Catalytic Promiscuity of Ancestral Esterases and Hydroxynitrile Lyases

Article abstract of DOI:10.1021/jacs.5b12209

Catalytic promiscuity is a useful, but accidental, enzyme property, so finding catalytically promiscuous enzymes in nature is inefficient. Some ancestral enzymes were branch points in the evolution of new enzymes and are hypothesized to have been promiscuous. To test the hypothesis that ancestral enzymes were more promiscuous than their modern descendants, we reconstructed ancestral enzymes at four branch points in the divergence hydroxynitrile lyases (HNL's) from esterases ～100 million years ago. Both enzyme types are α/β-hydrolase-fold enzymes and have the same catalytic triad, but differ in reaction type and mechanism. Esterases catalyze hydrolysis via an acyl enzyme intermediate, while lyases catalyze an elimination without an intermediate. Screening ancestral enzymes and their modern descendants with six esterase substrates and six lyase substrates found higher catalytic promiscuity among the ancestral enzymes (P < 0.01). Ancestral esterases were more likely to catalyze a lyase reaction than modern esterases, and the ancestral HNL was more likely to catalyze ester hydrolysis than modern HNL's. One ancestral enzyme (HNL1) along the path from esterase to hydroxynitrile lyases was especially promiscuous and catalyzed both hydrolysis and lyase reactions with many substrates. A broader screen tested mechanistically related reactions that were not selected for by evolution: decarboxylation, Michael addition, γ-lactam hydrolysis and 1,5-diketone hydrolysis. The ancestral enzymes were more promiscuous than their modern descendants (P = 0.04). Thus, these reconstructed ancestral enzymes are catalytically promiscuous, but HNL1 is especially so.

Full text of DOI:10.1021/jacs.5b12209

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Article
Catalytic promiscuity of ancestral esterases and hydroxynitrile lyases
Titu Devamani, Alissa M. Rauwerdink, Mark Lunzer, Bryan J. Jones, Joanna L. Mooney,
Maxilmilien Alaric O. Tan, Zhi-Jun Zhang, Jian-He Xu, Antony M. Dean, and Romas J. Kazlauskas
J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.5b12209 • Publication Date (Web): 06 Jan 2016
Downloaded from http://pubs.acs.org on January 14, 2016
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Catalytic promiscuity of ancestral esterases and hydroxynitrile lyases
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Titu Devamani, Alissa M. Rauwerdink, Mark Lunzer, Bryan J. Jones, Joanna L. Mooney,
c
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Maxilmilien Alaric O. Tan, ZhiꢀJun Zhang, JianꢀHe Xu, Antony M. Dean *, Romas J.
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a
Kazlauskas *
a
University of Minnesota, Department of Biochemistry, Molecular Biology & Biophysics and The
Biotechnology Institute, 1479 Gortner Avenue, Saint Paul, MN 55108 USA
b
University of Minnesota, Department of Ecology, Evolution & Behavior and The Biotechnology
Institute, 1479 Gortner Avenue, Saint Paul, MN 55108 USA
c
University of Minnesota, Department of Chemistry, 220 Pleasant Street, Minneapolis, MN
55455 USA
d
East China University of Science and Technology, School of Biotechnology, Meilong Road 130,
Shanghai 200237 P. R. China
e
Sun Yat-sen University, Institute of Ecology and Evolution, No.135, Xinggang West Road,
Guangzhou, 510275 P. R. China
Abstract: Catalytic promiscuity is a useful, but accidental, enzyme property, so finding catalytiꢀ
cally promiscuous enzymes in nature is inefficient. Some ancestral enzymes were branch points
in the evolution of new enzymes and are hypothesized to have been promiscuous. To test the hyꢀ
pothesis that ancestral enzymes were more promiscuous than their modern descendants, we reꢀ
constructed ancestral enzymes at four branch points in the divergence hydroxynitrile lyases
(
HNL’s) from esterases ~100 million years ago. Both enzyme types are α/βꢀhydrolaseꢀfold enꢀ
zymes and have the same catalytic triad, but differ in reaction type and mechanism. Esterases
catalyze hydrolysis via an acyl enzyme intermediate, while lyases catalyze an elimination withꢀ
out an intermediate. Screening ancestral enzymes and their modern descendants with six esterase
substrates and six lyase substrates found higher catalytic promiscuity among the ancestral enꢀ
zymes (P <0.01). Ancestral esterases were more likely to catalyze a lyase reaction than modern
esterases and the ancestral HNL was more likely to catalyze ester hydrolysis than modern HNL’s.
One ancestral enzyme (HNL1) along the path from esterase to hydroxynitrile lyases was espeꢀ
cially promiscuous and catalyzed both hydrolysis and lyase reactions with many substrates. A
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broader screen tested mechanistically related reactions that were not selected for by evolution:
decarboxylation, Michael addition, γꢀlactam hydrolysis and 1,5ꢀdiketone hydrolysis. The ancesꢀ
tral enzymes were more promiscuous than their modern descendants (P = 0.04). Thus, these reꢀ
constructed ancestral enzyme are catalytically promiscuous, but HNL1 is especially so.
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Introduction
Catalytic promiscuity is the ability of enzymes to catalyze additional reactions beyond
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those beneficial for life. Catalytic promiscuity differs from substrate promiscuity in that differꢀ
ent transition states, rather than the same transition state, must be stabilized. For example, the
natural function of carbonic anhydrase is to catalyze the reversible hydration of carbon dioxide,
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yet it also catalyzes the promiscuous hydrolysis of the ester pꢀnitrophenyl acetate.
Catalytic promiscuity is useful because it expands the range of available reactions for inꢀ
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dustrial biocatalysis applications. For example, halohydrin dehalogenase normally catalyzes the
hydrolysis of vicinal haloalcohols to vicinal diols and also catalyzes a promiscuous substitution
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of the halide with other nucleophiles. Fox and coworkers exploited this promiscuous reaction,
substituting chloride with cyanide to make a key intermediate in the synthesis of a cholesterolꢀ
lowering drug.
Catalytic promiscuity is difficult to search for because it is an accidental property of enꢀ
zymes. It is easier to search in places where the enzymatic property of interest is likely adaptive.
For example, searching high temperature environments will likely yield thermostable enzymes.
Searching chemical spill sites may yield enzymes that degrade environmental contaminants. Altꢀ
hough there is no location where catalytic promiscuity is adaptive, there may a time when it was
adaptive.
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Several groups have suggested that ancestral enzymes must have been catalytically
promiscuous. Some ancestral enzymes lie at key branch points where divergent evolution selectꢀ
ed new catalytic functions as new metabolic opportunities arose. These ancestors might also have
been promiscuous with regard to other mechanistically related reactions. If so, they would proꢀ
vide suitable places to search for catalytic promiscuity.
That ancestral enzymes no longer exist poses no problem – they can be resurrected.
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Analysis of modern sequences allows researchers to infer the sequences of ancestral enzymes.
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These analyses use the topology of phylogenetic trees, the likelihood of different amino acid
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replacements and various assumptions about evolution such as whether or not different sites in a
gene evolve at different rates. Chemical synthesis of the gene and expression of the protein in a
suitable host yields the ancestral enzyme for characterization. In cases where analyses suggest
several possible amino acids at a site, several ancestors can be made and characterized. In most
cases such ancestors have similar properties.
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Reconstructed ancestral enzymes show broader substrate ranges than their modern deꢀ
scendants. Voordeckers et al. showed that the reconstructed ancestors of modern αꢀglycosidases
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catalyzed hydrolysis of a wider range of substrates (maltose and isomaltose analogs). Likewise,
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reconstructed βꢀlactamases and carboxyl methyltransferases showed higher substrate promisꢀ
cuity. An ancestral protein kinase bound an inhibitor with an affinity intermediate between the
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modern descendants. Similarly, in vitro divergent evolution of glucuronidase to new substrates
specificity proceeded through nonꢀspecific intermediates. However, whether or not ancestral enꢀ
zymes were more catalytically promiscuous than modern enzymes has not been explored. In this
paper, we reconstruct ancestral hydroxynitrile lyases and esterases and assess their catalytic
promiscuity. The transition states for these two reaction types differ significantly, so finding a
single enzyme that catalyzes both reaction types efficiently would be remarkable.
Hydroxynitrile lyases (HNL's) are plant enzymes that catalyze the elimination of hydroꢀ
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gen cyanide from cyanohydrins as a defense against herbivorous insects. HNL's occur in at
least five protein folds; our focus here is on HNL's in the α/βꢀhydrolaseꢀfold superfamily. These
HNL's evolved from esterases ~100 million years ago when flowering plants and insects first diꢀ
versified. Presumably, a weak promiscuous ability to cleave cyanohydrins first arose serendipiꢀ
tously in an ancient esterase. Providing some protection against herbivorous insects, selection for
higher activity favored gene duplication, freeing the new copy to functionally diverge into the
modern specialized hydroxynitrile lyases.
The reaction mechanism for the hydrolysis reaction catalyzed by esterases (Enzyme Clasꢀ
sification (EC) group 3.1.1) differs in three key ways from the elimination reaction catalyzed by
hydroxynitrile lyases (EC 4.1.2). Hydrolysis follows the canonical serine hydrolase mechanism
with an acyl enzyme intermediate (Figure 1A). The elimination has no acyl enzyme intermediate
and uses only binding and general acidꢀbase chemistry (Figure 1B). Second, the two mechanisms
require conflicting substrate orientations. Hydrolysis requires the carbonyl oxygen to bind in the
oxyanion hole, while the lyase mechanism requires that it not bind in the oxyanion hole. Third,
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hydrolysis involves a hydrophobic leaving group, while the lyase reaction creates a polar leaving
group. These clearly different mechanisms provide a good test of the hypothesis that ancestral
enzymes were catalytically promiscuous. The prediction is that while modern enzymes catalyze
only one of these reaction types, ancestral α/βꢀhydrolases may catalyze both hydrolysis and elimꢀ
inations.
