Differential catalytic promiscuity of the alkaline phosphatase superfamily bimetallo core reveals mechanistic features underlying enzyme evolution

Article abstract of DOI:10.1074/jbc.M117.788240

Members of enzyme superfamilies specialize in different reactions but often exhibit catalytic promiscuity for one another's reactions, consistent with catalytic promiscuity as an important driver in the evolution of new enzymes. Wanting to understand how

Full text of DOI:10.1074/jbc.M117.788240

JBC Papers in Press. Published on October 25, 2017 as Manuscript M117.788240
The latest version is at http://www.jbc.org/cgi/doi/10.1074/jbc.M117.788240
Differential Catalytic Promiscuity of the Alkaline Phosphatase Superfamily
Bimetallo Core Reveals Mechanistic Features Underlying Enzyme
Evolution
Fanny Sunden1, Ishraq AlSadhan1, Artem Lyubimov2,3,4,5,6, Tzanko Doukov7,
Jeffrey Swan1 and Daniel Herschlag1,8*
1Department of Biochemistry, Beckman Center, Stanford University, Stanford, CA
94305 USA; 2Departments of Molecular and Cellular Physiology; 3Neurology and
Neurological Science; 4Structural Biology; 5Photon Science; 6Howard Hughes
Medical Institute, Stanford University, Stanford, CA 94305, USA;
7Macromolecular Crystallographic Group, The Stanford Synchrotron Radiation
Lightsource, National Accelerator Laboratory, Stanford University, Stanford, CA
94309; 8Departments of Chemical Engineering and Chemistry, Stanford
University, Stanford, California 94305, USA; 9Stanford ChEM-H (Chemistry,
Engineering, and Medicine for Human Health), Stanford University, Stanford,
California 94305, USA
*Corresponding author: herschla@stanford.edu
1
Copyright 2017 by The American Society for Biochemistry and Molecular Biology, Inc.

ABSTRACT
substrates through metabolic pathways and
permitting exquisite biological regulation
(1,2). But how did these remarkably efficient
enzymes arise? What are the mechanistic and
structural properties that determine the
catalytic proficiencies of a family of proteins
and thus the catalytic landscape that evolution
has traversed?
Members of enzyme superfamilies specialize
in different reactions but often exhibit
catalytic promiscuity for one another’s
reactions, consistent with catalytic promiscuity
as an important driver in the evolution of new
enzymes. Wanting to understand how catalytic
promiscuity and other factors may influence
evolution across a superfamily, we turned to
the well-studied Alkaline Phosphatase (AP)
superfamily, comparing three of its
members—two evolutionarily distinct
phosphatases and a phosphodiesterase. We
mutated distinguishing active-site residues to
generate enzymes that had a common Zn2+
bimetallo core, but little sequence similarity
and different auxiliary domains. We then
tested the catalytic capabilities of these pruned
enzymes with a series of substrates. A
substantial rate enhancement of ~1011-fold for
both phosphate mono- and diester hydrolysis
by each enzyme indicated that the Zn2+
bimetallo core is an effective mono/di-esterase
generalist and that the bimetallo cores were
not evolutionarily tuned to prefer their cognate
reactions. In contrast, our pruned enzymes
were ineffective sulfatases, and this limited
promiscuity may have provided a driving
force for founding the distinct one-metal-ion
branch that contains all known AP
Catalytic promiscuity, the ability of an
enzyme to carry out reactions other than its
evolutionary optimized reaction, is expected to
strongly bias the emergence of new enzymes,
as it can provide an evolutionary head start
following
gene
duplication
(3,4).
Correspondingly, the catalytic promiscuity of
modern day enzymes and their conserved
elements may be evolutionary vestiges and
may provide clues to catalytic capabilities and
mechanistic features that have influenced
evolutionary history and insights for the
engineering of new enzymes (4-13).
We can identify enzymes that are
members of superfamilies—i.e., enzymes with
similar overall folds and common underlying
catalytic features—and these enzymes very
likely emerged from a common evolutionary
precursor (3,8,10,11,14,15). Indeed, extant
superfamily members that catalyze different
reactions often exhibit cross promiscuity,
consistent with an evolutionary role for
promiscuity (3,11,16). In some cases, for
enzymes that have evolved subsequent to the
last universal common ancestor (LUCA) of
current cellular life, reconstructions can
superfamily sulfatases. Finally, our pruned
enzymes exhibited 107-108 fold
phosphotriesterase rate enhancements, despite
absence of such enzymes within the AP
superfamily. We speculate that the
superfamily active site architecture involved in
nucleophile positioning prevents
provide
information
about
potential
accommodation of the additional triester
substituent. Overall, we suggest that catalytic
promiscuity and the ease or difficulty of
remodeling and building onto existing protein
scaffolds have greatly influenced the course of
enzyme evolution and will provide lessons for
engineering new enzymes.
evolutionary pathways (17-21), whereas in
other cases reconstructions are stymied as
superfamily members were already present in
LUCA or emerged very early after the
common ancestor (22) (P.C. Babbitt, personal
communication).
In addition to our ability to create
models for the sequences of evolutionary
precursors (17-21), we would like to know,
from a mechanistic standpoint, why certain
enzymes have evolved from one another, and
conversely, why others did not. As noted
INTRODUCTION
Enzymes catalyze biological reactions
with enormous rate enhancements—up to
1029-fold—and do so with high specificity,
thereby ensuring rapid and controlled flux of
2

above, catalytic promiscuity is one factor that
has likely played an important role, and we
gain new insights into evolutionary pathways
from the determinations of catalytic
promiscuities herein. Nevertheless, catalytic
promiscuity does not ensure the development
and fixation of a new activity, even if it can
provide a selective advantage; in addition to
the inevitable role of chance in evolution,
mutational paths, to be followed, need to avoid
major sustained fitness dips that would
prevent or greatly hinder optimization.
predominantly catalyzing sulfatase reactions
and the other with enzymes containing the
canonical alkaline phosphatase bimetallo
center (38). We focused on the latter group,
which possesses two metal ions coordinated
by conserved residues in the active site and a
universally conserved serine or threonine
nucleophile (Fig. 1). This group includes
phosphatases,
phosphate
diesterases,
phosphomutases, among other enzymes (Chart
1)(30-34,39). The enzymes studied here typify
a cross-section of the structural diversity seen
across the AP superfamily bimetallo branch
while maintaining Zn2+ ions in their bimetallo
cores, which facilitates the comparisons herein
(Fig. 2). EcAP and Chryseobacterium
meningosepticum Alkaline Phosphatase PafA
(PafA) are both phosphate monoesterases
(Table 1A), but are evolutionarily distant as
suggested by a low sequence identity (8.9%;
Supplemental Fig. 1, (40)) and distinct
inserted elements (Fig. 2A & B)(34,40).
Thus, information about the physical
and mechanistic features that determine
catalytic efficiency and catalytic promiscuity
will help us understand how enzymes have
emerged throughout evolution and the
evolutionary potential and evolvability of
enzymes. Ultimately these insights may be
harnessed to engineer new, beneficial enzymes
((4-13,23,24), and references herein (23) e.g.
(25-27)).
Xanthomonas
citri
Nucleotide
Pyrophosphatase/Phosphodiesterase (NPP) has
a subset of active site and scaffold features in
common with PafA but is evolutionary distant
from PafA (10% sequence identity;
Supplemental Fig. 1, (40)) and is a phosphate
diesterase rather than a monoesterase (Table
1A), related to a large set of AP superfamily
diesterases (Fig. 2A & B)(34,40-42).
We have taken advantage of the well-
established
functional,
structural,
and
evolutionary connections within the Alkaline
Phosphatase (AP) superfamily to relate the
catalytic promiscuity of the superfamily’s Zn2+
bimetallo core to the elaboration of enzyme
classes across this superfamily (6,28-37). To
ensure that our results would be representative
rather than idiosyncratic we compared three
Alkaline Phosphatase (AP) superfamily
members, mutating the distinguishing active
site residues to create mutant enzymes with a
common Zn2+ bimetallo core but pervasive
sequence differences and distinct auxiliary
domains. Determination of the catalysis of
these three ‘pruned’ enzymes across a series of
reaction classes and consideration of catalytic
and structural features allowed us to relate
intrinsic catalytic prowess of the Zn2+
bimetallo core to evolutionary choices made
across this superfamily.
The active site residues common to
these enzymes reside in analogous positions
within conserved secondary structure elements
(Fig. 2A, white & gray), and these secondary
structure elements comprise highly similar
tertiary
gray)(30,31,33,34,38-40,43).
enzymes have
structures
(Fig.
All
2B,
three
a
serine or threonine
nucleophile, two Zn2+ ions, and identical Zn2+
ligands (Figs. 2A & C, black). Mechanistic
similarities are also readily apparent. The
conserved seryl/threonyl residue is a ligand of
Zn2 and serves as a nucleophile that attacks the
phosphorus atom of the phosphate monoester
or diester substrate; the leaving group oxygen
atom, which develops negative charge in the
transition state, is stabilized by Zn1; and one of
the oxygen atoms of the transferred
phosphoryl group situates between the Zn2+
RESULTS
The Bimetallo Branch of the AP Superfamily
The AP superfamily subdivides into
two main subgroups, one with enzymes
containing
a
single metal ion and
3

