Diversification of function in the haloacid dehalogenase enzyme superfamily: The role of the cap domain in hydrolytic phosphorus{single bond}carbon bond cleavage

Article abstract of DOI:10.1016/j.bioorg.2006.09.007

Phosphonatase functions in the 2-aminoethylphosphonate (AEP) degradation pathway of bacteria, catalyzing the hydrolysis of the C{single bond}P bond in phosphonoacetaldehyde (Pald) via formation of a bi-covalent Lys53ethylenamine/Asp12 aspartylphosphate intermediate. Because phosphonatase is a member of the haloacid dehalogenase superfamily, a family predominantly comprised of phosphatases, the question arises as to how this new catalytic activity evolved. The source of general acid-base catalysis for Schiff-base formation and aspartylphosphate hydrolysis was probed using pH-rate profile analysis of active-site mutants and X-ray crystallographic analysis of modified forms of the enzyme. The 2.9 A X-ray crystal structure of the mutant Lys53Arg complexed with Mg²⁺ and phosphate shows that the equilibrium between the open and the closed conformation is disrupted, favoring the open conformation. Thus, proton dissociation from the cap domain Lys53 is required for cap domain-core domain closure. The likely recipient of the Lys53 proton is a water-His56 pair that serves to relay the proton to the carbonyl oxygen of the phosphonoacetaldehyde (Pald) substrate upon addition of the Lys53. The pH-rate profile analysis of active-site mutants was carried out to test this proposal. The proximal core domain residues Cys22 and Tyr128 were ruled out, and the role of cap domain His56 was supported by the results. The X-ray crystallographic structure of wild-type phosphonatase reduced with NaBH₄ in the presence of Pald was determined at 2.4 A resolution to reveal Nε-ethyl-Lys53 juxtaposed with a sulfate ligand bound in the phosphate site. The position of the C(2) of the N-ethyl group in this structure is consistent with the hypothesis that the cap domain Nε-ethylenamine-Lys53 functions as a general base in the hydrolysis of the aspartylphosphate bi-covalent enzyme intermediate. Because the enzyme residues proposed to play a key role in P{single bond}C bond cleavage are localized on the cap domain, this domain appears to have evolved to support the diversification of the HAD phosphatase core domain for catalysis of hydrolytic P{single bond}C bond cleavage.

Full text of DOI:10.1016/j.bioorg.2006.09.007

Bioorganic Chemistry 34 (2006) 394–409
www.elsevier.com/locate/bioorg
Diversification of function in the
haloacid dehalogenase enzyme superfamily: The
role of the cap domain in hydrolytic
q,qq
phosphorusAcarbon bond cleavage
a
b
Sushmita D. Lahiri , Guofeng Zhang ,
b,
a,
*
Debra Dunaway-Mariano *, Karen N. Allen
a
Department of Physiology and Biophysics, Boston University School of Medicine, Boston, MA 02118-2394, USA
b
Department of Chemistry, University of New Mexico, Albuquerque, NM 87131, USA
Received 30 August 2006
Abstract
Phosphonatase functions in the 2-aminoethylphosphonate (AEP) degradation pathway of bacte-
ria, catalyzing the hydrolysis of the CAP bond in phosphonoacetaldehyde (Pald) via formation of a
bi-covalent Lys53ethylenamine/Asp12 aspartylphosphate intermediate. Because phosphonatase is a
member of the haloacid dehalogenase superfamily, a family predominantly comprised of phospha-
tases, the question arises as to how this new catalytic activity evolved. The source of general acid-
base catalysis for Schiff-base formation and aspartylphosphate hydrolysis was probed using pH-rate
profile analysis of active-site mutants and X-ray crystallographic analysis of modified forms of the
˚
2+
enzyme. The 2.9 A X-ray crystal structure of the mutant Lys53Arg complexed with Mg and phos-
phate shows that the equilibrium between the open and the closed conformation is disrupted, favor-
ing the open conformation. Thus, proton dissociation from the cap domain Lys53 is required for cap
domain–core domain closure. The likely recipient of the Lys53 proton is a water-His56 pair that
serves to relay the proton to the carbonyl oxygen of the phosphonoacetaldehyde (Pald) substrate
upon addition of the Lys53. The pH-rate profile analysis of active-site mutants was carried out to
q
This work was supported by NIH Grant GM61099 to K.N.A. and D.D.-M.
The coordinates of the refined structures have been deposited with the Protein Data Bank, accession codes
qq
2
4
IOH and 2IOF for K53R and NaBH reduced phosphonatase, respectively.
Corresponding authors. Fax: +1 617 638 4273 (K.N. Allen), +1 505 277 6202 (D. Dunaway-Mariano).
*
E-mail addresses: dd39@unm.edu (D. Dunaway-Mariano), allen@med-xtal.bu.edu (K.N. Allen).
0
045-2068/$ - see front matter ꢀ 2006 Elsevier Inc. All rights reserved.
doi:10.1016/j.bioorg.2006.09.007

