.
This work was supported by NIH grant GM73220. We
would like to thank the staff at NE-CAT for advice with data
collection and processing. Data collected at the Northeastern
Collaborative Access Team beamlines of the Advanced
Photon Source are supported by award RR-15301 from the
National Center for Research Resources at the National
Institutes of Health. Use of the Advanced Photon Source is
supported by the US Department of Energy, Office of Basic
Energy Sciences under Contract No. DE-AC02-06CH11357.
We would like to acknowledge Cynthia Kinsland of the
Protein Production Facility in the Department of Chemistry
and Chemical Biology for help with molecular biology and
Leslie Kinsland for help with manuscript preparation. JBF
would like to acknowledge the Tri-Institutional Training
Program in Chemical Biology for financial support.
Altschul, S. F., Gish, W., Miller, W., Myers, E. W. & Lipman, D. J.
(1990). J. Mol. Biol. 215, 403–410.
Cendron, L., Ramazzina, I., Percudani, R., Rasore, C., Zanotti, G. &
Berni, R. (2011). J. Mol. Biol., doi:10.1016/j.jmb.2011.04.022.
DeLano, W. L. (2002). PyMOL. http://www.pymol.org.
Emsley, P. & Cowtan, K. (2004). Acta Cryst. D60, 2126–2132.
Eneqvist, T., Lundberg, E., Karlsson, A., Huang, S., Santos, C. R.,
Power, D. M. & Sauer-Eriksson, A. E. (2004). J. Biol. Chem. 279,
French, J. B., Cen, Y., Vrablik, T. L., Xu, P., Allen, E., Hanna-Rose, W.
& Sauve, A. A. (2010). Biochemistry, 49, 10421–10439.
French, J. B. & Ealick, S. E. (2010a). Biochemistry, 49, 5975–5977.
French, J. B. & Ealick, S. E. (2010b). J. Biol. Chem. 285, 35446–35454.
Gouet, P., Robert, X. & Courcelle, E. (2003). Nucleic Acids Res. 31,
Hennebry, S. C., Wright, H. M.,Likic,V. A.& Richardson, S. J. (2006).
Proteins, 64, 1024–1045.
Holm, L., Ka ¨a ¨ria ¨inen, S., Rosenstro ¨m, P. & Schenkel, A. (2008).
Bioinformatics, 24, 2780–2781.
Hornberg, A., Eneqvist, T., Olofsson, A., Lundgren, E. & Sauer-
Eriksson, A. E. (2000). J. Mol. Biol. 302, 649–669.
French & Ealick
? 5-Hydroxyisourate hydrolase
Acta Cryst. (2011). D67, 671–677
A mechanistic proposal for the conversion of 5-hydroxyisourate to 2-oxo-4-hydroxy-4-carboxy-5-
ureidoimidazoline by K. pneumoniae HIUH.
Jung, D.-K., Lee, Y., Park, S. G., Park, B. C., Kim, G.-H. & Rhee, S.
(2006). Proc. Natl Acad. Sci. USA, 103, 9790–9795.
Kahn, K. & Tipton, P. A. (1997). Biochemistry, 36, 4731–4738.
Kahn, K. & Tipton, P. A. (1998). Biochemistry, 37, 11651–
Lederman, E. R. & Crum, N. F. (2005). Am. J. Gastroenterol. 100,
Lundberg, E., Ba ¨ckstro ¨m, S., Sauer, U. H. & Sauer-Eriksson, A. E.
(2006). J. Struct. Biol. 155, 445–457.
Mohamadi, F., Richards, N. G. J., Guida, W. C., Liskamp, R., Lipton,
M., Caufield, C., Chang, G., Hendrickson, T. & Still, W. C. (1990). J.
Comput. Chem. 11, 440–467.
Murshudov, G. N., Skuba ´k, P., Lebedev, A. A., Pannu, N. S., Steiner,
R. A., Nicholls, R. A., Winn, M. D., Long, F. & Vagin, A. A. (2011).
Acta Cryst. D67, 355–367.
Muziol, T., Cody, V. & Wojtczak, A. (2001). Acta Biochim. Pol. 48,
O’Leary, S. E., Hicks, K. A., Ealick, S. E. & Begley, T. P. (2009).
Biochemistry, 48, 3033–3035.
Otwinowski, Z. & Minor, W. (1997). Methods Enzymol. 276, 307–326.
Podschun, R. & Ullmann, U. (1998). Clin. Microbiol. Rev. 11,
Ponder, J. W. & Richards, F. M. (1987). J. Comput. Chem. 8, 1016–
Pope, S. D., Chen, L.-L. & Stewart, V. (2009). J. Bacteriol. 191, 1006–
Ramazzina, I., Folli, C., Secchi, A., Berni, R. & Percudani, R. (2006).
Nature Chem. Biol. 2, 144–148.
Raychaudhuri, A. & Tipton, P. A. (2003). Biochemistry, 42, 6848–
Riva, L. de la, Badia, J., Aguilar, J., Bender, R. A. & Baldoma, L.
(2008). J. Bacteriol. 190, 7892–7903.
Sarma, A. D., Serfozo, P., Kahn, K. & Tipton, P. A. (1999). J. Biol.
Chem. 274, 33863–33865.
Schormann, N., Murrell, J. R. & Benson, M. D. (1998). Amyloid, 5,
Sunde, M., Richardson, S. J., Chang, L., Pettersson, T. M., Schreiber,
G. & Blake, C. C. (1996). Eur. J. Biochem. 236, 491–499.
Vagin, A. & Teplyakov, A. (2010). Acta Cryst. D66, 22–25.
Vogels, G. D. & Van der Drift, C. (1976). Bacteriol. Rev. 40, 403–468.
Weiner, S. J., Kollman, P. A., Case, D. A., Singh, U. C., Ghio, C.,
Alagona, G., Profeta, S. Jr & Weiner, P. (1984). J. Am. Chem. Soc.
Yun, S.-L. & Suelter, C. H. (1977). Biochim. Biophys. Acta, 480, 1–13.
Zanotti, G., Cendron, L., Ramazzina, I., Folli, C., Percudani, R. &
Berni, R. (2006). J. Mol. Biol. 363, 1–9.
Zrenner, R., Stitt, M., Sonnewald, U. & Boldt, R. (2006). Annu. Rev.
Plant Biol. 57, 805–836.
Acta Cryst. (2011). D67, 671–677French & Ealick
? 5-Hydroxyisourate hydrolase