7.17 C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

7.17 C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes PA Dalby, JM Ward, and HC Hailes, University College London, London, UK r 2012 Elsevier Ltd. All rights reserved.

7.17.1 7.17.1.1 7.17.1.2 7.17.1.2.1 7.17.1.2.2 7.17.1.3 7.17.1.4 7.17.2 7.17.2.1 7.17.2.1.1 7.17.2.1.2 7.17.2.1.3 7.17.2.2 7.17.2.3 7.17.2.4 7.17.2.5 7.17.2.6 7.17.3 7.17.3.1 7.17.3.1.1 7.17.3.1.2 7.17.3.2 7.17.3.2.1 7.17.3.2.2 7.17.3.2.3 7.17.3.3 7.17.3.4 7.17.3.4.1 7.17.3.4.2 7.17.3.5 7.17.4 7.17.4.1 7.17.4.2 7.17.4.3 7.17.5 References

TDP Enzymes Overview Mechanism Natural Reactions of Some TDP-Dependent Enzymes Transferases Lyases Structure and Evolution General Considerations for Using TDP Enzymes as Biocatalysts Transferases Transketolase (TK) Synthetic potential of transketolases Enzyme engineering of transketolase Biocatalytic process engineering of transketolase Dihydroxyacetone Synthase (DHAS)/Formaldehyde Transketolase (FTK) Acetoin-Ribose-5-Phosphate Transaldolase Glyoxylate Carboligase (GXC)/2-Hydroxy-3-Oxoadipate Synthase (HOAS) Acetolactate Synthase (ALS)/Acetohydroxyacid Synthase (AHAS) 1-Deoxy-D-Xylulose-5-Phosphate Synthase (DXPS) (EC 2.2.1.7) Lyases Benzoylformate Decarboxylase (BFDC) Synthetic potential of BFDC Enzyme engineering of BFDC Benzoin Aldolase/Benzaldehyde Lyase (BAL) Synthetic potential of BAL Enzyme engineering of BAL Biocatalytic process engineering of BAL Branched-Chain 2-Ketoacid Decarboxylase Pyruvate Decarboxylase (PDC) Synthetic potential of PDC Biocatalytic process engineering of PDC Phosphoketolase Outlook Protein Engineering Bioprospecting Process Engineering Conclusions

Glossary Bioprospecting The search for novel enzyme activities from environmental sources. Directed evolution Process of artificially evolving a gene or enzyme using recombinant DNA techniques.

7.17.1 7.17.1.1

372 372 373 373 373 376 376 376 376 376 378 379 380 380 381 381 382 382 382 382 383 384 384 384 385 385 385 385 386 386 386 386 387 387 387 388

Metagenomics Process of obtaining genetic material directly from environmental samples. Phylogeny n. pl. phylogenies The evolutionary development and history of a grouping of genes or species.

TDP Enzymes Overview Mechanism

To date, 19 distinct Enzyme Commission (EC) numbers are allocated for enzymes that utilize thiamin diphosphate (TDP) as a coenzyme.1 The reactions catalyzed are diverse, and include nonoxidative decarboxylation of a-keto acids, oxidative

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decarboxylation of a-keto acids, carboligations, and cleavage of C–C bonds.2 They fall within 4 of the 6 classes of enzyme, namely oxidoreductases, transferases, hydrolases, and lyases. Many, such as the oxidative decarboxylation of pyruvate by pyruvate dehydrogenase to form acetylCoA, require the additional use of other coenzymes and larger enzyme complexes, and will not be covered here. However, all TDP-catalyzed reactions make use of the low pKa for deprotonation at C2 of the thiazole ring, to readily form an ylide, and usually proceed via an enamine intermediate that is chemically equivalent to an activated aldehyde (Scheme 1). A review of the mechanisms in more detail is available elsewhere,3 and further insights have also been gained more recently.4–6

R1

R4 O

R1 N+

−

R

+H

S

OH

4

+

R2

HO

R2

S 3

OH

R

TDP Ylide

R3

N+

R3

R4

PDC: =O TK: CHOH-R BAL: Ar or R

PDC: Me TK: CH2OH BAL: Ar

R3-CHO (TK, BAL) CO2 (PDC)

O R1 R4 HO

R5

N+

H R4

S

HO

R1

R2

R5

TK

R1 + R4 N

+H+

N S

R2

HO Activated aldehyde/ enamine intermediate

PDC or BAL

HO H

O

R2

S

O OH

R4

R4

H

R5 Scheme 1 Mechanisms of TDP-dependent enzymes. Variations for pyruvate decarboxylase (PDC), transketolase (TK), and benzaldehyde lyase (BAL) are shown.

7.17.1.2 7.17.1.2.1

Natural Reactions of Some TDP-Dependent Enzymes Transferases

Some TDP-dependent transferases catalyze the transfer of a ketol unit onto an aldeyhde, to form new ketol and aldehyde products of different chain lengths. For example, transketolase (TK) catalyzes two steps in the nonoxidative branch of the pentose phosphate pathway,7 whereby a two-carbon ketol unit from D-xylulose-5-phosphate is reversibly transferred to either D-ribose-5phosphate or D-erythrose-4-phosphate to form D-sedoheptulose-7-phosphate or D-fructose-6-phosphate, respectively (Scheme 2). Similarly, dihydroxyacetone synthase (DHAS) catalyzes transfer of the ketol unit from D-xylulose-5-phosphate onto formaldehyde to produce glycerone (dihydroxyacetone). Other transferases such as 1-deoxy-D-xylulose-5-phosphate synthase (DXPS) transfer a ketone unit from an a-ketoacid, for example pyruvate, to an aldehyde and are made effectively irreversible by the nonoxidative decarboxylation of the a-ketoacid8 (Scheme 2). TK and DHAS have also been shown to catalyze this type of reaction in vitro, when using hydroxypyruvate (HPA) as the ketol donor.9,10 Another variant of this reaction can be found in which both substrates are a-ketoacids, such as the conversion of two pyruvate molecules to acetolactate by acetolactate synthase (ALS),11 or in the synthesis of 2-hydroxy-3-oxoadipate from 2-oxoglutarate and glyoxylate by glyoxylate carboligase (GXC), which is also known as 2-hydroxy-3-oxoadipate synthase (HOAS)12 (Scheme 2). In this case, the acceptor is a second a-ketoacid rather than an aldehyde, and hence the final product retains one carboxylate moiety. The acceptor can also be an a,b-unsaturated carboxylic acid via a Michael addition, as found with 2-succinyl-5-enolpyruvyl-6-hydroxy-3cyclohexene-1-carboxylic acid synthase (SEPHCHCS), which is involved in menaquinone biosynthesis in Escherichia coli13 (Scheme 2). Finally, an unusual TDP-catalyzed mechanism has been proposed for N2-(2-carboxyethyl)arginine synthase (CAS),14 the first enzyme in the clavulanic acid biosynthesis pathway, which forms a new carbon–nitrogen bond in the conversion of D-glyceraldehyde 3-phosphate and L-arginine into N2-(2-carboxyethyl)-L-arginine with the release of phosphate (Scheme 2). In this case, nucleophilic attack on glyceraldehyde-3-phosphate (G3P) proceeds as expected, but then the usual enamine intermediate is not formed. Instead, a b-elimination of water is postulated to lead to an enolate-TDP intermediate, which then undergoes a second elimination to form an acryloyl-TDP intermediate with the release of phosphate.15

7.17.1.2.2

Lyases

TDP-dependent lyases generally split acyloins or a-ketoacids to form new aldehydes. These reactions are similar to those catalyzed by the transferases, except that an aldehyde product is formed directly from the enamine intermediate. This also means that the

374

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

HO HO

H O

O

HO

O OH

OH

+

HO

H

OH

TK

HO

HO

O

+

OH

HO

HO 2−O PO 3

2−O

2−O

3PO

3PO

2−O PO 3

Xylulose-5-P

Sedoheptulose-7-P (or fructose-6-P)

Ribose-5-P (or erythrose-4-P)

Glyceraldehyde-3-P

HO HO H

O OH

+

HO

O

DHAS

O

H

O

+

H

2−O

HO

2−O PO 3

Formaldehyde

Xylulose-5-P

Glycerone

O O HO

2−O

OH HO 2−O

3PO

O

OH

Acetolactate

O

O O OH

O

OH

CO2

OH

Pyruvate

O

+

3PO

O

HO

HO

CO2

O

ALS O

2

+

1-Deoxy-xylulose-5-P

Glyceraldehyde-3-P

Pyruvate

Glyceraldehyde-3-P

DXPS

HO

+

OH

HO SEPHCHCS

+

OH O

O

O O

HO

HO

HO Isochorismate

2-Oxoglutarate

H

2-Succinyl-5-enolpyruvyl-6-hydroxy3-cyclohexene-1-carboxylic acid

HO

O

HO O

OH

OH

CO2

+

O

O

H2N

OH

H N

CAS

+

O

3PO

O

H O

O P

HO

+

HO O HN

HN NH

H2N D-Glyceraldehyde-

O

Arginine

NH H2N N2-(2-carboxyethyl)-L-arginine

3-phosphate Scheme 2 Naturally catalyzed reactions of various TDP-dependent transferases and lyases.