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Figure 1. Esterases catalyze hydrolysis of carboxylic acid esters while hydroxynitrile lyases
catalyze an elimination of hydrogen cyanide from cyanohydrins. Both active sites contain a serꢀ
ineꢀ histidineꢀaspartate catalytic triad. Aspartate not shown for clarity. A) Salicylic acid binding
protein 2 (SABP2) catalyzes the hydrolysis of methyl salicylate. The substrate ester binds with
the carbonyl oxygen in the oxyanion hole. The first step of the reaction is the nucleophilic attack
by the serine Oγ on the carbonyl carbon. The mechanism involves an acyl enzyme intermediate
(not shown). B) Hydroxynitrile lyase from Hevea brasiliensis (rubber tree) catalyzes the cleavꢀ
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age of acetone cyanohydrin. The substrate oxygen binds outside the oxyanion hole, which is
blocked by Thr11. Elimination proceeds in one step without an acyl enzyme intermediate.
Experimental section
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General. Enzymes and chemicals were bought from commercial suppliers and used directly
without further purification. Racemic mandelonitrile (Sigma Aldrich, St. Louis, MO) was
aliquoted in 10ꢀmL portions and stored at –18 °C. 2ꢀHydroxyꢀ6ꢀoxoꢀ6ꢀphenylhexaꢀ2, 4ꢀdienoic
acid (HOPDA) – was a gift from Lindsay Eltis’ research group at the University of British
Columbia and 2ꢀazabicyclo[2.2.1]heptꢀ5ꢀenꢀ3ꢀone (Vince lactam) was a gift from Robert Vince’s
research group at the University of Minnesota. Protein concentrations were determined from the
absorbance at 280 nm using extinction coefficients obtained from the ExPASy Bioinformatics
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Resource Portal. Protein gels were run on sodium dodecyl sulfate polyacrylamide gradient gels
NuPage 4–12% BisꢀTris gel from Invitrogen) using the BenchMark protein ladder (5 ꢁL/lane)
(
as a standard. DNA gels were run using 0.7% ultrapure agarose with 1×TAE buffer and 1 kb
DNA ladder as a standard. Micro titer plate assays were performed in triplicate and the mean of
the measurements is reported. One unit of enzyme activity corresponds to the amount of protein
required to release of 1 µmol of product per minute. Steady state kinetic data were fitted to the
MichaelisꢀMenten equation using the nonlinear fit routine in the Solver function of Microsoft
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Excel. NiNTA resin was regenerated according to Qiagen protocol. HꢀNMR spectra were run at
400 MHz in deuteriochloroform. Thin layer chromatography on silica gel was eluted with
hexanes: ethylacetate (8:2). Lysogeny broth (LB) and terrific broth (TB) were prepared
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according to Sambrook et al.
Ancestral enzyme reconstruction
Phylogenetics. Five thousand protein sequences, 150ꢀ600 amino acids long and sharing a miniꢀ
mum 30% sequence identity with Hevea brasiliensis acetone cyanohydrin lyase (GI: 1223884),
were obtained from the NCBI protein sequence database. Identical copies, mutant peptides and
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all patents were removed and the remaining sequences aligned using Muscle in SeaView. A
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preliminary neighbor joining tree was used to identify a cluster of 1285 sequences between
30% and 99% identical, and that included the ACLs and salicylic acid binding protein 2 from
Nicotiana tabacum, for further analysis.
The 1285 amino acid sequence alignment was adjusted manually guided by superꢀ
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positioned protein structures (H. brasiliensis 1QJ4, Manihot esculenta 1E8D, Arabidopsis thali-
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ana 3DQZ, Nicotiana tabacum 1XKL) obtained with DeepView/Swiss PDBꢀViewer to adjust
insertions and deletions into surface loops. The final alignment of 1285 sequences is available
upon request from the authors.
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Bootstrapped (1000 replicates) maximum likelihood trees were obtained using RAxML
using the ML + Bootstrap + GAMMAPROT + LG settings. Maximum likelihood uses a probaꢀ
bilistic model of sequence evolution to construct a tree from a given alignment. The GAMMAPꢀ
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ROT setting to allow some sites to evolve faster than others and the LG setting provides an
empirical amino acid exchange matrix to accommodate differences in the rates of exchange beꢀ
tween different amino acids. The most likely tree is one with topology, branch lengths and other
parameters that maximize the likelihood of the observed alignment. An advantage of the maxiꢀ
mum likelihood approach is that it provides a natural means to test alternative hypotheses ꢀ
branch lengths, topologies and ancestral states. Similar trees were obtained using FastML with
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GAMMAPROT and a JTT amino acid exchange matrix in RAxML with SH support and
bootstrapped neighbor joining with a Poisson correction in Seaview.
Ancestral sequences (Table S1) were inferred using maximum likelihood as implemented
in RAxML using the ML + Bootstrap + GAMMAPROT + LG tree obtained above. Maximum
likelihood calculates the most likely amino acid at each site at each node using an empirical
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Bayes approach. The likelihood of observing a particular amino acid, x, at a particular site at a
particular node is given as
where a is one of twenty amino acids at the focal site, t is the topology of the phylogenetic tree,
m is an evolutionary model, q represents the various model parameters (rates of amino acid exꢀ
change, variable rates across sites etc.) and p is the prior probability of observing x. Maximum
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likelihood resolves ambiguities in favour of the most likely model. For example, in the case of
tree (((Leu,Leu),(Met,Met)),Arg) the most likely ancestor of sequences had an Met because exꢀ
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changes between Met and Leu and between Met and Arg are commonplace while those between
Leu and Arg are rare. At branch points EST3 and HNL1, ancestral sequences were also inferred
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for the neighbor joining tree using maximum likelihood as implemented in MEGA.
Gene synthesis and cloning. Genes for the ancestral enzymes were synthesized by GenScript
and subcloned into a pET21a(+) vector at NdeI and XhoI restriction sites resulting in an upstream
T7 promoter and lac operator and an inꢀframe Cꢀterminal six Hisꢀtag. Fidelity of cloning and of
the gene synthesis was confirmed by DNA sequencing the entire gene (ACGT, Wheeling, IL).
Protein expression and purification. Lysogeny broth media containing 100 µg/mL ampicillin
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(
LBꢀamp, 5 mL) was inoculated with a single bacterial colony from an agar plate and incubated
in an orbital shaker at 37 °C and 200 rpm for 15 h. This culture was used to inoculate terrific
broth (TB)ꢀamp media (500 mL) in 2ꢀL baffled flasks, which was incubated at 37 °C and 250
rpm until the absorbance at 600 nm reached 1.0 (approximately 3ꢀ4 h). This culture was
transferred to 17 °C and 200 rpm for 1 h to cool, and then isopropyl βꢀDꢀ1ꢀthiogalactopyranoside
(
IPTG, 1 mM) was added to induce the protein expression. Cultivation was continued for 20 h.
The cells were harvested by centrifugation (8000 rpm, 10 min at 4 °C) and resuspended in buffer
A (20 mM imidazole, 50 mM NaH PO , 300 mM NaCl, pH 7.2, 20 mL). The cells were
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disrupted by sonication (400 W, 40% amplitude for 5 min) and centrifuged (4 °C, 12,000 rpm 45
min). The supernatant was loaded onto a column containing NiꢀNTA resin (1 mL, Qiagen) preꢀ
equilibrated with buffer A (10 mL). The column was washed buffer A (50 mL) followed by
buffer B (50 mM imidazole, 50 mM NaH PO , 300 mM NaCl, pH 7.2, 50 mL). The Hisꢀtagged
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protein was eluted with elution buffer (250 mM imidazole, 50 mM NaH PO , 300 mM NaCl, pH
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7.2, 15 mL) and concentrated to ~2 mL with an Amicon ultrafiltration centrifuge tube (10 kDa
cutoff). The imidazole buffer was exchanged by four successive additions of BES buffer (5 mM,
pH 7.0, 10 mL) to <15 ꢁM imidazole remaining, followed by concentration to ~2 mL with the
centrifuge tube. Typical yields were 20ꢀ100 mg protein from a 500ꢀmL culture.
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Racemic 2-nitro-1-phenylethanol was prepared according to a literature procedure.
Purification by silicaꢀgel column chromatography eluted with hexanes: ethyl acetate (85:15)
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yielded 1.22 g (70%) of a colorless oil. R 0.44, HꢀNMR: 2.86 (br, 1H), 4.45 (dd, J = 13.2, 2.9
f
Hz, 1H), 4.54 (dd, J = 13.2, 9.6 Hz, 1H), 5.43 ( dd , J = 9.8, 2.9 Hz, 1H), 7.33ꢀ7.39 (m, 5H).
Racemic 3-(2-nitro-1-phenylethyl)pentane-2,4-dione was prepared according to a literature
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procedure. Acidic alumina (1 g) was added to a stirred solution of acetylacetone (0.10 mL, 1.0
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mmol) and transꢀβꢀnitrostyrene (0.15 g, 1.0 mmol) in Et O (1 mL). The suspension was stirred at
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25 °C for 2 h, concentrated under vacuum, and purified by silicaꢀgel column chromatography
eluted with hexanes: ethyl acetate (80:20) yielding 0.106 g (43%) as pale yellow solid. mp 107ꢀ
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13 °C; lit. 110ꢀ112 °C; R 0.22, HꢀNMR: 1.92 (s, 3H), 2.28 (s, 3H), 4.18 (m, 1H), 4.38 (d, J =
f
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11.1 Hz, 1H), 4.61 (m, 2H) and 7.1ꢀ7.3 (m, 5H).
