ARTICLE Controlling Enantioselectivity of Esterase in Asymmetric Hydrolysis of Aryl Prochiral Diesters by Introducing Aromatic Interactions Fei Guo,1 Stefan Franzen,2 Lidan Ye,1 Jiali Gu,1 Hongwei Yu1 1

Institute of Bioengineering, Department of Chemical and Biological Engineering, Zhejiang University, Hangzhou, PR, China; telephone: 86-571-8795-1873; fax: 86-571-8795-1873; e-mail: [email protected]; 2 Department of Chemistry, Zhejiang University, Hangzhou, PR, China

ABSTRACT: Aromatic interactions specific to aryl radicals were introduced into two esterases, BioH from Escherichia coli and RspE from Rhodobacter sphaeroides to control their enantioselectivity in the asymmetric hydrolysis of prochiral aryl glutaric acid diesters. As a result, the enantiomeric excess (ee) of the S-product of dimethyl 3-phenylglutarate was increased from 25% (BioH wild type) to 96% (B_L83F/ L86F) and from 13% (RspE wild type) to >99% (R_Y27R), respectively, while another variant of RspE R_M121F gave a reversed ee of 50% (R-product). Similar enhancement or reversion of enantioselectivity were also observed in the hydrolysis of three other prochiral aryl diesters (dimethyl 3-(4-flouro)-phenylglutarate, dimethyl 3-(4-cholo)-phenylglutarate and dimethyl 3-(3,4-dicholo)-phenylglutarate). Especially, the mutant R_Y27R was shown to be an excellent S-selective hydrolase for prochiral aryl diesters, with ee of all S-products >99%. In the mutants with altered enantioselectivity, the successful introduction of designed aromatic interactions was confirmed by molecular dynamics simulations and binding free energy analysis. These results demonstrate that aromatic interaction is one of the origins of enzyme enantioselectivity, the tuning of which leads to dramatic change in enantioselectivity. Besides, the successful engineering of the enantioselectivity in two different proteins toward four different substrates suggests that the introduction of aromatic interactions is a generally applicable strategy in the control of enantioselectivity toward aryl substrates. Biotechnol. Bioeng. 2014;111: 1729–1739. ß 2014 Wiley Periodicals, Inc.

Correspondence to: H. Yu Contract grant sponsor: Natural Science Foundation of China Grant numbers: 21176215; 21176102 Contract grant sponsor: Open Funding Project of the State Key Laboratory of Bioreactor Engineering, Outstanding Young Scholar of Zhejiang Province Contract grant number: R4110092 Contract grant sponsor: Program for Zhejiang Leading Team of S&T Innovation Contract grant number: 2011R50007 Received 19 January 2014; Revision received 3 March 2014; Accepted 24 March 2014 Accepted manuscript online 16 April 2014; Article first published online 25 April 2014 in Wiley Online Library (http://onlinelibrary.wiley.com/doi/10.1002/bit.25249/abstract). DOI 10.1002/bit.25249

ß 2014 Wiley Periodicals, Inc.

KEYWORDS: enantioselectivity; esterase; rational design; aromatic interactions; molecular dynamics simulation

Introduction Enantioselectivity is one of the most significant properties of enzymes, which makes biocatalysts the preferred choice in the pharmaceutical, fine, and food chemical industries. However, the moderate enantiomeric excess (ee) produced by most naturally occurring enzymes is insufficient for industrial applications, especially toward non-native substrates (Fig. 1a) (Reetz, 2011; Tracewell and Arnold, 2009). A number of methods have been applied for improving enzyme enantioselectivity, including substrate engineering (Mats, 1998), solvent variation (Parida and Dordick, 1993), immobilization (Krebsfänger et al., 1998), and protein engineering (Rui et al., 2005). Especially, techniques such as directed evolution, semi-rational and rational design used in protein engineering have achieved great success in improving the enantioselectivity of many enzymes (Guo et al., 2013; Ma et al., 2013; May et al., 2000; Polyak et al., 2013; Zhang et al., 2012). Enantioselectivity refers to the capacity of an enzyme to produce an excess of one enantiomeric product over the other. The mechanism of enantioselectivity is usually attributed to the energy difference between the two enzyme–substrate complexes of the enantiomeric products. Thus the rational design of enzyme enantioselectivity is to reinforce this distinction by means of regulating the interactions between proteins and their ligands. A means of distinguishing enantiomeric ligands for lipases based on the size of the binding pockets in the enzyme was proposed in the 1990s derived from the Kazlauskas rule (Kazlauskas et al., 1991). Since then, the steric interaction of the substrate has been regarded as the main source of the enantioselectivity of lipases in the resolution of racemic secondary alcohol (Gascoyne et al., 2001; Rotticci et al., 2001).