H3PO4

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

HO

HO O O

HO O O

O GXC (transferase)

HO O

+ O

H

CO2

+

CO2

+

CO2

+

CO2

OH

2-Oxoglutarate

2-Hydroxy-3oxoadipate HO

HO O 2

+ O

HO Glyoxylate

375

GXC (lyase)

O H

O HO O

Glyoxylate

Tartronate semialdehyde

H

O

BAL

O OH

Benzaldehyde

Benzoin

PDC

O O

O H

HO

Acetaldehyde

Pyruvate

H

O O

O

BFDC OH

Benzylformate

Benzaldehyde

Scheme 2 Continued.

reverse reaction begins with the direct nucleophilic attack by the TDP ylide on an aldehyde substrate, rather than the a-ketoacids or a-hydroxy ketones typical for the transferase reactions. One family of TDP-dependent lyases (EC 4.1.2.X) cleaves a-hydroxy ketones (acyloins) to form two aldehyde molecules. For example, BAL catalyzes the reversible cleavage of the benzoin to form two molecules of benzaldehyde (Scheme 2). Interestingly, TK has also been shown to carry out a similar reaction, albeit slowly, in which two glycolaldehyde molecules are used to form L-erythulose.16 A second family of TDP-dependent lyases, the decarboxylases (EC 4.1.1.X), catalyzes the nonoxidative decarboxylation of a-ketoacids (Scheme 2). The reaction generates carbon dioxide, which renders them effectively irreversible in vitro under normal conditions, and ends with the release of an aldehyde from the TDP-enamine intermediate. Examples include pyruvate decarboxylase (PDC), sulfopyruvate decarboxylase (SPDC), indolepyruvate decarboxylase (IPDC), phenylpyruvate decarboxylase (PhPDC), benzoylformate decarboxylase (BFDC), and branched-chain ketoacid decarboxylase (KdcA).17–20 GXC can also act as a decarboxylating lyase by catalyzing the conversion of two glyoxylate molecules to form tartronate semialdehyde.21 TK (EC 2.2.1.1), DHAS (EC 2.2.1.3), DXPS (EC 2.2.1.7), ALS (EC 2.2.1.6), SEPHCHCS (EC 2.2.1.9), CAS (EC 2.5.1.66), GXC (EC 2.2.1.5 and EC 4.1.1.47), benzaldehyde lyase (BAL) (EC 4.1.2.38), PDC (EC 4.1.1.1), BFDC (EC 4.1.1.7).

376

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

7.17.1.3

Structure and Evolution

The structures of many TDP enzymes have been obtained, and all bind the TDP at the interface between pyrophosphate (PP) and pyrimidine (Pyr) domains, either within the same subunit as for pyruvate ferredoxin reductase (PFRD)22 and DXPS,23 or more commonly between different subunits.24 Many have additional domains that relate to their overall function, although the specific functions of many of these domains remain unclear. The three-dimensional structures of the PP and Pyr domains that comprise the TDP-containing active site are very well conserved. For example, those of TK, pyruvate oxidase (PO), and PDC align well, although their primary sequences and domain order vary considerably.24 The evolution of TDP-dependent enzymes has been reconstructed from both structural and sequence comparisons.25–28 Dimerization of an ancestral TDP-binding domain (ATB), then the formation of an (ATB)4 homotetramer, preceded gene duplication and divergence to form a PP2Pyr2 heterotetramer, still observed for the Sulpholobus solfataricus TK,29 and also within the PP6Pyr6 heterododecamer of modern SPDC.18 Gene fusion then appears to have led to the more prevalent (PP-Pyr)2 homodimeric and (PP-Pyr)4 homotetrameric forms of TDP-dependent enzymes, and further diversity in their chemistries resulting from domain rearrangement, additional domain recruitment, incremental mutations, and for some, the formation of multienzyme complexes. Gene splitting has also most likely led to the formation of the PP2Pyr2 heterotetrameric 2-oxoisovalerate dehydrogenase-like (2OXO) family. The ready availability of many TDP-dependent enzyme structures, and their well-characterized phylogenies, lend themselves well to the possibility of identifying new variants for improved biocatalysis via bioinformatics or protein engineering routes. As the diversity of chemical reactions, selectivities, and substrate specificities become more defined experimentally for each type of enzyme, the growing knowledge should also make engineering of new biocatalysts a more efficient process, for example using directed evolution techniques overviewed in Chapter 6.3.

7.17.1.4

General Considerations for Using TDP Enzymes as Biocatalysts

The potential for using TDP-dependent enzymes in chemical synthesis and larger scale biocatalytic processes is significant, as exemplified in the sections below (see Chapter 9.14). Several features of these enzymes make them attractive as biocatalysts. The requirement for cofactors in many enzymes can be economically limiting, such as for oxidoreductases, where the cofactor must be regenerated in a coupled enzyme reaction to avoid the expense of adding it stoichiometrically with the reactants. For TDPdependent enzymes, the TDP is regenerated in a single enzyme reaction cycle, avoiding the need to regenerate it by a coupled mechanism. However, there may be a need to retain an excess of cofactor in solution during biocatalytic processes, as the cofactor may otherwise partially dissociate and lead to loss of enzyme stability over time. Such an effect has indeed been observed for TK30 as will be discussed further below. Access to a range of carbon–carbon (C–C) bond-forming reactions is also very attractive, particularly where new stereocenters are formed along with asymmetric C–C bonds. For example, transferases such as TK and lyases such as BAL can be used to form asymmetric C–C bonds and a chiral center with a high enantioselectivity.31 TDP-enzyme catalyzed reactions are typically reversible, although the decarboxylating lyases and some transferases (e.g., ALS and DXPS) are rendered effectively irreversible by the generation of CO2. Some of the other transferase enzymes can be made to use this same strategy by the judicious use of a keto-acid substrate such as pyruvate or HPA.9 Making such reactions irreversible ensures that the reactions are driven to completion, thus achieving maximum efficiency and lower costs for biocatalysis. Most TDP-dependent enzymes have a broad phylogenetic diversity due to their presence in many widely differing organisms. Many are key housekeeping enzymes, such that they are present in all but a few exceptional organisms. This diversity, coupled with the breadth of the TDP-enzyme family,27,28 ensures that a wide variation in substrate, pH, temperature, and ionic strength preference exists already in Nature for a given enzyme. This creates a greater possibility of finding a biocatalyst that is suitable for a specific substrate or at a pH range governed by the stability of the reactants or products, using metagenomics, directed evolution, or more direct bioprospecting methods.

7.17.2

Transferases

7.17.2.1 7.17.2.1.1

Transketolase (TK) Synthetic potential of transketolases

TK (EC 2.2.1.1) has proven to be an attractive enzyme for the synthesis of a range of a,a0 -dihydroxy ketones that are of interest themselves or are synthetically useful intermediates to a range of other molecules including ketosugars and 2-amino-1,3-diols. Nonbiocatalytic routes to a,a0 -dihydroxy ketones have been reported, although these are normally multistep procedures with poor atom efficiencies, such as a five-step approach to aromatic 1,3-dihydroxyketones32 and an asymmetric strategy using the chiral auxiliary SAMP.33 A one-step biomimetic synthesis of racemic 1,3-dihydroxyketones has also been described.34 However, TK offers a stereoselective one-step approach and its ability to accept a wide range of aldehyde acceptors has made it particularly useful. TK has been used for some time as a catalyst in enzymatic and chemoenzymatic synthesis and several reviews have recorded this progress.2,31,35–37 Notably, using b-HPA as a ketol donor, a large range of aliphatic aldehyde acceptors, including

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

377

nonphosphorylated, a-hydroxylated, and non-a-hydroxylated compounds, have been used with E. coli, yeast, and spinach TK (Scheme 3). See Sprenger and Pohl for a summary of some of these data.2 HPA (free acid or Li-HPA salt) as a donor is useful synthetically as carbon dioxide is generated in the reaction, rendering the reaction irreversible9 as discussed in Section 7.17.1.4. For a-hydroxylated aldehydes, only the (2R)-isomer is accepted by wild-type TK enzymes to generate ketoses with (3S,4R) stereochemistries (1), where R1 is, for example, CH2CN, CH2NO2, and CH2CH3. For non-a-hydroxylated aldehydes with wildtype TKs, a variety of aldehydes with different groups at R2 have also been utilized, and the (3S)-product 2 is formed (Scheme 3).

OH

O O−

H

R1

+

O

TK

OH

ThDP,

O

R1

Mg 2+

OH O 3S 4R

+

O

CO2

+

CO2

1

O O−

H

+

OH

-Hydroxypyruvate

R2

OH

O

TK

OH

ThDP,

O

3S

R2

Mg2+

OH

OH

-Hydroxypyruvate

2

Scheme 3 Enantiospecificity and stereoselectivity using a-hydroxylated and non-a-hydroxylated aldehyde donors.

Some enzymatic and chemoenzymatic syntheses using TKs involve one step, whereas others are part of a multistep strategy. These include from earlier work a route to (þ)-exo-Brevicomin from racemic 2-hydroxybutanal and HPA, with six subsequent synthetic manipulations (Scheme 4). TK crucially established the D-threo configuration in the vicinal diol product 3 in a 45% yield.38 Related to this, spinach TK has been used with 2,3-dihydroxybutanal and HPA to yield 6-deoxy-L-sorbose, an intermediate used in the synthesis of furaneol, a flavor compound.39 Interestingly, a multienzymatic approach was used, producing the HPA from L-serine using serine glyoxylate aminotransferase (SGAT), and also the 2,3-dihydroxybutanal 4 from ethanal and HPA via 4-deoxy-L-erythrulose 5, which underwent subsequent microbial isomerization (Scheme 4).

OH

O H

+

O−

O

OH

TK

OH

NH2 OH O

L-Serine

SGAT

OH

OH OH

O Glyoxylate Glycine -Hydroxypyruvate

O

O

(+)-exo-Brevicomin

3 O

O−

H

OH

-Hydroxypyruvate

O−

6 steps OH

ThDP, Mg2+

O

O

H

+

OH

O

TK ThDP,

O

OH

Mg2+

4

OH

OH

6-Deoxy-L-sorbose

O

O OH

OH 5

H + O−

TK

ThDP, Mg2+

O

OH O

-Hydroxypyruvate

Scheme 4 Use of TK to prepare (þ)-exo-Brevicomin and 6-deoxy-L-sorbose.

TK has been used for the preparation of other ketoses using one- or two-step enzymatic procedures.40,41 Interest has also been expressed in industrial applications with wild-type TKs including the production of L-erythrulose (R1 ¼ H, Scheme 1) from glycolaldehyde and HPA using overexpressed E. coli TK. The reaction was characterized for reactor evaluation, and also continuous production in a membrane reactor was carried out.30,42,43 In recent work, recombinant Saccharomyces cerevisiae TK has been used to generate D-sedoheptulose-7-phosphate from HPA and D-ribose-5-phosphate in an 81% overall yield (Scheme 5) on a multigram scale.44

378

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

O OH

2−

OPO3 O

OH OH

D-Ribose-5-

+

TK

HO

ThDP, Mg2+

O HO

OH

O HPA

2−

phosphate

OH

HO

OPO3

OH

D-Sedoheptulose-7phosphate

OH

Scheme 5 Use of recombinant Saccharomyces cerevisiae TK to prepare D-sedoheptulose-7-phosphate.