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Benzoylacetic acid: Benzoylacetic acid was prepared according to the literature procedure. To
a solution of acetophenone (12 g, 0.10 mol), dry tetrahydrofuran (50 mL) in 250 mL overnightꢀ
ovenꢀdried roundꢀbottomed flask, NaH (60%) in oil (10.5 g, 0.25 mol) and dimethylcarbonate
(
16.2 g, 0.18 mol) was added. The suspension was heated to reflux for 2 h and cooled to room
temperature. The reaction was quenched with ice water (100 mL), acidified with 3 M HCl to pH
ꢀ3 and extracted with ethyl acetate (3 × 25 mL). The organic layer was dried over sodium
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sulfate and concentrated under vacuum. Purification by silicaꢀgel column chromatography eluted
with hexanes: ethyl acetate (95:5) yielded 11.5 g (65% yield) of methyl 3ꢀoxoꢀ3ꢀphenyl
proponoate. Hydrolysis of methyl 3ꢀoxoꢀ3ꢀphenyl propanoate (1.0 g) in sodium hydroxide
solution (0.5 M) at room temperature. After 12 h the reaction was acidified with HCl to pH 2ꢀ3,
extracted with ethyl acetate (3 × 10 mL), dried and concentrated under vacuum. Benzoyl acetic
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acid obtained in 55% (0.5 g) yield as a white solid. R 0.33; mp 85 °C; lit. 99 °C; HNMR: 4.1
f
(s, 2H), 5.7 (s, vinyl 1H), 7.2ꢀ8.05 (m, 5H), 9.75 (br, 1H).
Activity and enantioselectivity of enzyme-catalyzed reactions
Hydroxynitrile lyase activity. Hydroxynitrile lyase activity was assayed as described previously
3
4
–1
–
with minor modifications. The assay monitors the release of benzaldehyde (ε = 1352 M cm
2
80
1
)
from mandelonitrile spectrophotometrically and was corrected for the spontaneous cleavage of
mandelonitrile. The assay solution contained mandelonitrile (2.0 mM), citrate buffer (pH 5.0, 50
mM) and enzyme (0.05ꢀ8 µM from a stock in 5 mM BES buffer, pH 7) in a total volume of 200
µL (path length = 0.60 cm). Steady state kinetic constants were determined under identical
conditions with mandelonitrile concentrations from 1ꢀ20 mM.
The enantioselectivity was measured using the reverse reaction: addition of HCN to
benzaldehyde. The reaction (0.5 mL total volume) contained benzaldehyde (1 mM, from a stock
solution of 50 mM of benzaldehyde in 50 mM in sodium citrate buffer pH 5.0) and HCN (50
mM, from a stock solution of 1.0 M of HCN in tertꢀbutyl methyl ether) in sodium citrate buffer
3
5
(
50 mM, pH 5.0). Caution: HCN is toxic and requires careful handling. Enzyme (0.002 ꢀ 0.1
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mg protein in 5 mM of BES buffer, 100 µL, pH 7.0) was added to start the reaction. After 2 h
shaking at 600 rpm at 20 °C, the tert-butyl methyl ether layer was separated, evaporated by
stream of nitrogen, and dissolved in isopropanol. The enantiomeric purity of the product
mandelonitrile was determined by HPLC using a Chiralcel ODꢀH column (Diacel) eluted with
hexane: isopropanol (98: 2) at flow rate of 1.25 mL/min: The S and R enantiomers elute at 31
and 34 min, respectively. Absolute configuration of mandelonitrile was established by
comparison with a commercial sample of (R)ꢀmandelonitrile (Alfa Aesar, Ward Hill, MA).
Cleavage of acetone cyanohydrin, lactonitrile, 2ꢀhydroxypentanenitrile and 2ꢀ
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hydroxyhexanenitrile was assayed using a modified Kӧnig reaction.
Substrate (1.28 mM to
10 mM in 0.1 M citric acid) was assayed with up to 8 µM enzyme in 5 mM citrate phosphate
buffer, pH 5. After up to 20 min, the reaction was quenched by addition of aqueous Nꢀ
chlorosuccinimide (2 mM, 62.5 µL, also containing 20 mM succinimide). After 2 min, barbituric
acid (230 mM in 30% pyridine, 12.5 µL) was added and after 10 min, the absorbance at 580 nm
was measured and compared to a calibration curve constructed using K [Zn(CN) ] with
2
4
concentrations of HCN ranging from 2.5ꢀ100 µM.
Nitroaldolase (Henry reaction) activity. Nitroaldolase activity was assayed by monitoring the
release of benzaldehyde from 2ꢀnitroꢀ1ꢀphenylethanol as above for the hydroxynitrile lyase
activity. The assay mixture (200 µL total volume, path length 0.60 cm) contained 2ꢀnitroꢀ1ꢀ
phenylethanol (2 mM from stock of 50 mM 2ꢀnitroꢀ1ꢀphenylethanol in 1 mM HCl), 50 mM
citrate phosphate buffer, pH 5.5, and up to 5 µM enzyme in BES buffer (5 mM, pH 7.2). A blank
reaction to monitor the spontaneous cleavage of 2ꢀnitroꢀ1ꢀphenylethanol was identical, but the
enzyme solution was replaced by BES buffer. Steady state kinetics used the same assay, but
varied the concentration of 2ꢀnitroꢀ1ꢀphenylethanol (0.5ꢀ6 mM).
The enantioselectivity was measured using the reverse reaction: addition of nitromethane
to benzaldehyde. The reaction (0.5 mL total volume) contained benzaldehyde (1 mM from 50
mM of benzaldehyde in diisopropyl ether (DIPE) and nitromethane (1.0 M) in 50 mM in citrate:
phosphate buffer (pH 6.0) and enzyme (0.1ꢀ0.7 ꢁM) in 5 mM of BES buffer. After 2 h shaking at
500 rpm at room temperature, the organic layer was separated and evaporated by stream of
nitrogen gas and dissolved in isopropanol. The enantiomeric purity of the product 2ꢀnitroꢀ1ꢀ
phenylethanol was determined by HPLC using a Chiralcel ODꢀH column (Diacel) eluted with
hexane: isopropanol (95: 5) at flow rate of 1.0 mL/min. The R and S enantiomers elute at 23 and
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6 min respectively and it was reported to elute at 13.5 and 16.3 min with hexane: isopropanol
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90: 10) at flow rate of 1.0 mL/min in Chiralcel ODꢀH column.
Esterase activity. Esterase activity was measured by hydrolysis of pꢀnitrophenyl acetate
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(
pNPAc) and was corrected for spontaneous hydrolysis. The assay mixture (100 µL total
volume; path length 0.29 cm) contained 6 mM of pNPAc, 8 vol% acetonitrile, 5 mM BES, pH
.0, and up to 5 µM enzyme in BES buffer. The increase in absorbance at 404 nm was monitored
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spectrophotometrically. The extinction coefficient used for calculations (ε404 = 11.4 × 10 M
1
–1
cm ) takes into account the incomplete ionization of pꢀnitrophenol at this pH. Steady state
kinetics used the same assay, but varied the concentration of pNPAc (0.5ꢀ6 mM). Methyl
salicylate and methyl mandelate were assayed at 500 µM with up to 10 µM enzyme in 5 mM
BES, pH 7. Methyl pentanoate, 1ꢀnaphthyl acetate, and 2ꢀnaphthyl acetate were assayed at 2 mM
in 5 mM BES buffer pH 7 with up to 50 µM enzyme. Control reactions were performed by
adding equal volumes buffer that was removed from the enzyme by centrifugal filtration.
Reactions were shaken at room temperature for up to 18 hours before enzyme was removed by
centrifugation using Amicon Ultra 0.5 mL regenerated cellulose 10 kDa cutoff centrifugal filters.
Reactions with methyl salicylate, 1ꢀnaphthyl acetate and 2ꢀnaphthyl acetate were analyzed by
HPLC on an Agilent Eclipse Plus C18 column eluted with methanol: water + 0.1% formic acid
(80: 20) at 1.0 mL/min. Methyl mandelate reactions were analyzed on the same column but with
a 60: 40 mix of solvents. The enantiomeric purity of unreacted methyl mandelate was measured
using a Chiralpak ASꢀRH column (Diacel) eluted with acetonitrile: water + 0.1% trifluroacetic
acid (30: 70) at 1.0 mL/min. The S and R enantiomers elute at 6.4 and 6.8 min, respectively. The
absolute configuration was assigned by comparison to sample of Rꢀmethyl mandelate prepared
from commercial Rꢀmandelic acid. Methyl pentanoate hydrolysis was quantified pꢀnitrophenol as
4
0
a pHꢀindicator as described previously.
Lactonase activity. Lactonase activity corresponds to the hydrolysis of 4ꢀphenylꢀ4ꢀ
butryolactone. The reaction mixture (0.50 mL total volume) contained 4ꢀphenylꢀ4ꢀbutyrolactone
(0.1 mM), BES buffer (5 mM, pH 7.0), and enzyme (7ꢀ75 ꢁM). Reaction was stirred at 500 rpm
at 25 °C for 24 h. The enzyme was removed by filtration through a centrifugal filter (Amicon
Ultra 0.5 mL regenerated cellulose 10 kDa cutoff spun at 4000 rpm) and the filtrate analyzed by
HPLC. The conversion was measured using an Agilent Eclipse Plus C18 column eluted with aceꢀ
tonitrile: H O + 0.1% formic acid (40: 60) where 4ꢀhydroxyꢀ4ꢀphenylbutyric acid and 4ꢀphenylꢀ
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ꢀbutyrolactone eluted at 3.4 and 5.4 min respectively. A blank without enzyme showed ~5 %
spontaneous hydrolysis. The amount was subtracted to determine the rate and conversion of the
reaction. The enantiomeric purity of unreacted 4ꢀphenylꢀ4ꢀbutyrolactone was determined by
HPLC using a Chiralpak ASꢀRH column (Diacel) eluted with acetonitrile: water + 0.1% of forꢀ
mic acid (35: 65) at 1.0 mL/min. The R and S lactone enantiomers elute at 10.4 and 12.5 min,
respectively. This configuration was established by comparison with an HPLC of a hydrolysis of
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the same lactone with pig liver esterase, which favors the (+)ꢀenantiomer. The (+)ꢀenantiomer
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was later identified as (R).