S.D. Lahiri et al. / Bioorganic Chemistry 34 (2006) 394–409
395
test this proposal. The proximal core domain residues Cys22 and Tyr128 were ruled out, and the role
of cap domain His56 was supported by the results. The X-ray crystallographic structure of wild-type
˚
phosphonatase reduced with NaBH in the presence of Pald was determined at 2.4 A resolution to
4
reveal Ne-ethyl-Lys53 juxtaposed with a sulfate ligand bound in the phosphate site. The position of
the C(2) of the N-ethyl group in this structure is consistent with the hypothesis that the cap domain
Ne-ethylenamine-Lys53 functions as a general base in the hydrolysis of the aspartylphosphate bi-co-
valent enzyme intermediate. Because the enzyme residues proposed to play a key role in PAC bond
cleavage are localized on the cap domain, this domain appears to have evolved to support the diver-
sification of the HAD phosphatase core domain for catalysis of hydrolytic PAC bond cleavage.
ꢀ
2006 Elsevier Inc. All rights reserved.
Keywords: Schiff-base; Phosphoryl transfer; Phosphoenzyme; Phosphoaspartate; Structural enzymology; HAD
superfamily; General acid catalysis; General base catalysis; Enamine; Phosphate ester hydrolysis; Phosphonatase;
Phosphonate; Cap domain; Core domain; Electrophilic catalysis
1
. Introduction
Phosphonoacetaldehyde hydrolase (phosphonatase) catalyzes the hydrolysis of phos-
phonoacetaldehyde (Pald) to acetaldehyde and orthophosphate (Fig. 1) [1]. Phosphona-
tase functions in the 2-aminoethylphosphonate (AEP) degradation pathway operative in
strains of bacteria adapted for the use of the natural phosphonate AEP as a source of car-
bon, phosphorus, and nitrogen [2–4]. Phosphonates, which contain a PAC bond, differ in
chemical stability from the more common phosphate monoesters, which possess a
PAOAC bond linkage [5–7]. The catalytic pathways for their hydrolysis also differ [8,9].
We were thus intrigued by the discovery that phosphonatase is a member of the haloacid
dehalogenase superfamily (HADSF) [10], a family predominantly comprised of
phosphatases.
The vast majority of the enzymes in the HAD family are phosphatases[11,12], followed
by ATPases [13,14] and phosphomutases [15]. The HAD phosphotransferases conserve a
core catalytic domain that positions an Asp residue to function in nucleophilic cataly-
sis[15,16], and a second Asp residue to function in general acid/base catalysis. The second
Asp protonates the leaving group oxygen in the first half of the reaction and deprotonates
the water or alcohol nucleophile for reaction with the phospho-enzyme intermediate in the
second half of the reaction. In addition, the catalytic domain provides a Lys/Arg residue
to bind the substrate phosphoryl group and position the Asp nucleophile, and two carbox-
2+
2+
ylate groups to assist in the binding of the Mg cofactor (Fig. 2). The Mg cofactor
forms a coordinate with the Asp nucleophile and the substrate phosphoryl group, thereby
orienting and shielding the negative charges in the ground state and transition state [17].
Fig. 1. The bacterial AEP degradation pathway.

396
S.D. Lahiri et al. / Bioorganic Chemistry 34 (2006) 394–409
Fig. 2. Schematic depicting the function of the conserved catalytic scaffold of the HAD phosphatases in
phosphoryl transfer from substrate (left panel) and in the transfer of the phosphoryl group from the
phosphoenzyme intermediate (E-P) to water (right panel).
The ATPase members of the HADSF, which do not require protonation of the ADP leav-
ing group, and which require a comparatively slow rate of hydrolysis of the phosphoen-
zyme intermediate (allowing a hydrolysis coupled conformational change), have become
specialized through the replacement of the Asp general acid/base by a Thr residue [18].
In this paper, we examine the extent to which the HAD phosphatase catalytic scaffold
has diverged to support catalysis of hydrolytic cleavage of the phosphonate PAC bond
by the enzyme phosphonatase.
Earlier work showed that catalysis of PAC bond cleavage of Pald by phosphonatase
proceeds via an iminium ion (Schiff base) intermediate formed between the enzyme
Lys53 and the substrate carbonyl group [8,10,19]. Thus, it was known that phosphona-
tase possesses the catalytic machinery for Schiff-base formation. The Schiff-base form-
ing reaction between a substrate carbonyl and an enzyme active-site Lys residue is a
common form of electrophilic catalysis employed by enzymes catalyzing aldol conden-
sations or transketolization reactions [20]. The X-ray structure of phosphonatase bound
with substrate or product analogs revealed that this HADSF member [17,21], like the
HADSF members phosphomutase b-PGM [22,23] and phosphatase phosphoserine
hydrolase [24–28], possesses a common phosphatase core domain. Moreover, the cata-
lytic scaffold of the core domain is highly conserved among these enzymes. The active
site is formed at the interface of the core domain and the cap domain, tethered by flex-
ible linkers. The cap domain associates with the core domain to enclose the active site
for catalysis and dissociates to allow product dissociation and substrate-binding
(Fig. 3).
The Schiff-base forming Lys53 of phosphonatase is located on the surface of the cap
domain, at the interface with the core domain. Inspection of the phosphonatase core
domain confirmed that the Asp general acid/base, conserved among the HAD phospha-
tases, is replaced with Ala14. Thus, whereas the phosphonatase cap domain acquired
the Lys for Schiff-base formation, the core domain lost the Asp for protonation of the
leaving group, and deprotonation of the water nucleophile. Remarkably, the catalytic
ꢀ
1
turnover rate for phosphonatase (15 s ) [19] is comparable to that of the ATPases and
the phosphatases reported to date (ꢁ10 sꢀ ) [29–36].
1