Chemoenzymatic approaches using TK have been described to ketoses such as 5-thio-D-xylopyranose or ulosonic acid analogs, where key steps have been performed using TK or aldolases.45,46 An interesting two-step enzymatic procedure using a b-alanine: pyruvate transaminase (TAm) together with a wild-type TK overexpressed in a single E. coli host from separate plasmids enabled the formation of 2-amino-1,3,4-butanetriol in a 21% yield from glycolaldehyde and HPA (Scheme 6).47 This was carried out as a one-pot process using either unlysed cells of the dual plasmid or lysed cells.

OH

O H

+

HO

TK OH

-TAm

O OH

ThDP, Mg2+

O HPA

O

OH

OH

PLP, amine donor

OH

NH2 OH

OH (2S,3R)-2-amino1,3,4-butanetriol

O H O

+

TK D469E/T OH

HO

NH2

OH

ThDP, Mg2+

O HPA

CV2025 -TAm

O

OH

PLP, amine donor

OH OH (2S,3R)-2-aminopentane-1,3-diol

Scheme 6 Use of wild-type E. coli TK and a b-alanine:pyruvate transaminase to prepare (2S,3R)-2-amino-1,3,4-butanetriol, or E. coli TK D469E/ T and CV2025 o-transaminase to prepare (2S,3S)-2-aminopentane-1,3-diol.

7.17.2.1.2

Enzyme engineering of transketolase

The activity, substrate specificity, and enantioselectivity of TKs have all been extensively investigated and altered by site-directed mutagenesis of the yeast enzyme and also by both site-directed mutagenesis and directed evolution for the E. coli enzyme. Sitedirected mutagenesis of key active site residues identified from crystal structures of the yeast TK bound to substrate has confirmed many of their functional roles. For example, mutations R359A, R528A, and H469A at sites known to interact with the phosphate group of substrates increased the Km for substrates although only slightly decreased the catalytic efficiency to between 17 and 77%.48 In the same work, D477A was found to significantly impair catalytic efficiency, and a later study of the same mutant showed that this residue was also a key determinant of enantioselectivity in yeast TK.49 Similar mutations were carried out in E. coli TK,50 where S386N (yeast TK numbering), R359K, and R528K were all found to impair catalytic activity, particularly for the phosphorylated substrates with longer chain lengths, highlighting their role in stabilizing the phosphate moiety of substrates. D477E significantly decreased the enzyme activity toward natural phosphorylated substrates and glycolaldehyde, and yet retained 60% of the wild-type kcat/Km with formaldehyde, highlighting its role in hydrogen bonding to hydroxylated acceptor substrates. Mutations E162A and E162D close to the TDP cofactor and dimer interface of the yeast TK lowered the affinity for TDP and also slowed the reconstitution of the holo enzyme, confirming the role of this residue in cofactor binding and potentially also in dimerization of the enzyme.51 In the same study, mutation of the conserved D382 residue to both asparagine and alanine resulted in decreased TDP affinity and a poorly active enzyme, indicating that this residue is involved in both TDP binding and catalysis. The yeast TK active site mutation, H263A, led to a loss of activity, confirming its role in catalysis.52 However, it was also found to increase the rate constant for the release of glycolaldehyde from the enamine intermediate state. In a separate study, the yeast TK active site mutation, H103A, slowed the two-substrate reaction between xylulose-5-phosphate (X5P) and ribose-5-phosphate (R5P).53 However, the mutation increased the one-substrate reaction of X5P that forms G3P, as obtained in the two-substrate reaction, but also glycolaldehyde. The two reaction rates converged, indicating the role of H103 for stabilizing the enamine intermediate. More recently, directed evolution has been used to alter the activity, substrate specificity, and enantioselectivity of E. coli TK. Twenty E. coli TK active site residues were targeted for saturation mutagenesis.54 Half were guided by the E. coli TK crystal structure55 and in part, the studies above in yeast TK. The other ten sites were chosen using phylogenetic information27 to identify

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

379

second shell active site residues that varied in natural sequences. Using a fast HPLC assay,56 this work revealed a number of mutations (H461S, A29E, and R520V) that increased the specific activity of the enzyme for the conversion of the nonphosphorylated glycolaldehyde and HPA substrates into L-erythrulose.54 H461S and R520V were at residues equivalent to H469 and R528 in yeast TK and were known from crystal structures to interact with the phosphates of natural sugar substrates.48 These residues were relatively unimportant for binding to the nonphosphorylated substrates screened, and therefore they became free to mutate and so H461S and R520V presumably created greater access to the active site. Using the same saturation mutagenesis library as that described in the previous paragraph, and a colorimetric assay,57 a second series of mutants were identified with improved activity and substrate specificity for the non a-hydroxylated aliphatic acceptor propionaldehyde (propanal).58 Mutants at residues D469, A29, and R520 were particularly noteworthy, with D469T resulting in a fivefold increase in activity and a 22-fold shift in substrate specificity away from the conversion of glycolaldehyde. The equivalent residue in yeast TK (D477) had already been implicated in the determination of enantioselectivity and a hydrogen bond to the a-hydroxyl group in natural acceptor substrates, no longer present in propionaldehyde. Therefore, the saturation library for this residue, along with H26 and H100, was rescreened with a chiral gas-chromatography assay to specifically identify changes in enantioselectivity for the reaction of propionaldehyde and HPA.59 This screen identified mutants with enhanced (D469E) and reversed (H26Y) enantioselectivities for 90% 3S and 88% 3R 1,3-dihydroxypentan-2-one product, respectively. The D469E/T/K/L and H26Y mutants obtained from the directed evolution toward propionaldehyde were subsequently characterized for their activity and enantioselectivity with a series of nonnatural linear aldehydes from C4 to C8 chain lengths and then also C3, C5, and C6 cyclic carboxaldehydes.60 This revealed excellent ees (86–99%) in the products, particularly with the D469E mutant (Scheme 7), and therefore provides a valuable resource for expanding the synthetic potential of TK with lipophilic compounds. Furthermore, the E. coli TK mutants D469E/T/K and an additional new mutant F434A designed to widen the active site cavity have also been characterized for their activity and enantioselectivity toward a series of aromatic aldehydes such as benzaldehyde (Scheme 7) for which the wild-type enzyme showed no activity.61 This demonstrated the first examples of a TK variant able to achieve high yields and high ee (3R-isomers) with phenylacetaldehydes and also to achieve a productive reaction with benzaldehyde (F434A: 10% yield, 82% ee for 3R). The ability to use these substrates in further enzymatic conversions is an exciting prospect.

O

O H +

O−

O

OH

TK ThDP,

O

Mg2+

OH

WT TK 75% ee (3S) D469E TK 98% ee (3S) H26Y TK 92% ee (3R)

OH

WT TK no reaction F434A TK 82% ee (3R)

OH

-Hydroxypyruvate

O H +

O

O−

O OH

O

TK ThDP,

Mg2+

OH

-Hydroxypyruvate Scheme 7 Use of butanal and benzaldehyde with TK mutants and the stereoselectivities observed.

The D469T TK variant has also been coupled to a second enzyme CV20205 o-transaminase in a two-step synthetic process for chiral aminodiols in which the two enzyme steps were carried out with an intermediate ketodiol purification step.62 The 2-amino1,3-diol functionality formed is found in a range of biologically active compounds including the sphingosine lipids and broadspectrum antibiotics chloramphenicol. Using propanal and HPA, (3S)-1,3-dihydroxypentan-2-one was formed in the first step. The o-transaminase from Chromobacterium violaceum performed the second conversion using isopropylamine as the amine donor to yield (2S,3S)-2-aminopentane-1,3-diol on a preparative scale.63,64 Using a mix-and-match plasmid approach, the D469E and D469T variants of E. coli TK have also been recently reintegrated into the de novo pathway that couples TK to the CV20205 o-transaminase within E. coli. This improved the previous pathway that used wild-type TK47 to achieve a direct synthesis of (2S,3S)-2-aminopentane-1,3-diol in a 90% yield and (2S,3R)-2-amino-1,3,4butanetriol (ABT) in an 87% yield65 (Scheme 6). With a broad range of engineered TK variants now available, simple molecular biology methods for combining them with other enzymes such as transaminases, and tools at the microscale to rapidly assess reaction parameters, the number of chemo-enzymatic and multienzymatic uses of TK in synthesis is likely to expand rapidly.

7.17.2.1.3

Biocatalytic process engineering of transketolase

The use of TKs in biocatalytic processes has been extensively investigated,30,66–68 and it is currently used in a biocatalytic process for X5P synthesis43 and other sugar derivatives.69 The asymmetric formation of a new carbon–carbon bond, and the tendency to stereoselectively form keto-diol products with the S-configuration,70 makes it potentially very attractive for industrial syntheses.47,62,71–73 In particular, the ability of TK to use b-HPA as the ketol donor in place of X5P, and thus render the reaction effectively irreversible, makes its reactions more atom efficient and also potentially simplifies isolation of the product.