Decarboxylase activity. Decarboxylase activity corresponds to the decarboxylation of
benzoylacetic acid to acetophenone. The reaction mixture (0.50 mL total volume) contained
benzoylacetic acid (0.25 mM), BES buffer (5 mM, pH 7.3) and enzyme (13ꢀ75 ꢁM). The
reaction mixture was stirred at 500 rpm at 25 °C for 6 h, then the enzyme was removed by
filtration through an Amicon ultrafiltration centrifuge tube (10 kDa cutoff) at 4000 rpm and the
filtrate analyzed by HPLC on an Agilent Eclipse Plus C18 column eluted with acetonitrile: H O
2
+
0.1% formic acid (50: 50) monitored at 254 nm where benzoylacetic acid and acetophenone
elute at 4.3 and 7.4 min respectively. The molar absorbance of benzoylacetic acid is 3.03ꢀfold
larger than that for acetophenone and the peak areas were corrected by this factor. A blank
without enzyme showed ~0.1% spontaneous cleavage.
Michael addition. A twoꢀphase mixture of diꢀisopropyl ether (250 µL) and citrate: phosphate
buffer (50 mM, pH 5.5, 250 µL) containing transꢀβꢀnitrostyrene (2 mM), acetyl acetone (100
mM) and enzyme (80ꢀ140 ꢁM) was stirred at 500 rpm at 25 °C for 8 h. The diꢀisopropyl ether
layer was separated, flushed with stream of nitrogen and analyzed by HPLC using a Chiralpak
OJꢀR column (Diacel) eluted with acetonitrile: H O (40: 60) at 1.0 mL/min. The S and R enantiꢀ
2
omers of the product 3ꢀ(2ꢀnitroꢀ1ꢀphenylethyl)pentaneꢀ2,4 dione elute at 7.1 and 9.1 min, respecꢀ
tively and the starting material transꢀβꢀnitrostyrene elutes at 19.7 min. The absolute configuraꢀ
tion was assigned with a Chiralcel ODꢀH column eluted with hexane: isopropanol (95: 5) where
43
the S and R enantiomers were reported to elute at 25 and 27 min, respectively. A blank reaction
without enzyme showed ~ 6% spontaneous formation of racemic product. The reported rates and
enantiomeric purities were corrected for this spontaneous reaction.
Lactamase activity. Lactamase activity corresponds to the hydrolysis of 2ꢀ
azabicyclo[2.2.1]heptꢀ5ꢀenꢀ3ꢀone. The reaction mixture (0.50 mL total volume) contained 2ꢀ
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azabicyclo[2.2.1]heptꢀ5ꢀenꢀ3ꢀone (0.50 mM), pꢀamino benzoic acid (0.1 mM, internal standard),
Tris buffer (5 mM, pH 8.5), and enzyme (7ꢀ75 ꢁM). The reaction was stirred at 500 rpm at 25 °C
for 24 h. The enzyme was removed by filtration through an Amicon ultrafiltration centrifuge
tube (10 kDa cutoff) at 4000 rpm and the filtrate analyzed by HPLC to determine conversion and
enantioselectivity. The amount of substrate reacted was determined from the relative peaks area
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upon HPLC analysis on Agilent Eclipse Plus C18 column eluted with acetonitrile: H O + 0.1%
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formic acid (40:60). pꢀAminobenzoic acid and 2ꢀazabicyclo [2.2.1] heptꢀenꢀ3ꢀone elute at 3.9
and 4.4 min respectively (relative absorbance = 4.37). A reaction with no enzyme showed ~0.1%
conversion. The enantiomeric purity of unreacted 2ꢀazabicyclo [2.2.1] heptꢀenꢀ3ꢀone as deterꢀ
mined by HPLC using a Chiralpak ASꢀRH column (Diacel) eluted with acetonitrile: water +
0.1% formic acid (20: 80) at 1.0 mL/min. The (2S, 4R) and (2R, 4S) enantiomers eluted at 3.4
and 3.9 min, respectively. The absolute configuration was established by comparison of HPLC
traces with a sample of the (2R, 4S) enantiomer (Sigma Aldrich, St. Louis, MO).
C-C Bond hydrolysis activity. Hydrolysis of CꢀC bonds was determined with 2ꢀhydroxyꢀ6ꢀoxoꢀ
6ꢀphenylhexaꢀ2, 4ꢀdienoic acid (HOPDA) as the substrate to yield benzoic acid and 2ꢀ
4
4
hydroxypentaꢀ2, 4ꢀdienoic acid. This substrate was a gift from the Eltis group, who synthesized
it enzymatically. The assays were performed at pH 7.5 in 100 mM phosphate buffer with 100
µM HOPDA and up to 50 µM enzyme. The decrease in HOPDA was monitored at 434 nm (ε =
–1
–1
2
5.7 mM cm ).
Epoxide hydrolase activity. Epoxide hydrolase activity corresponds to the hydrolysis 2ꢀ(4ꢀ
nitrophenyl)oxirane to the corresponding 1ꢀ(4ꢀnitrophenyl)ethaneꢀ1,2ꢀdiol. The reaction mixture
(0.50 mL total volume) contained 2ꢀ(4ꢀnitrophenyl)oxirane (0.1 mM), BES buffer (5 mM, pH
7.0), and enzyme (7ꢀ75 ꢁM). Reaction was stirred at 500 rpm at 25 °C for 12 h. The enzyme was
removed by filtration through a centrifugal filter (Amicon Ultra 0.5 mL regenerated cellulose 10
kDa cutoff spun at 4000 rpm) and the filtrate analyzed by HPLC on an Agilent Eclipse Plus C18
column eluted with acetonitrile: H O + (40: 60). 1ꢀ(4ꢀNitrophenyl)ethaneꢀ1,2ꢀdiol and 2ꢀ(4ꢀ
2
nitrophenyl)oxirane elute at 3.4 and 10.7 min, respectively. Reactions without enzyme showed
~6 % spontaneous hydrolysis. No additional hydrolysis was detected in the reactions containing
enzyme.
Aldol addition activity. Aldol activity was measured by the formation of 4ꢀhydroxyꢀ4ꢀ
(nitrophenyl)butanꢀ2ꢀone from pꢀnitrobenzaldehdye and acetone. The reaction mixture (1 mL
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total volume) contained pꢀnitrobenzaldehdye (0.5 mM), acetone (10 mM), BES buffer (5 mM,
pH 7.0), and enzyme (3ꢀ37 ꢁM). Reaction was stirred at 500 rpm at 25 °C for 24 h. The enzyme
was removed by filtration through a centrifugal filter (Amicon Ultra 0.5 mL regenerated celluꢀ
lose 10,000 NMWL spun at 4000 rpm) and the filtrate analyzed by HPLC on an Agilent Eclipse
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Plus C18 column eluted with acetonitrile: H O + (50: 50). 4ꢀHydroxyꢀ4ꢀ(nitrophenyl)butanꢀ2ꢀ
2
one and pꢀnitrobenzaldehyde elute at 4.5 and 6.1 min, respectively. A blank without enzyme
showed ~ 0.1 % spontaneous addition. No additional product was detected in the reactions conꢀ
taining enzyme.
Baylis-Hillman reaction. BaylisꢀHillman reaction activity was measured by the formation of 3ꢀ
(hydroxyꢀ4ꢀ(nitrophenyl)methyl)butꢀ3ꢀenꢀ2ꢀone from pꢀnitrobenzaldehdye and methyl vinyl keꢀ
tone. The reaction mixture (1 mL total volume) contained pꢀnitrobenzaldehdye (0.5 mM), methyl
vinyl ketone (10 mM), BES buffer (5 mM, pH 7.0), and enzyme (3ꢀ37 ꢁM). Reaction was stirred
at 500 rpm at 25 °C for 24 h. The enzyme was removed by filtration through a centrifugal filter
(Amicon Ultra 0.5 mL regenerated cellulose 10 kDa cutoff spun at 4000 rpm) and the filtrate anꢀ
alyzed by HPLC on an Agilent Eclipse Plus C18 column eluted with acetonitrile: H O + (50: 50).
2
3ꢀ(Hydroxyꢀ4ꢀ(nitrophenyl)methyl)butꢀ3ꢀenꢀ2ꢀone and nitrobenzaldehyde elute at 4.1 and 6.1
min, respectively. A blank without enzyme showed ~ 0.1 % spontaneous addition. No additional
product was detected in the reactions containing enzyme.
Pixel plots in Figures 3 and 4. The data in Tables S2 and S3 were used to create Figures 3 and 4
4
5
using the approach similar to Wahler et al. To show the wide range of reaction rates, the pixel
darkness is proportional to the log of the rates, but color is linearly proportional to the
1
0
enantioselectivity. Green indicates Rꢀselectivity, red indicates Sꢀselectivity and gray indicates no
enantioselectivity data or no enantioselectivity. The RGB color values for an achiral substrate or
for chiral substrate where enantioselectivity was not measured were: R = G = B = 255 – RATE
–1
where RATE = log (rate in min ) scaled to a whole number between 255 (for the fastest rate
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–1
1
2,600 min for the MeHNLꢀcatalyzed cleavage of acetone cyanohydrin in Table S2) and 0 (for
–1
the slowest rate 0.00023 min for the EST1ꢀcatalyzed hydrolysis of the lactone in Table S3).
The RATE values are subtracted from 255 because color values closer to zero give darker colors.
For Rꢀselective enantioselective reactions, the color values were R = 255 – (E*RATE)/(E+1), G
= 255 – RATE/(E+1), B = (R + G)/2 and for Sꢀselective reactions R = 255 – RATE/(E+1), G =
255 – (E*RATE)/(E+1), B = 255 – (R + G)/2 where E is the enantioselectivity.
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Statistical analysis. Fisher’s exact tests were used to compare the substrate promiscuity and
catalytic promiscuity of ancestral and modern enzymes. First, all enzymesꢀsubstrate
combinations were classified as active or not active. Tables in supporting information.