S.D. Lahiri et al. / Bioorganic Chemistry 34 (2006) 394–409
397
2
+
Fig. 3. Ribbon diagram of Bacillus cereus phosphonatase [17] depicting a monomer from the (A) enzyme-Mg
2+
complex in the ‘‘open conformation’’ and (B) the enzyme-Mg -tungstate complex in the ‘‘closed conformation’’.
The protein is differentially colored as follows: red, cap domain; blue, core domain; green, linkers; yellow, hinges;
2
+
gray, Lys53; cyan, Mg ; and orange, tungstate. (For interpretation of the references to color in this figure legend,
the reader is referred to the web version of this paper.)
Schiff-base formation, which proceeds via a carbinolamine intermediate, requires
deprotonation of the Lys53 and protonation of the Pald C@O [37]. Based on inspection
of the phosphonatase active site modeled with Pald (Fig. 4, the Pald position is based
on the position of the inhibitor vinyl sulfonate since the position of Pald in complex with
the Asp12Ala mutant is slightly shifted [21]), a hydrogen bond network existing between
Lys53, a water molecule, His56 and the Pald C@O was suggested as a possible means for
proton transfer between enzyme and substrate. Attack of the Asp12 nucleophile at the
phosphorus of the Schiff-base intermediate displaces Lys53N-ethyleneamine as the leaving
group (Fig. 5). Protonation of the leaving group is therefore not required in this step,
Fig. 4. The active site model generated by replacing vinyl sulfonate with Pald in the structure of wild-type
2
+
phosphonatase-Mg -vinyl sulfonate and then docking W111, W120 from the active site of the tungstate-bound
2+
wild-type enzyme (W405 is a Mg ligand).

398
S.D. Lahiri et al. / Bioorganic Chemistry 34 (2006) 394–409
Fig. 5. Schematic of the proposed phosphonatase catalytic mechanism.
consistent with presence of Ala in the general acid/base position of the HAD catalytic scaf-
fold. However, general base catalysis of the hydrolysis of the aspartylphosphate interme-
diate must therefore be performed by the Lys53N-ethylenamine as there is no other residue
properly positioned to fulfill this function.
Whereas, the mechanism of proton shuttling outlined in Fig. 5 is reasonable, it
remained unsupported by experimentation. Herein, we report the results of pH-rate profile
analysis of active-site mutants and the X-ray crystal structure of Lys53Arg phosphonatase
2+
complexed with Mg and phosphate, which evidence Schiff-base formation via proton
relay between the Lys53 and the Pald C@O. In addition, we report the structure of the
borohydride-reduced product Schiff-base intermediate, which evidences the role of the
Lys53N-ethyleneamine in activation of the water nucleophile for aspartylphosphate
hydrolysis. These results support the hypothesis that the phosphonatase cap domain is
responsible for the successful adaptation of the HAD phosphatase core domain for catal-
ysis of hydrolytic PAC bond cleavage in Pald.
2
. Materials and methods
2
.1. Materials
Escherichia coli JM109 competent cells were purchased from Promega and the expres-
sion vector pKK223-3 was purchased from Pharmacia Biotech. DNA manipulation

S.D. Lahiri et al. / Bioorganic Chemistry 34 (2006) 394–409
399
enzymes were purchased from New England Biolabs. The phosphonatase mutants were
generated using site-directed mutagenesis as previously described [21]. DNA sequencing
was carried out at the Center for Genetics in Medicine, University of New Mexico School
of Medicine. The wild-type and mutant phosphonatase proteins were purified as previous-
ly described [10]. Pald was synthesized according to the published procedure [38]. Yeast
alcohol dehydrogenase, NADH, and all buffers were purchased from Sigma.
2
.2. Enzyme assay
Phosphonatase activity was measured at 25 ꢁC using the coupled, spectrophotometric
assay described previously [21]. In this assay, acetaldehyde formation is coupled to NADH
oxidation via the alcohol dehydrogenase (ADH) catalyzed reaction. Each 1 ml reaction
+
mixture contained 50 mM K Hepes (pH 7.5), 5 mM MgCl , 0.15 mM NADH, 10 U
2
ADH, and various concentrations of Pald (0.5 to 5–10 K ). The reactions were monitored
m
ꢀ1
ꢀ1
at 340 nm (De = 6.2 mM cm ). The steady-state kinetic parameters, kcat and K , were
m
obtained by analyzing the initial velocity data using Eq. (1) and the KinetAsystI computer
program developed by Cleland [39]:
V
0
¼ V max½Sꢂ=ð½Sꢂ þ K
m
Þ
ð1Þ
where V is the initial velocity, K is the Michealis constant for the substrate, V
is the
0
m
max
maximal velocity and [S] is the concentration of the substrate. The kcat was calculated from
Eq. (2):
kcat ¼ V max=½Eꢂ
ð2Þ
The enzyme concentration [E] was determined using the Bradford method [40].
2
.3. pH rate profile determinations
All reactions were carried out at 25 ꢁC using a universal buffer containing 20 mM MES,
0 mM Hepes, 30 mM CHES, and 20 mM TAPS at a constant ionic strength attained
3
using 0.2 M NaCl. The initial reaction rates were measured using the coupled assay as
described above. The kcat and Km values were obtained using Eqs. (1) and (2). At low
pH (5.0–6.0) the background reaction of NADH was measured before initiating the phos-
phonatase reaction. The pH rate profile data were fitted to the Eq. (3).
log Y ¼ log½C=ð1 þ H=K
a
þ K
b
=HÞꢂ
ð3Þ
where Y is the kcat/K , H is the proton concentration of the reaction buffer, K and K are
m
a
b
the apparent ionization constants of the acidic and basic groups, and C is the constant val-
ue of Y where it does not change with the solution pH.
2
.4. Capture of the phosphonatase Schiff-base by borohydride reduction
Phosphonatase (25 lM) was incubated with 2 mM Pald in 100 mM Hepes (pH 8.0) con-
taining 5 mM MgCl and 1 mM NaBH for 10 min at 0 ꢁC. The procedure was repeated to
2
4
ensure complete inactivation of the enzyme (the first treatment caused 75% inactivation).
The protein was concentrated in 1 mM Hepes pH 7.5, 0.1 mM DTT, and 10 mM MgCl in
2
a Millipore Ultra-free concentrator to 15 mg/ml for crystallization.