Biotechnology and Bioengineering, Vol. 111, No. 9, September, 2014

1729

Figure 1. Strategies for the adjustment of enantioselectivity. a: Protein without enantioselectivity toward enantiomeric substrates. b: Enantioselectivity arising from the steric hindrance of the nonpreferred substrates. c: Selectivity loss due to the weakened effects on steric hindrance when applied to flexible motifs. d, e: Enantioselectivity resulted from the introduction of electronic interactions.

Introducing bulky mutations that collide with the non-preferred substrates decreases the size of the substrate-binding pocket and thus prevents the binding of non-preferred substrates (Fig. 1b) (Ema et al., 2005; Henke et al., 2003; Knoll et al., 2006; Magnusson et al., 2005; Park et al., 2003). Moreover, the preference and the magnitude of the enantioselectivity regulated by steric effects were proposed to be related to the volumes of the substituted amino acid residues (Bartsch et al., 2008). The strategy of manipulating the space around the substrates and regulating the repulsive steric effects has therefore been proven to be effective; however, it may result in loss of efficacy when applied to the flexible motifs of enzymes (Fig. 1c). In recent years, a number of more specific molecular interactions between proteins and ligands, such as hydrogen bonds, dipole interactions, and dispersion interactions, were proposed to study molecular recognition between the proteins and their ligands and to investigate enzyme enantioselectivity (Gartler et al., 2007; Havranek, 2010; Zhu and Hua, 2010). Due to the specificity of these

1730

Biotechnology and Bioengineering, Vol. 111, No. 9, September, 2014

interactions, enzyme enantioselectivity originated from their introduction relies on the particular structure of the substrates and is consequently more substrate-specific (Carballeira et al., 2007; Giri et al., 2010; Lavecchia et al., 2007; Sakakura and Ishihara, 2011). Among these interactions, aromatic interactions are specific forces to stabilize the phenyl ring of aryl substrates. A strategy for the rational design of enantioselectivity specific to aryl ligand is therefore proposed based on the introduction of aromatic interactions between the protein and the aryl ligand. If the interactions solely stabilize the protein–ligand complex of the preferred optical isomer, the enantioselectivity toward this product will be enhanced, while interactions favorable for the other complex will result in a reversed selectivity (Fig. 1d and e). Chiral 3-arylglutaric acid derivatives are essential synthetic blocks of some commercial medicines and biologically active compounds. It is of great interest to produce optically pure 3arylgultaric acid monoesters through asymmetric hydrolysis of prochiral 3-arylgultaric acid diesters (Ch^enevert and Desjardins, 1994; Fryszkowska et al., 2005; Yu et al., 2000). Previous studies have focused on the screening of commercial lipases and esterases without discussing the origins of the enantioselectivity in this reaction (Cabrera et al., 2008; Cabrera and Palomo, 2011; Homann et al., 2001). To reveal the origin of enantioselectivity and to illustrate the general applicability of the aromatic interaction tuning strategy in controlling enantioselectivity in the asymmetric hydrolysis of aryl substrates, the BioH esterase from Escherichia coli and the RspE esterase from Rhodobacter sphaeroides were chosen for rational design and used as the catalysts for asymmetrically hydrolyzing dimethyl 3-phenylglutarate and its analogues. The effects of introducing aromatic interactions through point mutations on the enzyme enantioselectivity were studied and the molecular details of interactions between the mutants and the substrate were investigated by molecular dynamics simulations.

Materials and Methods Chemicals and Enzymes 3-Phenylglutaric acid (3a) was purchased from Aladdin Ltd. (Shanghai, China) and the esterification reaction used to produce 1a was conducted using SOCl2 as the catalyst (Hosangadi and Dave, 1996). Substrate 1(b–d) was synthesized from corresponding aromatic aldehyde as described previously (Perregaard et al., 1995). Other chemicals were commercially available and of analytical grade purity. PrimerSTARTM (DNA polymerase) and restriction endonuclease Dpn I were purchased from TAKARA Ltd. (Dalian, China) and markers of DNA and protein were purchased from Sangon Biotech (Shanghai) Co.Ltd. (Shanghai, China). Construction of Mutants The pET-30a plasmids containing the recombinant BioH WT and RspE WT gene were used as the templates for polymerase