Classifications for resurrected ancestral enzymes at nodes EST3 and HNL1 were each averaged
and rounded (e.g. a mean of 2/3 active is classified as active and a mean of 1/3 active is
classified as not active). Substrate promiscuity compared the ability of ancestral and modern
enzyme to catalyze their natural reactions ꢀ ester hydrolysis for esterases and hydroxynitrile
cleavage for the HNLs. Catalytic promiscuity compared the ability of the ancestral and modern
enzyme to catalyze unnatural reactions ꢀ hydroxynitrile cleavage for esterases and ester
hydrolysis for HNL’s. The reactions in Table 3 were classified as unnatural reactions for both
esterase and HNL’s.
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Results
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The α/βꢀhydrolaseꢀfold superfamily contains >60,000 proteins
with wideꢀranging
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catalytic activities.
Most are hydrolases (Enzyme Classification group 3), although the family
also includes hydroxynitrile lyases (Enzyme Classification group 4) that catalyze an elimination
reaction.
These hydroxynitrile lyases cluster within a larger group of plant esterases (Figure S1;
selected sequence alignments in Figure S2, also see reference 52) suggesting that they diverged
from esterases ~100 million years ago when flowering plants (angiosperms) diversified.
5
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Hydroxynitrile lyases are plant enzymes involved in defense from insects. The hydroxynitrile
lyases include enzymes from the rubber tree, Hevea brasiliensis, (HbHNL), cassava, Manihot
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esculenta, (MeHNL)
and wild castor, Baliospermum montanum. The esterases include
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8
salicylic acid binding protein 2 (SABP2) from tobacco (Nicotiana tabacum) and polyneuridineꢀ
5
9
aldehyde esterase from snakeroot (Rauvolfia serpentina) (RsEST). This group of plant esterases
and hydroxynitrile lyases share >40% amino acid identity (Table S1 in the supporting
information lists pairwise sequence identities).
60
Although esterase from Arabidopsis thaliana (AtEST) catalyses ester hydrolysis, it also
6
1
catalyzes fast promiscuous cleavage of mandelonitrile. It does not catalyze cleavage of acetone
cyanohydrin, the natural cyanohydrin substrate in most plants. Cruciferous plants, which include
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Arabidopsis, lack the metabolic pathway to make cyanogenic glucosides, so do not need
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hydroxynitrile lyases. However, Arabidopsis does contain other cyanogenic metabolites like an
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acyl cyanide, where a hydrolase may contribute to cyanogenesis and plant defense.
More distantly related α/βꢀhydrolaseꢀfold enzymes include more esterases (e.g. esterase
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from R. communis (RcEST) as well as enzymes that catalyze even more diverse reactions. The
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decarboxylase methylketone synthase I from tomato has 35ꢀ50% aa identity to the HNL/EST
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group
found in various bacteria or animals are C–C hydrolases (e.g., BphD from Burkholderia
in Figure S1. More divergent α/βꢀhydrolaseꢀfold enzymes (only 10ꢀ20% aa identify)
4
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cepacia ), epoxide ring hydrolases (e.g., human soluble epoxide hydrolase
), and haloalkane
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hydrolases (e.g., LinB from Sphingobium sp. ). These are too distantly related to be included in
the tree in Figure S1.
Ancestral enzyme reconstruction. Many of these plant esterases and hydroxynitrile lyase share
>40% amino acid identity making it feasible to infer ancestral enzyme sequences. We predicted
the sequences of the most recent common ancestors at four different divergence points (Figures 2
and S1; sequences in Figure S2). These sequences were inferred from the sequences of the
modern descendant enzymes using either neighbor joining, maximum parsimony, maximum
1
1
likelihood, or a combination of maximum parsimony & maximum likelihood (Table 1). Some
sites, typically at the protein surface, evolve rapidly because they do not affect protein function
and their ancestral composition is less certain. The different algorithms predict slightly different
sequences. If all reconstructed ancestors at a node behave similarly, then we assume that these
reconstructions are a close approximation to the actual historical ancestor.
Figure 2. Simplified phylogenetic tree of esterases and HNL’s identifies different types of ancesꢀ
tral enzymes. Ancestral enzymes EST1 and EST2 are conserved function enzymes. They lie
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along a path from ancient to modern esterases and are expected to be esterases. Ancestral enꢀ
zyme EST3 is a functional branch point enzyme. It lies between ancient esterases and modern
HNL’s and is expected to be an esterase with promiscuous HNL activity. Ancestral enzyme
HNL1 is a transitional ancestral enzyme because it leads only to modern HNL’s. This is expected
to be an HNL, but may show remaining esterase activity.
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Table 1. Extant and ancestral enzymes in this study and their amino acid sequence identity with
a
hydroxynitrile lyase HbHNL and esterase SABP2.
b
Enzyme
Construct Origin
% ID to HbHNL % ID to SABP2
HbHNL
MeHNL
Hevea brasiliensis
Manihot esculenta
100
76
44
41
HNL1
max parsimony & likelihood
maximum likelihood
neighbor joining
79
75
79
49
51
50
HNL1ꢀML
HNL1ꢀNJ
EST3
max parsimony & likelihood
maximum likelihood
neighbor joining
67
58
56
55
60
59
EST3ꢀML
EST3ꢀNJ
max parsimony & likelihood
EST2
48
71
EST1
AtEST
RcEST
RsEST
SABP2
max parsimony & likelihood
Arabidopsis thaliana
Ricinus communis
49
47
21
41
44
70
50
28
Rauvolfia serpentina
Nicotinia tabacum
56
100
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a
Supporting information Table S1 compares the sequence identity for each enzyme pair and
b
Figure S1 shows the aligned amino acid sequences. Plant species origin for extant enzymes and
methods used to derive the ancestral sequences.
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The sequences of the ancestral enzymes differ substantially from the modern
descendants, but these differences are similar to the differences between modern enzymes. For
example, the amino acids in the modern enzyme MeHNL are 76% identical to those in HbHNL;
similarly, the ancestral enzymes at node HNL1 are 72ꢀ79% identical to HbHNL. Modern
esterases are 41ꢀ47% identical to HbHNL; ancestral enzymes at nodes EST1 and EST2 are 48ꢀ
49% identical to HbHNL, while ancestral enzymes at node EST3 are 56ꢀ67% identical to
HbHNL. These differences correspond to large numbers of amino acid substitutions; HNL1 and
HbHNL differ by 54ꢀ72 amino acid substitutions. Such large numbers of substitutions can be
71
expected to substantially change their properties. Nakano and Asano created hybrid HNL’s
using a consensus sequence approach, but these cannot correspond to ancestral enzymes. No
information about the catalytic promiscuity of these hybrids is available.
Ancestral enzymes are branch points in the evolution of modern enzymes. We classify
these branch points into three types as related to divergent evolution. Some ancestral enzymes lie
at points where all connecting branches lead to enzymes with the same catalytic function. These,
which we call conserved function ancestors, are expected to have the ancestral function. Second,
other ancestral enzymes are branch points to younger nodes with different functions. These,
which we call function branch point ancestors, are also expected to have the ancestral function,
since one of the branches retains the ancestral function. However these functional branch points
may have already undergone selection toward both activities, and thus were likely to be
bifunctional or promiscuous for the new function as well as active for the ancestral function. The
third type of ancestral enzyme is the branch point from an older node with the ancestral activity
to a clade of modern enzymes, all with the new function. These, which we call transitional
ancestors, are expected to be specialized for the new function, but may retain vestigial activity
toward the ancestral reaction.
In this classification, EST1 and EST2 are conserved function ancestors, predicted to be
esterases since all descendant branches from theses enzymes contain esterases. EST3 is a branch
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point ancestor, also predicted to be an esterase, likely with promiscuous HNL activity, since
some descendant enzymes are esterases (ancestral function) and some are hydroxynitrile lyases.
Finally, HNL1 is a transitional ancestor, since all modern descendants are HNL’s, and the node
immediately ancestral to HNL1, (i.e. EST3) is predicted to be an esterase. HNL1 is predicted to
be an HNL like its descendants, yet to possibly retain some of the esterase activity of its ancesꢀ
tors.
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Esterase & hydroxynitrile lyase activities of modern enzymes. All predicted ancestral enzyme
sequences conserve the catalytic triad of SerꢀHisꢀAsp found in modern descendants. The
ancestral enzymes also contain active site differences that match their modern descendants.
Esterases contain a glycine on the oxyanion loop that allows access to the oxyanion hole.
Ancestral enzymes EST1 and EST2 also contain a glycine at this location. Modern
hydroxynitrile lyases contain a threonine as this location; ancestral enzymes at HNL1 also
contain a threonine. Esterases contain a hydrophobic site to bind the alcohol (often phenylalanine
or histidine, which would be uncharged with a non polar region), while HNL's contain a polar
site – a lysineꢀglutamate pair. The ancestral enzymes EST1 and EST2 contain histidine, while
enzymes at HNL1 contain the lysineꢀglutamate pair. EST3 is an exception to this generalization.
Three of the four reconstructions contain an asparagine on the oxyanion loop, suggesting an
HNLꢀlike block of the oxyanion hole, but nonꢀpolar residues in the alcohol binding site suggest
an ESTꢀlike active site. The modern enzyme AtEST has a similar mix of residues in the active
site. Based upon these sequence differences, we expect HNL1 to be a hydroxynitrile lyase, EST1
and EST2 to be esterases, but the function of EST3 is not easily predicted from its sequence.
Since it is most similar to AtEST, it may be, like AtEST, an esterase with a promiscuous
hydroxynitrile lyase activity.