ions, presumably making close electrostatic
interactions that stabilize the trigonal
bipyramidal transition state (Fig. 2D)(44).
Superfamily Members. We first asked how
effective the AP superfamily Zn2+ bimetallo
center is in catalyzing phosphate monoester
and diester hydrolysis when embedded in each
of the three superfamily scaffolds after
truncation of the active site side chains
particular to each active site (Fig. 2C, colored
residues interacting with the substrate non-
bridging oxygen atoms facing away from the
Zn2+ bimetallo center). It has been suggested
that the scaffold tunes the Zn2+ bimetallo site
to favor monoesterase over diesterase activity
(34), and the ample sequence differences
throughout these scaffolds could cause these
or other catalytic differences. Further, while it
is reasonable that the scaffolds make the same
functional interactions in phosphate monoester
and diester transition states, the charges of
reactants and transition state structures for
reactions vary, so there is no reason a priori to
expect the same strength for each catalytic
interaction and same catalytic contribution; in
other words, we cannot predict the relative
rate constants for reactions with different
substrates, even ones making the same atom-
to-atom interactions.
These structural and mechanistic
similarities contrast with local and global
differences (Fig. 2). Figure 2C shows the
residues that sit opposite to the bimetallo
center
(colored
residues).
The
two
monoesterases (EcAP and PafA) donate
multiple hydrogen bonds to each of the two
oxygen atoms on the transferred phosphoryl
group that face away from the Zn2+ bimetallo
site, interactions that are unique to each
enzyme and suggestive of evolution along
distinct pathways within the AP superfamily
(40,41). The diesterase (NPP) makes
interactions with the nonesterified outward-
facing phosphoryl oxygen atom that are the
same as for the PafA monoesterase, apparently
reflecting a shared evolutionary origin (Fig.
2C: NPP, N111 (magenta); PafA, N100
(green), (37,40,41) whereas the other outward
facing phosphoryl oxygen makes different
interactions, in its esterified form with an NPP
hydrophobic pocket and in its nonesterified
form with a PafA hydrogen bond network
(Fig. 2B & 2C: NPP, magenta; PafA, green)
(34,37,45).
We created “pruned” versions of each
enzyme by subtractive mutagenesis, denoted
throughout with asterisks (i.e., EcAP*, NPP*,
PafA*), by mutating the side chains directly
contacting and within 4 Å of the substrate
non-bridging oxygen atoms and, for NPP, side
chains that interact with the methyl substituent
of me-pNPP used herein (Fig. 2C, color-coded
side chains; Chart 2)(45). Subtractive
mutagenesis was used in all cases except for
mutations of the nucleophile, T79S and T90S
for PafA and NPP, respectively, so that all
three pruned enzymes contained the same
nucleophile. The pruned constructs had
mutations as follows: EcAP*: D101A, R166S,
D153A, E322A, K328A; NPP*: T90S
(nucleophile), F91A, N111A, L123A, Y205A;
and PafA*: T79S (nucleophile), N100A,
K162A, R164A (Table 1B). Prior studies
showed that the only non-alanine mutant,
R166S, gives similar kinetics as the
corresponding alanine mutation (36), and for
each enzyme, we created at least one
additional mutated version to control for
Mechanistic studies have shown, as
expected, that mutations of groups making
specialized interactions with a phosphoryl
oxygen atom (in the monoesterases) or with
the ester group attached to the transferred
phosphoryl group (in the diesterase)
preferentially lower activity of the cognate
reaction and thereby decrease specificity
(6,36,40,45). In addition to the local active site
differences
conservation
and
within
very
low
their
sequence
structurally
conserved cores (sequence identities: 6.0% for
EcAP/NPP, 8.9% for EcAP/PafA, and 10% for
NPP/PafA for the structurally conserved cores;
Supplemental Fig. 1), these enzymes contain
different sets of added helices and sheets (Fig.
2A & 2B, color coded regions)(40).
The Phosphate Monoesterase and Diesterase
Activities of the Zn2+ Bimetallo Scaffold of AP
4

unintended consequences from the specific
mutations made. For E322, mutation to
tyrosine was employed as prior studies
showed equivalent effects for it and alanine
and as the homologous residue in NPP is a
tyrosine (Y205)(6,46). The ratio between the
rate constants for monoester and diester
hydrolysis of each enzyme set fell within 3-
fold of one another (Table 2), providing
evidence against unintended or idiosyncratic
effects.
Figure 4A compares the phosphate
monoesterase activity for the WT and pruned
versions of each enzyme (gray and black bars,
respectively). As expected, the pruned
monoesterases (EcAP* & PafA*) are more
diminished in monoester activity than the
pruned diesterase (NPP*). Intriguingly, the
monoesterase activities of the three pruned
enzymes were within 5-fold.
Figure 4B shows the corresponding
comparisons for phosphate diesterase activity.
As expected, converse effects on the two
reactions are observed—i.e., the diesterase
pruned construct (NPP*) gave the largest
effect. Remarkably, as for the monoesterase
reaction, all three enzymes had very similar
diesterase activity, again within a 5-fold range.
We expected, most simply, that the
bimetallo cores would not rearrange in the
mutated enzymes, as numerous EcAP variants
with mutations in these active site residues
exhibit no measureable change in the
bimetallo center or in more remote parts of the
structure (6,36,47-50). Nevertheless, as we
mutated
four
or
five
side
chains
The phosphate monoesterase and
diesterase results are summarized together in
Figure 4C as the ratios of these activities.
There is an enormous difference in the
substrate preferences of the WT enzymes of
~12 orders of magnitude, and the intrinsic
discrimination is greater still (i.e., the
chemical step is not rate-limiting in all
cases)(Fig. 4C, gray bars)(40,51). In contrast,
simultaneously in each active site, we tested
for structural rearrangements.
For each enzyme, its wild-type (WT)
and pruned form gave CD spectra that were
identical, within error (Supplemental Fig. 2).
To obtain a more detailed view of possible
structural rearrangements, crystals of PafA*
and EcAP* were obtained and their structures
for
the
pruned
constructs
the
were determined to 2.03 and 2.45
Å
monoesterase:diesterase ratios are uniform, all
within 2-fold (Fig. 4C, black bars (near axis)).
Further, the pruned enzymes provide
substantial rate enhancements of ~1010–1011
fold rate for the two reactions, indicating that
catalysis from the bimetallo scaffold rivals or
exceeds that of many fully evolved enzymes
(Fig. 5)(2-4).
resolution, respectively (Tables 3 and 4,
Supplemental Fig. 3). The overall structures of
WT EcAP and EcAP* were superimposed
with a backbone RMSD of 0.42 Å over 801
atoms (Fig. 3A), and WT PafA and PafA*
were superimposed with a backbone RMSD of
0.18 Å over 449 atoms (Fig. 3B). The
conserved active site Zn2+ ions and ligands
were identical for the two sets of wild-type
and pruned enzymes (Fig. 3C), with the only
exception being the serine nucleophile of
PafA* (T79S), which was not coordinated to
Zn2 and appeared to be interacting with
another ion in the active site (Supplemental
Fig. 3). This difference presumably arose
because the PafA* crystals were grown at pH
4.3, a pH at which the enzyme is not active
and the serine residue is likely neutral (Fig.
3C; data not shown; see also ref (51)).
The similarity of the catalytic ratios
for pruned enzymes derived from two
different monoesterases and
a diesterase
strongly suggests that the bimetallo site is
functionally conserved across the AP
superfamily, despite the very low sequence
identity of the surrounding scaffold and
despite the different additional structural
elements (Fig. 2A & B)(33,39,40). Thus,
specialization for the individual cognate
reactions arises from the residues that have
been pruned from each construct, and
evolution has not independently tuned the
5

common Zn2+ bimetallo center to favor
phosphate monoester or diester hydrolysis.
Most pertinent for understanding catalysis and
evolution in the AP superfamily, the Zn2+
bimetallo center appears to be a potent and
general enzyme, as elaborated in the next
section and in the Discussion.
from the bimetallo scaffold would provide a
considerable head start toward the evolution of
fully optimized monoesterases and diesterases
or
allow
significant
catalysis
while
progressing from one activity to the other (Fig.
5)(2-4). The observation of different
interactions in the EcAP and PafA
monoesterase active sites (Fig. 2C, colored
residues) indicates an evolutionary divergence,
perhaps to provide distinct catalytic properties
best suited to growth conditions of each parent
organism (40,42,55).
Enzymatic Activity of the Zn2+ Bimetallo
Scaffold as a Potential Evolutionary Driver
Across the AP Superfamily. Interestingly, the
monoesterase:diesterase ratios for the pruned
enzymes were not just similar, but were also
near unity (Fig. 4C; Tables 1B and 2). To test
whether this similarity was deep-seated or
coincidental due to the particular phosphate
ester leaving group used, we carried out
analogous measurements with phosphate
mono- and diesters across a range of intrinsic
reactivities, using phosphate mono- and
diesters with a 3-nitrophenoxide or 3,4-
dinitrophenoxide instead of p-nitrophenoxide
leaving group and thereby spanning a range of
leaving group pKa values of three units and
reactivity of ~103 fold (52-54) (& unpub
results). Monoesterase:diesterase activity
ratios near unity were observed for these
substrates as well (Supplemental Table 1). We
do not know if this identity represents an
Weak Sulfatase Activity of the Zn2+ Bimetallo
Scaffold May Have Restricted Evolutionary
Diversification within this AP Superfamily
Branch. Highly promiscuous phosphate
monoesterase and diesterase activities of the
ancient Zn2+ bimetallo core may have
promoted the evolution of AP-superfamily
phosphate monoesterases and diesterases.
Conversely, activities not found in extant AP
superfamily bimetallo enzymes may be absent
because they are not efficiently catalyzed by
this motif. The AP superfamily sulfatases
provide a particularly incisive test of this
prediction, as the sulfatase and phosphatase
substrates and transition states are highly
analogous
geometrically
(56,57),
but
equivalence
contributions for the different reactions or
differential individual effects that
coincidentally sum to the same value.
of
individual
catalytic
sulfatases are not found in the bimetallo
branch, falling instead solely in the one metal-
ion active site subgroup (38,43,58-60).
The sulfatase activities of our
bimetallo scaffolds were extremely low, so
low that only upper limits could be determined
(Fig. 5, pNPS; Table 1B). Our most sensitive
measurements revealed that the bimetallo
scaffold is at least 105 fold less effective as a
sulfatase than as a phosphate monoesterase or
diesterase. Furthermore, the very low intrinsic
reactivity of sulfate esters necessitates highly
efficient catalysis (16,61). Thus, our results
support a mechanistic mandate for the absence
of bimetallo cores among sulfatases.
Intriguingly, the AP superfamily appears to
have overcome this limitation of bimetallo
scaffold catalysis by utilizing only one metal
ion for sulfatases, instead of two. While the
ancient origin of the AP superfamily members
prevents us from tracing possible ancestral
The similarity of the monoesterase
and diesterase activities of the AP superfamily
bimetallo scaffold suggests a model in which
this promiscuous scaffold provided a strong
physical and functional foundation that Nature
built upon to make distinct, more efficient, and
specialized monoesterases and diesterases.
Physically, two of the three phosphoryl
oxygen atoms face away from the bimetallo
center—these atoms are accessible to the local
active site and serve as functional handles
providing evolutionary opportunities to sculpt
new interactions, or to co-opt existing residues
for new purposes, along the pathway from a
generalist to specialist or from a monoesterase
to diesterase (or vice versa). Functionally, the
substantial ~1010–1011 fold rate enhancements
6