380

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

The acceptance of a broad range of nonphosphorylated substrates also simplifies the downstream product purification, with no additional need to remove phosphate groups from the product.71,72 TKs are also available from all known organisms, providing a diverse range of potential turnover rates and enzyme stabilities. For example, E. coli TK has been found to have 6 and 30 times greater specific enzyme activity toward HPA, than the yeast or spinach orthologs, respectively.2 A number of advances have been made to improve the usefulness of E. coli TK as a biocatalyst. Cost-effective biotransformations preferably use a low-cost biocatalyst, which typically requires improvement of the fermentation process for enzyme production. Exploration of different E. coli strains and high copy number vectors increased the production levels of TK to 43% of the total cell protein in a fed-batch 1000 l fermentation.66,74 Having obtained sufficient enzyme biocatalyst, the biotransformation process was then also optimized as it was initially found to be subject to a number of potential limitations. These could mostly be overcome using process engineering approaches. For example, the HPA and glycolaldehyde substrates, as well as the corresponding L-erythulose product were found to be most stable below pH 6, whereas the enzyme was most stable at pH 7.0 and most active at pH 6.5–8.0.30 The use of aldehyde substrates was also found to gradually decrease the enzyme activity,42 which, along with the lower substrate stability at pH 7–8,30 indicated that a substrate feeding strategy would be beneficial. A strategy of enzyme immobilization has also been found to stabilize TK by increasing the half-life for deactivation by aldehydes, as well as providing a readily reusable form of the biocatalyst.75 An alternative approach is to use membrane reactors shown previously to allow the continuous production of L-erythulose by TK.42 Kinetic modelling of the reaction suggested that a cascade of such reactors could be used to limit the deactivation by glycolaldehyde.76 The L-erythulose product was also found to inhibit TK at 4100 mM, and thus a strategy of in situ product removal using an immobilized phenylboronate resin was implemented to achieve a complete reaction at high concentrations.77 E. coli TK is potentially prone to several additional slow deactivation processes during biocatalysis, including oxidation,75 irreversible denaturation at low pH,30 and TDP dissociation.30 The addition of a reducing agent such as mercaptoethanol75 or the use of a nonoxygenated headspace gas such as nitrogen30 was able to minimize enzyme oxidation. The conformational stabilities of both yeast78,79 and E. coli80–82 TKs have been characterized extensively by denaturation studies, and also in combination with site-directed mutagenesis at residues close to both the active site and the dimer interface.51,80 The dissociation of TDP under mild chemical denaturing conditions was found to proceed via an intermediate inactive enzyme–TDP complex where the TDP is potentially not in the preferred active conformation,82 thus mirroring the intermediate state formed during yeast holoTK reconstitution from apoTK.78,79 Although the addition of excess TDP to biocatalytic processes can counter the cofactor dissociation, it is a potentially expensive route. An alternative future route to more stable TK biocatalysts could be via protein engineering of the cofactor-binding site.

7.17.2.2

Dihydroxyacetone Synthase (DHAS)/Formaldehyde Transketolase (FTK)

DHAS (EC 2.2.1.3) transfers a ketol C2 unit from X5P to formaldehyde, and like TK, can also use HPA as a donor group.2,36 The main synthetic applications have been for the preparation of isotopically labeled dihydroxyacetone 6 (Scheme 8).10 In a second step, the labeled dihydroxyacetone was phosphorylated by dihydroxyacetone kinase (DHAK) to yield dihydroxyacetonephosphate (DHAP), an aldolase substrate. O

O H

C

H

+

O C=

O−

OH

O -Hydroxypyruvate

13C

DHAS ThDP,

HO

Mg2+

C

C

ATP

6

O

DHAK

OH

HO

C

C

ADP

OPO32−

DHAP

Scheme 8 Use of DHAS to prepare isotopically labeled dihydroxyacetone 6 and DHAP.

7.17.2.3

Acetoin-Ribose-5-Phosphate Transaldolase

Acetoin-ribose-5-phosphate transaldolase (EC 2.2.1.4) has been used for the preparation of the monosaccharide 1-deoxy-D-altroheptulose 7-phosphate.83 The enzyme naturally catalyzes the reaction between racemic 3-hydroxybutan-2-one (acetoin) and D-ribose 5-phosphate to yield ethanal and the 1-deoxy-D-altro-heptulose 7-phosphate (Scheme 9).

O

OH

OH 2−

H

OPO3

+ OH 3-Hydroxybutan-2-one

O

OH

acetoin-ribose5-phosphate transaldolase ThDP, Mg2+

D-Ribose-5-phosphate

Scheme 9 The reaction catalyzed by acetoin-ribose-5-phosphate transaldolase.

O

OH

OH 2−

H

OPO3

+ O

OH

OH

1-Deoxy-D-altroheptulose 7-phosphate

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes 7.17.2.4

381

Glyoxylate Carboligase (GXC)/2-Hydroxy-3-Oxoadipate Synthase (HOAS)

As described in Section 7.17.1.2.1, GXC or HOAS (EC 2.2.1.5) catalyzes the addition of glyoxylate to 2-oxoglutarate to yield 2-hydroxy-3-oxoadipate (Scheme 10, R is CO2H).12 Early work also indicated that it can catalyze the decarboxylation of 2-oxoglutarate, and from 2-oxoglutarate and ethanal, generate 5-hydroxy-4-ketohexanoic acid (Scheme 10, R is CH3).84 When R is CO2H, HOAS can also be used to form 5-hydroxy-4-ketovaleric acid via the decarboxylation of 2-hydroxy-3-oxoadipate, and 2,3-dihydroxy-4-ketopimelic acid has also been isolated. O

O R

H

O

OH

O

HOAS

+

HO

OH O

R = CO2H or CH3

ThDP, Mg2+

2-Oxoglutarate O

HO

O OH

OH

R

O

HO

O 5-Hydroxy-4-ketovaleric acid

OH

O R = CO2H or CH3

OH

OH O 2,3-Dihydroxy-4-ketopimelic acid

Scheme 10 HOAS-catalyzed reactions using 2-oxoglutarate and two further products formed when R is CO2H.

7.17.2.5

Acetolactate Synthase (ALS)/Acetohydroxyacid Synthase (AHAS)

ALS (EC 2.2.1.6 – previous EC 4.1.3.18 is now obsolete), also referred to as AHAS, catalyzes the addition of pyruvate (R ¼ H) to a further pyruvate molecule (as mentioned in Section 7.17.1.2.1) to yield (S)-a-acetolactate 7 (R ¼ R1 ¼ H) or a-ketoglutarate (R ¼ CH3) to yield (S)-a-aceto-a-hydroxybutyrate 8 (R ¼ H, R1 ¼ CH3).11 It also catalyzes the addition of two a-ketoglutarates to yield 9 (R ¼ R1 ¼ CH3) (Scheme 11).

O HO

O R

+

R is H or CH3

AHAS

R

ThDP, Mg

O

O

O

R1

HO

2+

R1

HO CO −2

R1 is H or CH3

7 R = R1 = H 8 R = H, R1 = CH3 9 R = R1 = CH3

Scheme 11 AHAS reactions using pyruvate (R ¼ H) or a-ketoglutarate (R ¼ CH3).

In more recent studies, it has been established that three AHAS isozymes in E. coli catalyze the addition of pyruvate to benzaldehyde to yield (R)-phenylacetylcarbinol ((R)-PAC), a reaction that is also catalyzed by PDC.85 Notably, some of the AHAS isozymes produced only negligible amounts of ethanal during the reaction whereas the (R)-PAC product was generated with high ees (498%) (Scheme 12).

O

OH

O H +

AHAS

HO

ThDP, Mg2+ O

O (R)-Phenylacetylcarbinol (R)-PAC

Scheme 12 Formation of (R)-PAC using AHAS.

The substrate range was explored using the most effective isozyme, AHAS I, with pyruvate and a range of substituted benzaldehydes, which were accepted in up to a 99% yield. Heteroaromatic aldehydes such as the pyridine carboxaldehydes and thiophene carboxaldehyde were also readily accepted, together with the aliphatic substrate cyclohexane carboxaldehyde, highlighting its versatility in synthesis.86 In later work, AHAS I was used in a continuous flow reactor with pyruvate and benzaldehyde for (R)-PAC scale-up studies.87

382

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

7.17.2.6

1-Deoxy-D-Xylulose-5-Phosphate Synthase (DXPS) (EC 2.2.1.7)

DXPS is a biosynthetic precursor of isopentenyl diphosphate via the nonmevalonate pathway, and also of TDP and pyridoxal phosphate in bacteria.2,31,36 Since the identification of DXPS, which uses pyruvate and D-glyceraldehyde 5-phosphate as substrates,8 there has been interest in synthetic applications. For example, using overexpressed E. coli DXPS D-erythrose 4-phosphate and D-ribose 5-phosphate have been used as alternative acceptors to readily generate 1-deoxy-D-fructose 6-phosphate and 1-deoxy88,89 D-sedoheptulose-7-phosphate, respectively, on a semipreparative scale (Scheme 13). DXPS also accepts nonphosphorylated 90 aldehydes, which is useful synthetically as it eliminates the need to remove phosphate later.

O

O OH +

OH

OH

H

O

OH

DXPS

OPO32−

OPO32−

ThDP, Mg2+

OH

O

OH

1-deoxy-D-fructose 6phosphate

D-Erythrose

4-phosphate O

O OH +

O

OH

OH OPO3

H OH D-Ribose

OH

OH

DXPS

2−

OPO32−

ThDP, Mg2+

O

OH

OH

1-deoxy-D-sedoheptulose 7-phosphate

5-phosphate

Scheme 13 Use of DXPS to yield 1-deoxy-D-fructose 6-phosphate and 1-deoxy-D-sedoheptulose 7-phosphate.

Recently, the substrate promiscuity of DXPS has been investigated using aliphatic aldehydes such as propanal and butanal, which were accepted, although much less readily than D-glyceraldehyde 5-phosphate.91 In the absence of D-glyceraldehyde 5-phosphate DXPS will use pyruvate alone and form acetoin and acetolactate.91 The crystal structure of the E. coli DXPS with two segments missing due to fungal proteolysis and a Deinococcus radiodurans DXPS have been determined and show an unusual arrangement of the domains compared with TK.23 In DXPS, which, like TK, is a dimer, domain I is directly above domains II and III of the same monomer whereas in TK domain I is positioned above domains II and III of the other monomer. This means that each of the two active sites in DXPS is composed of residues from only one monomer compared with TK, where the active site has contributions from both monomers.

7.17.3

Lyases

7.17.3.1 7.17.3.1.1

Benzoylformate Decarboxylase (BFDC) Synthetic potential of BFDC

The BFDC (EC 4.1.1.7) enzyme from Pseudomonas putida is a homotetramer of 528 amino acid subunits that catalyzes the removal of CO2 from benzoylformate to generate benzaldehyde in the mandelate pathway (see Chapter 7.19).19 It can be used to prepare a range of hydroxyketones when provided with two aldehydes such as benzaldehyde and acetaldehyde.92 BFDC accepted a range of linear and branched aliphatic aldehydes to produce very high yields (often 490%) of the 2-hydroxyketones, and a range of ees. For example, acetaldehyde was converted into (R)-acetoin with 34% ee, and isovaleraldehyde was converted into (R)-5-hydroxy2,7-dimethyloctan-4-one with 85% ee (Scheme 14).93 However, in the same study, pivaldehyde (trimethyl acetaldehyde)

O

O

BFDC H

ThDP, Mg2+ OH Acetoin

Acetaldehyde O

O

BFDC H

ThDP, Mg2+

Isovaleraldehyde Scheme 14 Use of BFDC with aliphatic aldehydes to yield acyloins.

