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Catalytic Promiscuity in Biocatalysis Using Old Enzymes to Form New Bonds and Follow New Pathways.

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Minireviews
U. T. Bornscheuer and R. J. Kazlauskas
Reaction Specificity of Enzymes
Catalytic Promiscuity in Biocatalysis: Using Old
Enzymes to Form New Bonds and Follow New Pathways
Uwe T. Bornscheuer* and Romas J. Kazlauskas*
Keywords:
biosynthesis · biotransformations · enzyme catalysis ·
molecular modeling · proteins
Biocatalysis has expanded rapidly in the last decades with the
discoveries of highly stereoselective enzymes with broad substrate
specificity. A new frontier for biocatalysis is broad reaction specificity,
where enzymes catalyze alternate reactions. Although often underappreciated, catalytic promiscuity has a natural role in evolution and
occasionally in the biosynthesis of secondary metabolites. Examples of
catalytic promiscuity with current or potential applications in synthesis
are reviewed here. Combined with protein engineering, the catalytic
promiscuity of enzymes may broadly extend their usefulness in
organic synthesis.
1. Introduction
The realization that many enzymes have broad substrate
specificity fueled much of the growth in biocatalysis over the
last twenty years, especially in organic synthesis. Identifying a
few enzymes that show high stereoselectivity toward a broad
range of synthetically useful molecules enabled organic
chemists to rapidly develop new synthetic applications for
these enzymes.
One current frontier for biocatalysis is reaction specificity.
Can a single active site catalyze more than one distinct
chemical transformation? Can small changes in the active site
enable new chemistry in that active site? Over the last few
years evidence has mounted that such catalytic promiscuity
exists not just among a few enzymes but is rather common.[1]
This Minireview will focus on catalytic promiscuity related to
biocatalysis—enzyme-catalyzed reactions that are or might be
[*] Prof. Dr. U. T. Bornscheuer
Institute of Chemistry and Biochemistry
Department of Technical Chemistry and Biotechnology
Greifswald University
Soldmannstrasse 16, 17487 Greifswald (Germany)
Fax: (+ 49) 3834-86-80066
E-mail: uwe.bornscheuer@uni-greifswald.de
Prof. Dr. R. J. Kazlauskas
University of Minnesota
Department of Biochemistry, Molecular Biology and Biophysics
1479 Gortner Avenue, 174 Gortner Lab, St. Paul, MN 55108 (USA)
Fax: (+ 1) 612-625-5780
E-mail: rjk@umn.edu
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useful in organic synthesis. We will give
examples of individual proteins with
several catalytic abilities and also examples in which small changes (typically metal-ion substitutions or sitedirected mutagenesis) introduce new catalytic activity. The
most successful examples are carbon–carbon bond-forming
reactions, oxidations catalyzed by hydrolytic enzymes, and
glycosyl transfer reactions.
2. Classifying Catalytic Promiscuity
Catalytic promiscuity in enzymes is the ability of enzyme
active sites to catalyze distinctly different chemical transformations. The chemical transformations may differ in the
functional group involved, that is, the type of bond formed or
cleaved during the reaction and/or may differ in the catalytic
mechanism or path of bond making and breaking. Most
examples of catalytic promiscuity include both changes. For
example, adding a vanadium ion to a phosphatase converts it
into an oxidase capable of the enantioselective oxidation of
sulfides (this is discussed in more detail in Section 4.2). These
two reactions involve different functional groups—breaking
the O O bond in hydrogen peroxide instead of the P O bond
in a phosphate ester. In addition, the reaction mechanism
differs significantly because one reaction is a hydrolysis while
the other is an oxidation.
In Figure 1 catalytic promiscuity is organized according to
differences in the functional groups involved and differences
in the mechanisms of catalysis (for selected examples see
Table 1). The placement of a reaction on this graph is
subjective because the degree of similarity of functional
groups and reaction pathways, whose details are likely
unknown, is a subjective judgment. Nevertheless, this classi-
DOI: 10.1002/anie.200460416
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Pyruvate kinase, which catalyzes phosphoryl group
transfer, can also catalyze sulfuryl group transfer.[9]
Some phosphatases also catalyze sulfate ester
hydrolysis.[10] Enzyme inhibitors such as phosphonate inhibitors of serine hydrolases will not be
included because they rarely involve a complete
catalytic turnover.
Other reaction classes must involve changes in
mechanism. For example, removal of a catalytically
essential amino acid residue dramatically slows the
reaction but does not eliminate it. The remaining,
less efficient reaction must follow a different path.
For example, a Cys-to-Asp mutation in a phosphatase retains some activity.[11] Another group of
reactions involve substrate-assisted catalysis, where
the only substrates converted are those that restore
the missing functional group so that it can actively
participate in catalysis.[12] In some reactions stereochemistry reveals alternate paths. For example,
epoxide hydrolase converts one enantiomer by
inversion of the stereocenter, while the other reacts
by retention.[13] These pathways must differ, even
though the details are not clear.
Figure 1. Classifying catalytic promiscuity. Catalytic promiscuity may involve reacBinding proteins can sometimes catalyze reaction of a different functional group, a change in the catalytic mechanism, or both.