sulfatases and possible pathways for this
conversion (P.C. Babbitt, personal
communication), our results reveal
enzymatic activities is lower still. But once a
structural motif evolves, gene duplication
allows it to be put to multiple uses, and
a
fundamental underlying mechanistic property
that Nature faced as it navigated these
evolutionary processes.
promiscuous
catalytic
activity
greatly
increases the probability that evolution will be
able to find, utilize, and optimize a potentially
beneficial activity (3-5,64). These factors have
presumably led to the preponderance of
enzymes with shared folds and shared
catalytic features (3,10,11).
Phosphotriesterase Activity of the Zn2+
Bimetallo Scaffold? As all known cognate
phosphate triesterases exist outside of the AP
superfamily (62), we wondered if the AP
superfamily bimetallo scaffold is also less
efficient as a triesterase. Indeed, the triesterase
rate enhancements were ~103–104 fold lower
than those for reactions of phosphate diesters
and monoesters (Fig. 5, paraoxon; Table 1B,
Supplemental Fig. 4). The lower catalysis
could arise from differences in substrate
charge or transition state charge distribution,
due to the tighter triester transition state
structures, although such differences did not
manifest for the mono- and di-esters despite
their charge and transition state differences
(44).
We have posed two fundamental
questions at the interface of catalytic function
and evolution, using the Zn2+ bimetallo
scaffold of the AP superfamily: (1) Does the
superfamily bimetallo core intrinsically favor
certain reactions over others and, if so, are
these preferences mirrored in evolutionary
history? (2) Did evolution, subsequent to
divergence in the AP superfamily, “tune”
individual bimetallo cores and scaffolds to
favor cognate over noncognate reactions?
We addressed the second question by
determining the activity of three AP
superfamily Zn2+ bimetallo scaffolds, absent
of the specialized interactions specific to each
subset of superfamily members. The AP
superfamily members studied herein—E. coli
The bimetallo scaffold is at least 2–3
orders of magnitude more effective at
catalyzing the phosphate triester reaction than
sulfatase reaction, and the greater intrinsic
reactivity of phosphate triesters than sulfate
esters (63) would render it more probable for
Nature to be able to take advantage of this
catalytic promiscuity. Nevertheless, there is an
apparent absence of an AP superfamily
triesterase, and this may arise from the lower
catalytic effectiveness of the bimetallo center
for this reaction relative to phosphate mono-
and diester reactions. However, it may also, or
instead, arise from the rarity of phosphate
triesters in biology and/or from factors
intrinsic to the AP superfamily structural
scaffold that render it difficult to sculpt
binding sites for both substituents of the
transferred phosphoryl group, as elaborated in
the Discussion.
Alkaline
Phosphatase
(EcAP),
an
evolutionarily distinct alkaline phosphatase
from Chryseobacterium meningosepticum
(PafA), and a phosphate diesterase from
Xanthamonas citri (NPP)—have very low
overall sequence identity and distinct
architectural differences within their common
bimetallo scaffolds (Fig. 2A & B, gray and
white; Supplemental Fig. 1). Nevertheless,
these enzymes, with a small set of divergent
active site side chains pruned away (Fig. 2C,
color-coded), have highly similar phosphate
monoesterase and phosphate diesterase
activities, each within a 5-fold range (Fig. 4A
& 4B; Fig. 5). This observation is consistent
with the common Zn2+–Zn2+ distance observed
for AP superfamily members that catalyze
hydrolysis of phosphate mono- and diesters
(65), and these similarities suggest that
electrostatic or geometric effects rooted in the
Zn2+ bimetallo site did not specialize the active
site Zn2+ bimetallo center for cognate over
DISCUSSION
The probability of de novo evolution
of proteins that adopt specific structures is
extremely low, and the probability that such
proteins exhibit substantial and specific
7

noncognate reactions. Rather, active site
features other than the bimetallo core are
responsible for monoester versus diester
specialization. As illustrated in Figs. 2 and 6,
the bimetallo core provides a scaffold for these
specializing auxiliary domains and side chains
(see also Ref. (40,45,46)).
comparisons of EcAP reactions reveal a strong
dependence of catalytic power on substrate
charge (56). Thus, to achieve sulfatase
activity, the Alkaline Phosphatase superfamily
may have had to undergo a more involved
remodeling of its core, in addition to taking on
auxiliary domains (38). We speculate that this
extensive remodeling might have been
evolutionary allowed if metal ligand mutations
led to loss of the second metal ion and use of
one or more of the freed ligands to aid
With this information in hand, we turn
to the broad question of the interplay of
catalytic promiscuity and the selection and
optimization of enzymes through evolution.
We found that the rate enhancements of our
pruned enzymes for both phosphate monoester
sulfatase
catalysis.
Alternatively,
the
sulfatases may have arisen from a highly
evolved AP superfamily phosphatase, using
the catalytic interactions beyond the bimetallo
core to provide modest and selectable sulfate
ester catalysis while the second metal site was
repurpose; it is also possible that the
phosphatases arose from the addition of a
second metal ion site to a sulfatase precursor.
and diester hydrolysis are substantial, 1010
–
1011 fold (Fig. 5), rivaling or exceeding the
catalytic power of many natural enzymes (2).
Thus, an AP superfamily scaffold with the
Zn2+ bimetallo center alone is a highly
effective catalyst that could have provided an
effective starting point for the evolution of the
extant enzymes that specialize in each of these
reactions, as depicted by the central generalist
“bimetallo core” in Fig. 6 (see also simplified
version of Fig. 6 in Supplemental Fig. 6).
While the ancient origin of the Alkaline
Phosphatase superfamily prevents tracing its
evolutionary history, the generalist ability of
the bimetallo core seems likely to have
contributed to its use in both phosphate mono-
and di-esterases, whether these activities arose
from a common generalist or from one
another.
We also determined the promiscuous
phosphotriesterase activity of the AP
superfamily bimetallo scaffold, a biological
reaction that is catalyzed by members of the
Amidohydrolase superfamily (66-68), but not
known to be catalyzed within the AP
superfamily. Catalysis by the pruned AP
superfamily enzymes is substantial, and
considerably greater than that for a sulfate
ester, although still ~103 fold less than that for
phosphate monoesters and diesters (Fig. 5).
So why did Nature chose structural
and catalytic motifs other than those of the AP
superfamily to catalyze phosphate triester
reactions (62,66,69)? The simplest models are
probabilistic, especially given the small
number of phosphotriesterases found in Nature
((70)). When the selective advantage for a
phosphotriesterase arose, random duplications
may have occurred for members of the
Amidohydrolase superfamily or duplications
in that family may have had higher
promiscuity and thus been favored early in the
evolution of these new enzymes. However,
there is another possibility, rooted in the
structural architecture of the AP superfamily.
Below we describe how the architecture has
been used and remodeled through the
evolution of phosphate mono- and diesterases,
In contrast to this phosphate mono-/di-
esterase generalist activity, the pruned
enzymes are many orders of magnitude less
effective in catalyzing reactions of sulfate
esters (Fig. 5). Intriguingly, sulfatase reactions
are catalyzed exclusively by enzymes on a
distinct one-metal ion branch of the AP
superfamily (38,43,58-60). These observations
suggest a mechanistic imperative for the
absence of AP superfamily bimetallo
sulfatases and a functional driving force for
the formation of
a
distinct sulfatase
superfamily branch with just one divalent
metal ion (Fig. 6, orange) (59). While the
mechanistic origin of the inability of the
bimetallo centers to catalyze sulfuryl transfer
remains to be determined, linear free energy
8

and we then return to how architectural
features may have limited the ability to evolve
an AP superfamily triesterase.
can be built (Fig. 6). What about the potential
to build an Alkaline Phosphatase superfamily
active site that is optimal for a phosphate
triester substrate? Stereochemical studies
indicate that the second diester substituent has
a strong preference for the O2 position (Fig.
2D) in diesterase reactions catalyzed by EcAP
—i.e., in the same position it occupies in NPP
reactions—despite the presence of the
phosphatase helix and absence of a substituent
binding pocket (37). This observation implies
inhibition of positioning of the substituent in
the opposite position. Indeed, inspection of
active site interactions of EcAP and PafA
reveals that the backbone amide of the serine
or threonine nucleophile donates a hydrogen
bond to this oxygen atom (O1, Fig. 2D; Fig. 6).
Remarkably, the monoesterase PafA
and diesterase NPP share a common auxiliary
domain that is distinct from any in EcAP,
despite PafA and EcAP both functioning as
monoesterases (Fig. 6, Asn auxiliary domain,
magenta) ((40,41)). This element is inserted in
a common position in the bimetallo core and is
structurally homologous (Fig. 2A). Its role
appears to be to position the asparagine
residue (Fig. 2C, N111 and N100 in NPP and
PafA, respectively) that interacts with one of
the phosphoryl oxygen atoms that faces away
from the bimetallo core (Fig. 2D, O1). This
element provides an equivalent mono- and di-
esterase enhancement, consistent with
interaction with an unesterified oxygen atom
for mono- and diester substrates (40).
Architecturally, there is enough room and
there are sufficient interactions to build this
auxiliary domain onto the bimetallo core to
enhance catalysis.
Given that this backbone interaction is
located right at the site of nucleophilic attack,
we speculate that resculpting an AP
superfamily member to replace this interaction
and accommodate an additional triesterase
substituent would be difficult, and this
difficulty may account for the absence of
known AP superfamily triesterases (Fig. 6,
brown “X”). These observations underscore
the role of idiosyncratic structural and
chemical properties of each protein scaffold in
determining evolvability for new enzymatic
functions (73-77). Analogously, these features
will determine which scaffolds can be
efficiently engineered to effectively carry out
new reactions.
How, then, are monoester and diester
substrates distinguished? As highlighted in
Fig. 6, PafA, EcAP, and all known Alkaline
Phosphatase
superfamily
monoesterases
contain a common active site alpha helix (Fig.
6, EcAP & PafA, “phosphatase helix,” light
blue)(40). This helix appears to position active
site residues that interact with the other
outward-facing phosphoryl oxygen atom (Fig.
2D, O2). However, the phosphatase helix sits
in the area where the second phosphodiester
substituent is situated, and it is absent in all
known diesterases (40). Instead, residues that
line the region exposed by the absence of the
phosphatase helix are used to create a binding
pocket for this substituent (Fig. 6, NPP, diester
pocket, red), and different NPP subfamilies
utilize different residues in this area and in
nearby auxiliary domains to achieve distinct
substituent specificities (34,71,72).
Overall, our results suggest
a
profound connection between catalytic
mechanism and evolution and support a
prominent role for catalytic promiscuity in
deciding which enzymes and scaffolds were
adopted for particular catalytic roles and how
they were adapted to take on new catalytic
roles throughout evolution
(Fig. 6).
Furthermore, our results, combined with prior
mechanistic and structural studies, provide
perspective on how AP superfamily structural
elements and domains may have been used to
enhance and specialize catalysis (Fig. 6). Once
a scaffold like the AP superfamily bimetallo
core is in place, its further evolutionary
elaboration will depend on what reactions are
The different sets of auxiliary domains
in EcAP, PafA, and NPP highlight the
versatility of the bimetallo core as a scaffold
onto which additional domains and elements
9