Since the substrate specificities of the various enzymes are unknown, we used six esters
with varying structure to measure the carboxylic acid esterase activities (E.C. group 3.1.1):
methyl salicylate, methyl mandelate, 1ꢀ and 2ꢀnaphthyl acetate, methyl pentanoate and 4ꢀphenylꢀ
4ꢀbutyrolactone. We omitted the oftenꢀused pꢀnitrophenyl acetate because its high reactivity
makes it different from typical esters. Similarly, we used six substrates of varying structure to
measure aldehyde lyase activity (E.C. group 4.1.2): acetone cyanohydrin (natural substrate for
hydroxynitrile lyases), mandelonitrile, lactonitrile, 2ꢀhydroypentanenitrile, 2ꢀ
hydroxyhexanenitrile and 2ꢀnitroꢀ1ꢀphenylethanol. This last substrate involves a nitroꢀaldol
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cleavage, whose mechanism appears similar to that for cyanohydrin cleavage. The modern
esterases catalyzed hydrolysis of almost all of the ester substrates and similarly, the modern
hydroxynitrile lyases catalyzed cleavage of almost all the hydroxynitrile lyase substrates. Thus,
this range of substrate cannot test for increased substrate promiscuity since the ancestral enzymes
cannot do more than the modern enzyme with this set of substrates.
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The modern enzymes showed little catalytic promiscuity; that is, the hydroxynitrile lyases
catalyzed mainly hydroxynitrile cleavage and the modern esterases catalyzed mainly ester
hydrolysis, Figure 3, Table S2. The modern hydroxynitrile lyases, HbHNL and MeHNL,
catalyzed cleavage of all five cyanohydrins and the nitroaldol compound. The best substrate is
–1
the natural one, acetone cyanohydrin, with k values of 2,400 and 12,600 min , respectively.
cat
–
They also efficiently cleaved their unnatural substrate mandelonitrile (k = 1530 and 1340 min
cat
1
73, 74
, respectively) with high enantioselectivity for the (S)ꢀenantiomer, as previously reported.
These two HNL’s catalyzed the slower cleavage of three other cyanohydrins and the nitroaldol
72, 75ꢀ77
–1
compound
with k values ranging from 0.3 to 50 min . The enantioselectivity of the
cat
nitroaldol reaction was measured in the synthesis direction. Both MeHNL and HbHNL favored
the (S)ꢀenantiomer, as was reported previously.
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Figure 3. Aldehyde lyase and ester hydrolase activities of modern and ancestral hydroxynitrile
lyases and esterases. Columns correspond to different enzymes, while rows correspond to the
reaction with the substrate at the left. Modern esterases catalyze mainly hydrolysis of esters and
modern lyases catalyze mainly elimination reactions, while ancestral enzymes often catalyze
both reactions. Darker squares correspond to faster reaction. For enantioselective reactions, green
indicates Rꢀselective reactions and red Sꢀselective reactions. Color intensity reflects enantioselecꢀ
tivity. Supporting information Table S2 contains the data from which this figure was derived.
The modern hydroxynitrile lyases showed little promiscuous esterase activity. HbHNL
–1
catalyzed the slow hydrolysis of methyl salicylate, 0.07 min while MeHNL showed no
detectable esterase activity.
The modern esterases all catalyzed ester hydrolysis. SABP2 catalyzed hydrolysis of all
six esters tested. The natural function of SABP2 is to hydrolyze methyl salicylate to salicylic
–
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acid with a reported k value of 27 min and K of 8.6 µM. We measured a lower value of
cat
M
–1
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conditions. Among the other five esters, the best substrate was methyl pentanoate, with a rate of
–1
1
8,000 min . RcEST is a predicted, but not experimentally confirmed, polyneuridine aldehyde
esterase from sequence similarities. RsEST is experimentally confirmed as a polyneuridine
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aldehyde esterase, but closely related analogs of polyneuridine aldehyde did not react. Both
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–1
esterases catalyzed hydrolysis of all five esters tested, but some rates were as low as 0.01 min .
–
1
The good ester substrates for RcEST were methyl salicylate (7.5 min ) and methyl pentanoate
–
1
–
(260 min ), while the good esterase substrate for RsEST was only methyl pentanoate (650 min
1
–1
–1
). RsEST also catalyzed very slow hydrolysis of the lactone (0.00077 min or 1 d ). AtEST
–1
catalyzed hydrolysis of three of the five esters with rates ranging from 0.04 to 1.7 min and the
–1
–1
very slow hydrolysis of the lactone (0.0005 min or 0.7 d ) confirming its assignment as an
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esterase, albeit not an efficient one. Koo et al. also reported the AtEST catalyzes hydrolysis of
esters. Neither SABP2, RcEST, or RsEST catalyzed cleavage of any of the hydroxynitriles or of
the nitrolaldol substrate. As reported previously, AtEST is unusual in that it showed efficient
–1
cleavage of mandelonitrile (2,530 min ) and the slow cleavage of the similar nitroaldol
–
1
compound (5 min ). In both cases, the (R)ꢀenantiomer was cleaved in contrast to the (S)ꢀ
enantiomer favored by HbHNL and MeHNL.
Esterase & hydroxynitrile lyase activities of ancestral enzymes. In contrast, almost all of the
ancestral enzymes catalyzed both an ester hydrolysis and a cyanohydrin cleavage, Figure 3,
Table S2. At the HNL1 node, all three reconstructions catalyzed both cyanohydrin cleavage and
ester hydrolysis. Cyanohydrins were better substrates than esters and acetone cyanohydrin is the
best substrate. HNL1 has detectable activity with nine of the twelve substrates; HNL1ꢀML with
ten substrates and HNL1ꢀNJ with eleven substrates. HNL1 showed the highest cyanohydrin
–1
cleavage activity (880 min with acetone cyanohydrin) and HNL1ꢀNJ showed the highest ester
–1
hydrolysis activity (0.9 min with methyl pentanoate).
Ancestral esterases, EST1, EST2, and one of the three reconstructions at node EST3
(EST3ꢀML) also catalyzed both cyanohydrin cleavage and ester hydrolysis. EST1 and EST2
–1
show high activity (>1 min ) toward the majority of ester substrates, whereas the modern
esterases have high activity with only one or two of the substrates. Both EST1 and EST2 also
have activity with mandelonitrile and, other than AtEST, the modern esterases had no detectable
hydroxynitrile cleavage activity. The best substrate for EST3ꢀML was mandelonitrile, not an
–
1
ester. Ester hydrolysis was very slow: 0.008 min for EST3ꢀML. The other two ancestral
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enzyme reconstructions at EST3 catalyzed only ester hydrolysis. Methyl pentanoate was the best
–
1
–1
substrate: 42 min for EST3ꢀNJ; only 0.6 min for EST3. Chemical steps are rateꢀlimiting for
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both esterases and hydroxynitrile lyases. We assume that mechanisms of the ancestral
enzymes are similar to those of modern descendants and that chemical steps are also rateꢀ
limiting.
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Statistical comparison with Fisher’s exact test indicates that these ancestral and modern
enzymes do not differ significantly in their substrate promiscuity, but do differ in their catalytic
promiscuity. The 2x2 Fisher exact test with the natural reactions (ester hydrolysis catalyzed by
esterases and hydroxynitrile cleavage by HNL’s) is not significant (P = 0.67) indicating that anꢀ
cestral enzymes are no more substrate promiscuous than their modern descendants. While the
ancestral enzymes accepted almost all of the substrates, so did the modern enzymes. The ancesꢀ
tral enzymes were not given the opportunity to demonstrate substrate promiscuity with this selecꢀ
tion of substrates. Further testing with a wider range of substrate might show increased substrate
promiscuity of these enzymes as has been observed for other ancestral enzymes. The 2x2 Fisher
exact test with the unnatural reactions (ester hydrolysis catalyzed by HNL’s and hydroxynitrile
cleavage catalyzed by esterases) is highly significant (P < 0.01) indicating that ancestral enzymes
are more likely to be catalytically promiscuous than modern enzymes.
Among the ancestral enzymes, HNL1 is a transitional enzyme between true esterases and
true HNLs. It shows good HNL activity while retaining some esterase activity. It can best cataꢀ
lyze both hydrolysis and elimination reactions. The other ancestors (EST1, EST2, EST3) were,
in their time, modern specialist esterases. A statistical comparison of HNL1 to all other enzymes
shows that HNL1 is more catalytically promiscuous than the group of modern and other ancestral
enzymes (P = 0.012). The enhanced catalytic promiscuity of HNL1 does not come at the cost of
stability. Its unfolding temperature (~80 °C) is higher than that for the modern descendants
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HbHNL and MeHNL (54ꢀ70 °C).
Other researchers also found increased stability for many
8
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ancestral enzymes.
Other hydrolase and lyase activities of modern and ancestral enzymes. To test the ability of
these enzymes to catalyze an even broader range of reactions, we tested two other hydrolyses
(lactam hydrolysis, EC 3.5.2, and carbonꢀcarbon bond hydrolysis, EC 3.7.1) and three other lyase
reactions (decarboxylation, EC 4.1.1, Michael addition, EC 4.1.99) Figure 4, Table S3. All four
are reactions catalyzed by enzymes in the α/βꢀhydrolaseꢀfold family, but the branch points for
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these enzymes occur outside the region of the reconstructed ancestral enzymes. We refer to these
as outside reactions. Two of the modern esterases catalyzed a slow hydrolysis reaction (SABP2,
–1
–1
–1
0.003 min for lactam hydrolysis; RsEST, 0.0004 min or 0.5 d , for carbonꢀcarbon bond
hydrolysis), but none of the remaining four modern enzyme catalyzed any of these four
reactions. In contrast, seven of the ten ancestral enzymes catalyzed at least one of the reactions.
EST2 catalyzed two reactions, EST3ꢀNJ and HNL1ꢀNJ catalyzed three reactions. The reaction
rates were all slow; the fastest reaction was the EST3ꢀcatalyzed hydrolysis of the carbonꢀcarbon
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–
1
–1
bond (0.03 min or ~40 d ). Both modern and ancestral enzymes were enantioselective.