OH (5 R)-5-hydroxy-2,7-dimethyl octan-4-one

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

383

isobutyraldehyde and tert-butyl acetaldehyde led to no conversion to products. These bulky, short aldehydes are blocked in some way from either forming the intermediate adduct with the TDP or the adduct blocks the next stage of the reaction. When aromatic aldehydes based on benzaldehyde are used, benzoin products are formed. The recombinant P. putida BFDC synthesizes good yields of (R)-benzoins 60–70%94 and with very high ee obtained in most cases (Scheme 15). For example, 4-methyl benzaldehyde, 2-fluoro benzaldehyde both yield their respective (R)-benzoins with an ee of 499%. High yields were also observed with 2-furyl benzaldehyde (62% yield, ee 94%) and 2-thiophenyl benzaldehyde (65% yield, ee 95%) but not with cyano, bromo, or methoxy moieties at the ortho position (0, o2%, and o2% yield, respectively). Interestingly, halogens at the para position of benzaldehyde produced decreasing yields of product with increasing size, for example, the 4-fluoro, 4-chloro, and 4-bromo benzaldehydes produced their benzoins in 25%, 17%, and 13% yields, respectively. All of these products were obtained as the (R) enantiomer.

O R

O

BFDC H ThDP,

R

Mg2+ R

OH (R)-Benzoin

R is H or 2- or 4substituted

R 2-F 2-furyl 2-SH 2-Br 2-OMe

Yield (%) 68 62 65 <2 <2

R 4-F 4-Cl 4-Br 4-Me 4-OMe

Yield (%) 25 17 13 69 12

Scheme 15 Use of BFDC with substituted aromatic aldehydes to yield benzoins.

Recently, an interesting reversible effect of high carbon dioxide pressure on the enantioselectivity of P. putida BFDC has been observed.95 Increasing the pressure up to 250 MPa led to a shift in the ee% of 2-hydroxy-propriophenone from 490% (S) with carbon dioxide at ambient pressure, to a greater amount of the (R) form. The wild-type enzyme shows a relatively small change in the ee, but one particular double mutant, A460I-F464I, induces a significant shift to 75% of the (R) form at 250 MPa. This highlights an interesting form of process control that can be exerted to obtain the desired ee of product.

7.17.3.1.2

Enzyme engineering of BFDC

The P. putida BDFC (PpBFDC) has been subjected to directed evolution in two independent studies. The first study applied a single round of error-prone PCR (epPCR) to identify mutants with fivefold improved carboligase activity to convert benzaldehyde and acetaldehyde into (S)-2-hydroxy-1-phenyl-propanone (HPP) as shown in Scheme 16.96 Beneficial mutations were found at residue L476(P/Q), which is found in a surface loop that forms a hydrophobic network at the enzyme–dimer interface, and is also in close proximity to the TDP cofactor. Interestingly, the mutation L476Q also increased the tolerance of the enzyme to organic solvents ethanol (1.5 M) and DMSO (20% v/v). The L476 residue was proposed to act as a gatekeeper for substrates and product in and out of the PpBFDC active site. Further saturation mutagenesis of this residue revealed that nine different substitutions could improve the carboligase activity, all with improved enantioselectivity (93–97% ee) over the wild type (85% ee), although none induced any significant improvement in activity or solvent tolerance over the original L476Q mutation. Later, the same group altered the substrate specificity of the PpBFDC, again using epPCR.97 L476Q was again identified, along with new mutant M365L-L461S, to convert ortho-substituted benzaldehydes with typically 499% ee for the S-isomer HPP derivative products, where no previous reaction was observed for the wild-type enzyme. O

H

BFDC/BAL

+

O H

Benzaldehyde

Acetaldehyde

ThDP, Mg2+

O

OH (S)-2-hydroxy-1-phenyl propanone

Scheme 16 Benzoylformate decarboxylase (BFDC)- and benzaldehyde lyase (BAL)-catalyzed carboligase reaction.

In the second study, a crystal structure of the PpBFDC was used to design three active-site mutations (S26A, H70A, and H281A)98 examined for their impact on BFDC activity. S26A resulted in a 23-fold increase in the Km for benzoylformate and a 100-fold increase in the Ki for the competitive inhibitor (R)-mandelate, implicating it in substrate binding. The kcat for S26A also decreased 54-fold. H70A decreased the kcat 3400-fold and increased Km fivefold, whereas H281A decreased the kcat 171-fold and increased Km fourfold; thus, all three residues were identified as catalytically important. Saturation mutagenesis of these putative catalytic residues was later performed.99 This demonstrated a surprising tolerance at all three sites to residues that would remove apparently important catalytic interactions, although the wild-type residue was optimal for both kcat and Km in all cases. A further 12 active-site residues within 5 A˚ of the mandelate inhibitor-binding site were subjected to saturation mutagenesis and screened

384

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

for the decarboxylation of benzoylformate and four other substrates including pyruvate.100 The mutants T377L and A460Y showed the most significant shifts in activity to the new substrates, notably pyruvate, and these two sites were recombined in a second round of saturation mutagenesis. The double mutant T377L/A460Y proved to result in the greatest (11 000-fold) shift in substrate preference toward pyruvate from benzoylformate, and closely approached the kcat/Km of PDC from Zymomonas mobilis (ZmPDC). Interestingly, the mutations representing the highly conserved residues that occur in the equivalent positions for ZmPDC (A460V/I and T377G) did not significantly improve the activity of PpBFDC toward pyruvate. Rather, residues that did not appear in natural PDCs proved to yield the best PDC-like activity in PpBFDC, demonstrating how the use of sequence homology comparisons to determine the specific mutations to make can be very misleading, although their identification of potential sites for saturation mutagenesis is still of value.101 These experiments also demonstrate the relative ease with which the substrate specificity of TDP-dependent decarboxylases can be altered, which may become useful in creating novel biocatalysts for the reverse reactions of CO2 addition at high pressures.

7.17.3.2 7.17.3.2.1

Benzoin Aldolase/Benzaldehyde Lyase (BAL) Synthetic potential of BAL

BAL (EC.4.1.2.38), also known as benzoin aldolase, was originally characterized from Pseudomonas fluorescens102 as an enzyme possibly involved in the degradative assimilation of benzoin and anisoin that are potential lignin breakdown products. It was later shown to be able to catalyze the reverse reaction of aromatic 2-hydroxyketone formation by coupling two molecules of benzaldehyde to yield benzoin, and also the coupling of other aromatic aldehydes,103,104 and linear aldehydes such as acetaldehyde, which yields 2-HPP. If benzoin is cleaved in the presence of acetaldehyde, a replacement reaction occurs to yield 2-HPP. BAL will also couple formaldehyde to benzaldehyde and other aromatic aldehydes, to produce hydroxyacetophenones in a high yield when excess formaldehyde is used.105 Linear aldehydes such as butanal or pentanal alone can be coupled to form their respective acyloins (Scheme 17).106 Similarly, propanal can be coupled to yield proprion,107 and branched aliphatic aldehydes such as 3-methylbutanal (isovaleraldehyde) generate (R)-5-hydroxy-2,7-dimethyloctan-4-one with a high yield (490%) and an ee of 89% (R).93

O

O

BAL H

ThDP,

Mg2+ OH

Propanal

(R)-Proprion O

O

BAL H

ThDP,

Mg2+ OH

Pentanal

(6 R)-6-Hydroxy-decan-5-one Scheme 17 Use of BAL to generate linear acyloins.

Complex polyols such as protected 1,3,4-trihydroxy-2-butanone can be accessed using BAL in direct and cross-coupling reactions of benzyloxyacetaldehyde with dimethoxyacetaldehyde to form 1-(benzyloxy)-3-hydroxy-4,4-dimethoxybutan-2-one108 (Scheme 18). The synthesis of benzoin has even been attempted using benzaldehyde in the gas phase,109 and although substrates were consumed, the product, being nonvolatile, remained bound to the beads containing the enzyme and so could not be easily quantified.

O H

O

O

H

+ O

Benzyloxyacetaldehyde

OH

BAL

O Dimethoxy acetaldehyde

ThDP, Mg2+

O

O O

O

1-(benzyloxy)-3-hydroxy-4,4dimethoxybutan-2-one

Scheme 18 Use of BAL in an asymmetric cross condensation.

7.17.3.2.2

Enzyme engineering of BAL

The factors that distinguish the activity of BAL from P. fluorescens (PfBAL) from that of PpBFDC, which can also catalyze the carboligation of benzaldehyde to benzoin, were initially explored by homology modeling of the protein structure to that of

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

385

PpBFDC and then making key active site mutations.110 Three potentially important sites in BAL, A28, H29, and Q113, which showed differences to the equivalent sites in BFDC, were chosen and mutated independently to A28S, H29A, and Q113H/A. These were compared with the BFDC mutations S26A (equivalent to A28 in BAL) and H70A/Q, two of which are described above. None of the mutations in BAL significantly affected the Km for benzoin, with Q113A decreasing Km the greatest, by a factor of two. However, all four mutations in BAL resulted in between 13-fold (A28S) and 350-fold (Q113H) decreases in kcat for the lyase activity with benzoin, indicating some role in catalysis for each of the three sites. The same group has more recently confirmed that H29A in BAL decreases both the lyase and the ligase activities,111 suggesting that H29 plays a role in water-mediated general acid–base catalysis. They also found that the W163A mutation exerted a long-distance (10 A˚) decrease in TDP affinity, but also decreased the benzoin ligase activity. This was speculated to be mediated by a potential p-stacking arrangement with H29. S26 in BFDC (A28 in BAL) was previously implicated in benzoylformate substrate binding, but also in catalysis. Interestingly, the A28S mutation in BAL introduced BFDC activity that was not detected in the wild-type BAL enzyme. Furthermore, the converse mutation in BFDC did not introduce the lyase activity with benzoin. Serine at this position is therefore important for the catalysis of benzoylformate decarboxylation in either enzyme, but neither serine nor alanine was sufficient to create a productive binding site for benzoin in BFDC. A recently obtained crystal structure for BAL112 enabled a more detailed mutagenic study of the BAL active site113 that also compared equivalent or converse mutations in PpBFDC. It was found that the primary difference between BAL and BFDC is steric, with the much wider binding site in BAL. The active site entrance is also slightly shifted between the two enzymes. The H281A mutation that increased the active site cavity size in BFDC notably increased the ligase activity to form benzoin from benzaldehyde. However, the equivalent mutation H286A in BAL diminished the ligase activity but had little effect on the benzoin lyase activity. The lesser effect in BAL was attributed to the greater distance of H286 from the active site due to the shifted position of the active site entrance. The difference in active site sizes between BAL and BDFC was also found to influence the enantioselectivity of the two enzymes for the ligase activity, with BAL forming (R)-2-HPP and BFDC forming the S isomer.