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S.D. Lahiri et al. / Bioorganic Chemistry 34 (2006) 394–409
2
.5. Crystallization
Crystals of the reduced Schiff-base phosphonatase adduct were grown in solution con-
taining 0.1 M NaCl, 0.1 M Hepes, pH 7.5, and 1.6 M ammonium sulfate by the hanging-
drop vapor diffusion method. Crystals of the K53R mutant were grown in solution
containing 50 mM KH PO and 17% PEG 8000 by the hanging-drop vapor diffusion
2
4
method.
˚
The data sets were collected to 2.9 and 2.5 A, respectively, using a Rigaku RU-300 X-
ray generator equipped with an RAXIS-II area detector. Data collection for the K53R
phosphonatase crystals was carried out at ambient temperature. Data collection for the
reduced Schiff base-phosphonatase crystals was carried out at ꢀ180 ꢁC using 20% glycerol
as a cryoprotectant. The data collection statistics are summarized in Table 1. The DENZO
and SCALEPACK packages [41] were used for data indexing, reduction, and scaling. The
K53R phosphonatase crystals are rhombohedral (R3 space group) with unit-cell dimen-
2
sions a = 251.73, b = 251.73, c = 156.34. The reduced Schiff-base phosphonatase crystals
are monoclinic belonging to space group C(2) with unit-cell dimensions a = 159.82,
b = 64.02, c = 98.34, b = 123.517ꢁ. A Matthews’s coefficient calculation suggested five
and two monomers in the asymmetric units, respectively.
Table 1
Crystallographic data collection and refinement statistics
K53R
4
NaBH reduced protein
˚
Unit-cell dimensions (A)
a = b = 251.73
c = 156.34
a = 159.82
b = 64.02
c = 98.34
b = 123.517
C2
Space group
R32
1.54
˚
Wavelength (A)
1.54
˚
Resolution range (A)
100–2.9
368969/51994
99.9 (99.5)
9 (2)
12 (30)
10260
35598/4000
25.27/26.6
38
100–2.4
238424/28942
92.8 (54)
8 (2)
19 (32)
4082
17839/1945
23.0/27.2
27.9
Total/unique reflections
a
Completeness (%)
I/r (I)
b
R
merge (%)
Number of protein atoms/asymmetric unit
Number of reflections (working/free set)
c
d
R
work /Rfree (%)
˚
2
Average B-factors (A )
Amino acid residues
36
23
21
2
2
+
Mg
Sulfate/phosphate
Water
Bond length (A)
30
37
0.007
31
31
0.008
˚
Angles (ꢁ)
1.5
1.4
a
Statistics for the outermost resolution shell are shown in parentheses.
P
P
P
P
b
R
merge
=
hkl
i
j Ihkl,iꢀÆIhklæ Ij/ hkl
i
jIhkl,ij, where ÆIhklæ is the mean intensity of the multiple Ihkl,i
,
observations for symmetry related reflections.
c
P
P
R
R
work
=
=
hkljFobs ꢀ Fcalcj/ hkljFobsj.
P
P
d
free
hkl RTjFobs ꢀ Fcalcj/ hkljFobsj, where the test set T includes 10% of the data.