chain reaction (PCR)-based site-directed mutagenesis using the QuikChangeTM method (Stratagene, La Jolla, CA). The PCR primers are listed in Table I. The PCR mixture contained 15 ng of template plasmid, 5 mL of 5
PrimerSTARTM Buffer (Mg2þ plus), 2 mL of dNTP (2.5 mM each), 1 mL of each primer (10 mM), and 0.5 mL of PrimerSTARTM HS polymerase (2.5 U/mL) in a total volume of 25 mL. The PCR reaction was carried out under the following conditions: 98 C denaturation for 1 min, 20 cycles of 98 C denaturation for 10 s, 55 C annealing for 10 s, and 72 C extension for 7 min. The PCR product was purified and then digested with Dpn I, which specifically cleaves the Adenine methylated template. The digestion mixture contained 17 mL of purified PCR product, 2 mL of 10
T Buffer and 1 mL of Dpn I. The digestion was conducted at 37 C for 1 h. Then the mixture was transformed into competent cells (E.coli BL21 (DE3)) using the heat shock method at 42 C for 90 s. Finally, colonies were cultured on Luria-Bertani (LB) (Sambrook and David, 2001) agar plates. The entire mutant gene was sequenced to verify that no other alterations were introduced into the nucleotide sequence. Overexpression and Purification of Enzymes All mutants were cultured in 50 mL LB media supplemented with 50 mg/mL of kanamycin at 37 C. Esterase expression was induced when isopropyl-b-d-thiogalactoside (IPTG, final concentration 100 mM) was added at the early exponential phase (OD600 0.4–0.6), and the cultivation was continued for 4 h. Cells were collected by centrifugation (3,723g, 20 min), and washed with phosphate buffered saline (PBS, pH 7.4). The cell precipitation was resuspended in Tris– HCl buffer (50 mM, pH 8.0) and then lysed by sonication. Enzymes were purified by HisTrap affinity columns (GE Healthcare, Beijing, China) based on the affinity between Ni2þ on the column and the His-tag of the cloned protein, which is a sequence of six histidine residues. Then the enzymes were desalted by desalting columns (GE Healthcare) and redissolved in 0.1 M phosphate buffer (pH 8.0).

The expression of the mutated gene was examined by SDS– polyacrylamide gel electrophoresis (SDS–PAGE) and the protein concentration was determined by the Bradford method (Bradford, 1976). Enzyme Assay and Analytical Methods Enzymatic hydrolysis was carried out at 30 C in phosphate buffer (0.1 M, pH 8.0). The final concentration of the prochiral diester substrates was 1.0 mM (5% acetonitrile as cosolvent) and the final concentration of the purified and desalted enzymes was 0.5 mg/mL. The reactions were stopped by adding HCl (0.5%, v/v). Then the reaction mixture was extracted using ethyl acetate. The samples were dried and redissolved in isopropanol for analysis. The chemical yield was calculated from the results of HPLC using a C18 column and a UV detector at 254 nm. The mobile phase was composed of water and acetonitrile (50/50, v/v) at a flow rate of 1 mL/min. The enantiomeric excess of product (eep) was analyzed via HPLC using an AD-H chiral column and a UV detector at 254 nm. The mobile phase was composed of nhexane and iso-propanol (95/5, v/v) with a flow rate as 1 mL/ min. The enantiomeric ratio (E) and corresponding difference of binding free energy (DDGexperiment) were calculated according to: E¼

1 þ eep 1  eep

DDGexperiment ¼ RT ln E

ð1Þ ð2Þ

With R ¼ 1.99 cal/K1mol1, T ¼ 303 K (Straathof and Jongejan, 1997). Computational Methods The initial geometries of the wild-type esterases were obtained from the protein data bank (BioH PDB ID: 1M33; RspE PDB ID: 4FHZ) and all subsequent structures

Table I. Mutated sites in the mutants and PCR primers used for mutant construction. Mutants

Template

Mutated sites

B_L83F

BioH

Leu83Phe

B_L86F

BioH

Leu86Phe

B_L83FL86F

BioH

Leu83Phe/Leu86Phe

R_Y27R

RspE

Tyr27Arg

R_I71F

RspE

Ile71Phe

R_I71R

RspE

Ile71Arg

R_M121F

RspE

Met121Phe

Primersa GGTTAGGCTGGAGTTTTGGCGGGCTGGTGGCAAGCCAGA GCCACCAGCCCGCCAAAACTCCAGCCTAACCAAATGGCT GGAGTCTGGGCGGGTTTGTGGCAAGCCAGATTGCGTTAA ATCTGGCTTGCCACAAACCCGCCCAGACTCCAGCCTAAC GGTTAGGCTGGAGTTTTGGCGGGTTTGTGGCAAGCCAGA GCCACAAACCCGCCAAAACTCCAGCCTAACCAAATGGCT TCTTCCTCCACGGCCGCGGCGCGGACGGGGCGGATCTCT GCCCCGTCCGCGCCGCGGCCGTGGAGGAAGACGACGAGG TCCAGTGGTTTCCGTTCCCCTGGCTCGACGGCTCCTCGG CCGTCGAGCCAGGGGAACGGAAACCACTGGAAGCCGAAG TCCAGTGGTTTCCGCGCCCCTGGCTCGACGGCTCCTCGG CCGTCGAGCCAGGGGCGCGGAAACCACTGGAAGCCGAAG TCTCGCAGGGCACGTTCATGGCGCTTCATGTGGCGCCGC ACATGAAGCGCCATGAACGTGCCCTGCGAGAAGCCCACG

a

The mutated sites are underlined.