“needed,” which reactions can be catalyzed by
that scaffold, and how readily that scaffold can
accommodate new structural and functional
elements. The insights herein and elsewhere
into the structural and mechanistic properties
of AP and other superfamilies should help
guide efforts to select, redesign existing, and
design de novo enzymes for practical
applications in chemistry, medicine, and
engineering.
described (45). Briefly, cells containing NPP
were lysed with osmotic shock and run over a
Q-sepharose column. After extensive washing,
the protein was eluted and protein-containing
fractions were pooled. StrepTactin® resin (5
mL, IBA Life Sciences, Göttingen, Germany)
was washed extensively with 0.5 M sodium
hydroxide, and was packed onto a gravity
column. Following neutralization of the resin
with 1 M Tris-HCl, pH 8.0, and washing with
several column volumes of column buffer
(100 mM Tris-HCl, pH 8.0, and 150 mM
NaCl), the pooled enzyme fractions were
loaded over the resin. Following washing the
resin with column buffer (100 mM Tris-HCl,
pH 8.0, 150 mM NaCl) the enzyme was eluted
with 5 mM desthiobiotin in the same buffer
and buffer exchanged into storage buffer (100
mM Tris-HCl, pH 8.0, 150 mM NaCl, and 100
µM ZnCl2).
EXPERIMENTAL PROCEDURES
Protein Expression and Purification.
Mutants of AP from Escherichia coli (EcAP),
NPP from Xanthomonas axonopodis pv. Citri,
and
PafA
from
Chryseobacterium
meningosepticum were expressed in E. coli
SM547λ (DE3) cells. Cells were grown to an
optical density of 0.6–0.8 in rich medium and
glucose (10 g tryptone, 5 g yeast extract, 5 g
NaCl, and 2 g glucose per liter) with 50 µg/mL
carbenicillin at 37 °C. Protein expression was
induced by adding 0.3 mM Isopropyl
thiogalactopyranoside. Cultures were grown at
30 °C for 16–20 h and were harvested by
centrifugation.
PafA was purified from a fusion pET-
22b construct containing a C-terminal StrepII
tag. The cell pellets were resuspended in
column buffer (100 mM Tris-HCl, pH 8.0, and
150 mM NaCl) and either frozen for later
purification or lysed by passing the suspension
through an Emulsiflex (Avestin, Ottawa, ON)
three times. The lysate was centrifuged to
remove cell debris (20,000g for 20 min), and
the supernatant was filtered through a 0.45 µm
filter. StrepTactin® resin (5 mL, IBA Life
Sciences) was washed with sodium hydroxide
as described above, and following neutralizing
the resin, the filtrate was loaded over the resin.
The resin was washed with 6 column volumes
of column buffer and the protein was eluted
with 5 mM desthiobiotin in column buffer.
Fractions containing purified PafA were
pooled and buffer exchanged into storage
buffer (10 mM sodium MOPS, pH 8.0, 50 mM
NaCl, and 100 µM ZnCl2).
EcAP was purified from a fusion
pMAL-p2X construct containing an N-
terminal maltose binding protein (MBP) with
a Factor Xa cleavage site between it and the
natural N-terminal and a C-terminal StrepII
tag end as previously described (6,47). Briefly,
pelleted cells with EcAP were lysed with
osmotic shock and purified over amylose
resin. The resin was washed extensively and
the protein was eluted with 10 mM maltose.
The enzyme was buffer exchanged into
storage buffer (10 mM sodium MOPS, pH 8.0,
50 mM NaCl, and 100 µM ZnCl2). Constructs
with and without the MBP tag used for
purification for NPP* and EcAP* were shown
to have the same activity, as did PafA* with
and without the Strep tag.
Protein purity was assessed with
sodium dodecyl sulfate−polyacrylamide gel
electrophoresis and was estimated to be >95%
for all enzymes by staining with Coomassie
Blue.
NPP was purified from a similar
pMAL-p2X fusion construct as EcAP,
containing an N-terminal maltose binding
protein (MBP) and C-terminal StrepII tags
with a Factor Xa cleavage site between it and
the natural N-terminal end, as previously
Synthesis of phosphomonoester and
diester substrates. Substrate 3,4-dinitrophenyl
10

and 3-nitrophenyl phosphate monoesters were
synthesized according to previously described
experimental procedures (3,4-diNPP and 3-
NPP respectively)(6).
in all cases. Reported errors were estimated
from at least two independent kinetic
measurements. Comparisons with independent
enzyme preparations for each mutant gave
values within error.
Aryl methyl diesters were prepared
using the method originally reported by Ba-
Saif et al. (78), using commercially available
dimethyl chlorophosphate (me-3,4-diNPP and
me-3-NPP respectively). After demethylation,
the acid form of the methyl diester was
isolated by resuspending the lithium salt in 5
M HCl and extracting into diethyl ether. This
method resulted in approximately 1%
contamination with aryl monoester. The
material was further purified over a RediSep
Rf Gold C18 (Teledyne Isco, Lincoln, NE)
The following buffers were used in pH
dependences that established that kinetics
were followed in a pH-independent region for
each of the pruned enzymes: sodium MES, pH
5.5 and 6.0; sodium MOPS, pH 7.0; sodium
CHES, pH 9.0; sodium CAPS, pH 10, each at
100 mM and in the presence of 500 mM NaCl
and 100 µM ZnCl2 (Supplemental Fig. 4). The
pH dependency for pNPP and me-pNPP is
similar in each enzyme, suggesting that the
dianionic form of pNPP is reacting and not the
monoionic species. The pH-dependency for
pNPP and me-pNPP for EcAP* is also similar
to the paraoxon pH dependency, suggesting
that paraoxon is hydrolyzed in the same active
site.
column
equilibrated
with
100
mM
triethylammonium chloride, pH 5.4. Gradient
chromatography was run with this buffer and a
0-40% acetonitrile gradient at 2% per min.
Fractions containing the aryl methyl diester
were pooled, dried by rotary evaporation, and
the acid form of the aryl methyl diester
isolated as described above. Using the isolated
acid form, concentrations for kinetic assays
were determined gravimetrically and by
quantification by absorbance of the leaving
group after total hydrolysis and values agreed
within 20% in all cases.
As the mutant activities are orders of
magnitude lower than those for the wild type
enzymes with their cognate substrates, we
carried out inhibition experiments to ensure
that the measured activities represent the
pruned enzymes and not a trace contaminant
and that phosphate monoester and diester
hydrolysis reactions occurred at the same
active site for each enzyme. Inhibition
experiments with inorganic phosphate (Pi)
were carried out with subsaturating amounts
of substrate. The Pi concentration range was
0.1–100 mM for EcAP, 0.5–100 mM for NPP,
and 0.13–100 mM for PafA (Supplemental
Fig. 5). The observation of Ki values that are
the same for the monoester and diester
substrates and distinct from the Ki values for
the wild type enzymes suggests that the same
enzyme is catalyzing both reactions in each
Kinetic
measurements were performed at 25 °C in a
Perkin-Elmer UV/bis Lambda 25
Assays.
Activity
spectrophotometer (Perkin Elmer, Waltham,
MA) in 0.1 M sodium MOPS, pH 8.0, 0.5 M
NaCl, 100 µM ZnCl2, unless otherwise noted.
For substrates with a p-nitrophenolate (Chart
2) or 3,4-dinitrophenolate leaving group, the
formation of free p-nitrophenolate or 3,4-
dinitrophenolate from hydrolysis of the
substrates was monitored continuously at 400
nm. For substrates with an 3-nitrophenolate
leaving group, the formation of free m-
nitrophenolate was monitored continuously at
394 nm. Rate constants were determined from
initial rates, and the activity of the free
enzyme, kcat/KM, was determined. The kinetic
parameters were shown to be first-order in
both enzyme and substrate concentration, with
concentrations of each varied over at least a 5-
fold range. Linear fits had R2 values of >0.98
case and that this enzyme is not
a
contaminating wild-type enzyme. To further
test the origin of the observed catalysis we
purified pruned versions of each enzyme with
the nucleophilic Ser or Thr residue mutated to
Gly:
S102G/D153A/R166S/E322Y/K328A; NPP:
T90G/F91A/N111A/L123A/Y205A; and
EcAP:
PafA: T79G/N100A/K162A/R164A. In each
11