S.D. Lahiri et al. / Bioorganic Chemistry 34 (2006) 394–409
401
2
.6. Structure determination and refinement
The molecular replacement method was used to determine the phases for both struc-
tures using wild-type phosphonatase as the model (1FEZ.pdb) in the program AMoRE
[
‘
‘
42]. The lowest Rfactor and highest correlation coefficients were obtained using all
‘open’’ monomers to model the dimer of K53R phosphonatase and by using one
‘closed’’ monomer and one ‘‘open’’ monomer to model the dimer of the NaBH reduced
4
Schiff-base phosphonatase. The resulting model solutions were subsequently subjected to
rigid-body refinement in the CNS program suite [43], allowing independent movement of
the cap and the core domains. Iterative rounds of minimization and simulated annealing
using slow-cool torsional molecular dynamics followed by B-domain refinement and
manual rebuilding were used to refine the structure in CNS. For statistical cross-valida-
tion purposes (calculation of Rfree, [44]) 10% of the data was excluded from refinement.
Manual fitting was done using sigmaa weighted 2F ꢀ F and F ꢀ F electron density
o
c
o
c
maps [45] in the graphics program O [46]. Refinement, using a MLF target was per-
formed until Rfree ceased to decrease. Waters were added using the automated water-
picking program in CNS. The model was checked using a 2F ꢀ F composite-omit
o
c
map. The final model incorporated 257 out of 267 possible amino acids. Residues 1–4
at the N-terminus and residue 262–267 at the C-terminus were not visible in either struc-
ture in the electron-density map (also disordered in the wild-type model), and were omit-
ted from the final model.
For the K53R phosphonatase structure refinement the Lys53 from the wild-type model
was mutated to Ala for unbiased identification of the residue. In the later stages of refine-
ment, based on the density, this residue was replaced with Arg53. The active site also
contained strong density for one phosphate ion (from the crystallization conditions)
2
+
and a Mg cofactor from the protein solution. In the case of the NaBH reduced phos-
4
phonatase Schiff-base structure the electron density (contoured at 2r) connected to the
Schiff-base Lys53 was observed in the active site of one monomer only. Additional densi-
2
+
ties corresponding to a sulfate ion (from the crystallization conditions) and a Mg
cofactor, respectively, were present in the active site of both monomers. The final Rfactor
for the model was 27.5% (Rfree) and 25.9% (Rcryst) for the K53R structure and 26.8%
(
Rfree) and 22.7% (Rcryst) for the NaBH reduced structure. Analysis of the Ramachandran
4
plot, as defined by PROCHECK [47], showed a good final model for both structures.
Refinement and final model statistics are compiled in Table 1.
3
. Results and discussion
3
.1. Schiff-base formation: pH rate profile analysis
The proposed model for Schiff-base formation in phosphonatase catalysis is depicted in
Fig. 5. In this model, the proton that is carried into the active site by Lys53 (pK = 9.3;
a
[
48]) upon cap closure becomes delocalized by His56 and ultimately transferred to the Pald
C@O via a hydrogen bond network that incorporates water120. The active configuration
of the free enzyme should thus include the protonated form of Lys53 and the neutral
(
unprotonated) form of His56. A priori, it could be argued that the logkcat/Km versus
pH profile for phosphonatase catalysis (first reported in reference [18]; and again in
Fig. 6A) reflects loss of activity at low pH due to the protonation of His56 and loss of

402
S.D. Lahiri et al. / Bioorganic Chemistry 34 (2006) 394–409
1
A 2
B 0
C
1
0
1
0
-1
-2
-3
-4
-
-
-
-
1
2
3
4
-
-
-
2
3
5
6
7
8
9
10
5
6
7
8
9
10
5
6
7
8
9
10
pH
pH
pH
D -1
-1
-2
-3
-4
E
-
-
-
2
3
4
5
-
-
-5
6 5
6
7
8
9
10
-65
6
7
8
9
10
pH
Fig. 6. The kcat/K
m
pH profile for catalysis of Pald hydrolysis by wild-type (A) phosphonatase and the Cys22Ser
(apparent pKa1 = 6.2 and apparent pKa2 = 8.1) (B), Cys22Ser/Tyr128Phe (apparent pKa1 = 6.4 and apparent
pKa2 = 8.5) (C), His56Gln (apparent pKa1 = 6.6 and apparent pKa2 = 8.2) (D), and His56Ala (apparent
pKa1 = 5.3 and apparent pKa2 = 8.7) (E) active-site mutants.
activity at high pH due to the deprotonation of the Lys53. However, the log kcat/K versus
m
pH profiles measured for phosphonatase active site mutants (described below) indicate
that this is a simplistic interpretation.
A more comprehensive view of the pH dependence of the kcat/Km is derived from a
composite of the ionization of several groups (substrate and enzyme). Therefore, the
apparent pK values calculated from these data using Eq. (3) (reported in the legend to
a
Fig. 6) cannot be assigned to a specific acid group. On the other hand, we might expect
that the replacement of an essential enzyme acid or base residue by site-directed mutagen-
esis would noticeably perturb the log kcat/K versus pH profile. Whereas Table 2 provides
m
the kcat/K values for wild-type and mutant phosphonatase measured at a single pH value,
m
the pH profiles shown in Fig. 6, provide a comparison over the entire pH range. The kinet-
ic constants measured for the Cys22Ser, Cys22Ser/Tyr128Phe, His56Gln, and His56Ala
active-site mutants at pH 7.5 indicate that the core domain active site residues Cys22
and Tyr128 contribute to the function of the catalytic site but do not appear, based on
only a 10-fold reduction in kcat and kcat/Km values to participate in acid-base catalysis
(Table 2). The log kcat/Km versus pH profiles of the Cys22Ser and Cys22Ser/Tyr128Phe
mutants shown in Figs. 6B and C, respectively, are similar, but not identical, to that of
the wild-type enzyme.
The replacement of the cap domain His56 with Ala or Gln impairs catalytic efficiency
to a greater extent than does the replacement of the core-domain active-site residues
Cys22 and Tyr128 (Table 2). The kcat of the His56Ala mutant is 200-fold less than that