Guo et al.: Control of Enantioselectivity Biotechnology and Bioengineering

1731

of the mutants were set up using Swiss-PDB Viewer. Nonpolar hydrogen atoms were added to the enzyme structure using the AMBER 11 simulation package. For the neutral histidine residues, the e-N and d-N were protonated depending on the probability to form hydrogen bonds. Partial charges and force field parameters of the substrates were automatically generated via the antechamber program using AMBER force field (Laskowski et al., 1993; Wang et al., 2004). Sodium was used to neutralize the system. The whole system was immersed in a rectangular box of TIP3P water molecules. The box was extended 9 A in all three dimensions from the edges of the protein structure. Molecular dynamics (MD) simulations were employed to investigate the transition state (TS) complexes of the enzymes and the model substrate dimethyl 3-phenylglutarate. R- and S-complexes were constructed for each mutant as the TS for the R- and S-products. These conformations formed based on two restrained distances: 2 A between the OG of the catalytic serine and the catalytic carbonyl carbon of the substrate, and  3 A between the Hd of the catalytic histidine and the catalytic ester oxygen of the substrate. Although the substrate molecules in R- and S-complex were the same, the different binding modes led to different conformations and energies. The MD simulations were conducted as described previously (Gu and Yu, 2012), including steepest descent and conjugate gradient minimization, heating and density equilibration, constant-pressure and constant-temperature

(NPT) equilibration, and a 5 ns productive NPT simulation. The temperature was maintained by coupling to the Langevin thermostat. The SHAKE algorithm was used to constrain bond distances of the hydrogen atoms. The nonbonding cut off distance was set at 8.0 A and simulation snapshots were saved every 2 ps for analysis. Conformation analysis of each simulation was performed using the ptraj program in the AMBER package, and the mmpbsa program was employed to calculate the binding free energy of each complex and the decomposition of the binding free energy to individual residues.

Results Decision of Hot-Spots and Design of Mutants Although cloned from different organisms, BioH and RspE belong to the same hydrolase family and have the same SerHis-Asp catalytic triads. In the asymmetrical hydrolysis of model substrate dimethyl 3-phenylglutarate (Fig. 2, 1a) into monomethyl S-3-phenylglutarate (Fig. 2, S-2a), both wildtype BioH and RspE (B_WT and R_WT) exhibited moderate enantioselectivity and good chemical yield (Table III, entry 1 and 5). Substrate docking was conducted by molecular dynamics simulation and the TS complexes for enantiomeric products were aligned (Fig. 3). In the TS complexes of B_WT, the ester

Figure 2. Hydrolysis of prochiral dimethyl 3-arylglutarate (1(a–d)). The reaction contains two steps, the asymmetric production of chiral monoester (R-2(a–d) and S-2(a–d)) and the production of 3-arylglutaric acid (3(a–d)).

1732

Biotechnology and Bioengineering, Vol. 111, No. 9, September, 2014

Table II. Decomposition of binding free energy into individual residues in BioH and RspE.a Binding free energy (kcal/mol)b Key residue number BioH 22 81 82c 83 111 183 209 235d RspE 27 71 116 117c 118 121 200d

S

R

2.55 1.02 20.88 2.62 1.05 2.21 0.90 0.49

2.15 0.12 20.19 1.82 1.22 1.08 1.34 1.01

2.37 1.31 0.17 20.74 2.91 0.66 0.92

2.25 1.21 0.30 21.27 1.34 0.53 0.68

a

Only key residues are listed in the table. The binding free energy decompositions were calculated by the mmpbsa program of the AMBER software package. c,d The catalytic serine and histidine. b

bonds to be hydrolyzed in both substrates overlapped in a similar conformation with the catalytic (Ser82 and His235) and oxyanion-hole (Leu83 and Trp22) residues, while the orientation of the non-hydrolyzed half of the C-chain was slightly different. The dominating distinction between the two substrates was the orientation of the benzene rings vertical to the C-chain. In the TS complex for the S-product (S-complex), the benzene ring was deeply embedded into the pocket and faced inwards while the other benzene ring oriented outwards with fewer residues surrounding it. This difference between the two complexes might explain the S-selectivity. Unlike the small and highly hydrophobic