Figure 4. Other eliminations and hydrolyses catalyzed by modern and ancestral hydroxynitrile
lyases and esterases. Columns correspond to different enzymes, while rows correspond to reacꢀ
tions with the substrate at the left. Only two of the six modern enzymes catalyze one of these reꢀ
actions, while seven of the ten ancestral enzymes catalyze at least one of these reactions. Darker
squares correspond to faster reaction. For enantioselective reactions, green indicates Rꢀselective
reactions and red Sꢀselective reactions. Color intensity reflects enantioselectivity. Supporting inꢀ
formation Table S3 contains the data from which this figure was derived. The substrates for these
reaction were: decarboxylation, 3ꢀoxoꢀ3ꢀphenylpropanoic acid; Michael addition, acetylacetone
and 2ꢀnitrovinylbenzene; CꢀC hydrolysis, 2ꢀhydroxyꢀ6ꢀoxoꢀ6ꢀphenylhexaꢀ2,4ꢀdienoic acid; lacꢀ
tam hydrolysis, 2ꢀazabicylo[2.2.1]heptꢀ5ꢀenꢀ3ꢀone (Vince lactam).
The 2x2 Fisher’s exact test indicates that the ancestral enzymes are more likely (P = 0.04) to
catalyze one of these mechanistically related reactions than is a modern descendant even though
they were never subject to selection for this ability. Ancestral enzymes HNL1 and EST3 are tranꢀ
sitional and functional branch point enzymes for HNL activity, but not for these novel reactions,
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so this catalytic promiscuity is accidental. Nevertheless, ancestral enzymes are more likely than
modern enzymes to possess these accidental abilities.
None of the ancestral enzymes or modern enzymes catalyzed epoxide hydrolysis
–1
(
hydrolysis of 2ꢀ(4ꢀnitrophenyl)oxirane; <0.0001 min ). This result is not surprising since the
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expoxide hydrolase mechanism requires an aspartate nucleophile, while all the ancestral and
modern enzymes contain a serine as the nucleophile in the active site. None of the ancestral
enzymes or modern enzymes catalyzed the aldol addition of acetone to pꢀnitrobenzaldehyde or
the BaylisꢀHillman reaction, addition of methyl vinyl ketone to pꢀnitrobenzaldehdye (<0.0001
–1
min ). We tested the aldol and BaylisꢀHillman reactions because catalysis involves an amine
and HNL’s contain a lysine residue near the active site. No α/βꢀhydrolaseꢀfold enzyme naturally
catalyzes these additions, but two groups reported weak promiscuous activity of α/β hydrolaseꢀ
8
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fold enzymes toward these reactions.
Discussion
Ancestral enzymes at branch points for new catalytic activities are good places to search
for catalytic promiscuity. Random variation can create promiscuity both in modern and ancestral
enzymes, but along branches for new catalytic function, there is an expectation of catalytic promꢀ
iscuity and our results support this expectation. Modern catalytically promiscuous enzymes also
exist, but there is no systematic method to find them. In contrast, examination of phylogenetic
trees can identify which ancestral enzymes are most like to be promiscuous.
The major catalytic behavior of ancestral enzymes is consistent with our understanding of
the hydrolysis versus lyase reaction mechanisms. The ancestral esterases, EST1, EST2, and
EST3 have esteraseꢀlike active sites and are efficient hydrolases. The ancestral hydroxynitrile
lyases at HNL1 have a hydroxynitrileꢀlyaseꢀlike active site and are efficient lyases. They also
show significant esterase activity. We did not find such catalytic promiscuity when we engiꢀ
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neered esterase variants for HNL activity. Site directed mutagenesis added three catalytic HNL
residues to the esterases PFE and SABP2. In both cases, esterase activity was abolished. In the
case of PFE, no HNL activity was detected, while for SABP2 the HNL activity was weak (0.09
–1
–1
min vs. 60ꢀ170 min for HNL1). Other amino acid must be responsible for the higher HNL
activity of HNL1 and its ability to catalyze both reaction types.
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Substrate promiscuity has been correlated with protein flexibility. Ubiquitin is a small
protein (76 amino acids) that binds to many different proteins as part of its regulatory function.
In solution and in the absence of protein ligands, ubiquitin is flexible and the range of observed
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conformations matches those seen in the crystal structures. Molecular dynamics simulations of
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glutathione Sꢀtransferases and cytochrome P450 monooxygenases correlated higher flexibilꢀ
ity with broader substrate specificity. As substrates bind, they stabilize the conformation that
creates the best contacts. For example, xꢀray structures of an enzyme that catalyzes an isomerizaꢀ
tion in both histidine and tryptophan biosynthesis shows different active site conformations when
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bound to the two different substrates. In another example, xꢀray structures of two amino glycoꢀ
side antibiotics bound to a promiscuous kinase show different loop conformation for the two anꢀ
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tibiotics.
Conformational flexibility may also contribute to catalytic promiscuity. Chemically difꢀ
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ferent reactions likely proceed via different protein conformations. The increase in the flexibilꢀ
94
ity of a loop in a lactonase variant correlated with increased catalytic promiscuity. As the subꢀ
strate orientations for the hydrolysis and lyase reactions differ, so an enzyme that catalyzes both
reactions must bind the two substrates in at least two different orientations. Although the details
are unknown, conformational flexibility of the ancestral enzymes may also account for their cataꢀ
lytic promiscuity.
Whatever the mechanism generating catalytic promiscuity it likely involves residues outꢀ
side the active sites. The modern and ancestral enzymes differ in approximately fifty residues,
only a few of which are located in the active site. Distant residues can change the binding affinity
of proteins. For example, the protein kinases Abl and Src differ in their affinity for the inhibitor
16
Gleevec. The difference stems from the shift of a loop in Abl to wrap around Gleevec. In conꢀ
trast a hydrogen bond network involving distant residues prevents a similar shift in Src. Distant
residues might similarly create several possible catalytic conformations.
Associated Content
Supporting Information
Cladogram showing the hydroxynitrile lyases within a larger group of plant esterases in the α/βꢀ
hydrolase fold superfamily. Amino acid sequence alignment of selected modern and reconstructꢀ
ed ancestral α/βꢀhydrolases. Pairwise sequence identities of modern and ancestral enzymes. Cyaꢀ
nohydrin cleavage and ester hydrolysis catalyzed by modern and ancestral enzymes. Rates and
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enantioselectivity of other nucleophilic additions and other hydrolyses catalyze by modern and
ancestral hydroxynitrile lyases and esterases. Data (conversion and enantiomeric excess) to
measure the enantioselectivity of modern and ancestral enzymes. Representative HPLC chromaꢀ
tograms to measure enantioselectivity of modern and ancestral enzymes. Categorial grouping of
data from Tables S2 and S3 for Fisher’s exact test. This material is available free of charge via
the Internet at http:// pubs.acs.org.
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Author Information
Corresponding Authors
rjk@umn.edu; deanx024@umn.edu; dean@mail.sysu.edu.cn
Author Contributions
T. D. and A. M. R. contributed equally. All authors approved the final version of the manuscript.
Notes
The authors declare no competing financial interest.
Acknowledgements
This work was supported by NIH grant 5R01GM102205 and NSF grant 1152804 to AMD and
RJK.
References
1
. O’Brien, P. J.; Herschlag, D. Chem. Biol. 1999, 6, R91.
. Humble, M. S.; Berglund, P. Eur. J. Org. Chem. 2011, 3391.
2
3. Pocker, Y.; Stone, J. T. Biochemistry 1967, 6, 668.
4. Bornscheuer, U. T.; Kazlauskas, R. J. Angew. Chem. Intl. Ed. 2004, 43, 6032.
5. Nakamura, T.; Nagasawa, T.; Yu, F.; Watanabe, I.; Yamada, H. Biochem. Biophys. Res.
Commun. 1991, 180, 124.
6. Fox, R. J.; Davis, S. C.; Mundorff, E. C.; Newman, L. M.; Gavrilovic, V.; Ma, S. K.; Chung,
L. M.; Ching, C.; Tam, S.; Muley, S.; Grate, J.; Gruber, J.; Whitman, J. C.; Sheldon, R. A.;
Huisman, G. W. Nat. Biotechnol. 2007, 25, 338.
7
. Waley, S. G. Comput. Biochem. Physiol. 1969, 30, 1.
. Ycas, M. J. Theor. Biol. 1974, 44, 145.
8
9. Jensen, R. A. Annu. Rev. Microbiol. 1976, 30, 409.
0. Weng, J.ꢀK.; Philippe, R. N.; Noel, J. P. Science 2012, 336, 1667.
1
2
6
ACS Paragon Plus Environment

Page 27 of 36
Journal of the American Chemical Society
1
2
3
4
5
6
7
8
9
1
1. Thornton, J. W. Nat. Rev. Genet. 2004, 5, 366.
2. Harms, M. J.; Thornton, J. W. Nat. Rev. Genet. 2013, 14, 559.
1
13. Voordeckers, K.; Vanneste, K.; van der Zande, E.; Voet, A.; Maere, S.; Verstrepen, K. J.
PLoS Biol. 2012, 10, e1001446.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
1
4. Risso, V.; Gavira, J.; MejiaꢀCarmona, D. F.; Gaucher, E.; SanchezꢀRuiz, J. M. J. Am. Chem.
Soc. 2013, 135, 2899.
15. Huang, R.; Hippauf, F.; Rohrbeck, D.; Haustein, M.; Wenke, K.; Feike, J.; Sorrelle, N.;
Piechulla, B.; Barkman, T, J. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 2966.
1
6. Wilson, C.; Agafonov, R. V.; Hoemberger, M.; Kutter, S.; Zorba, A.; Halpin, J.; Buosi, V.;
Otten, R.; Waterman, D.; Theobald, D. L.; Kern, D. Science 2015, 347, 882.
7. Poultan, J. E. Plant Physiol. 1990, 94, 401.
1
18. Gasteiger, E.; Hoogland, C.; Gattiker, A.; Duvaud, S.; Wilkins, M. R.; Appel, R. D.; Bairoch
A. in The Proteomics Protocols Handbook, Walker, J. G, Ed., Humana Press: Totowa, NJ, 2005,
5
71.