The balloons indicate the types of catalytic promiscuity labeled in italics. Table 1
tions, which is clearly a change in mechanism—
lists selected examples, a few of which are also included in this figure.
from no bond breaking to some bond breaking. For
example, bovine serum albumin catalyzes Kemp
elimination, b-elimination of 3-ketobutyl umbelliferyl ethers (a useful reaction in enzyme assays),[14] and
fication helps distinguish different types of catalytic promiscuity.
moderately enantioselective oxidation of amines to amine
The largest group of reactions involves functional group
oxides with sodium periodate.[15] Myoglobin (an oxygenanalogues. For example, many proteases also catalyze ester
carrying iron heme protein) catalyzes slow oxidation in the
hydrolysis. The bonds broken in the two cases (C N vs. C O)
presence of hydrogen peroxide.[16] Site-directed mutagenesis
differ, but the catalytic mechanism is likely very similar.
to shift the position of the distal histidine (Leu29His/
Several metalloproteases also catalyze the hydrolysis of P
His64Leu) increased the rate of reaction more than 20-fold
O[2] or P F[3] bonds. Conversely, several esterases cleave the
and enantioselectivity significantly. Oxidation of thioanisole
yielded the sulfoxide with 97 % ee, and oxidation of cis-bC N bond in b-lactams,[4] and proteases can cleave the S O[5]
methylstyrene gave the epoxide in 99 % ee.[17]
in sulfites or the S N bond in sulfinamides.[6] On a commercial scale, BASF uses a lipase, which normally cleaves C O
All catalytic antibodies are examples of binding molecules
bonds in triglycerides, to resolve amines by enantioselective
that can catalyze a reaction. Interestingly, a catalytic antibody
acylation, which forms a C N bond.[7] Asparaginase, which
that catalyzes an aldol addition by nucleophilic catalysis
(formation of an imine between the substrate carbonyl and a
cleaves the primary amide in the side chain of asparagine, also
lysine residue) also catalyzes the Kemp elimination, which
cleaves a nitrile in an analogous substrate, b-cyanoalanine.[8]
Romas Kazlauskas was born in 1956 and
studied chemistry at the Massachusetts Institute of Technology (PhD) and Harvard
University (postdoc with George Whitesides). He worked at General Electric Company (1985–1988) and McGill University,
Montreal, Canada (1988–2003) and is currently an associate professor in Biochemistry,
Molecular Biology, and Biophysics at the
University of Minnesota, Twin Cities. He is
an expert in enantioselective organic synthesis using enzymes.
Angew. Chem. Int. Ed. 2004, 43, 6032 –6040
Uwe Bornscheuer was born in 1964 and
studied chemistry at the University of Hannover, Germany, where he graduated with his
Diploma in 1990. After receiving his PhD in
Chemistry in 1993 at the Institute of Technical Chemistry at the same university, he
spent a postdoctoral year at the University
of Nagoya, Japan. He then returned to Germany and joined the Institute of Technical
Biochemistry at the University of Stuttgart,
where he finished his Habilitation in 1998.
Since 1999 he has been Professor for Technical Chemistry and Biotechnology at the
University of Greifswald. His main research interest is the application of
biocatalysts in the synthesis of optically active compounds and in lipid
modification.
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Table 1: Selected examples of catalytic promiscuity in a single enzyme.
Enzyme
Enzyme class
Normal activity
Promiscuous activity
Ref.
proline aminopeptidase
metallohydrolase (two
Mn2+ centers)
metallohydrolase (two
Zn2+ centers)
metalloenzyme (Mn2+,
K+, Mg2+ centers)
C N hydrolysis in proline amides
P F hydrolysis in diisopropyl fluorophosphate
[3]
C N hydrolysis in amides
P O hydrolysis in bis-p-nitrophenylphosphate
[2]
phosphoryl transfer from phosphoenolpyruvate
[10]
metalloenzyme (Mn2+
center)
non-heme diiron
dehydration of 2-hydroxy-6-succinyl2,4-cyclohexadiene carboxylate
hydroxylation of methane
sulfuryl transfer from sulfoenolpyruvate; also
phosphoryl transfer to fluoride, hydroxylamine,
or a-hydroxycarboxylic acids
racemization of N-acylamino acids
non-heme diiron
desaturation of the C9 C10 link in
stearic acid to give oleic acid
aminopeptidase
pyruvate kinase
o-succinylbenzoate synthase
methane monooxygenase
plant steroyl acyl carrier
protein D9 desaturase
cephalosporin C synthase metalloenzyme (nonheme Fe center, 2-oxoglutarate-dependant)
lipase, esterase
serine hydrolase
lipase, chymotrypsin
serine hydrolase
oxidative ring expansion of the fivemembered ring to a six-membered,
hydroxylation of a methyl group
ester hydrolysis
triglyceride or peptide hydrolysis
subtilisin
lipase, trypsin
serine hydrolase
serine esterase
peptide hydrolysis
triglyceride or peptide hydrolysis
pepsin
asparaginase
aspartate hydrolase
Thr-Lys-Asp triad
epoxide hydrolase
Asp-His-Asp triad
oxynitrilase
aldolase catalytic antibody
serine hydroxymethyltransferase
Ser-His-Asp triad
Lys
amide hydrolysis
C N hydrolysis in asparagine to give
aspartate
hydrolysis of epoxides with inversion
of configuration
addition of cyanide to aldehydes
aldol reaction
pyruvate decarboxylase
thiamine-dependent
pyridoxal-dependent
epoxidation, N-oxide formation, dehalogenation,
desaturation of benzylic substrates
sulfoxidation of 9-thia or 10-thia analogues of
stearate and the hydroxylation of 9-fluoro analogues
one of the two normal activities
[36]
[48]
[46,
47]
[43]
b-lactam hydrolysis
aldol addition or Michael addition
[5]
[27,
30]
sulfinamide hydrolysis
[7]
[32,
oligomerization of (Si(CH3)2(OEt)2), dimerization of Si(CH3)3OCH3
33]
sulfite hydrolysis
[4]
CN hydrolysis in b-cyanoalanine to give aspar- [9]
tate
hydrolysis of epoxides with retention of config- [14]
uration
addition of cyanide to imines
[35]
Kemp elimination
[19]
transfer of Cb of serine to tetrahydrop- threonine retroaldol reaction, decarboxylation of [26]
teroylglutamate
aminomalonate, racemization of alanine, transamination of alanine and pyruvate
decarboxylation of pyruvate
acyloin condensation of acetaldehyde and ben- [22–
zaldehyde
24]
requires the lysine to act as a base.[18] Another catalytic
antibody that catalyzes decarboxylation also catalyzes ester
hydrolysis.[19]
In principle, changes in substrate specificity cause subtle
changes in the electron distribution in the transition state and
could be considered as examples of catalytic promiscuity.