substrate-binding pocket of BioH, the substrate of RspE was embedded into a half-open pocket with several hydrophilic residues. Although the substrate-binding pocket showed different geometry and polarity from BioH, in the TS complexes of R_WT the situation of the substrate binding was similar to that of BioH. The catalytic part of the substrates overlapped and the residual part of the substrate oriented differently. Therefore, reinforcing the difference in substrate binding might be a common way to improve the enantioselectivity for both enzymes. Decomposition of the binding free energy of the TS complexes revealed the residues with major contribution (Table II). Besides the catalytic residues that contributed a lot, other residues may be considered as the key residues that interact with the substrate. Among them, three residues (Phe111, Leu86, and Leu83) of BioH and two residues (Tyr27 and Ile71) of RspE were pointed out as the key residues that may interact solely with the S-benzene ring, while two phenylalanines (Phe143 and Phe128) of BioH and one residue (Met121) of RspE were regarded as key residues for the Rbenzene ring Therefore, targeting at these residues, introduction of two patterns of aromatic interactions was designed, namely, p–p stacks and cation–p interactions (Table I). The non-aryl residues were substituted by aryl residues to introduce p–p interactions, creating mutants B_L83F, B_L86F, R_I71F, and R_M121F, while mutants R_Y27R and R_I71R were created by introducing cation–p interactions. Positively charged arginine was not employed as a substituting amino acid in BioH since it might lead to destabilization of the highly hydrophobic substrate binding pocket and misfolding of the protein. Asymmetric Hydrolysis of Model Substrates by BioH and RspE Mutants Mutants of BioH and RspE were constructed and overexpressed in E. coli. The point mutation slightly influenced the

Figure 3. Protein and substrate complexes of wild type BioH (a) and RspE (b) with the model substrate. The S-complex is shown in green ribbons with the S-substrate in light green, while the R-complex is shown in orange ribbons with the R-substrate in orange yellow. The residues around the substrates are indicated by fonts.

Guo et al.: Control of Enantioselectivity Biotechnology and Bioengineering

1733

Table III. Asymmetric hydrolysis of aryl prochiral esters by BioH and RspE mutants.a Entry 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

Substrates

Mutants

Time (H)

Conversion (%)b

Yield (%)c

B_WT B_L83F B_L86F B_L83FL86F R_WT R_Y27R

24 10 10 10 8 4 15 12 3 8 12 30 10 10 4 15 10 20 17 4 5 1.8 2 7.5 9 30 10 10 12 1 3 12 15

>99 >99 >99 >99 >99 80 >99 >99 80 >99 >99 >99 >99 >99 85 >99 >99 >99 >99 90 >99 90 >99 89 >99 >99 >99 95 >99 87 >99 90 >99

>99 >99 >99 >99 >99 >99 90 >99 >99 94 >99 >99 >99 >99 >99 90 >99 >99 >99 >99 95 >99 96 >99 97 >99 >99 >99 93 >99 89 >99 95

R_I71F R_I71R R_M121F B_WT B_L86F R_WT R_Y27R R_M121F B_WT B_L83F R_WT R_Y27R R_M121F B_WT B_L83FL86F R_WT R_Y27R R_M121F

eep (%) 25.3 61.1 92.4 96.2 13.2 87.2 >99 61.7 70.2 >99 50.3 73.5 93.4 8.4 84.1 >99 49.3 18.7 38.1 50.3 56.4 90.5 >99 17.2 17.6 7.4 52.9 46.8 48.7 92.0 >99 14.5 17.5

(S) (S) (S) (S) (S) (S) (S) (S) (S) (S) (R) (S) (S) (S) (S) (S) (R) (R) (R) (S) (S) (S) (S) (R) (R) (R) (R) (S) (S) (S) (S) (S) (S)

Ed 1.68 4.14 25.32 51.63 1.30 14.63 — 4.22 5.71 — 3.02 6.55 29.30 1.18 11.58 — 2.94 1.46 2.23 3.02 — 20.05 — 1.42 — 1.16 3.25 2.76 — 24.00 — 1.34 —

The reactions were conducted at 30 C in phosphate buffer (0.1 M, pH 8.0). The initial concentration of the prochiral diester substrates was 1.0 mM (5% acetonitrile as cosolvent) and the concentration of the purified and desalted enzymes was 0.5 mg/mL. b Conversion was defined as the consumption of diester substrate. c Yield was defined as the monoester ratio of the consumed diester substrate. d Enantiomeric ratio was calculated as E ¼ (1 þ eep)/(1–eep). Product enantiomeric excess in the early phase of the reaction (yield > 99%) was employed in the calculation. a