7.17.3.2.3

Biocatalytic process engineering of BAL

Process engineering aspects of biocatalysis with BAL have begun to be addressed for potential industrial-scale syntheses. For example, reaction engineering of PfBAL led to a continuous process for the synthesis of (R)-2-HPP from benzaldehyde and acetaldehyde114 that minimized the production and hence precipitation of the unwanted benzoin side-product. This process used an enzyme membrane reactor, with regenerated cellulose to retain the enzyme, and to produce 2-HPP with a space–time yield of 1120 g l1 d1, and also (R)-(3-chorlophenyl)-2-hydroxy-1-propanone at 1214 g l1 d1, both with ee 499%. This far outperformed the author’s previous use of a batch reactor, which achieved a space–time yield of only 36 g l1 d1 for 2-HPP. Application of the enzyme membrane reactor configuration to the synthesis of other 2-HPP has also been demonstrated more recently.115 In an alternative approach, His-tagged BAL has been immobilized on a super-paramagnetic solid support (maghemitesilica) that was surface derivatized with Co2 þ -nitriloacetic acid (NTA). This heterogeneous biocatalyst form was able to perform carboligations to 2-HPP and also another derivative, while also facilitating the reuse of the enzyme.116

7.17.3.3

Branched-Chain 2-Ketoacid Decarboxylase

The branched-chain 2- KdcA from Lactococcus lactis (LlKdcA) was recently overexpressed in E. coli and subjected to active-site mutagenesis based on a homology model.20 The kinetics for wild-type KdcA were also determined for a range of ketoacid substrates. Although the kcat values for various branched chain ketoacids were most prominent, the kcat/Km for the enzyme was greatest by a factor two for phenylpyruvate. Based on the sequence and structure comparisons, the mutants V461I, S286Y, F381W, and M538W were made to reduce the substrate binding cavity toward that of ZmPDC. These mutations increased the substrate specificity by varying degrees toward pyruvate, whereas they mostly (except F381W) resulted in a significant decrease in the acceptance of phenylpyruvate. The results broadly indicated that sterics and the active site cavity played a significant role in determining the substrate preference of both the KdcA and the PDC enzymes.

7.17.3.4 7.17.3.4.1

Pyruvate Decarboxylase (PDC) Synthetic potential of PDC

PDC (EC 4.1.1.1) is present in yeast and many bacteria, where it decarboxylates pyruvate to yield acetaldehyde. The acetaldehyde is often then reduced by alcohol dehydrogenase to make ethanol in organisms such as S. cerevisiae and Z. mobilis. It has long been known that useful side reactions can occur with PDC. For example, acetolactate can be generated if a second molecule of pyruvate enters the active site when the 2-carbon unit is still on the TDP. Similarly, acetoin can be formed if acetaldehyde is present, and PAC is produced if benzaldehyde is present in the reaction mixture. PDC will also react b-HPA and glycolaldehyde to produce 1,3,4-trihydroxy-2-butanone (erythrulose), as catalyzed also by TK.17 The production of (R)-PAC is important as it is a precursor for ephedrine and pseudoephedrine, and several processes using PDC have been described (Scheme 19), similar to that described in Section 7.17.2.5 for AHAS (Scheme 12). Several use whole cells of S. cerevisiae or Candida utilis and these have become increasingly sophisticated.117–119 Others use crude enzyme preparations from C. utilis120 or recombinant E. coli whole-cell biocatalysts expressing the Z. mobilis PDC enzyme.121 PDC and aldehyde dehydrogenase II genes (pdc and adhB) from Z. mobilis have

386

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

O

O H

H

+

OH

PDC ThDP, Mg2+

O (R)-Phenylacetylcarbinol (R)-PAC Scheme 19 Use of PDC to prepare (R)-PAC.

also been coexpressed in an engineered Hansenula polymorpha strain to improve the production of ethanol from glycerol by more than threefold.122

7.17.3.4.2

Biocatalytic process engineering of PDC

In addition to examining the various biocatalyst forms of PDC for synthetic potential, different process options have been explored. A mutant (W392M) of PDC from Z. mobilis with greater activity for the synthesis of (R)-PAC was first used in a continuous fed-batch process. The same synthesis was then also operated in an enzyme membrane reactor for the continuous production of PAC.123 Experimental exploration and reaction modelling of this process led to the optimization of the substrate feeding regime to obtain the product with a space–time yield of 81 g l1 d1. The final concentration of the product could also be doubled by cascading two enzyme membrane reactors and feeding an additional substrate after the first reactor step. The PAC synthesis above has also been operated as an aqueous/organic two-phase reaction using the same mutant PDC enzyme,124 with the aim of increasing the process intensity and reducing the toxicity of the aldehyde substrates to the enzyme. The aldehyde substrates were fed in the organic phase and the PAC product partitioned back into it for easier product removal. After testing a wide range of organic solvents, various alcohols were found to yield the highest specific production of PAC and almost a 10-fold improvement over that in the absence of an organic solvent.

7.17.3.5

Phosphoketolase

Phosphoketolase (EC 4.1.2.9) is a key enzyme in the Bifidobacterial pathway of glucose assimilation and is an atypical TDP-containing enzyme as it carries out a dehydration step on the TDP enamine adduct, which is then cleaved by inorganic phosphate to yield acetyl phosphate. A crystal structure of phosphoketolase125 reveals the location of the phosphate group and the structures of the TDP intermediates before and after the dehydration step. In this manner, phosphoketolase cleaves fructose-6-phosphate to yield acetyl phosphate and erythrose-4-phosphate (Scheme 20). X5P is also cleaved in the same pathway to yield acetyl phosphate and G3P.

OH

OH

OH

Phosphoketolase ThDP, Mg 2+

OPO32−

OPO32− O

OH

Fructose-6-phosphate

O +

O PO32−

Acetylphosphate

OH OPO32−

H OH

Erythrose-4-phosphate

Scheme 20 Phosphoketolase cleavage of fructose-6-phosphate.

The phosphoketolase enzyme is largely confined to the Bifidobacteria and certain Bacillus species such as Bacillus coagulans. It has not been used as a standalone enzyme for biocatalysis as yet and it has received prominence as the key enzyme in bacteria such as B. coagulans, which can grow at high temperatures of 50 1C, and also be used to ferment waste plant and biomass material to generate lactate and ethanol.126

7.17.4 7.17.4.1

Outlook Protein Engineering

At the sequence level, TDP enzymes are conserved most strictly at those residues that form the TDP-binding motif, but vary considerably at residues important for their catalysis and substrate binding.26,27 Along with mutagenic and structural studies, this indicates that the natural evolution of such a wide range of activities and substrate acceptance within each class of TDP-dependent enzymes has mostly been influenced by the organization of the active site in proximity to the cofactor. The wild-type enzymes exhibit a significant degree of promiscuity in terms of substrate acceptance, but also notably for their catalytic mechanisms and the resulting classes of activity. A particular example is demonstrated by the close interrelatedness of the benzoin lyase, benzoin ligase, and benzoylformate decarboxylation activities of BAL and BFDC enzymes. That single mutations can significantly alter these

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

387

relative activities, even introducing one anew where the wild type showed no detectable activity, is also a testament to the versatility of these enzymes. Meanwhile, many of the active sites appear to be remarkably tolerant to mutations. It therefore seems that TDP enzymes could have evolved their different functionalities with relatively few mutations within the active sites while utilizing the same basic chemical properties of the cofactor held in place by a highly conserved binding pocket. This raises the distinct possibility that there is still much greater unexplored potential for organic synthesis that these enzymes can bring via more extensive protein engineering. For altering their substrate specificity, there is considerable scope for enzyme engineering, where the existing enzymes already show that they can accept a wide range of substrates with different size, steric bulk, polarity, and aromaticity. The large active sites of many of these enzymes also seem to present few potential limits to increasing substrate size. Several studies described above have aimed to interconvert the substrate specificity between those of the naturally identified enzymes using rational mutagenesis guided by sequence and structure information. Meanwhile, a few examples of directed evolution have shown that many nonnatural substrate derivatives can be accessed with relatively few mutations. Such studies are actually only just beginning to explore the potential diversity, and there has been no indication as yet that any limits (if they exist) are being reached in terms of the range of substrates they can be engineered to utilize. The diversity of TDP-dependent enzyme catalysis continues to surprise with the addition of a new and unusual mechanism in the case of CAS (Scheme 2). Perhaps this too is suggestive of the potential to engineer novel reactions in TDP-dependent enzymes or for the activated aldehyde to accept alternative electrophiles to the proton or carbonyl. All too little has been achieved as of yet to obtain or engineer TDP-dependent enzymes that are more stable under biocatalytic process conditions, such as at higher temperatures or in the presence of cosolvents. Although a single mutation in PpBFDC was found to increase the tolerance of the enzyme to some organic cosolvents, as described above, this was a serendipitous event and such mutations have not been actively pursued in these enzymes. This in part may be a reflection of the increased challenge associated with the relatively large size and complexity of TDP enzymes as they are multimeric and require two or more cofactors. Enzyme engineering might therefore need to be fairly extensive to achieve notable gains in stability.

7.17.4.2

Bioprospecting

Metagenomics and the experimental screening of microorganisms have continued to identify new variants of the TDP-dependent enzymes, even including those with the potential to catalyze new types of reaction as observed for CAS. This approach is likely to continue as a favored route to new enzymes that may be useful in synthetic applications. Broadly, a much greater variation has been observed with the substrate spectrum of the lyases than it has with the transferases. This most likely stems from the role of the lyases in catabolic activities, and where their evolution toward novel substrates provides access to new carbon sources that enable cellular growth. The transferases, by contrast, are more involved in central anabolic metabolism in ways that do not vary considerably between different organisms. It might therefore be expected that the search for transferases with novel substrate specificities in different organisms would be in vain. However, protein engineering of transferases has shown significant potential for broadened substrate specificity, and so the appearance of more promiscuous transferases in nature cannot be ruled out. Beyond substrate specificity, the search for both lyases and transferases from other organisms has additional potential to obtain enzyme variants with improved properties for biocatalysis. In particular, enzymes from extremophiles that are more thermostable or solvent tolerant would be useful for the industrial-scale synthesis using substrates with poor aqueous solubility. Furthermore, the extension of their pH range or optimum, especially to lower pH, would prove useful where substrates or products are pH labile.