However, these differences are usually much smaller than the
examples considered here.
3. Catalytic Promiscuity within the Same Protein
A classic example of catalytic promiscuity is yeast
pyruvate decarboxylase, which not only decarboxylates
pyruvate but also links acetaldehyde and benzaldehyde (a
lyase activity) to form (R)-phenylacetylcarbinol, a precursor
for ephedrine manufacture (Scheme 1).[20] This acyloin condensation involves an additional step—formation of a carbon–carbon bond—that does not occur in the natural
reaction. Although Neuberg and Hirsch discovered this
reaction in whole yeast cells in 1921,[21] researchers more
recently identified pyruvate decarboxylase as the responsible
enzyme.[22] This reaction also demonstrates that the alternate
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Scheme 1. Pyruvate decarboxylase, a thiamine-dependent enzyme, also
catalyzes the enantioselective acyloin condensation of acetaldehyde
and benzaldehyde.
substrates (acetaldehyde plus benzaldehyde) can be much
larger than the natural substrate (pyruvate only). More
recently, a single amino acid substitution in the more stable
pyruvate decarboxylase from Xymomonas mobilis, which
does not catalyze the lyase reaction, added this lyase ability.[23]
The pyridoxal-dependent enzymes are another classic
example of catalytic promiscuity. In most pyridoxal-dependent enzymes the additional functional groups in the active site
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direct the aldimine intermediate toward a single pathway.[24]
But in some cases, the aldimine can react by multiple paths.
For example, serine hydroxymethyltransferase also catalyzes
threonine retroaldol reaction, decarboxylation of aminomalonate, and racemization of alanine.[25] Site-directed mutagenesis of alanine racemase decreased its racemase activity
and enhanced its minor transamination activity.[26]
Another carbon–carbon bond-forming reaction is an aldol
addition of hexanal catalyzed by lipase B from Candida antarctica (Scheme 2).[27] Although the reaction was not enan-
Scheme 3. Michael addition of alternate nucleophiles by O-acetylserine
sulfhydrylase to give an amino acrylate intermediate yields unnatural
amino acids.
Scheme 4. Trypsin-catalyzed hydrolysis and condensation of trimethylethoxysilane to give hexamethyldisiloxane in water.
Scheme 2. Lipase B from Candida antarctica (CAL-B) catalyzes an aldol
addition of hexanal. Although this side reaction is > 105 times slower
than the normal reaction (hydrolysis of triglycerides), it is at least ten
times faster than aldol additions catalyzed by a catalytic antibody with
aldolase activity. The calculated transition-state structure for enolate
formation is on the right.
though silanols and alkoxysilanes are inherently reactive and
can undergo spontaneous condensation or peptide-promoted
condensation,[33] the trypsin-catalyzed reaction was at least
ten times faster than the spontaneous reaction. The condensation involves the trypsin active site because addition of
trypsin-specific inhibitors eliminated catalysis and because
not all trypsins catalyze this reaction: porcine trypsin was
effective, but trypsin from Atlantic cod was not.
Oxynitrilase, which catalyzes addition of cyanide to
aldehydes, also catalyzes the addition of cyanide to imines
with moderate stereoselectivity (3:1–4:1).[34]
A case of mistaken identity due to catalytic promiscuity is
an enzyme originally identified as a N-acyl amino acid
racemase.[35] Gerlt and co-workers recently discovered that
this enzyme is 1000 times more efficient as a catalyst for a
dehydration to form o-succinylbenzoate, suggesting that
succinylbenzoate formation is its true role (Scheme 5). By
changing the N-acyl amino acid from N-acetyl methionine
(the previous best substrate for racemase activity) to Nsuccinyl phenylglycine, which better resembles the succinyl-
tioselective, the diastereoselectivity differed from that of the
spontaneous reaction. The authors hypothesized that the
aldol addition did not require the active site serine, and
indeed, replacement with alanine (Ser105Ala) increased the
aldol addition approximately twofold.
One reaction that forms several new types of bonds is a
Michael addition with alternate nucleophiles catalyzed by Oacetylserine sulfhydrylase (Scheme 3). The normal role of this
pyridoxal-containing enzyme is cysteine biosynthesis from Oacetylserine by the elimination of acetate to give an amino
acrylate intermediate. Michael addition of the nucleophile
sulfide to this intermediate yields cysteine. However, other
nucleophiles also react including thiols, selenols, azide,
cyanide, and some aromatic N-heterocycles yielding unnatural amino acids. Thus, besides catalyzing formation of a C S bond, this enzyme can also
catalyze formation of C Se, C C, and C N
bonds using a similar mechanism.[28] Recently,
Maier engineered an E. coli strain for these
unnatural reactions. Normal fermentation produced the starting material O-acetylserine and
the enzyme; addition of nucleophile yielded
the unnatural amino acids in 45–91 % yield.[29]
Lipases also catalyze Michael addition of
various nucleophiles to 2-(trifluoromethyl)propenoic acid.[30]
Lipase[31] and trypsin[32] catalyze the condensation of silanols and alkoxysilanes, respectively, which involves formation of Si O
Si bonds. Trypsin catalyzed the hydrolysis and
Scheme 5. An enzyme discovered as an N-acyl amino acid racemase is 1000-fold more efcondensation of trimethylethoxysilane to
ficient in the dehydration to form o-succinylbenzoate. Both reaction mechanisms involve
hexamethylsiloxane in water (Scheme 4). Ala similar anionic intermediate.