expression level, but no large difference was observed in the molecular weight of the protein. Asymmetric hydrolyses of the model substrate by these mutants were conducted under the same condition as the wild type and the results are shown in Table III. The enzymatic assay verified that the substitution of leucine by phenylalanine in BioH enhanced the S-enantioselectivity, which was in accordance with our design. L83F showed a good enantioselectivity in the asymmetric hydrolysis (Table III, entry 2). Moreover, the introduction of phenylalanine at site 86 turned out to generate a highly effective mutant with remarkably improved enantioselectivity (Table III, entry 3). Based on the insights from the single mutations, we further tested the combination of mutations L83F and L86F, which resulted in an excellent double mutant that can asymmetrically hydrolyze the model substrate, giving >99% conversion, >99% yield, 96.2% enantiomeric excess of S-monoester and 51.63 E (Table III, entry 4).

1734

Biotechnology and Bioengineering, Vol. 111, No. 9, September, 2014

Similar results were also obtained in the hydrolysis mediated by RspE mutants. Phenylalanine substitution at site 71 created a good mutant with improved S-enantioselectivity (Table III, entry 8). And the introduction of aryl residue at site 121 resulted in a mutant with reversed enantioselectivity (Table III, entry 11). Besides mutants with aryl mutations, mutants with positively charged substitutions were also testified in the hydrolysis reactions. Both mutant R_Y27R and mutant R_I71R showed good S-enantioselectivity in the early phase of the reaction, in which the monoester was not converted into diacid (Table III, entry 6 and 9). Moreover, with the proceeding of the reaction, the eep of residual S-monoester approached to 100% (Table III, entry 7 and 10). High enantiomeric excess of the monoester can be achieved by sacrificing the chemical yield when the enzyme has good enantioselectivity but low ability to hydrolyze the monoester, since the monoester preferred by the enzyme is the nonpreferred product of diester hydrolysis.

Table IV. Binding free energies of protein–ligand complexes with dimethyl 3-phenylglutarate.a

Entry 1 2 3 4 5 6 7 8 9

Mutants

DGR (kcal/mol)b

DGS (kcal/mol)b

DDGS–R (kcal/mol)b

B_WT B_L83F B_L86F B_FF R_WT R_Y27R R_I71F R_I71R R_M121F

17.3  4.6 17.0  5.0 19.6  4.1 19.9  4.8 24.4  5.8 24.7  4.8 24.9  4.7 23.4  5.8 22.1  3.9

15.7  4.9 14.4  4.5 15.5  4.9 16.5  4.9 23.5  4.4 21.5  4.4 22.4  5.2 21.5  4.4 24.5  4.7

1.6 2.6 4.1 3.4 0.9 3.2 2.5 1.9 2.4

a

The binding free energies were calculated by the mmpbsa program of the AMBER software package. b DGR and DGS indicate the binding free energy of R- and S-complex. DDGS–R indicates the difference of binding free energy. Negative and positive values represent the S-selectivity and the reversed R-selectivity, respectively.

Binding Free Energy Analysis of Protein–Ligand Complexes Molecular dynamics (MD) simulations were conducted on the TS complexes of the mutants and the substrates for the enantiomeric products, and the binding free energies (DG) were investigated. The results of the binding free energies calculated for each complex and the difference of binding free energies between the S- and R-complexes are shown in Table IV. Introduction of phenylalanine into BioH increased the difference in binding free energies (DDGS–R) (Table IV, entry 1–4), implying that the interaction between the protein and the substrate was changed by the introduced substitutions. This result was consistent with the improved E. The

introduced aromatic interaction by the mutations can be further verified by the decomposition of the binding free energies (Table V). Compared to the leucine in the wild-type enzyme, Phe83 and Phe86 increased the protein–substrate interaction in the S-complexes, confirming the introduction of the p–p interaction as designed. Meanwhile, the interaction between the substrate and Phe111 was also increased in the S-complex of B_L86F, which indicated the aryl substitution at site 86 may form a tri-membered aromatic interaction group together with the substrate and Phe111. The geometric conformations of the aromatic interactions in B_L83F and B_L86F are shown in Figure 5a and b. Similarly, the binding free energy analysis for the complexes of the RspE mutants with aryl substitutions also confirmed the introduction of additional aromatic interactions between the protein and the substrate. The difference in the binding free energies of R_I71F and R_M121F was in accordance with the change of the enantioselectivity (Table IV, entry 7 and 9). Energy decomposition also confirmed the introduction of aromatic interactions. The binding free energy contributions of the mutated residues were increased in the S- and R-complexes, respectively in those two mutants (Table V). The geometric conformations of the aromatic interactions in R_I71F and R_M121F are shown in Figure 5c and d. Cation–p interactions were expected to be introduced by the arginine substitutions. Although no proper conformation of cation–p interactions was obtained by the MD simulations (Fig. 5e and f), the introduction of the interactions between the substitution and the substrate were confirmed by analysis of binding free energy (Table IV, entry 6 and 8). The decomposition of the binding free energy also indicated the interactions introduced by arginine were preferred in Scomplexes (Table V).