S.D. Lahiri et al. / Bioorganic Chemistry 34 (2006) 394–409
403
Table 2
The steady-state kinetic constants for wild-type and mutant phosphonatase measured at pH 7.5 and 25 ꢁC
a
ꢀ1
ꢀ1
ꢀ1
Enzyme
kcat (s
)
K
m
(lM)
k
cat/K
m
(s lM
)
b
1
ꢀ1
ꢀ2
ꢀ2
ꢀ4
ꢀ3
Wild-type
C22S
1.5 ± 0.1 · 10
2.26 ± 0.09
2.11 ± 0.03
33 ± 2
33 ± 5
33 ± 2
145 ± 6
170 ± 10
5.0 · 10
6.9 · 10
6.4 · 10
5.2 · 10
2.0 · 10
b
b
Y128F/C22S
b
ꢀ2
H56A
7.5 ± 0.1 · 10
3.6 ± 0.1 · 10
ꢀ
1
H56O
a
2
Reactions contained Pald, 10 mM MgCl , 0.15 mM NADH, and 5 U alcohol dehydrogenase in 50 mM
+
K Hepes (pH 7.5, 25 ꢁC). The concentration of phosphonatase used was in the range of 0.02–2 lM, depending on
the mutant studied.
b
Reported in Morais et al. [21].
of the wild-type enzyme and the kcat/Km is 1000-fold less. The His56Gln mutant is also
less active than the wild-type phosphonatase: the kcat is decreased 42-fold and the
kcat/Km is decreased 250-fold. The log kcat/Km versus pH profiles of the His56Gln vs
His56Ala mutants are shown in Figs. 6D and E, respectively. The pH-rate profile of
the His56Ala mutant differs to the greatest extent from that of the wild-type phospho-
natase. In particular, the drop in activity below pH 7 observed in the wild-type enzyme
and the Cys22Ser, Cys22SerTyr128Phe, and His56Gln mutants, is minimal in the
His56Ala mutant. The suppression of the response to increasing acidity might be attrib-
uted to the loss of that function carried out by His56, which can be maintained by Gln
but not by Ala.
In Fig. 7, we illustrate the hydrogen-bond network that surrounds the His56 imidazole
side chain in the substrate bound, catalytically active form of the enzyme as suggested by
the X-ray crystallographic data. Substitution of His56 with Gln might not necessarily
interrupt this hydrogen bond network because the functions of the His imidazole nitrogens
could conceivably be assumed by the Gln side chain carbonyl and amide nitrogen, respec-
tively. On the other hand, the methyl side chain of the substituted Ala in the His56Ala
mutant would certainly not support a hydrogen bond network. To the extent that this
putative network is important for catalysis in the wild-type phosphonatase, we anticipate
a loss in catalytic activity upon its disruption.
Fig. 7. Schematic representation of the hydrogen-bond network in wild-type phosphonatase and in the His56Gln
and His56Ala mutants.

404
S.D. Lahiri et al. / Bioorganic Chemistry 34 (2006) 394–409
3
complexed with phosphate and Mg
.2. Impact of Lys53 substitution on cap closure: structure of the Lys53Arg mutant
2
+
The importance of delocalization of the proton on Lys53 in catalysis was probed by
substituting Lys53 with Arg and determining whether or not the substitution destabilizes
the cap closed conformation required for catalytic turnover. As illustrated in Fig. 3 of the
Introduction, phosphonatase cycles between an open conformation that allows substrate
to bind and product to dissociate, and a closed conformation that is required for catalysis.
When the cap closes over the active site, Lys53 enters the catalytic site and engages in the
hydrogen bond network represented in Figs. 5 and 7. Previous studies have shown that
bound substrate or product ligands stabilize the cap closed conformation by an electro-
statically driven induced-fit mechanism [48]. Crystallographic analyses of liganded and
unliganded phosphonatase have captured the open conformation in the apoenzyme but
only the closed conformation in the liganded enzyme [17,21]. If the delocalization of the
Lys53 proton is an integral part of the catalytically active conformation then we might
expect that the substitution of Lys53 with Arg will disrupt the electrostatic balance and
thus destabilize the cap closed conformation. Accordingly, we carried out the structure
determination of the phosphonatase Lys53Arg mutant crystallized in the presence of
2
+
the product phosphate and the Mg cofactor. As might be expected, substitution of
the Schiff-base forming Lys with Arg diminished catalytic activity to background levels
in the enzymatic assay.
Crystals of Lys53Arg were non-isomorphous with the wild-type crystals. There are five
monomers (2 1/2 dimers) in the asymmetric unit and the physiological dimer interface is
retained (one dimer is formed by crystallographic symmetry). The overall fold of the
K53R mutant is no different from that of the wild-type enzyme, which is characterized
by a distinct a-helical cap domain and a/b core domain. NCS restraints were applied dur-
ing crystallographic refinement, thus all monomers are the same in their overall conforma-
˚
tion (rmsd of 0.07 A over all Ca atoms). Fig. 8A depicts three structures: wild-type open,
wild-type closed and the K53R mutant wherein the core domains are superimposed to
show the relative dispositions of the cap and core domains. The calculated values for
the rmsd between the K53R monomer Ca positions, and those of the wild-type open
˚
and closed conformation monomers, are 0.77 and 1.02 A, respectively.
The only significant difference in the wild-type and mutant structures is the angle by
which the cap and core domains are disposed relative to one another via rigid-body move-
ments. Specifically, the angle of rotation through the hinge axis (the linker connecting the
cap and the core domain) is 22.35ꢁ between the wild-type open and closed monomers,
while it is 42.88ꢁ between the K53R and wild-type closed monomer (calculated with DYN-
DOM [49]). There is a slight change in the relative orientation of helices 2 and 3 of the cap
domain. Because these two helices are connected by the ‘‘substrate-specificity loop’’ (res-
idues 51–54) [50], the disposition of the nearby residues that contribute to the domain–do-
main interface, and that contribute functional groups to the catalytic site, were examined.
Importantly, mutation of Lys53 to Arg results in only a slight structural rearrangement of
the residues in the loop region, and all interactions between the protein and the ligands
2
+
(
Mg and phosphate) are maintained in the K53R mutant as compared to the wild-type
enzyme. Thus, although the open conformation observed with the K53R mutant is more
open than that observed for the wild-type phosphonatase there is no evidence of a
gross change in the active site structure. The loss of activity stems from the inability of