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benzoate precursor, the efficiency of the racemization reaction increased 1000-fold, making it similar to the succinylbenzoate reaction.
Esterases and lipases, which catalyze hydrolysis of esters,
have overlapping catalytic activities with non-heme haloperoxidases, which catalyze oxidations by hydrogen peroxide via
a peroxycarboxylic acid (Scheme 6). For example, esterase
from Pseudomonas fluorescens,[36] a lactonase,[37] and many
lipases,[38] show low peroxidase activity in the presence of a
carboxylic acid and hydrogen peroxide. Conversely, at least
one haloperoxidase shows low esterase activity.[39]
The overlapping catalytic activity of these enzymes stems
from common transition states and acyl enzyme intermediates
in both reactions (Scheme 7). In esterases, the acyl enzyme
Scheme 6. Non-heme haloperoxidases catalyze the formation of a peroxocarboxylic acid by means of an esterase-like mechanism. The subsequent oxidation of substrates with the peracid may not be enzyme
catalyzed.
intermediate undergoes hydrolysis, while in haloperoxidases,
it undergoes perhydrolysis to yield a peroxycarboxylic acid.
The subsequent oxidation of halide to hypohalous acid by this
peroxycarboxylic acid may not be enzyme catalyzed. In spite
of this overlap, esterases and lipases are more efficient at ester
hydrolysis, while haloperoxidases are more efficient at generating peroxycarboxylic acids. Detailed structural analysis of
a related haloperoxidase[40] and an esterase[41] showed only
subtle differences in the two active sites and did not reveal
why one is a better haloperoxidase and the other a better
esterase.
Catalytic promiscuity has a natural role in the biosynthesis
of several secondary metabolites. For example, the synthesis
of the antibiotic cephalosporin C in eukaryotes uses a single
enzyme with a single active site to catalyze two different
oxidative reactions—an oxidative ring expansion of the fivemembered ring to a six-membered and a hydroxylation of a
methyl group (Scheme 8). Single amino acid substitutions can
inactivate either activity.[42] In contrast cephalosporin synthesis in prokaryotes uses separate enzymes for the two steps,
but both of these enzymes are closely related to the bifunctional one in eukaryotes.
Scheme 8. A non-heme iron(ii) and 2-oxoglutarate-dependent cephalosporin C synthase in eukaryotes catalyzes two different catalytic steps
with the same active site.
These non-heme iron(ii) and 2-oxoglutarate-dependent
oxidative enzymes can even have trifunctional roles. Gibberellin 20-oxidase catalyzes three successive oxidations of the
C20 methyl group to the alcohol, aldehyde and finally to the
carboxylate.[43] Clavaminic acid synthase catalyzes a hydroxylation, an oxidative cyclization, and desaturation.[44]
Scheme 7. Mechanisms for ester hydrolysis and peroxidation by non-heme haloperoxidases both involve an acyl enzyme intermediate. In esterases
(X = H), the H2O attacks this intermediate to complete a hydrolysis. In haloperoxidases (X = OH), the acyl enzyme intermediate comes from acetate added to the reaction mixture. Hydrogen peroxide attacks this intermediate to form a peracid. R = Ph for esterase, H for haloperoxidase.
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Other non-heme diiron oxidative enzymes also catalyze a
wide range of oxidations. For example, the plant steroyl acyl
carrier protein D9 desaturase normally catalyzes the desaturation of stearic acid to oleic acid but also catalyzes
sulfoxidation of 9-thia or 10-thia stearate analogues[45] and
the hydroxylation of 9-fluoro analogues.[46] Methane monooxygenase, another non-heme diiron enzyme, catalyzes
methane hydroxylation as well as a wide range of other
oxidations including epoxidation, N-oxide formation, dehalogenation, and desaturation of benzylic substrates.[47] Degradation enzymes, such as cytochrome P450 enzymes, which
contain a heme iron, also catalyze a wide range of oxidations.
4. Catalytic Promiscuity within Modified Proteins
4.1. Natural Evolution of New Catalytic Activity
Divergent evolution is a natural process that creates
different species from a common ancestor. This process also
works on a molecular scale to create enzymes with new
catalytic activities. New enzymatic activities arise by gene
duplication followed by evolution of new activity for the
copy.[48] Two examples of divergent evolution are the a/bhydrolase-fold superfamily[49] and the enolase superfamily.[50]
The a/b-hydrolase-fold enzymes all involve nucleophilic
catalysis but include a wide range of substrates and reaction
types including ester or peptide hydrolysis (serine nucleophile), dehalogenase, and epoxide hydrolase (aspartate
nucleophile). The enolase enzymes have similar active sites
and catalyze divalent-metal-assisted general-base-catalyzed
removal of a proton a to a carboxylic acid to form an enolic
intermediate. Examples of different reactions catalyzed by
the enolase superfamily include racemization (mandelate, Nacyl amino acids) and b-eliminations (o-succinyl benzoate
synthase). An example of a “misassigned” enolase enzyme
was mentioned in Section 3.