Table V. Binding free energy decomposition of key residues in the complex of mutants and dimethyl 3-phenylglutarate.a Key residues Mutants B_WT B_L83F B_L86F

R_WT R_Y27R R_I71F R_I71R R_M121F

Complexesb

Leu/Phe83

Leu/Phe86

Phe111

R S R S R S

1.82  4.11 2.62  4.56 1.74  4.94 2.82  5.21 — — Tyr/Arg27 2.25  5.30 2.37  5.07 2.29  9.38 2.59  11.03 — — — — — —

0.02  5.14 0.02  6.93 — — 0.01  4.70 0.10  5.89 Ile/Phe/Arg71 1.21  5.70 1.32  5.57 — — 1.34  5.48 1.85  5.31 1.33  9.71 1.82  10.74 — —

1.22  5.65 1.05  5.21 — — 1.15  6.10 1.14  5.15 Met/Phe121 0.53  4.853 0.66  4.59 — — — — — — 0.68  5.31 0.31  4.95

R S R S R S R S R S

a

The binding free energy decompositions were calculated by the mmpbsa program of the AMBER software package. The unit was reported in kcal/mol. R- and S- complex are the TS complexes for R- and S-product, respectively.

b

Guo et al.: Control of Enantioselectivity Biotechnology and Bioengineering

1735

Figure 4. Difference of binding free energy calculated from the experimental E (DDGexperiment) plotted against the absolute value of difference of binding free energy calculated in the molecular dynamics simulations (DDGcalculation). The diagonal line indicates the full consistency between DDGexperiment and DDGcalculation. The mutants represented by the dots are indicated by fonts.

Furthermore, the diagram of DDGexperiment plotted against the absolute value of DDGS–R (DDGcalculation) revealed a good agreement between the experimental data and modeling results despite the overestimated difference of the binding free energy between R- and S-complexes by computational calculation (Fig. 4). Asymmetric Hydrolysis of Model Substrate Analogues by Selected Mutants To further verify the general applicability of this aromatic interaction tuning strategy in controlling enantioselectivity, several model substrate analogues were investigated as the substrates of the BioH and RspE mutants mediated asymmetric hydrolysis. Dimethyl 3-(4-fluoro)-phenylglutarate (1b), dimethyl 3-(4-chloro)-phenylglutarate (1c) and dimethyl 3-(3,4-dichloro)-phenylglutarate (1d) were chosen because (i) the substitution on the phenyl ring influences the ring’s property and consequently affects the aromatic interactions; (ii) the chiral monoester of these substrates are synthetic blocks of some commercial medicines and biologically active compounds. The reaction results provided the evidence that the aromatic interactions introduced could affect the enantioselectivity of the mutants in the hydrolysis of aryl substrates. With the selected mutants shown in Table III, both improved and reversed enantioselectivity were obtained toward these model substrate analogues. Among those mutants, R_Y27R was an outstanding S-enantioselective hydrolase toward the prochiral aryl glutaric acid diesters with >99% eep for all aryl substrates. Meanwhile, the large volume of the chlorine atom brought up additional steric hindrance which resulted in the different

1736

Biotechnology and Bioengineering, Vol. 111, No. 9, September, 2014

preference between the model substrate and the fluorosubstrate. The improved enantioselectivity in BioH mutants toward the chloro-substrates verified the function of the aromatic interactions introduced in spite of the negative influence imposed by the presence of additional steric effect. In the case of RspE, due to the open and large substratebinding pocket, this negative influence caused by steric effect was relatively weak.

Discussion In the present work, aromatic interactions were introduced into two esterases, leading to successful enhancement and reversion of enantioselectivity in the asymmetric hydrolysis of prochiral diesters. The successful introduction of the designed aromatic interactions was confirmed by molecular dynamics simulations and binding free energy analysis. The origins of the enantioselectivity that the wild type esterases and the mutants showed in the asymmetric hydrolysis consisted of several components. First of all, the asymmetric substrate-binding pocket provides the wild type enzyme with moderate enantioselectivity. The prochiral diester substrates do not have asymmetry; however, once these prochiral molecules bind to an enzyme, they are embedded in the microenvironment of the asymmetric substrate-binding site. The bound substrates of the enantiomeric products could have different conformations, which are favored to a different extent by the active site. Secondly, the aromatic interactions introduced by the mutations granted the mutants dramatically increased or reversed enantioselectivity as indicated by the experimental and simulation results. The interactions between the aryl or positively charged substitutions and the aryl ring of the

Figure 5.