S.D. Lahiri et al. / Bioorganic Chemistry 34 (2006) 394–409
405
Fig. 8. Superposition of the mutant and modified Ca traces of phosphonatase with wild-type. (A) K53R Ca trace
yellow) with the open (red) and closed (blue) wild-type monomers. (B) The NaBH reduced phosphonatase
(
4
backbone (green) with the open (red) and closed (blue) wild-type monomers. (For interpretation of the references
to color in this figure legend, the reader is referred to the web version of this paper.)
the guanidinium group of the Arg residue in the K53R mutant to participate in Schiff-base
formation, and not from large alterations in the enzyme conformation.
The fact that the phosphate bound K53R phosphonatase is observed in the cap-open
conformation in the crystal structure is of special interest. In wild-type phosphonatase,
the binding of anionic ligands (e.g., phosphate, sulfate, vinyl sulfate or tungstate) which
mimic the substrate cause the monomer to attain a closed conformation in both the crystal
[
11,17,21] and in solution [48]. Thus, the open conformation observed with the K53R
mutant suggests that the replacement of the Lys53 primary ammonium group with the
Arg guanidinium group interferes with the ligand induced domain closure. This finding
is consistent with the predicted dependence of cap closure and catalytic turnover on the
delocalization of the Lys53 charge via the hydrogen-bond network depicted in Fig. 5.
3
.3. Structure of the Lys53ethylenamine adduct
The hydrolysis of the bi-covalent Lys53ethylenamine/Asp12 aspartylphosphate inter-
mediate to product requires that the Lys53ethylenamine undergoes protonation at the
C(2) position (Fig. 5). In parallel, a water molecule must be activated by deprotonation
for in-line attack at the phosphorus of the aspartylphosphate. The structure of the product
Schiff-base was examined to identify the putative acid/base residues involved in these
events. In the presence of Pald, NaBH4 inactivates phosphonatase by reducing the
Lys53ethylenamine group of the reaction intermediate accumulated at steady-state (the
product Schiff-base intermediate), thereby preventing its hydrolysis [7]. Structure determi-
nation of crystals of the NaBH reduced phosphonatase indeed revealed the Lys53ethyl
4
residue. Fortuitously, the crystallization medium (which contained 1.6 M ammonium sul-
fate but no added phosphate) allowed the binding of a sulfate ion to the phosphate-bind-
ing subsite of the active site, affording a view of an analogue of the bi-covalent