7.17.4.3

Process Engineering

To date, relatively few examples of TDP-dependent enzymes have been fully exploited in terms of reaction and process engineering, with TK, BAL, and PDC receiving the most attention as described in the relevant sections above. Common approaches have been explored with each, such as the use of enzyme immobilization and also the optimization of continuous enzyme membrane reactor configurations. The challenges of substrate solubility, toxicity of aldehydes, cofactor dissociation, and product inhibition are likely to be similar for many of the different TDP-dependent enzyme classes. Therefore, the lessons learnt so far with these few enzyme examples should have significant potential for more rapidly designing suitably scalable biocatalytic processes for other systems.

7.17.5

Conclusions

The range of synthetic applications of TDP-dependent enzymes has grown rapidly due to the continual discovery of new types of reaction catalyzed, promiscuous activities, and also via metagenomic and mutagenic exploration of substrate specificities and activities. Coupled with some solid examples of reaction engineering and modeling, for the optimized design of biocatalytic processes, the utility of these enzymes for the manufacturing of high-value molecules at scale is ripe for the picking.

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References 1. 2. 3. 4. 5. 6. 7. 8. 9. 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. 61. 62. 63. 64. 65. 66. 67. 68. 69.

Kanehisa, M.; Goto, S. Nucleic Acids Res. 2000, 28(1), 27–30. Sprenger, G. A.; Pohl, M. J. Mol. Catal. B: Enzym. 1999, 6(3), 145–159. Jordan, F. Nat. Prod. Rep. 2003, 20(2), 184–201. Asztalos, P.; Parthier, C.; Golbik, R.; et al. Biochemistry 2007, 46(43), 12037–12052. Meshalkina, L. E.; Kochetov, G. A.; Hubner, G.; Tittmann, K.; Golbik, R. Biochemistry (Mosc.) 2009, 74(3), 293–300. Pei, X. Y.; Erixon, K. M.; Luisi, B. F.; Leeper, F. J. Biochemistry 2010, 49(8), 1727–1736. Sprenger, G. A.; Schorken, U.; Sprenger, G.; Sahm, H. Eur. J. Biochem. 1995, 230(2), 525–532. Sprenger, G. A.; Schorken, U.; Wiegert, T.; et al. Proc. Natl. Acad. Sci. USA 1997, 94(24), 12857–12862. Datta, A. G.; Racker, E. J. Biol. Chem. 1961, 236, 617–623. Yanase, H.; Okuda, M.; Kita, K.; et al. Appl. Microbiol. Biotechnol. 1995, 43(2), 228–234. Barak, Z.; Calvo, J. M.; Schloss, J. V. Methods Enzymol. 1988, 166, 455–458. Schlossberg, M. A.; Richert, D. A.; Bloom, R. J.; Westerfeld, W. W. Biochemistry 1968, 7(1), 333–337. Jiang, M.; Cao, Y.; Guo, Z. F.; et al. Biochemistry 2007, 46(38), 10979–10989. Caines, M. E. C.; Elkins, J. M.; Hewitson, K. S.; Schofield, C. J. J. Biol. Chem. 2004, 279(7), 5685–5692. Caines, M. E. C.; Sorensen, J. L.; Schofield, C. J. Biochem. Biophys. Res. Commun. 2009, 385(4), 512–517. Sevostyanova, I. A.; Solovjeva, O. N.; Kochetov, G. A. Biochem. Biophys. Res. Commun. 2004, 313(3), 771–774. Baykal, A.; Chakraborty, S.; Dodoo, A.; Jordan, F. Bioorg. Chem. 2006, 34(6), 380–393. Graupner, M.; Xu, H.; White, R. H. J. Bacteriol. 2000, 182(17), 4862–4867. McLeish, M. J.; Kneen, M. M.; Gopalakrishna, K. N.; et al. J. Bacteriol. 2003, 185(8), 2451–2456. Yep, A.; Kenyon, G. L.; McLeish, M. J. Bioorg. Chem. 2006, 34(6), 325–336. Chang, Y. Y.; Wang, A. Y.; Cronan, J. E. J. Biol. Chem. 1993, 268(6), 3911–3919. Chabriere, E.; Charon, M. H.; Volbeda, A.; et al. Nat. Struct. Biol. 1999, 6(2), 182–190. Xiang, S.; Usunow, G.; Lange, G.; Busch, M.; Tong, L. J. Biol. Chem. 2007, 282(4), 2676–2682. Muller, Y. A.; Lindqvist, Y.; Furey, W.; et al. Structure 1993, 1(2), 95–103. Duggleby, R. G. Acc. Chem. Res. 2006, 39(8), 550–557. Schenk, G.; Layfield, R.; Candy, J. M.; Duggleby, R. G.; Nixon, P. F. J. Mol. Evol. 1997, 44(5), 552–572. Costelloe, S. J.; Ward, J. M.; Dalby, P. A. J. Mol. Evol. 2008, 66(1), 36–49. Widmann, M.; Radloff, R.; Pleiss, J. BMC Biochem. 2010, 9. Bommer, M. J. S. Accelerating the Discovery of Transaminase Biocatalysts; University College London: London, UK, 2007. Mitra, R. K.; Woodley, J. M.; Lilly, M. D. Enzyme Microb. Technol. 1998, 22(1), 64–70. Pohl, M.; Lingen, B.; Muller, M. Chem. Eur. J. 2002, 8(23), 5288–5295. Fetizon, M.; Goulaouic, P.; Hanna, I. Tetrahedron Lett. 1985, 26(40), 4925–4928. Enders, D.; Voith, M.; Ince, S. J. Synthesis – Stuttgart 2002, (12), 1775–1779. Smith, M. E. B.; Smithies, K.; Senussi, T.; Dalby, P. A.; Hailes, H. C. Eur. J. Org. Chem. 2006, (5), 1121–1123. Takayama, S.; McGarvey, G. J.; Wong, C. H. Annu. Rev. Microbiol. 1997, 51, 285–310. Schorken, U.; Sprenger, G. A. Biochim. Biophys. Acta – Proteins Proteomics 1998, 1385(2), 229–243. Muller, M.; Gocke, D.; Pohl, M. FEBS J. 2009, 276(11), 2894–2904. Myles, D. C.; Andrulis, P. J.; Whitesides, G. M. Tetrahedron Lett. 1991, 32(37), 4835–4838. Hecquet, L.; Bolte, J.; Demuynck, C. Tetrahedron 1996, 52(24), 8223–8232. Andre, C.; Demuynck, C.; Gefflaut, T.; et al. J. Mol. Catal. B: Enzym. 1998, 5(1–4), 113–118. Hecquet, L.; Demuynck, C.; Schneider, G.; Bolte, J. J. Mol. Catal. B – Enzym. 2001, 11(4–6), 771–776. Bongs, J.; Hahn, D.; Schorken, U.; et al. Biotechnol. Lett. 1997, 19(3), 213–215. Shaeri, J.; Wohlgemuth, R.; Woodley, J. M. Org. Process Res. Dev. 2006, 10(3), 605–610. Charmantray, F.; Helaine, V.; Legeret, B.; Hecquet, L. J. Mol. Catal. B – Enzym. 2009, 57(1–4), 6–9. Crestia, D.; Demuynck, C.; Bolte, J. Tetrahedron 2004, 60(10), 2417–2425. Charmantray, F.; Dellis, P.; Helaine, V.; Samreth, S.; Hecquet, L. Eur. J. Org. Chem. 2006, (24), 5526–5532. Ingram, C. U.; Bommer, M.; Smith, M. E.; et al. Biotechnol. Bioeng. 2007, 96(3), 559–569. Nilsson, U.; Meshalkina, L.; Lindqvist, Y.; Schneider, G. J. Biol. Chem. 1997, 272(3), 1864–1869. Nilsson, U.; Hecquet, L.; Gefflaut, T.; Guerard, C.; Schneider, G. FEBS Lett. 1998, 424(1–2), 49–52. Schorken, U.; Sahm, H.; Sprenger, G. A. In Substrate Specificity; Site Directed Mutagenesis and Modelling of the Substrate Channel and Preliminary X-ray Crystallographic Data of E. Coli Transketolase ; Bisswanger, H., Schellenberger, A., Eds.; Intemann: Prien, 1996, pp 543–554. Meshalkina, L.; Nilsson, U.; Wikner, C.; Kostikowa, T.; Schneider, G. Eur. J. Biochem. 1997, 244(2), 646–652. Fiedler, E.; Golbik, R.; Schneider, G.; et al. J. Biol. Chem. 2001, 276(19), 16051–16058. Selivanov, V. A.; Kovina, M. V.; Kochevova, N. V.; Meshalkina, L. E.; Kochetov, G. A. FEBS Lett. 2004, 567(2–3), 270–274. Hibbert, E. G.; Senussi, T.; Costelloe, S. J.; et al. J. Biotechnol. 2007, 131(4), 425–432. Littlechild, J.; Turner, N.; Hobbs, G.; et al. Acta Crystallogr. Sect D: Biol. Crystallogr. 1995, 51(Pt 6), 1074–1076. Miller, O. J.; Hibbert, E. G.; Ingram, C. U.; Lye, G. J.; Dalby, P. A. Biotechnol. Lett. 2007, 29(11), 1759–1770. Smith, M. E.; Kaulmann, U.; Ward, J. M.; Hailes, H. C. Bioorg. Med. Chem. 2006, 14(20), 7062–7065. Hibbert, E. G.; Senussi, T.; Smith, M. E.; et al. J. Biotechnol. 2008, 134(3–4), 240–245. Smith, M. E. B.; Hibbert, E. G.; Jones, A. B.; Dalby, P. A.; Hailes, H. C. Adv. Synth. Catal. 2008, 350(16), 2631–2638. Cazares, A.; Galman, J. L.; Crago, L. G.; et al. Org. Biomol. Chem. 2010, 8(6), 1301–1309. Galman, J. L.; Steadman, D.; Bacon, S.; et al. Chem. Commun. 2010, 46(40), 7608–7610. Smith, M. E. B.; Chen, B. H.; Hibbert, E. G.; et al. Org. Process Res. Dev. 2010, 14(1), 99–107. Hailes, H. C.; Dalby, P. A.; Lye, G. J.; Ward, J. M. Chim. Oggi – Chem. Today 2009, 27(4), 28–31. Hailes, H. C.; Dalby, P. A.; Lye, G. J.; et al. Curr. Org. Chem. 2010, 14(17), 1883–1893. Rios-Solis, L.; Halim, M.; Cazares, A.; et al. Biotechnol. Bioeng. 2011 (in press). Hobbs, G. R.; Mitra, R. K.; Chauhan, R. P.; Woodley, J. M.; Lilly, M. D. J. Biotechnol. 1996, 45(2), 173–179. Hobbs, G. R.; Lilly, M. D.; Turner, N. J.; et al. J. Chem. Soc. Perkin Trans. 1 1993, (2), 165–166. Morris, K. G.; Smith, M. E. B.; Turner, N. J.; et al. Tetrahedron: Asymmetry 1996, 7(8), 2185–2188. Wohlgemuth, R. J. Mol. Catal. B: Enzym. 2009, 61(1–2), 23–29.