One example of a surprisingly rapid natural evolution of
new catalytic activity is an atrazine chlorohydrolase
(Scheme 9).[51] Researchers initially found that atrazine, an
herbicide widely used since the late 1950s, did not readily
degrade in soils, but since 1993 a number of groups have
reported rapid degradation. The key enzyme in this biodegradation—atrazine chlorohydrolase, which cleaves the C Cl
bond—differs by only eight amino acid substitutions from
melamine hydrolase, which catalyzes the hydrolysis of a C N
bond in melamine. Melamine hydrolase has low atrazine
chlorohydrolase activity, but the new enzyme has no detectable melamine hydrolase activity. Directed evolution further
expanded the substrate range of this atrazine chlorohydrolases to include C S and C O bond cleavage.[52] Another
example of rapid natural evolution is the evolution of a
phosphotriesterase that degrades the insecticide paraoxon.[53]
4.2. Changing the Metal Ion
Metal substitutions can also change catalytic activity. One
of the earliest examples from 1976 shows that replacement of
the Zn2+ ion in the active site of a carboxypeptidase with a
Cu2+ ion converted this peptidase into a slow oxidase.[54]
Replacement of the Zn2+ ion in the active site of thermolysin
with much larger ions such as tungstate, molybdate, and
selenate created enzymes that catalyze oxidation of thioethers to sulfoxides with hydrogen peroxide.[55] Replacing the
active-site serine in subtilisin with selenomethionine resulted
in peroxidase activity.[56]
Other examples of overlapping catalytic activity are acid
phosphatases and vanadate-dependent haloperoxidases.[57, 58]
The amino acid sequence, three-dimensional structure, and
active site are similar in both classes of enzymes. Vanadate
binds to the same site as a phosphate ester presumably
because it readily adopts a five-coordinate structure that
resembles the transition state for phosphate ester hydrolysis.
The vanadate ion catalyzes peroxidation by binding peroxide
to the vanadium center, thereby increasing its electrophilicity.
Further support for the similarity of the two active sites is the
ability of vanadate to inhibit phosphatases and the ability of
phosphate to inactivate vanadate-dependent haloperoxidases
by displacing the vanadate. This exchange of active sites also
exchanges the catalytic activity of these two classes of
enzymes. Several acid phosphatases show low haloperoxidase
activity upon addition of vanadate,[59] and conversely, apohaloperoxidases show low phosphatase activity.[60] Sheldon
and co-workers reported an enantioselective oxidation of
sulfides to sulfoxides using a vanadate-substituted phytase.[61]
However, the altered enzymes were much less effective
catalysts than the true enzyme: the turnover numbers were
103–104 times lower for the haloperoxidase activity of a
vanadate-containing phosphatase than for a true haloperoxidase, or for the phosphatase activity of apo-haloperoxidase as
compared to that of a true phosphatase. This large difference
shows that each enzyme is optimized for the reaction it
catalyzes. However, like the esterase/non-heme haloperoxidase case mentioned above, even with the available X-ray
crystal structures, it is not clear which structural features are
responsible for the different optimized activity.
4.3. Engineering of Enzymes
Scheme 9. An atrazine chlorohydrolase recently evolved naturally from
a melamine hydrolase. The starting melamine hydrolase (C N bond
cleavage) has low atrazine chlorohydrolase activity (C Cl bond cleavage), but the new atrazine chlorohydrolase has lost its melamine hydrolase activity.
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A number of groups mimicked divergent evolution by
using site-directed mutagenesis. By comparing related enzymes with different catalytic activity, they identified substitutions that change the catalytic activity. Making these
changes in one of the enzymes altered the catalytic activity.
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For example, four amino acid substitutions in a fatty acid
desaturase by their equivalents in a fatty acid hydroxylase
yielded an efficient hydroxylase (Scheme 10).[62] In another
glycosidases use a double displacement mechanism with a
catalytic acid/base and a catalytic nucleophile. In the initial
step an a-linked covalent intermediate is formed by attack of
the catalytic nucleophile on the starting b-glycoside. The
catalytic acid assists this step by protonating the leaving
group. In the second step this covalent intermediate is
released by the catalytic-base-assisted attack of the incoming
nucleophile—water or a new glycoside.
By disabling key steps in the mechanism, Withers and
colleagues made three new catalysts (Scheme 12). The first,
Scheme 10. Both desaturases and hydroxylases have a diiron center
and oxidize C12 of oleic acid. Mutating four amino acids in a desaturase turned it into a hydroxylase.
example, single amino acid changes in l-Ala-d/l-Glu epimerase introduced ortho-succinoyl benzoate synthase activity or
muconate-lactonizing enzyme activity.[63] Within the family of
glutathione transferases, mutations changed a transferase that
catalyzes a Michael addition into one that catalyzes a
nucleophilic aromatic substitution.[64] Similarly, a glutathione
transferase with peroxidase activity gained steroid isomerase
activity after five mutations to mimic the active site in a
related steroid isomerase.[65] Mutations within an oxidosqualene cyclase changed the site of proton loss, thereby yielding
different steroids.[66] A much more challenging task is
converting a noncatalytic protein into a dehydratase with
low activity.[67]
Scheme 12. Disabling key mechanistic steps in a retaining b-glycosidase creates new catalytic activities. Removal of the catalytic nucleophile creates a glycosynthases where only a-glycosyl fluorides react
presumably by a single displacement mechanism. Removal of the catalytic acid/base creates a thioglycoligase (only strong incoming nucleophiles such as thiols react), and removal of both catalytic nucleophile
and catalytic acid/base creates a thioglycosynthase (only a-glycosyl fluorides and strong incoming nucleophiles react). DNP = 2,4-dinitrophenyl.
4.4. Glycosynthases, Thioglycoligases, and Thioglycosynthases
Retaining b-glycosidases normally catalyze the hydrolysis
of b-glycosidic links, but they also catalyze glycoside exchange
under conditions of low water concentration. The reaction
involves a starting b-glycoside (sugar-OR) reacting with an
incoming nucleophile (HOR’, Scheme 11). The incoming
nucleophile is water in the case of hydrolysis and a second
glycoside in the case of glycoside exchange. Retaining
Scheme 11. Glycoside exchange using retaining b-glycosidases involves
a double displacement. A glycosyl donor (b-sugar-OR) forms an alinked glycosyl enzyme, which then reacts with an incoming nucleophile or acceptor (HOR’) to make a new b-glycosidic link (sugar-OR’).