406
S.D. Lahiri et al. / Bioorganic Chemistry 34 (2006) 394–409
Lys53ethylenamine/Asp12aspartylphosphate intermediate. This provided for the assess-
ment of the overall conformational state of this intermediate complex as well as identifi-
cation of the active site residues proximal to the Lys53ethylenamine and
Asp12aspartylphosphate groups. In addition, the structure revealed ordered solvent mol-
ecules positioned to participate in the hydrolysis of the Lys53ethylenamine and
Asp12aspartylphosphate groups, and thus an overall picture of the active site poised for
the conversion of the bi-covalent intermediate to the acetaldehyde and phosphate
products.
2
+
˚
The 2.5 A structure of the Lys53ethylphosphonatase-sulfate–Mg complex revealed a
single physiological dimer in each asymmetric unit, with one subunit assuming the cap-
open conformation and the other subunit the cap-closed conformation. The closed confor-
mation does not differ significantly from that previously determined for the unliganded
˚
closed enzyme, with an rmsd of 0.16 A. The open monomer, on the other hand, is more
open than previously observed for the open conformation of unliganded phosphonatase
˚
(Fig. 8B), with an overall rmsd of 1.24 A. The interactions at the dimer interface are
unchanged from those of the wild-type mixed dimer.
2
+
The active site of the open monomer contained the N-ethylLys53, the Mg cofactor
and sulfate ion (from the crystallization conditions) (Fig. 8B). The position of the sulfate
(
which mimics phosphate) is identical to that of the tungstate bound in wild-type phospho-
natase (1FEZ.pdb) and the phosphate bound in the K53R phosphonatase (vide supra).
Thus, the sulfate interactions can be interpreted as mimicking the interaction with the
product phosphate in the reaction pathway. In addition to the sulfate electron density,
there is a strong electron density connected to the K53 (Fig. 9A) that can be fitted well
2
+
to N-ethyl lysine. The residues forming ligands to the Mg are the same as in the wild-
type phosphonatase and as described for the K53R mutant structure.
The ethyl moiety of the N-ethyl-Lys53 is surrounded by a hydrophobic environment
provided by the active-site residues Tyr128, Cys22, and Met49. It is interesting to note that
in the open conformation of the unliganded enzyme, Met49 projects away from the active
site but in the closed monomer it is positioned inside, to shield the catalytic site from sol-
vent. In the cap-open structure of the N-ethyl-Lys53 phosphonatase, the Met49 is also
positioned inside the active site where it forms hydrophobic contacts with the ethyl moiety.
˚
In addition, His56 (positioned within 6.8 A of the ethyl-lysine group, see Fig. 9B) forms a
hydrogen bond to a water molecule, W123, near the C(2) of the ethyl group. This water
molecule is also within hydrogen bonding distance of the Met49 sulfur atom and the back-
bone carbonyl of Leu51 (one of the residues of the linker connecting cap to core). Notably,
positioned in-line to the bound sulfate and in between the sulfate and C(1) of the ethyl
group of ethyl-Lys53 is the water molecule, W138. This water molecule is within hydrogen
˚
bonding distance of the sulfate and close to the terminal ethyl carbon (3.8 A).
The structure of the N-ethyl-Lys53 phosphonatase sheds light on the mode of hydroly-
sis of the product Schiff-base generated from the second partial reaction of Pald hydroly-
sis. After CAP bond cleavage the C(2) of the displaced Lys53ethlyenamine must be
protonated to form the Lys53ethyliminium ion prior to hydrolysis to acetaldehyde
(Fig. 5). Therefore, this step requires two catalysts: a general acid to protonate the C(2)
carbon and a general base to deprotonate a water for attack. Within the active-site archi-
tecture there are two possibilities. Tyr 128 can act as a general acid to protonate at C(2)
and the same Tyr 128, now ionized, may act as a general base and activate a water for
addition to the C(1) of the Lys53ethyliminium ion. Alternatively, the Lys53ethylenamine

S.D. Lahiri et al. / Bioorganic Chemistry 34 (2006) 394–409
407
Fig. 9. The active site of NaBH
modified Lys53. The active site residues (yellow) and ligands sulfate (orange) and cofactor Mg (cyan) are
depicted as ball-and -stick. The 2F electron density map (contoured at 1.5r) is shown as blue cages. (B)
4
reduced phosphonatase. (A) Depicted with the electron density surrounding the
2
+
o
ꢀ F
c
˚
Chemdraw depiction of N-ethy-Lys53 and surrounding active-site residues (distances in A). (For interpretation of
the references to color in this figure legend, the reader is referred to the web version of this paper.)
itself removes a proton from water activating it for attack at the phosphoaspartate. The
resulting Lys53ethyliminium might be hydrolyzed by the nucleophilic attack of a water
molecule activated by the His56 nearby.
Through examination of the neighboring interactions of the N-ethyl-Lys53 adduct in
˚
the reduced structure, it can be seen that the position of Tyr128 is nearly 7 A away from
the ethyl group of the Lys53N-ethyl moiety making it impossible for this residue to
participate in the Lys53ethyliminium hydrolysis. Substitution of Tyr128 with Phe does
not significantly alter catalytic efficiency (Table 2) nor the pH dependence of catalytic

408
S.D. Lahiri et al. / Bioorganic Chemistry 34 (2006) 394–409
efficiency (Fig. 6C). In fact, there is no enzymic residue in the correct position to deprot-
onate water 138 for hydrolysis of the phosphoenzyme.
Thus, it is most likely that the Lys53ethylenamine plays the key role in proton transfer
required for the hydrolysis steps. Accordingly, the well-positioned enamine would abstract
a proton from the water molecule oriented for in-line attack on the phosphorus of the
phosphoaspartate. In the subsequent step of the reaction, water123, activated by His56,
attacks the iminum carbon, thus initiating the release of acetaldehyde.
4
. Conclusions
The presented structural results, together with pH-rate profile analysis, evidence the
dependence of cap closure and catalytic turnover on the delocalization of the Lys53 charge
via a hydrogen bond network between Lys53 and the substrate carbonyl, via a water mol-
ecule and His56. Additionally, based on both biochemical and structural data, our work-
ing hypothesis is that the Lys53ethlyenamine acts as the general base for
Asp12aspartylphosphate hydrolysis. Such a mechanism would connect the reactivity of
the Schiff-base subsite with that of the phosphoenzyme and thereby render the core
domain Asp general acid/base of the HAD phosphotransferases obsolete. Indeed, in phos-
phonatase Ala replaces the Asp. We suggest that it is the specialization observed in the cap
domain that is responsible for the successful adaptation of the HAD phosphatase core
domain for catalysis of hydrolytic PAC bond cleavage.
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