C–X Bond Formation: C–C Bond Formation using TDP-Dependent Enzymes

70. 71. 72. 73. 74. 75. 76. 77. 78. 79. 80. 81. 82. 83. 84. 85. 86. 87. 88. 89. 90. 91. 92. 93. 94. 95. 96. 97. 98. 99. 100. 101. 102. 103. 104. 105. 106. 107. 108. 109. 110. 111. 112. 113. 114. 115. 116. 117. 118. 119. 120. 121. 122. 123. 124. 125. 126.

Galman, J. L.; Hailes, H. C. Tetrahedron: Asymmetry 2009, 20(15), 1828–1831. Schenk, G.; Duggleby, R. G.; Nixon, P. F. Int. J. Biochem. Cell Biol. 1998, 30(12), 1297–1318. Turner, N. J. Curr. Opin. Biotechnol. 2000, 11(6), 527–531. Shaeri, J.; Wright, I.; Rathbone, E. B.; Wohlgemuth, R.; Woodley, J. M. Biotechnol. Bioeng. 2008, 101(4), 761–767. French, C.; Ward, J. M. Ann. N.Y. Acad. Sci. 1996, 799, 11–18. Brocklebank, S. P.; Mitra, R. K.; Woodley, J. M.; Lilly, M. D. Enzyme Eng. Xiii 1996, 799, 729–736. Vasic-Racki, D.; Bongs, J.; Schorken, U.; Sprenger, G. A.; Liese, A. Bioprocess Biosyst. Eng. 2003, 25(5), 285–290. Chauhan, R. P.; Powell, L. W.; Woodley, J. M. Biotechnol. Bioeng. 1997, 56(3), 345–351. Selivanov, V. A.; Kovina, M. V.; Kochevova, N. V.; Meshalkina, L. E.; Kochetov, G. A. J. Mol. Catal. B: Enzym. 2003, 26(1–2), 33–40. Esakova, O. A.; Meshalkina, L. E.; Kochetov, G. A. Life Sci. 2005, 78(1), 8–13. Aucamp, J. P.; Martinez-Torres, R. J.; Hibbert, E. G.; Dalby, P. A. Biotechnol. Bioeng. 2008, 99(6), 1303–1310. Dalby, P. A.; Aucamp, J. P.; George, R.; Martinez-Torres, R. J. Biochem. Soc. Trans. 2007, 35(Pt 6), 1606–1609. Martinez-Torres, R. J.; Aucamp, J. P.; George, R.; Dalby, P. A. Enzyme Microb. Technol. 2007, 41(5), 653–662. Yokota, A.; Sasajima, K. Agric. Biol. Chem. 1983, 47(7), 1545–1553. Schlossberg, M. A.; Bloom, R. J.; Richert, D. A.; Westerfeld, W. W. Biochemistry 1970, 9(5), 1148–1153. Engel, S.; Vyazmensky, M.; Geresh, S.; Barak, Z.; Chipman, D. M. Biotechnol. Bioeng. 2003, 83(7), 833–840. Engel, S.; Vyazmensky, M.; Berkovich, D.; Barak, Z.; Chipman, D. M. Biotechnol. Bioeng. 2004, 88(7), 825–831. Engel, S.; Vyazmensky, M.; Berkovich, D.; et al. Biotechnol. Bioeng. 2005, 89(6), 733–740. Querol, J.; Grosdemange-Billiard, C.; Rohmer, M.; Boronat, A.; Imperial, S. Tetrahedron Lett. 2002, 43(46), 8265–8268. Schurmann, M.; Schurmann, M.; Sprenger, G. A. J. Mol. Catal. B: Enzym. 2002, 19, 247–252. Eubanks, L. M.; Poulter, C. D. Biochemistry 2003, 42(4), 1140–1149. Brammer, L. A.; Meyers, C. F. Org. Lett. 2009, 11(20), 4748–4751. Iding, H.; Dunnwald, T.; Greiner, L.; et al. Chem. – Eur. J. 2000, 6(8), 1483–1495. de Maria, P. D.; Pohl, M.; Gocke, D.; et al. Eur. J. Org. Chem. 2007, (18), 2940–2944. Demir, A. S.; Dunnwald, T.; Iding, H.; Pohl, M.; Muller, M. Tetrahedron: Asymmetry 1999, 10(24), 4769–4774. Berheide, M.; Peper, S.; Kara, S.; et al. Biotechnol. Bioeng. 2010, 106(1), 18–26. Lingen, B.; Grotzinger, J.; Kolter, D.; Kula, M. R.; Pohl, M. Protein Eng. 2002, 15(7), 585–593. Lingen, B.; Kolter-Jung, D.; Dunkelmann, P.; et al. ChemBioChem 2003, 4(8), 721–726. Polovnikova, E. S.; McLeish, M. J.; Sergienko, E. A.; et al. Biochemistry 2003, 42(7), 1820–1830. Yep, A.; Kenyon, G. L.; McLeish, M. J. Proc. Natl. Acad. Sci. USA 2008, 105(15), 5733–5738. Yep, A.; McLeish, M. J. Biochemistry 2009, 48(35), 8387–8395. Paramesvaran, J.; Hibbert, E. G.; Russell, A. J.; Dalby, P. A. Protein Eng. Des. Sel. 2009, 22(7), 401–411. Gonzalez, B.; Vicuna, R. J. Bacteriol. 1989, 171(5), 2401–2405. Demir, A. S.; Pohl, M.; Janzen, E.; Muller, M. J. Chem. Soc. Perkin Trans. 1 2001, (7), 633–635. Dunkelmann, P.; Kolter-Jung, D.; Nitsche, A.; et al. J. Am. Chem. Soc. 2002, 124(41), 12084–12085. Demir, A. S.; Ayhan, P.; Igdir, A. C.; Duygu, A. N. Tetrahedron 2004, 60(31), 6509–6512. Janzen, E.; Muller, M.; Kolter-Jung, D.; et al. Bioorg. Chem. 2006, 34(6), 345–361. Mikolajek, R. J.; Spiess, A. C.; Pohl, M.; Buches, J. Biotechnol. Progr. 2009, 25(1), 132–138. Ayhan, P.; Simsek, I.; Cifci, B.; Demir, A. S. Org. Biomol. Chem. 2011, 9(8), 2602–2605. Mikolajek, R.; Spiess, A. C.; Buchs, J. J. Biotechnol. 2007, 129(4), 723–725. Kneen, M. M.; Pogozheva, I. D.; Kenyon, G. L.; McLeish, M. J. Biochim. Biophys. Acta – Proteins and Proteomics 2005, 1753(2), 263–271. Brandt, G. S.; Nemeria, N.; Chakraborty, S.; et al. Biochemistry 2008, 47(29), 7734–7743. Mosbacher, T. G.; Mueller, M.; Schulz, G. E. FEBS J. 2005, 272(23), 6067–6076. Knoll, M.; Muller, M.; Pleiss, J.; Pohl, M. ChemBioChem 2006, 7(12), 1928–1934. Stillger, T.; Pohl, M.; Wandrey, C.; Liese, A. Org. Process Res. Dev. 2006, 10(6), 1172–1177. Hildebrand, F.; Kuhl, S.; Pohl, M.; et al. Biotechnol. Bioeng. 2007, 96(5), 835–843. Sopaci, S. B.; Simsek, I.; Tural, B.; Volkan, M.; Demir, A. S. Org. Biomol. Chem. 2009, 7(8), 1658–1664. Mandwal, A. K.; Tripathi, C. K. M.; Trivedi, P. D.; et al. Biotechnol. Lett. 2004, 26(3), 217–221. Khan, T. R.; Daugulis, A. J. Biotechnol. Bioeng. 2010, 107(4), 633–641. Wang, Z. L.; Liang, R.; Xu, J. H.; Liu, Y. B.; Qi, H. S. A. Appl. Biochem. Biotechnol. 2010, 160(6), 1865–1877. Leksawasdi, N.; Chow, Y. Y. S.; Breuer, M.; et al. J. Biotechnol. 2004, 111(2), 179–189. Yun, H.; Kim, B. G. Biotechnol. Bioprocess Eng. 2008, 13(3), 372–376. Hong, W. K.; Kim, C. H.; Heo, S. Y.; et al. Biotechnol. Lett. 2010, 32(8), 1077–1082. Goetz, G.; Iwan, P.; Hauer, B.; Breuer, M.; Pohl, M. Biotechnol. Bioeng. 2001, 74(4), 317–325. Rosche, B.; Breuer, M.; Hauer, B.; Rogers, P. L. Biotechnol. Bioeng. 2004, 86(7), 788–794. Suzuki, R.; Katayama, T.; Kim, B. J.; et al. J. Biol. Chem. 2010, 285(44), 34279–34287. Ou, M. S.; Ingram, L. O.; Shanmugam, K. T. J. Indust. Microbiol. Biotechnol. 2011, 38(5), 599–605.

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