case, the observed reaction rate was decreased
by one to two orders of magnitude relative to
the pruned enzymes and no reaction was
over a reservoir of 1 mL precipitant solution to
crystalize by the hanging drop method.
Crystals were harvested and frozen in liquid
observed
above
the
non-enzymatic
nitrogen
without
cryoprotectant,
background hydrolysis, strongly suggesting
that the observed activities of the pruned
constructs arose from the pruned enzymes
studied herein.
crystallographic data were collected at the
Stanford Linear Accelerator at beamline 11-1,
using the Pilatus 6M detector (Dectris) with
thin slice oscillations (0.2°) for a total 240°
rotation. Diffraction images were processed
with iMosflm and scaled with AIMLESS and
POINTLESS programs from the CCP4 suite
of software (79). The dataset was complete
(99.9%) and redundant (8.5-fold) to 2.03Å
resolution, with good merging statistics, which
are summarized in Table 3. Dataset resolution
was kept at the limit of the detector (2.03Å),
since strong merging statistics, high
completeness and high redundancy persisted
into the highest resolution bin (Table 3).
Metal occupancy. To test for metal ion
concentration
dependent
activation
of
expressed enzyme, PafA, NPP and EcAP were
incubated with the following metal
concentrations: 10 µM ZnCl2 µM, 100 µM
ZnCl2, 500 µM ZnCl2, 1.0 mM MgCl2 and 100
µM ZnCl2 in 10 mM sodium MOPS, pH 8.0,
and 50 mM NaCl at 25 °C. No activation was
observed for either PafA or NPP. The EcAP
pruned variant containing the E322Y mutation
displayed activation behavior, with a half time
for activation of ~4 days, while the E322A-
containing mutant maintained constant
activity. These results mirrored the activation
previously observed in the otherwise wild-type
context (6). The E322Y and E322A AP*
constructs had activities within 2-fold after
activation. The occupancy of metal ions in the
active site of all of the pruned enzymes and
WT enzymes were determined with atomic
emission spectroscopy (Supplemental Table
2), as described previously (6,47), following
overnight incubation in storage buffer with
100 µM ZnCl2, at room temperature.
The structure was solved by molecular
replacement (MR) using PHENIX(80) with
the WT PafA structure (PDB ID: 5TJ3(40)) as
a search model. Prior to MR, we modified the
search model by removing all solvent and
metal atoms, side-chains surrounding the two
active site metal sites, and the side-chains of
the mutated residues (T79S, N100A, K162A
and R164A). The molecular replacement
solution showed strong electron density peaks
for the missing active site side-chains and the
metal ions, a lack of density for the residues
mutated to alanines, and a clear change in the
position of the T79S loop. Additionally, a
strong positive density peak was found
coordinated between the two active site Zn2+
ions and S79, which we interpreted as Cl-.
After several rounds of refinement with
phenix.refine (80) and manual adjustments
with Coot (81), the final model had excellent
Circular dichroism (CD). CD spectra were
collected for the WT and pruned enzymes,
each with their MBP tags removed, in 10 mM
sodium MOPS, pH 8.0, 50 mM NaCl 100 µM
ZnCl2. The following enzyme concentrations
were used: EcAP WT and pruned 1.4 µM;
NPP WT and pruned, 1.0 µM; and PafA WT
and pruned, 1.4 µM.
geometry and stereochemistry, with Rwork
17.0% and Rfree = 20.9% (Table 3).
=
Crystallization
and
structure
EcAP* was buffer exchanged into 10 mM
sodium MOPS, pH 7.0, 50 mM NaCl 100 µM
ZnCl2 and concentrated to 5 mg/ml. Equal
volumes of EcAP* and precipitant solution
(23% PEG3350, 0.2 M NH3F, 0.2 M sodium
HEPES, pH 8.0) were mixed and placed over
a reservoir of 1 mL precipitant solution to
crystalize by the hanging drop method.
determination of PafA* and EcAP*. PafA*
was buffer exchanged into 10 mM Tris-HCl,
pH 8.0, 50 mM NaCl, 100 µM ZnCl2 and
concentrated to 2.5 mg/ml. Equal volumes of
PafA and precipitant solution (22% PEG3350,
0.1 M sodium acetate, pH 4.4, 0.2 M
ammonium sulfate) were mixed and placed
12

Crystals were harvested and frozen in liquid
nitrogen with paratone-N oil as cryoprotectant.
Crystallographic data were collected at the
Stanford Linear Accelerator at beamline 12-2
using the Pilatus 6M PAD detector with thin
slice oscillations and shutterless mode for
360° rotation, resulting in highly redundant
(Table 4), justifying the inclusion of the
weaker data in improving the resulting model.
ACKNOWLEDGEMENTS
We dedicate this paper to Patsy Babbitt on the
occasion of her retirement in honor of her
profound and creative insights into enzyme
evolution and function. This work was funded
by a grant from the US National Institutes of
Health to D.H. (GM64798). The SSRL
Structural Molecular Biology Program is
supported by the Department of Energy,
Office of Biological and Environmental
Research, and by the National Institutes of
Health, National Center for Research
Resources, Biomedical Technology Program,
and the National Institute of General Medical
Sciences. We thank Filip Yabukarski for
assistance with collecting crystallographic
data for PafA*. We thank Patsy Babbitt and
Jonathan Lassila for helpful discussions,
advice, and communication of unpublished
results. We thank Daniel Mokhtari and Ariana
Peck and members of the Herschlag lab for
discussions and comments on the manuscript.
2.45
Å
resolution dataset containing
anomalous signal from the Zn, despite the
energy offset compared to the Zn peak energy
(12658 eV versus 9668 eV). Diffraction
images were processed with the AUTOXDS
script (Ana Gonzalez, SSRL) using the current
XDS version, AIMLESS, and POINTLESS
programs from the CCP4 package (79).
Anomalous signal was kept during the
processing. Dataset resolution cutoff was
chosen based on the high completeness, >35
fold multiplicity, and >30% CC1/2 as
suggested by the scaling program AIMLESS.
The structure was solved by molecular
replacement with a single monomer (subunit
A) from the 3TG0 PDB entry in the PHASER
program(82) that produced a single solution in
space group P6322 (#182), with two
polypeptide chains in the unit cell. The
molecular replacement solution showed
negative peaks in the Fo-Fc map for the
mutated side chains (D153A, E322A, K328A)
or a lack of side chain density in the 2Fo-Fc
map for D101A and R166S. The following
features that were observed and incorporated
into the structural model: (a) a well-populated
metal site ligated by H162 and E164; (b) a
less-occupied metal site ligated by H125 and
D304; (c) the StrepII tag at the C-terminus
providing a histidine ligand (H452) to the
metal binding site (a). Anomalous difference
maps using the refined model suggest that all
metal binding sites have Zn2+ ions
incorporated. The model was updated and
checked in COOT (81), while being refined
initially with maximum likelihood program
REFMAC(83) and finally with BUSTER
(version 2.10.2 (84)). The final model had
excellent geometry and stereochemistry, in the
top 1% percent distribution for this resolution
range in MolProbity (85) and low residual
factors values: R = 16.46% and Rfree = 21.90%
AUTHOR CONTRIBUTION
F.S. and I.A. expressed and purified enzymes,
performed all characterization of the enzymes,
crystallized the enzymes, and mounted them
for X-ray studies.
T.D. and A.L. solved the crystal structures for
the PafA* and EcAP*.
J.S. Synthesized the substrates with alternative
leaving groups.
D.H. and F.S. designed the study, analyzed the
data and wrote the manuscript with
contributions from all authors.
CONFLICT OF INTEREST
The authors declare that they have no conflicts
of interest with the contents of this article.
REFERENCES:
1.
Edwards, D. R., Lohman, D.C.,
Wolfenden, R. (2012) Catalytic
proficiency: the extreme case of
S-O cleaving sulfatases. J. Am.
Chem. Soc. 134, 525-531
13

2.
3.
Wolfenden, R., Snide, M.J. (2001)
The depth of chemical time and
the power of enzymes as
catalysts. Acc. Chem. Res. 34, 938-
945
O'Brien, P. J., Herschlag, D.
(1999) Catalytic promiscuity and
the evolution of new enzymatic
activities. Chem. Biol. 6, R91-
R105
Jensen, R. A. (1976) Enzyme
recruitment in evolution of new
function. Annu. Rev. Microbiol.
30, 409-425
Khersonsky, O., Tawfik, D.S..
(2010) Enzyme promiscuity: a
mechanistic and evolutionary
perspective. Annu. Rev. Biochem.
79, 471-505
10.
Babbitt, P. C., Mrachko, G.T.,
Hasson, M.S., Huisman, G.W.,
Kolter, R., Ringe, D., Petsko, G.A.,
Kenyon, G.L., Gerlt, J.A. (1995) A
functionally divers enzyme
superfamily that abstracts the ac
protons of carboxylic acids.
Science 267, 1159-1161
Glasner, M. E., Gerlt, J.A., Babbitt,
P.C. (2006) Evolution of enzyme
superfamilies. Curr. Opin. Chem.
Biol. 10, 492-497
Pandya, C., Farelli, J.D., Dunaway-
Mariano, D., Allen, K.N. (2014)
Enzyme Promiscuity: Engine of
Evolutionary Innovation. J. Biol.
Chem. 289, 30229-30236
Allen, K. N., and Dunaway-
Mariano, D. (2004) Phosphoryl
group transfer: evolution of a
catalytic scaffold. Trends in
biochemical sciences 29, 495-503
Meng, Q., Chen, K., Ma, L., Hu, S.,
Yu, J. (2011) A systematic
11.
12.
4.
5.
13.
14.
6.
7.
Zalatan, J. G., Fenn, T.D.,
Herschlag, D. (2008)
Comparative enzymology in the
alkaline phosphatase
superfamily to determine the
catalytic role of an active-site
metal ion. J. Mol. Biol. 384, 1174-
1189
O'Brien, P. J., Herschlag, D.
(1999) Does the active site
arginine change the nature of the
transition state for alkaline
phosphatase-catalyzed
identification of Kolobok
superfamily transposons in
Trichomonas vaginalis and
sequence analysis on related
transposases. J. Genet. Genomics
38, 63-70
Gerlt, J. A., Allen, K.N., Almo, S.C.,
Armstrong, R.N., Babbitt, P.C.,
Cronan, J.E., Dunaway-Mariano,
D., Imker, H.J., Jacobson, M.P.,
Minor, W., Poulter, C.D., Raushel,
F.M., Sali, A., Shoichet, B.K.,
Sweedler, J.V. (2011) The
Enzyme Function Initiative.
Biochemistry 50, 9950-9962
O'Brien, P. J., Herschlag, D.
(1998) Sulfatase activity of E.coli
alkaline phosphatase
demonstrates a functional link to
arylsulfatases, an evolutionary
related enzymes family. J. Am.
Chem. Soc. 120, 12369-12370
15.
16.
phosphoryl transfer? J. Am. Chem.
Soc. 121, 11022-11023
8.
9.
Gerlt, J. A., Babbitt, P.C. (2001)
Divergent evolution of enzymatic
function: mechanistically diverse
superfamilies and functionally
distinct suprefamilies. Annu. Rev.
Biochem. 70, 209-246
O'Brien, P. J. (2006) Catalytic
Promiscuity and the Divergent
Evolution of DNA Repair
Enzymes. Chem. Rev. 106, 720-
752
14