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glycosynthase, results upon removing the catalytic nucleophile (e.g., a Glu-to-Ala mutation).[68] This removal prevents
formation of the key covalent intermediate, dramatically
altering the mechanism. Normal glycosides no longer react,
but a-glycosyl fluorides do react, likely by a direct displacement mechanism. Glycosynthases, like the starting enzyme,
form b-glycoside links. However, glycosynthases no longer
catalyze hydrolysis of product, which is a nonactivated
glycoside, and thus give higher yields. Five different glycosynthases have been reported with differing glycosyl fluoride
specificity and differing regioselectivity (formation of b-1,3 vs.
b-1,4 links). The second type of new catalyst, thioglycoligase,
results upon removing the catalytic acid/base.[69] One role of
this catalytic acid/base is activation of the incoming nucleophile. Absence of this activator precludes reaction with
normal incoming nucleophiles and requires strong nucleophiles such as thiols. Introducing single amino acid mutations
into b-glycosidases from Agrobacterium sp. Abg (mutation:
E171A) created a variant that formed S-glycosidic linkages in
high yield (Scheme 13). The wild-type enzyme gave no
product, possibly due to steric hindrance caused by the larger
sulfur atom. Finally, removing both the catalytic nucleophile
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Chemie
Enzyme Catalysis
Scheme 13. A thioglycoligase lacking the catalytic acid/base no longer
catalyzes hydrolysis. However, using an activated glycosyl donor overcomes the lack of a proton donor for the leaving group, and using a
nucleophilic thiol overcomes the lack of base to activate the incoming
nucleophile. Thus, the mutant now catalyzes formation of an S-glycosidic link.
and the catalytic acid/base creates a thioglycosynthase, which
requires both an a-glycosyl fluoride and a thiol acceptor.[70]
5. Summary and Outlook
Catalytic promiscuity in enzymes is more common than
generally appreciated. The dense collection of catalytic
groups in an active site can accept alternate functional groups
in the substrate and follow alternate reaction pathways. Small
modifications in an active site can further dramatically
expand the range of alternative pathways. In most, but not
all, cases these alternate reactions are slower than the natural
reactions. We hope that this review will encourage others to
both search more extensively for catalytic promiscuity in
existing enzymes and to use the current tools of protein
engineering and directed evolution to extend the useful
applications of enzymes.
We thank Bernhard Hauer (BASF, Ludwigshafen, Germany)
for helpful suggestions and discussion. U.T.B. thanks the Fonds
der Chemischen Industrie (Frankfurt, Germany) for financial
support.
Received: April 22, 2004
Published Online: November 2, 2004
[1] Reviews: a) S. D. Copley, Curr. Opin. Chem. Biol. 2003, 7, 265 –
272; b) A. Yarnell, Chem. Eng. News 2003, 81, 33 – 35; c) P. J.
OMBrian, D. Herschlag, Chem. Biol. 1999, 6, 91 – 105.
[2] a) H. I. Park, L.-J. Ming, Angew. Chem. 1999, 111, 3097 – 3100;
Angew. Chem. Int. Ed. 1999, 38, 2914 – 2916; b) A. Ercan, H. I.
Park, L.-J. Ming, Chem. Commun. 2000, 2501 – 2502.
[3] T. C. Cheng, S. P. Harvey, G. L. Chen, Appl. Environ. Microbiol.
1996, 62, 1636 – 1641.
[4] PLE: M. Jones, M. I. Page, J. Chem. Soc. Chem. Commun. 1991,
316 – 317; PFL: R. Brieva, J. Z. Crich, C. J. Sih, J. Org. Chem.
1993, 58, 1068 – 1075; CAL-B: a) W. Adam, P. Groer, H.-U.
Humpf, C. R. Saha-MOller, J. Org. Chem. 2000, 65, 4919 – 4922;
b) E. ForrP, F. FQlOp, Org. Lett. 2003, 5, 1209 – 1212; c) S. Park,
E. ForrP, H. Grewal, F. FQlOp, R. J. Kazlauskas, Adv. Synth.
Catal. 2003, 345, 986 – 995.
Angew. Chem. Int. Ed. 2004, 43, 6032 –6040
[5] T. W. Reid, D. Fahrney, J. Am. Chem. Soc. 1967, 89, 3941 – 3943.
[6] P. F. Mugford, V. P. Magloire, R. J. Kazlauskas, unpublished
results.
[7] Review: M. Breuer, K. Ditrich, T. Habicher, B. Hauer, M.
Kesseler, R. Stuermer, T. Zelinski, Angew. Chem. 2004, 116,
806 – 843; Angew. Chem. Int. Ed. 2004, 43, 788 – 824.
[8] R. C. Jackson, R. E. Handschumacher, Biochemistry 1970, 9,
3585 – 3590.
[9] J. A. Peliska, M. H. OMLeary, Biochemistry 1989, 28, 1604 – 1611.
[10] P. J. O’Brien, D. Herschlag, J. Am. Chem. Soc. 1998, 120, 12 369 –
12 370.
[11] Y. Romsicki, G. Scapin, V. Beaulieu-Audy, S. Patel, J. W. Becker,
B. P. Kennedy, E. Asante-Appiah, J. Biol. Chem. 2003, 278,
29 009 – 29 015.
[12] A recent example in biocatalysis: A. Magnusson, K. Hult, M.
Holmquist, J. Am. Chem. Soc. 2001, 123, 4354 – 4355.
[13] a) W. Kroutil, M. Mischitz, K. Faber, J. Chem. Soc. Perkin Trans.
1 1997, 3629 – 3636; b) K. Faber, W. Kroutil, Tetrahedron:
Asymmetry 2002, 13, 377 – 382.