9. Sambrook, J.; Fritsch, E. F.; Maniatis, T. Molecular Cloning: A Laboratory Manual, 2 ed.
nd
1
Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, 1989.
2
2
2
2
0. Edgar, R. C. Nucl. Acids. Res. 2004, 32, 1792.
1. Gouy, M.; Guindon, S.; Gascuel, O. Mol. Biol. Evol. 2010, 27, 221.
2. Saitou, N.; Nei, M. Mol. Biol. Evol. 1987, 4, 406.
3. Guex, N.; Peitsch, M. C. Electrophoresis 1997, 18, 2714.
24. Stamatakis, A. Bioinformatics 2006, 22, 2688.
2
2
2
5. Le, S. Q.; Gascuel, O. Mol. Biol. Evol. 2008, 25, 1307.
6. Jones, D. T.; Taylor, W. R.; Thornton, J. M. Comput. Applic. Biosci. 1992, 8, 275.
7. Shimodaira, H.; Hasegawa, M. Mol. Biol. Evol. 1999, 16, 1114.
28. Yang, Z.; Kumar, S.; Nei, M. Genetics 1995, 141, 1641.
2
2
3
2
9. Tamura, K.; Stecher, G.; Peterson, D.; Filipski, A.; Kumar, S. Mol. Biol. Evol. 2013, 30,
725.
0. Yuryev, R.; Briechle, S.; GruberꢀKhadjawi, M.; Griengl, H.; Liese, A. ChemCatChem 2010,
, 981.
31. Ballini, R.; Maggi, R.; Palmieri, A.; Sartori. G. Synthesis 2007, 19, 3017.
2. Liu, L.; Zou, Y. J. Appl. Polym. Sci. 2012, 123, 554.
3
2
7
ACS Paragon Plus Environment

Journal of the American Chemical Society
Page 28 of 36
1
2
3
4
5
6
7
8
9
1
1
1
1
1
1
1
1
1
1
2
2
2
2
2
2
2
2
2
2
3
3
3
3
3
3
3
3
3
3
4
4
4
4
4
4
4
4
4
4
5
5
5
5
5
5
5
5
5
5
6
3
3. Martin, S. J. Am. Chem. Soc. 1959, 89, 2598.
4. von Langermann, J.; Nedrud, D. M.; Kazlauskas, R. J. ChemBioChem 2014, 15, 1931.
3
35. Wolfsie, J. H.; Sheffer, C. B. J. Occup. Med. 1959, 281.
36. Selmar, D.; Carvalho, F. J. P.; Conn, E. E. Anal. Biochem. 1987, 166, 208.
37. Andexer, J.; Guterl, JꢀK.; Pohl, M.; Eggert, T. Chem. Commun. 2006, 40, 4201.
38. Fuhshuku, K.; Asano, Y. J. Biotech. 2011, 153, 153.
0
1
2
3
4
5
6
7
8
9
0
1
2
3
4
5
6
7
8
9
0
1
2
3
4
5
6
7
8
9
0
1
2
3
4
5
6
7
8
9
0
1
2
3
4
5
6
7
8
9
0
39. Bernhardt, P.; Hult, K.; Kazlauskas, R. J. Angew. Chem. Int. Ed. Engl. 2005, 44, 2742.
40. Janes, L. E.; Lӧwendahl, A. C.; Kazlauskas, R. J. Chem. Eur. J. 1998, 4, 2324.
4
4
4
1. Barton, P.; Page, M. I. J. Chem. Soc. Perkin. Trans. 1993, 2, 2317.
2. Corric, I.; Muller, S.; List, B. J. Am. Chem. Soc. 2010, 132, 17370.
3. Kasaplar, P.; Riente, P.; Hartmann, C.; Periquas, M. A. Adv. Synth. Catal. 2012, 354, 2905.
44. Seah, S. Y. K.; Labbé, G.; Nerdinger, S.; Johnson, M. R.; Snieckus, V.; Eltis, L. D. J. Biol.
Chem. 2000, 275, 15701.
4
5. Wahler, D.; Badalassi, F.; Crotti, P.; Reymond, J.ꢀL. Chem. Eur. J. 2002, 8, 3211.
6. Agresti, A. A. Stat. Sci. 1992, 7, 131.
4
47. Lenfant, N.; Hotelier, T.; Velluet, E.; Bourne, Y.; Marchot, P.; Chatonnet, A. Nucl. Acid.
Res. 2013, 41, D423.
4
8. Kourist, R.; Jochens, H.; Bartsch, S.; Kuipers, R.; Padhi, S. K.; Gall, M. G.; Böttcher, D.;
Joosten, H.ꢀJ.; Bornscheuer, U. T. ChemBioChem 2010, 11, 1635.
9. Ollis, D.; Cheah, E.; Cygler, M.; Dijkstra, B. W.; Frolow, F.; Franken, S. M.; Harel, M.;
Remington, S.; Silman, I.; Schrag, J. D. Protein Eng. Des. Sel. 1992, 5, 197.
4
5
5
5
0. Nardini, M.; Dijkstra, B. W. Curr. Opin. Struct. Biol. 1999, 9, 732.
1. Holmquist, M. Curr. Prot. Pept. Sci. 2000, 1, 209.
2. Rauwerdink, A. M.; Lunzer, M.; Devamani, T.; Jones, B. Mooney, J.; Zhang, Z.ꢀJ.; Xu, J.ꢀ
H.; Kazlauskas, R. J.; Dean, A. M. Mol. Biol. Evol. 2016, 33, 000.
http://doi.org/10.1093/molbev/msv338
5
5
5
3. Hickel, A.; Hasslacher, M.; Griengl, H. Physiol. Plant. 1996, 98, 891.
4. Selmar, D.; Lieberei, R.; Biehl, B.; Conn, E. E. Physiol. Plant. 1989, 75, 97.
5. Gruber, K.; Gartler, G.; Krammer, B.; Schwab, H.; Kratky, C. J. Biol. Chem. 2004, 279,
20501.
6. Hughes, J.; Carvalho, F. J. P.; Hughes, M. A. Arch. Biochem. Biophys. 1994, 311, 469.
5
2
8
ACS Paragon Plus Environment

Page 29 of 36
Journal of the American Chemical Society
1
2
3
4
5
6
7
8
9
5
7. Dadashipour, M.; Yamazaki, M.; Momonoi, K.; Tamura, K.; Fuhshuku, K.; Kanase, Y.;
Uchimura, E.; Kaiyun, G.; Asano, Y. J. Biotechnol. 2011, 153, 100.
58. Forouhar, F.; Yang, Y.; Kumar, D.; Chen, Y.; Fridman, E.; Park, S. W.; Chiang, Y.; Acton,
T. B.; Montelione, G. T.; Pichersky, E., Klessig, D. F.;Tong, L. Proc. Natl. Acad. Sci. U. S. A.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
2
005, 102, 1773.
59. MatternꢀDogru, E.; Ma, X.; Hartmann, J.; Decker, H.; Stӧckigt, J. Eur. J. Biochem. 2002,
269, 2889.
60. Koo, Y. J.; Yoon, E. S.; Seo, J. K.; Kim, J.ꢀK.; Choi, Y. D. J. Korean Soc. Appl. Biol. Chem.
2
013, 56, 27.
6
1. Andexer J. N.; von Langermann, J.; Mell, A.; Bocola, M.; Kragl, U.; Eggert, T.; Pohl, M.
Angew. Chem. Int. Ed. Engl. 2007, 46, 8679.
62. Tattersall, D. B.; Bak, S.; Jones, P. R.; Olsen, C. E.; Nielsen, J. K.; Hansen, M. L.; Høj, P. B.;
Møller, B. L. Science 2001, 293, 1826.
6
3. Rajniak, J.; Barco, B.; Clay, N. K.; Sattely, E. S. Nature 2015, 525, 376.
6
4. Chan, A. P.; Crabtree, J.; Zhao, Q.; Lorenzi, H.; Orvis, J.; Puiu, D.; MelakeꢀBerhan, A.;
Jones, K. M.; Redman, J.; Chen, G.; Cahoon, E. B.; Gedil, M.; Stanke, M.; Haas, B. J.;
Wortman, J. R.; FraserꢀLiggett, C. M.; Ravel, J.; Rabinowicz, P. D. Nat. Biotechnol. 2010, 28,
9
51.
6
5. Yu, G.; Nguyen, T. T.; Guo, Y.; Schauvinhold, I.; Auldridge, M. E.; Bhuiyan, N.; BenꢀIsrael,
I.; Iijima, Y.; Fridman, E.; Noel, J. P.; Pichersky, E. Plant Physiol. 2010, 154, 67–77.
66. Auldridge, M. E.; Guo, Y.; Austin, M. B.; Ramsey, J.; Fridman, E.; Pichersky, E.; Noel, J. P.
Plant Cell. 2012, 24, 1596.
6
7. Yu, Z.; Xu, F.; Huse, L. M.; Morisseau, C.; Draper, A. J.; Newman, J. W.; Parker, C.;
Graham, L.; Engler, M. M.; Hammock, B. D.; Zeldin, D. C.; Kroetz, D. L. Mol. Med. 2013, 87,
992.
6
8. Thalji, R. K.; McAtee, J. J.; Belyanskaya, S.; Brandt, M.; Brown, G. D.; Costell, M. H.;
Ding, Y.; Dodson, J. W.; Eisennagel, S. H.; Fries, R. E.; Gross, J. W.; Harpel, M. R.; Holt, D. A.;
Israel, D. I.; Jolivette, L. J.; Krosky, D.; Li, H.; Lu, Q.; Mandichak, T.; Roethke, T.;
Schnackenberg, C. G.; Schwartz, B.; Shewchuk, L. M.; Xie, W.; Behm, D. J.; Douglas, S. A.;
Shaw, A. L.; Marino, J. P., Jr. Bioorg. Med. Chem. Lett. 2013, 23, 3584.
6
9. Beetham, J. K.; Tian, T. G.; Hammock, B. D. Arch. Biochem. Biophys. 1993, 305, 197.
2
9
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