17.
Voordeckers, K., Brown, C.A.,
Vanneste, K., van der Zande, E.,
Voet, A., Maere, S., Verstrepen,
K.J. (2012) Reconstruction of
ancestral metabolic enzymes
reveals molecular mechanisms
underlying evolutionary
innovations through gene
duplication. PLoS biology 10, 1-
17
Thomson, J. M., Gaucher, E. A.,
Burgan, M. F., De Kee, D. W., Li, T.,
Aris, J. P., and Benner, S. A.
(2005) Resurrecting ancestral
alcohol dehydrogenases from
yeast. Nature genetics 37, 630-
635
Bridgham, J. T., Carroll, S. M., and
Thornton, J. W. (2006) Evolution
of hormone-receptor complexity
by molecular exploitation.
Science 312, 97-101
Ortlund, E. A., Bridgham, J. T.,
Redinbo, M. R., and Thornton, J.
W. (2007) Crystal structure of an
ancient protein: evolution by
conformational epistasis. Science
317, 1544-1548
Bar-Rogovsky, H., Stern, A., Penn,
O., Kobl, I., Pupko, T., Tawfik, D.S.
(2015) Assessing the prediction
fidelity of ancestral
relationships. Biochemistry 55, 6375-
6388
24.
Renata, H., Wang, Z.J., Arnold,
F.H. (2015) Expanding the
enzyme universe: accessing non-
natural reactions by mechanism-
guided directed evolution.
Angew. Chem. Int. Ed. 54, 3351-
3367
25.
26.
Babtie, A., Tokuriki, N.,
18.
Hollfelder, F. (2010) What makes
an enzyme promiscuous? Curr.
Opin. Chem. Biol. 14, 200-207
Furnham, N., Dawson, N.L.,
Rahman, S.A., Thornton, J.M.,
Orengo, C.A. (2016) Large-Scale
Analysis Exploring Evolution of
Catalytic Machineries and
19.
20.
Mechanisms in Enzyme Superfamilies. J.
Mol. Biol. 428, 253-267
27.
Das, S., Dawson, N.L., Orengo, C.A.
(2015) Diversity in protein
domain superfamilies. curr. opin.
genet. dev. 35, 40-49
28.
van Loo, B., et al. (2010) An
efficient, multiply promiscuous
hydrolase in the alkaline
phosphatase superfamily. Proc.
Natl. Acad. Sci. USA 107, 2740-
2745
21.
22.
23.
29.
30.
Kim, A., et al. (2011) Divergence
of chemical function in the
alkaline phosphatase
superfamily: structure and
mechanism of the P-C bond
cleaving enzyme
phosphonoacetate hydrolase.
Biochemistry 50, 3481-3494
Jonas, S., van Loo, B., Hyvonen,
M., Hollfelder, F. (2008) A new
member of the alkaline
reconstruction by a library
approach. Protein Eng. Des. Sel.
28, 1-12
Weiss, M. C., Sousa, F.L.,
Mrnjavac, N., Neukirchen, S.,
Roettger, M., Nelson-Sathi, S.,
Martin, F. (2016) The physiology
and habitat of the last universal
common ancestor. Nat. Microbiol.
1, 1-8
Baier, F., Copp, J.N., Tokuriki N.
(2016) Evolution of enzyme
superfamilies: comprehensive
exploration of sequence-function
phosphatase superfamily with a
formylglycine nucleophile:
structural and kinetic
characterisation of a
phosphonate monoester
15

hydrolase/phosphodiesterase
from Rhizobium leguminosarum.
J. Mol. Biol. 384, 120-136
Gijsbers, R., Ceulemans, H.,
Stalmans, W., Bollen, M. (2001)
Structural and catalytic
similarities between nucleotide
pyrophosphatases/phosphodiest
erases and alkaline
phosphatases. J. Biol. Chem. 276,
1361-1368
Chapman, E., Best, M.D., Hanson,
S.R., Wong, C.H. (2004)
evaluation of the effect of Arg166
mutations in Escherichia coli
alkaline phosphatase.
Biochemistry 47, 7663-7672
Lassila, J. K., Herschlag, D. (2008)
Promiscuous sulfatase activity
and thio-effects in a
phosphodiesterase of the
alkaline phosphatase
superfamily. Biochemistry 47,
12853-12859
Jonas, S., Hollfelder, F. (2009)
Mapping catalytic promiscuity in
the alkaline phosphatase
superfamily. Pure Appl. Chem. 81,
731-742
Galperin, M. Y., Bairoch, A.,
Koonin, E.V. (1998) A
31.
32.
37.
38.
39.
Sulfotransferases: structure,
mechanism, biological activity,
inhibition, and synthetic utility.
Angew. Chem. Int. Ed. Engl. 43,
3526-3548
33.
34.
Galperin, M. Y., Jedrzejas, M.J.
(2001) Conserved core structure
and active site residues in
alkaline phosphatase
superfamily enzymes. Proteins
45, 318-324
Zalatan, J. G., Fenn, T.D., Brunger,
A.T., Herschlag, D. (2006)
Structural and functional
comparisons of nucleotide
pyrophosphatase/phosphodieste
rase and alkaline phosphatase:
implications for mechanism and
evolution. Biochemistry 45,
9788-9803
superfamily of metalloenzymes
unifies phosphopentomutase and
cofactor-independent
phosphoglycerat mutase with
alkaline phosphatases and
sulfatases. Protein Sci. 7, 1829-
1835
40.
Sunden, F., AlSadhan, I.,
Lyubimov, A.Y., Ressl, S.,
Wiersma-Koch, H., Borland, J.,
Brown, C.L., Johnson, T.A., Singh,
Z., Herschlag, D. (2016)
Mechanistic and evolutionary
insights from comparative
enzymology of
35.
Olguin, L. F., Askew, S.E.,
phosphomonoesterases and
phosphodiesterases across the
alkaline phosphatase
O'Donoghue, A.C., Hollfelder, F.
(2008) Efficient catalytic
promiscuity in an enzyme
superfamily: an arylsulfatase
shows a rate acceleration of
10(13) for phosphate monoester
hydrolysis. J. Am. Chem. Soc. 130,
16547-16555
O'Brien, P. J., Lassila, J.K., Fenn,
T.D., Zalatan, J. G., Herschlag, D.
(2008) Arginine coordination in
enzymatic phosphoryl transfer:
superfamily. J. Am. Chem. Soc.
Bihani, S. C., Das, A., Nilgiriwala,
K. S., Prashar, V., Pirocchi, M.,
Apte, S. K., Ferrer, J., Hosur, M. V.
(2011) X-ray structure reveals a
new class and provides insight
into evolution of alkaline
phosphatases. PloS one 6, 1-10
Berlutti, F., Passariello, C., Selan,
L., Thaller, M. C., Rossolini, G. M.
41.
42.
36.
16

(2001) The Chryseobacterium
meningosepticum PafA enzyme:
prototype of a new enzyme
family of prokaryoitc phosphate-
irrepressible alkaline
active site: structural studies of
three highly active mutants of
Escherichia coli alkaline
phosphatase. J. Mol. Biol. 316,
941-953
phosphatases? Microbiology 147,
2831-2839
Bond, C. S., Clements, P.R., Ashby,
S.J., Collyer, C.A., Harrop, J.J.,
Guss, J.M. (1997) Structure of a
human lysosomal sulfatase.
Structure, 277-289
Lassila, J. K., Zalatan, J.G.,
Herschlag, D. (2011) Biological
phosphoryl-transfer reactions:
understanding mechanism and
catalysis. Annu. Rev. Biochem. 80,
669-702
Wiersma-Koch, H. I., Sunden, F.,
Herschlag, D. (2013) Site-
directed mutagenesis maps
interactions that enhance
cognate and limit promiscuous
catalysis by an alkaline
phosphatase superfamily
diesterase. Biochemistry 52,
9167-9176
Zalatan, J. G., Fenn, T. D.,
Herschlag, D. (2008)
Comparative enzymology in the
alkaline phosphatase
superfamily to determine the
catalytic role of an active-site
metal ion. J. Mol. Biol. 384, 1174-
1189
Sunden, F., Peck, A., Salzman, J.,
Ressl, S., Herschlag, D. (2015)
Extensive site-directed
49.
50.
Wang, J., Stieglitz, K., Kantrowitz,
E.R. (2005) Metal Specificity Is
Correlated with Two Crucial
Active Site Residues in
Escherichia coli Alkaline
Phosphatase. Biochemistry 44,
8378-8386
43.
44.
Murphy, J. E., Tibbits, T.T.,
Kantrowitz, E.R. (1995)
Mutations at positions 153 and
328 in Escherichia coli alkaline
phosphatase provide insight
towards the structure and
function of mammalian and yeast
alkaline phosphatases. J. Mol.
Biol. 253, 604-617
O'Brien, P. J., Herschlag, D.
(2002) Alkaline phosphatase
revisited: hydrolysis of alkyl
phosphates. Biochemistry 41,
3207-3225
45.
51.
52.
53.
Jencks, W. P., Regenstein, J.
(1976) Handbook of biochemistry
and molecular biology, Cleveland,
OH
46.
Tull, D., Withers, S.G. (1994)
Mechanisms of cellulases and
xylanases: A detailed kinetic
study of the exp- b -1,4-glycanase
from Cellulomonas fimi.
Biochemistry 33, 6363-6370
Kirby, A. J., Varvoglis, A.G. (1966)
The reactivity of phosphate
esters. Monoester hydrolysis. J.
Am. Chem. Soc. 89, 415-423
Ratner, H. (1984)
Flavobacterium
Meningosepticum. Infection
Control 5, 237-239
Nikolic-Hughes, I., Rees, D.C.,
Herschlag, D. (2004) Do
47.
48.
54.
55.
56.
mutagenesis reveals
interconnectedfunctional units in
the alkaline phosphatase active
site. eLife 4, e06492
Le Du, M. H., Lamoure, C., Muller,
B.H., Bulgakov, O.V., Lajeunesse,
E., Ménez, A., Boulain, J.C. (2002)
Artificial evolution of an enzyme
17

electrostatic interactions with
positively charged active site
groups tighten the transition
state for enzymatic phosphoryl
transfer? J. Am. Chem. Soc. 126,
11814-11819
Cleland, W. W., Hengge, A. (2006)
Enzymatic Mechanisms of
Phosphate and Sulfate Transfer.
Chem. Rev. 106, 3252-3278
Boltes, I., Czapinska, H., Kahnert,
A., von Bülow, R., Dierks, T.,
Schmidt, B., von Figura, K.,
Kertesz, M.A., Usón, I. (2001) 1.3
Å Structure of Arylsulfatase from
Pseudomonas aeruginosa
Establishes the Catalytic
PON1 suggest that its native
activity is lactonase. Biochemistry
Babtie, A. C., Bandyopadhyay, S.,
Olguin, L.F., Hollfelder, F. (2009)
Efficient catalytic promiscuity for
chemically distinct reactions.
Angew. Chem. Int. Ed. Engl. 48,
3692-3694
63.
57.
58.
64.
65.
Ohno, S. (1970) Evolution by gene
duplication, Springer-Verlag,
London
Bobyr, E., Lassila, J. K., Wiersma-
Koch, H. I., Fenn, T. D., Lee, J. J.,
Nikolic-Hughes, I., Hodgson, K. O.,
Rees, D. C., Hedman, B., and
Herschlag, D. (2012) High-
resolution analysis of Zn(2+)
coordination in the alkaline
phosphatase superfamily by
EXAFS and x-ray crystallography.
J Mol Biol 415, 102-117
Xiang, D. F., Kolb, P., Fedorov,
A.A., Meier, M.M., Fedorov, L.V.,
Nguyen, T.T., Sterner, R., Almo,
S.C., Shoichet, B.K., Raushel, F.M.
(2009) Functional annotation
and three-dimensional structure
of Dr0930 from Deinococcus
radiodurans, a close relative of
phosphotriesterase in the
Mechanism of Sulfate Ester
Cleavage in the Sulfatase Family.
Structure 9, 483-491
59.
Lukatela, G., Krauss, N., Theis, K.,
Selmer, T., Gieselmann, V., von
Figura, K., Saenger, W. (1998)
Crystal structure of human
arylsulfatase A: the aldehyde
function and the metal ion at the
active site suggest a novel
mechanism for sulfate ester
hydrolysis. Biochemistry 37,
3654-3664
66.
60.
61.
62.
Hanson, S. R., Best, M.D., Wong,
C.H. (2004) Sulfatases: structure,
mechanism, biological activity,
inhibition, and synthetic utility.
Angew. Chem. Int. Ed. Engl. 43,
5736-5763
Benkovic, S. J., Benkovic, P.A.
(1966) Studies on sulfate esters.
I. Nucleophilic reactions of
amines with p-nitrophenyl
sulfate. J. Am. Chem. Soc. 88,
5504-5511
amidohydrolase superfamily.
Biochemistry 48, 2237-2247
Seibert, C. M., Raushel, F.M.
(2005) Structural and Catalytic
Diversity within the
Amidohydrolase Superfamily.
Biochemistry 44, 6383-6391
Holm, L., and Sander, C. (1997)
An Evolutionary Treasure:
Unification of a Broad Set of
Amidohydrolases Related to
Urease. Proteins: Struct., Funct.,
Genet. 28, 72-82
67.
68.
Khersonsky, O., Tawfik, D.S.
(2005) Structure−reactivity
studies of serum paraoxonase
69.
Elias, M., Dupuy, J., Merone, L.,
Mandrich, L., Porzio, E., Moniot,
S., Rochu, D., Lecomte, C., Rossi,
18