[14] a) F. Hollfelder, A. J. Kirby, D. S. Tawfik, Nature 1996, 383, 60 –
62; b) G. Klein, J.-L. Reymond, Bioorg. Med. Chem. Lett. 1998,
8, 1113 – 1116.
[15] S. Colonna, N. Gaggero, J. Drabowicz, P. Lyzwa, M. Mikolajczyk,
Chem. Commun. 1999, 1787 – 1788.
[16] D. C. Levinger, J.-A. Stevenson, L.-L. Wong, J. Chem. Soc.
Chem. Commun. 1995, 2305 – 2306.
[17] S. Ozaki, T. Matsui, Y. Watanabe, J. Am. Chem. Soc. 1996, 118,
9784 – 9785;S. Ozaki, H.-J. Yang, T. Matsui, Y. Goto, Y.
Watanabe, Tetrahedron: Asymmetry 1999, 10, 183 – 192.
[18] L. C. James, D. S. Tawfik, Protein Sci. 2001, 10, 2600 – 2607.
[19] A. C. Backes, K. Hotta, D. Hilvert, Helv. Chim. Acta 2003, 86,
1167 – 1174.
[20] Review: O. P. Ward, A. Singh, Curr. Opin. Biotechnol. 2000, 11,
520 – 526.
[21] C. Neuberg, J. Hirsch, Biochem. Z. 1921, 115, 282 – 310.
[22] a) S. Bringer-Meyer, H. Sahm, Biocatalysis 1988, 1, 321 – 331;
b) D. H. G. Crout, H. Dalton, D. W. Hutchinson, M. Miyagoshi,
J. Chem. Soc. Perkin Trans. 1 1991, 1329 – 1334.
[23] G. Goetz, P. Iwan, B. Hauer, M. Breuer, M. Pohl, Biotechnol.
Bioeng. 2001, 74, 317 – 325.
[24] Review: H. Hayashi, J. Biochem. 1995, 118, 463 – 473.
[25] R. Contestabile, A. Paiardini, S. Pascarella, M. L. di Salvo, S.
DMAguanno, F. Bossa, Eur. J. Biochem. 2001, 268, 6508 – 6525.
[26] G.-Y. Yow, A. Watanabe, T. Yoshimura, N. Esaki, J. Mol. Catal.
B: Enzym. 2003, 23, 311 – 319.
[27] C. Branneby, P. Carlqvist, A. Magnusson, K. Hult, T. Brinck, P.
Berglund, J. Am. Chem. Soc. 2003, 125, 874 – 875.
[28] a) F. Ikegami, I. Murakoshi, Phytochemistry 1994, 35, 1089 –
1104; b) D. H. Flint, J. F. Tuminello T. J. Miller, J. Biol. Chem.
1996, 271, 16 053 – 16 067.
[29] T. H. P. Maier, Nat. Biotechnol. 2003, 21, 422 – 427.
[30] T. Kitazume, T. Ikeya, K. Murata, J. Chem. Soc. Chem. Commun.
1986, 1331 – 1333.
[31] H. Nishino, T. Mori, Y. Okahata Chem. Commun. 2002, 2684 –
2685.
[32] A. R. Bassindale, K. F. Brandstadt, T. H. Lane, P. G. Taylor, J.
Inorg. Biochem. 2003, 96, 401 – 406.
[33] N. KrOger, S. Lorenz, E. Brunner, M. Sumper, Science 2002, 298,
584 – 586.
[34] T. Lee, Y. Ahn, Bull. Korean Chem. Soc. 2002, 23, 1490 – 1492.
[35] a) D. R. J. Palmer, J. B. Garrett, V. Sharma, R. Meganathan, P. C.
Babbitt, J. A. Gerlt, Biochemistry 1999, 38, 4252 – 4258; b) E. A.
Taylor Ringia, J. B. Garrett, J. B. Thoden, H. M. Holden, I.
Rayment, J. A. Gerlt, Biochemistry 2004, 43, 224 – 229.
[36] I. Pelletier, J. Altenbuchner, Microbiology 1995, 141, 459 – 468.
[37] M. Kataoka, K. Honda, S. Shimizu, Eur. J. Biochem. 2000, 267,
3 – 10.
www.angewandte.org
2004 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
6039
Angewandte
Chemie
U. T. Bornscheuer and R. J. Kazlauskas
[38] a) F. BjOrkling, H. Frykman, S. E. Godtfredsen, O. Kirk,
Tetrahedron 1992, 48, 4587 – 4592; b) O. Kirk, L. S. Conrad,
Angew. Chem. 1999, 111, 1031 – 1033; Angew. Chem. Int. Ed.
1999, 38, 977 – 979.
[39] I. Pelletier, J. Altenbuchner, Microbiology 1995, 141, 459 – 468.
[40] B. Hofmann, S. TOlzer, S. , I. Pelletier, J. Altenbuchner, K.-H.
van PRe, H. J. Hecht, J. Mol. Biol. 1998, 279, 889 – 900.
[41] J. D. Cheeseman, A. Tocilj, S. Park, J. D. Schrag, R. J. Kazlauskas, Acta Crystallogr. D 2004, 60(7), 1237 – 1243.
[42] M. D. Lloyd, S. J. Lipscomb, K. S. Hewitson, C. M. H. Hensgens,
J. E. Baldwin, C. J. Schofield, J. Biol. Chem. 2004, 279, 15 420 –
15 426.
[43] T. Lange, P. Hedden, J. E. Graebe, Proc. Natl. Acad. Sci. USA
1994, 91, 8552 – 8556.
[44] M. D. Lloyd, K. D. Merritt, V. Lee, T. J. Sewell, B. Wha-Son, J. E.
Baldwin, C. J. Schofield, S. W. Elson, K. H. Baggaley, N. H.
Nicholson, Tetrahedron 1999, 55, 10 201 – 10 220.
[45] B. Behrouzian, P. H. Buist, J. Shanklin, Chem. Commun. 2001,
401 – 402.
[46] a) B. Behrouzian, B. Dawson, P. H. Buist, J. Shanklin, Chem.
Commun. 2001, 765 – 766; b) B. Behrouzian, C. K. Savile, B.
Dawson, P. H. Buist, J. Shanklin, J. Am. Chem. Soc. 2002, 124,
3277 – 3283.
[47] Y. Jin, J. D. Lipscomb, J. Biol. Inorg. Chem. 2001, 6, 717 – 725.
[48] J. A. Gerlt, P. C. Babbitt, Annu. Rev. Biochem. 2001, 70, 209 –
246.
[49] a) D. L. Ollis, E. Cheah, M. Cygler, B. Dijkstra, F. Frolow, S. M.
Franken, M. Harel, S. J. Remington, I. Silman, J. Schrag, J. L.
Sussman, K. H. G. Verschueren, A. Goldman, Protein Eng. 1992,
5, 197 – 211; b) M. Holmquist, Curr. Protein Pept. Sci. 2000, 1,
209 – 235.
[50] P. C. Babbitt, M. S. Hasson, J. E. Wedekind, D. R. J. Palmer,
W. C. Barrett, G. H. Reed, I. Rayment, D. Ringe, G. L. Kenyon,
J. A. Gerlt, Biochemistry 1996, 35, 16 489 – 16 501.
[51] J. L. Seffernick, L. P. Wackett, Biochemistry 2001, 40, 12 747 –
12750.
[52] S. Raillard, A. Krebber, Y. Chen, J. E. Ness, E. Bermudez, R.
Trinidad, R. Fullem, C. Davis, M. Welch, J. Seffernick, L. P.
Wackett, W. P. C. Stemmer, J. Minshull, Chem. Biol. 2001, 8,
891 – 898.
[53] T. S. Scanlan, R. C. Reid, Chem. Biol. 1995, 2, 71 – 75.
6040
2004 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
[54] K. Yamamura, E. T. Kaiser, J. Chem. Soc. Chem. Commun. 1976,
830 – 831.
[55] M. Bakker, F. van Rantwijk, R. A. Sheldon, Can. J. Chem. 2002,
80, 622 – 625.
[56] I. M. Bell, M. L. Fisher, Z. P. Wu, D. Hilvert, Biochemistry 1993,
32, 3754 – 3762.
[57] a) A. F. Neuwald, Protein Sci. 1997, 6, 469 – 472; b) J. Littlechild,
E. Garcia-Rodriguez, A. Dalby, M. Isupov, J. Mol. Recognit.
2002, 15, 291 – 296.
[58] W. Hemrika, R. Renirie, H. L. Dekker, P. Barnett, R. Wever,
Proc. Natl. Acad. Sci. USA 1997, 94, 2145 – 2149.
[59] N. Tanaka, V. Dumay, Q. Liao, A. J. Lange, R. Wever, Eur. J.
Biochem. 2002, 269, 2162 – 2167.
[60] R. Renirie, W. Hemrika, R. Wever, J. Biol. Chem. 2000, 275,
11 650 – 11 657.
[61] a) F. van de Velde, L. KOnemann, F. van Rantwijk, R. A. Sheldon, Chem. Commun. 1998, 1891 – 1892; b) F. van de Velde, L.
KOnemann, F. van Rantwijk, R. A. Sheldon, Biotechnol. Bioeng.
2000, 67, 87 – 96.
[62] a) P. Broun, J. Shanklin, E. Whittle, C. Somerville, Science 1998,
282, 1315 – 1317; b) J. A. Broadwater, E. Whittle, J. Shanklin, J.
Biol. Chem. 2002, 277, 15 613 – 15 620.
[63] D. M. Z. Schmidt, E. C. Mundorff, M. Dojka, E. Bermudez, J. E.
Ness, S. Govindarajan, P. C. Babbitt, J. Minshull, J. A. Gerlt,
Biochemistry 2003, 42, 8387 – 8393.
[64] L. O. Nilsson, A. Gustafsson, B. Mannervik, Proc. Natl. Acad.
Sci. USA 2000, 97, 9408 – 9412.
[65] P. L. Pettersson, A.-S. Johansson, B. Mannervik, J. Biol. Chem.
2002, 277, 30 019 – 30 022.
[66] a) E. A. Hart, L. Hua, L. B. Darr, W. K. Wilson, J. Pang, S. P. T.
Matsuda, J. Am. Chem. Soc. 1999, 121, 9887 – 9888; b) M. M.
Meyer, R. Xu, S. P. T. Matsuda, Org. Lett. 2002, 4, 1395 – 1398.
[67] A. E. Nixon, S. M. Firestine, F. G. Salinas, S. J. Benkovic, Proc.
Natl. Acad. Sci. USA 1999, 96, 3568 – 3571.
[68] S. J. Williams, S. G. Withers, Aust. J. Chem. 2002, 55, 3 – 12; for
earlier work on converting an inverting enzyme to a retaining
enzyme see: R. Kuroki, L. H. Weaver, B. M. Matthews, Nat.
Struct. Biol. 1995, 2, 1007 – 1011.
[69] M. Jahn, J. Marles, R. A. J. Warren, S. G. Withers, Angew. Chem.
2003, 115, 366 – 368; Angew. Chem. Int. Ed. 2003, 42, 352 – 354.
[70] M. Jahn, H. Chen, J. Muellegger, J. Marles, R. A. J. Warren, S. G.
Withers, Chem. Commun. 2004, 274 – 275.
www.angewandte.org
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