M., Masson, P., Manco, G.,
(2007) Empirical fitness
Chabriere, E. (2008) Structural
basis for natural lactonase and
promiscuous phosphotriesterase
activities. J. Mol. Biol. 379, 1017-
1028
Bigley, A. N., Raushel, F.M. (2013)
Catalytic Mechanisms for
Phosphotriesterases. Biochim.
Biophys. Acta. 1834, 443-453
Nishimasu, H., Okudaira, S.,
Hama, K., Mihara, E., Dohmae, N.,
Inoue, A., Ishitani, R., Takagi, J.,
Aoki, J., Nureki, O. (2011) Crystal
structure of autotaxin and
insight into GPCR activation by
lipid mediators. Nat. Struct. Biol.
18, 205-212
landscapes reveal accessible
evolutionary paths. Nature 445,
383-386
Basaif, S. A., Davis, A.M., Williams,
A. (1989) Effective Charge-
Distribution for Attack of
Phenoxide Ion on Aryl Methyl
Phosphate Monoanion - Studies
Related to the Action of
Ribonuclease. J. Org. Chem. 54,
5483-5486
Evans, P. R., Murshudov, G.N.
(2013) How good are my data
and what is the resolution? Acta
Crystallogr., Sect D: Biol.
78.
70.
71.
79.
80.
Crystallogr. 69, 1204-1214
Adams, P. D. (2010) PHENIX: a
comprehensive Python-based
system for macromolecular
structure solution. Acta
72.
Gorelik, A., Liu, F., Illes, K., Nagar,
B. (2017) Crystal structure of the
human alkaline
sphingomyelinase provides
insights into substrate
recognition. J. Biol. Chem. 292,
7087-7094
Crystallogr., Sect D: Biol.
Crystallogr. 66, 213-221
Emsley, P., Lohkamp, B., Scott,
W.G., Cowtan, K. (2010) Features
and development of Coot. Acta
Crystallogr., Sect D: Biol.
81.
82.
73.
74.
Kirschner, M. W., Gerhardt, J.C.
(2005) The Plausibility of Life:
Resolving Darwin’s Dilemma, Yale
University Press, Princeton
Feder, M. E. (2007) Evolvability
of physiological and biochemical
traits: evolutionary mechanisms
including and beyond single-
nucleotide mutation. J. Exp. Biol.
210, 1653-1660
Crystallogr. 66, 486-501
McCoy, A. J., Grosse-Kunstleve,
R.W., Adams, P.D., Winn, M.D.,
Storoni, L.C., Read, R.J. (2007)
Phaser crystallographic
software. J. Appl. Crystallogr. 40,
658-674
Murshudov, G. N., Vagin, A.A.,
Dodson, E.J. (1997) Refinement
of Macromolecular Structures by
the Maximum-Likelihood
method. Acta Crystallogr., Sect D:
Biol. crystallogr. 53, 240-255
Bricogne, G., Blanc, E., Brandl, M.,
Flensburg, C., Keller, P., Paciorek,
W., Roversi, P, Sharff, A., Smart,
O.S., Vonrhein, C., Womack, T.O.
(2016) BUSTER version 2.10.12.
83.
84.
75.
76.
Kirschner, M., Gerhardt, J. (1998)
Evolvability. Proc. Natl. Acad. Sci.
U.S.A. 95, 8420-8427
Aharoni, A., Gaidukov, L.,
Khersonsky, O., Mc, Q. G. S.,
Roodveldt, C., and Tawfik, D. S.
(2005) The 'evolvability' of
promiscuous protein functions.
Nature genetics 37, 73-76
Poelwijk, F. J., Kiviet, D.J.,
77.
Weinreich, D.M., Sander, T.
19

Global Phasing Ltd., Cambridge,
United Kingdom
Chen, V. B., Arendall, W. B. 3rd,
Headd, J. J., Keedy, D. A.,
Immormino, R. M., Kapral, G. J.,
Murray, L. W., Richardson, J. S.,
Richardson, D. C. (2010)
MolProbity: all-atom structure
validation for macromolecular
crystallography. Acta
protein tyrosine phosphatases.
Biochemistry 53, 6811-6819
Bobyr, E., Lassila, J. K., Wiersma-
Koch, H. I., Fenn, T. D., Lee, J. J.,
Nikolic-Hughes, I., Hodgson, K. O.,
Rees, D.C., Hedman, B.,
Herschlag, D. (2012) High-
resolution analysis of Zn(2+)
coordination in the alkaline
phosphatase superfamily by
EXAFS and x-ray crystallography.
J. Mol. Biol. 415, 102-117
85.
87.
Crystallogr., Sect D: Biol.
Crystallogr. 66, 12-21
86.
Andrews, L. D., Zalatan, J.G.,
Herschlag, D. (2014) Probing the
origins of catalytic
88.
Mohamed, M. F., Hollfelder, F.E.
(2013) Efficient, crosswise
catalytic promiscuity among
enzymes that catalyze
discrimination between
phosphate and sulfate monoester
hydrolysis: comparative analysis
of alkaline phosphatase and
phosphoryl transfer. Biochem.
Biophys. Acta. 1834, 417-424
20

Tables
Table 1. Rate Constants for Reactions of Wild-type and Pruned EcAP, NPP, and
PafA.
A
kcat/KM (M-1s-1)
Enzyme
WT EcAP
pNPP
me-pNPP
dT5’-pNPP
pNPS
paraoxon
3.3 × 107 b
[6.3 × 108]
1.0 e
1.8 × 101 c
<2.0 × 10-2
2.8 × 10-3 d
N.D.
2.3 × 102 e
1.6 × 106 e
2.0 × 10-5 f
<5.9 × 10-4
WT NPP
WT PafA
1.6(0.5) × 106 b
1.7(0.5) ×
1.6(0.3) × 10-3 1.6(0.1) × 10-4
N.D.
[4.3 × 108]
101 b
a Assay conditions: 0.1 M sodium MOPS, pH 8.0, 0.5 M NaCl, 100 µM ZnCl2, and 500
µM MgCl2 at 25 ºC, unless noted otherwise. Errors (in parenthesis) in activities are
standard deviations from a duplicate with the same or independent enzyme preparations.
b Data from ref. (51) and ref. (40). Diffusion rather than the chemical step is rate limiting
for pNPP hydrolysis by WT EcAP. The estimated second-order rate constant with the
chemical step rate limiting is given in the square brackets, calculated in ref. (40,47,86).
c Data from ref. (6).
d Data from ref. (86).
e Data from ref. (34). Assay conditions: 0.1 M Tris-HCl, pH 8.0, 0.5 M NaCl, 100 µM
ZnCl2, and at 25 ºC.
f Data from ref. (37). Assay conditions: 0.1 M sodium MES, pH 6.5, 0.5 M NaCl, 100 µM
ZnCl2.
B
kcat/KM (M-1s-1)
Enzyme
pNPPb
Me-pNPP
dT5’-pNPP
pNPS
paraoxon
1.7(0.4) × 10-1
1.3(0.3) × 10-1
6.8(0.3) × 10-3
<1 × 10-6
7.2(1.6) × 10-3
EcAP*
D101A/D153A/R166S/E322A/K328A
1.7(0.6) × 10-1
1.0(0.4) × 10-1
1.8(0.1)
<4 × 10-6
<4 × 10-6
8.6(1.8) × 10-3
5.4(1.3) × 10-2
NPP*
T90S/F91A/N111A/L123A/Y205A
7.7(0.8) × 10-1
4.7(0.2) × 10-1
7.4(0.1) × 10-1
PafA*
T79S/N100A/K162A/R164A
a Assay conditions: 0.1 M sodium MOPS, pH 8.0, 0.5 M NaCl, 100 µM ZnCl2 and 500
µM MgCl2 at 25 ºC. Errors (in parenthesis) in activities are standard deviations from a
duplicate with the same or independent enzyme preparations.
21

Table 2. Rate Constants for Alternative Pruned Constructs of EcAP, NPP and PafA
pNPP2-
kcat/KM (M-1 s-1)
me-pNPP
kcat/KM (M-1 s-1) krel
c
d
Mutant
krel
EcAP pruned constructs
D101 D153 R166 E322 K328
A
G
N
A
A
A
A
A
S
S
S
S
A
Y
Y
Y
A
A
A
A
0.17(0.04)
0.13(0.02)b
0.19(0.03)b
0.17(0.03)b
(1)
1.3
0.9
1.0
0.13(0.03)
0.12(0.01)
-
(1)
1.1
-
0.13(0.01)
1.0
NPP pruned constructs
T90
S
F91
A
N111 L123 Y205
A
G
A
A
A
A
0.17(0.06)
(1)
0.10(0.04)
(1)
S
A
0.086(0.005)
1.9
0.080(0.004)
1.3
PafA pruned constructs
T79
S
N100
A
K162
R164
A
A
0.77(0.08)
0.10(0.02)
(1)
0.47(0.02)
0.14(0.01)
(1)
T
A
A
A
7.7
3.4
a Assay conditions: 100 mM sodium MOPS, pH 8.0, 500 mM NaCl, 100 µM ZnCl2 at 25
ºC. Errors (in parenthesis) in activities are standard deviations from a duplicate with the
same or independent enzyme preparations.
b Rate constants from ref.(47).
c Calculated as the ratio of (kcat/KM)pNPP, pruned/ (kcat/KM)pNPP, alt. pruned
.
dCalculated as the ratio of (kcat/KM)me-pNPP, pruned/ (kcat/KM)me-pNPP, alt. pruned
.
22

Table 3. X-ray Crystallographic Data Collection and Refinement Statistics for
PafA*
Data Collection
Space group
Unit cell axes
a, b, c (Å)
I 4
113.33 113.33 69.790
90 90 90
α, β, γ (ο)
Resolution range (Å)
Rmerge (%)
Rpim (%)
40.07–2.03 (2.10–2.03)
16.8 (151.8)
6.1 (54.1)
<I>/<σI>
9.6 (2.7)
Completeness (%)
Multiplicity
CC1/2
99.9 (99.9)
8.5 (8.8)
0.995 (0.660)
Refinement
Resolution range (Å)
No. unique reflections
40.07 - 2.03
28559 (2831)
Rwork/Rfree (%)
17.04/20.85
4241
Number of atoms
Average B-factors (Å2)
protein
37.1
37.7
48.3
water
ligands
R.m.s. deviation from
ideality
Bond length (Å)
Bond angles (ο)
Ramachandran statistics
Favored regions (%)
Allowed regions (%)
Outliers (%)
0.013
1.18
95
4.3
0.39
5TOO
PDB code
23

Table 4. X-ray Crystallographic Data Collection and Refinement Statistics for
EcAP*
Data Collection
Space group
Unit cell axes
a, b, c (Å)
P 63 2 2
160.72 160.72 138.36
90 90 120
α, β, γ (ο)
Resolution range (Å)
Rmerge (%)
38.60–2.45 (2.54–2.45)
59.4 (560.0)
10.3 (106.0)
Rpim (%)c
<I>/<σI>
7.8 (1.2)
Completeness (%)
Multiplicity
CC1/2
99.5 (95.6)
35.5 (30.4)
0.995 (0.318)
Refinement
Resolution range (Å)
No. unique reflections
Rwork/Rfree (%)
Number of atoms
Average B-factors (Å2)
protein
38.60 – 2.45
39204
16.46/21.90
7042
56.0
54.4
68.5
water
ligands
R.m.s. deviation from
ideality
Bond length (Å)
Bond angles (ο)
Ramachandran statistics
Favored regions (%)
Allowed regions (%)
Outliers (%)
0.010
1.13
97.75
2.25
0.00
PDB code
5TPQ
24

FIGURES
Chart 1.
Chart 2.
Reactions of the AP Superfamily
Substrates Used in Study.
Phosphate)monoester)hydrolysis)
O
O
P
Phosphate-
R
O
P
O
H2O
O
OH
HO
R
+
+
Monoester-
O
P
O2N
O
O
O
Phosphate)diester)hydrolysis)
O
O
O
O
P
4"nitrophenyl-phosphate-
R1
R1
(pNPP2")!
O
P
O
+
H2O
O
O
+
R
HO R12
HO
H+
O
O
Diester
-
R2
O
O2N
Sulfate)ester)hydrolysis)
P
O
O
O
O
S
O
R
methyl"4"nitrophenyl-phosphate-
+
+
HO
R
O
S
O
H2O
O
O
(me"pNPP1")!
O
H+
O
O
O
O2N
NH
Phosphate)transfer)
P
N
O
O
O
P
O
O
O
R1
R2
O
+
O
P
O
HO R2
O
O
HO R1
+
OH
O
O
dT5'"4"nitrophenyl-phosphate-
O
O
(dT5'"pNPP1")!
R1
R3
+
HO R3
HO R1
O
P
O
+
O
P
O
Triester-
O
P
O2N
O
O
R2
R2
O
O
O
Phosphonoacetate)hydrolysis)
dimethyl"4"nitrophenyl-phosphate-
O
O
P
O
(paraoxon)!
H2O
O
P
+
O
OH
+
O
O
O
O
Sulfate-
O
Monoester-
O
S
O2N
O
O
4"nitrophenyl-Osulfate-
(pNPS1")!
25

Figure 1. AP superfamily bimetallo active site and reaction pathway. Generic active
site of AP bimetallo enzyme showing the alkoxide nucleophile (Ser or Thr), Zn2+ atoms,
and Zn2+ ligands with a depiction of the reaction’s transition state, and general reaction
mechanism for AP superfamily enzymes via a covalent intermediate (E-P), shown for
reaction of a phosphate monoester giving an inorganic phosphate (Pi) product.
H
His
D
Asp
Asp
D
D
Asp
H
His
NH
O
O
N
O
O
O
His
H
O
Zn21+
N
Zn22+
NH
N
O
P
HN
OR
O
Nu:
O
O
2
1
H
k2"
RO_"
k3
HO_"
"
k4"
k1"
E!ROP
E-P
E + ROP
E!Pi
E + Pi
k#1"
26

Figure 2. Overall structure and active sites of EcAP, PafA, and NPP. (A) Topology
diagrams showing structurally conserved domains in gray and white and auxiliary
domains that vary between enzymes in color, with blue for EcAP, magenta for NPP, and
green for PafA. Residues that were mutated in the pruned constructs are colored in
orange. (B) Structure of WT EcAP (3TG0(87)), NPP (2GSN(34)), and PafA (5TJ3(40))
with the conserved Zn2+ ligands and nucleophiles are shown in black, Zn2+ ions in gray,
and the auxiliary domains and residues mutated in the pruned constructs colored as in
part (A). (C) Active site schematics with a phosphoryl group and leaving group (red)
represented as in the reaction’s transition state and the residues mutated in each pruned
enzyme shown in color; blue for EcAP, magenta for NPP, and green for PafA. (D)
Schematic representation of the presumed ground and transition state interactions for
reactions of the pruned enzymes.
27

EcAP
NPP
PafA
A
B
C
Conserved"
α Helix
β Sheet
6 c
"
6
"
6
"
"
6 e
D"""""""
Zn"ligand"8Asp"
D"""""""
Zn"ligand"8His"
N"""""""
Nucleophile"
X"""""""
Ac-ve"site"residue"
"
"
N"
4g"
D
3"
N" F"
""101"102"
N"111"
3"
3a"
N"
3a"
3"
N"100"
H""
90"
91"
79"
L""123"
H""
1"
""328" K"
H""
H""
H""
D""
1"
1"
D""
H""
H""
D""
H""
H""
D""
7 e
D""
D""
D""
"
322" E"
D""
D""
"
4e
Y""205"
""153"
D
162" K"
164" R"
4d"
R"166"
7a"
C"
8b"
C"
C"
1a"
N"
N"
N"
H412
H486
H363
D369
D352
D257
D327
D51
D305
D38
D210
H331
NH
D54
H309
NH
H214
NH
NH
NH
O
O
O
O
O
O
E322
N
O
O
N
N
H370
O
N
O
O
O
O
H353
H258
O
O
O
Zn2+
N
Zn2+
NH
O
O
1
Zn2+
Zn2+
O
N
N
Zn2+
Zn2+
2
1
1
δ-
O3
2
2
N
O
δ-
δ-
O
H
N
Y205
HN
O
O
HN
HN
H
δ-
K328
H
H
H
Mg2+
X
δ-
δ-
δ-
δ-
δ-
OR
OR
P
O
S102
K162
OR
P
P
O
O
O
H
N
δ-
H
T90
T79
δ-
H
H
O2
O1
HO
R
O
O
O
O
δ-
O
H
O
O
O
δ-
δ-
H
H
NH
N
H
H
H
N
N
O
H
N
H
N
N
N
H
N
D101
H
R164
H
N
H
N
H
H
F91
D153
H
H
N
H
O
O
O
N
H
H
H
R166
L123
HN
O
N100
N111
O
Zn21+
Zn21+
Zn21+
OR
Zn22+
Zn22+
Zn22+
D
O
O
O
P
P
OR
O
P
O
O
+ OR
Serine
N
O
O
O
2
O
O
O
1
2
2
1
1
N
H
H
N
H
28

Figure 3. Structural comparisons of wild type and pruned PafA and EcAP. (A)
Superimposition of the overall structure of WT EcAP (3TG0(87); blue) and EcAP*
(5TPQ, light blue). (B) Superimposition of active site of WT PafA (5TJ3(40); dark green)
and PafA* (5TOO; pale green). (C) Overlay of active sites of WT and pruned EcAP and
PafA. Structures and coloring as in parts (A) and (B). Blue residue numbers correspond
to EcAP and green correspond to PafA; oxygen atoms are depicted in red.
A"
B"
D327/305
C"
D51/38
H331/309
D369/352
S102
T79
H370/353
H412/486
T79S
29

Figure 4. Comparison of phosphate monoester and diester reactions catalyzed by
WT and pruned EcAP, NPP, and PafA. (A) Values of kcat/KM for phosphate monoester,
pNPP, hydrolysis. For WT, values in dark gray are measured and values in light gray are
calculated to represent reactions with the chemical step rate limiting (values are from
Table 1A and ref. (40,47,51,86)). The values for the pruned enzymes are shown in black
(Table 1B). (B) Values of kcat/KM for hydrolysis of minimal diester substrate me-pNPP.
For WT, the value in light gray is for the full cognate diester substrate, dT5’-pNPP (Table
1A). The values for the pruned enzymes with the minimal diester are shown in black. (C)
Specificity of the enzymes as ratio of reaction rates of monoester and diester hydrolysis,
for WT (gray) and pruned enzymes (black). For the WT enzymes, the gray bars are
obtained from the corresponding bars in parts (A) and (B) for mono- and di-esters,
respectively. The values for the pruned enzymes are near unity and therefore difficult to
see. Ratios for the reactions are given in Supplemental Table 3.
A
B
C
monoester
diester
specificity
1010
108
106
104
102
100
10-2
1010
108
106
104
102
100
10-2
108
104
100
10-4
10-8
WT
WT
WT
WT
WT
WT
WT
WT
WT
*
*
*
*
*
*
*
*
*
NPP.
PafA.
EcAP# NPP. PafA.
NPP.
EcAP#
EcAP#
PafA.
EcAP NPP PafA
EcAP NPP PafA
EcAP NPP PafA
30