Conformation of the aromatic interactions formed by the model substrate and the aryl substitution introduced in mutants B_L83F (a), B_L86F (b), R_I71F (c), R_M121F (d), R_Y27R (e), and R_I71R (f).

substrates stabilized one of the TS complexes, resulting in the preference for that product. Aromatic interactions introduced at two sites from different sides of the substrate stabilized different TS complexes and thus constituted complementary enzymes (Mugford et al., 2008). Finally, the second step hydrolysis of monoester was also beneficial for the final enantiomeric excess of the monoester product. In case the enantioselectivity in the first step is high (>70%), optically pure monoester may be obtained. However, the hydrolysis activity toward monoester should be carefully controlled since it would decrease the chemical yield of the whole reaction.

The two proteins investigated in this study are esterases from different organisms. Their similarities include the catalytic triad and the reaction mechanism shared by all members of the hydrolase family. However, the substrate binding pockets of these two esterases differ. Substrates of BioH dock in a small pocket with hydrophobic residues in the vicinity, resulting in the rigid conformation of the substrates with low flexibility. On the other hand, the binding pocket of RspE is a cleft half-open to the solvent. Hydrophilic residues and solvent molecules interact with the substrates and the binding of the substrates is much less constrained by the steric effect compared to BioH. Despite the difference

Guo et al.: Control of Enantioselectivity Biotechnology and Bioengineering

1737

between these two enzymes, introduction of aromatic interactions exhibited solid effects on the enantioselectivity of both proteins, indicating the important role of aromatic interactions in enzyme enantioselectivity. These two model proteins along with four different aryl substrates provide evidence for the general applicability of the strategy to control enzyme enantioselectivity by introducing aromatic interactions. Although BioH and RspE are not optimal in their native form for application in the enantioselective hydrolysis of prochiral diesters due to limited enantioselectivity, mutants with satisfying enantioselectivity were constructed for both esterases by introduction of aromatic interactions via simply altering one or two amino acids. This example indicates the potential of proteins to evolve new functions under selection pressure by introduction of only a few mutations during gene duplication. References Bartsch S, Kourist R, Bornscheuer UT. 2008. Complete inversion of enantioselectivity towards acetylated tertiary alcohols by a double mutant of a Bacillus subtilis esterase. Angew Chem Int Ed 47(8): 1508–1511. Bradford MM. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72:248–254. Cabrera Z, Fernandez-Lorente G, Palomo JM, Guisan JM, FernandezLafuente R. 2008. Asymmetric hydrolysis of dimethyl 3-phenylglutarate catalyzed by Lecitase Ultra1. Enzyme Microb Technol 43(7): 531–536. Cabrera Z, Palomo JM. 2011. Enantioselective desymmetrization of prochiral diesters catalyzed by immobilized Rhizopus oryzae lipase. Tetrahedron Asymmetry 22(24):2080–2084. Carballeira JD, Krumlinde P, Bocola M, Vogel A, Reetz MT, Bäckvall J-E. 2007. Directed evolution and axial chirality: Optimization of the enantioselectivity of Pseudomonas aeruginosa lipase towards the kinetic resolution of a racemic allene. Chem Commun 21(19):1913. Ch^enevert R, Desjardins M. 1994. Chemoenzymatic enantioselective synthesis of baclofen. Can J Chem 72(11):2312–2317. Ema T, Fujii T, Ozaki M, Korenaga T, Sakai T. 2005. Rational control of enantioselectivity of lipase by site-directed mutagenesis based on the mechanism. Chem Commun (37):4650. Fryszkowska A, Komar M, Koszelewski D, Ostaszewski R. 2005. Enzymatic desymmetrization of 3-arylglutaric acid anhydrides. Tetrahedron Asymmetry 16(14):2475–2485. Gartler G, Kratky C, Gruber K. 2007. Structural determinants of the enantioselectivity of the hydroxynitrile lyase from Hevea brasiliensis. J Biotechnol 129(1):87–97. Gascoyne DG, Finkbeiner HL, Chan KP, Gordon JL, Stewart KR, Kazlauskas RJ. 2001. Molecular basis for enantioselectivity of lipase from Chromobacterium viscosum toward the diesters of 2,3-dihydro-3-(40 hydroxyphenyl)-1,1,3-trimethyl-1H-inden-5-ol. J Org Chem 66(9): 3041–3048. Giri S, Wang DZ, Chattaraj PK. 2010. Catalyst electronic polarizability and enantiomeric excess in asymmetric hydrogenation. Tetrahedron 66(25): 4560–4563. Gu J, Yu H. 2012. The role of residue S139 of mandelate racemase: Aynergistic effect of S139 and E317 on transition state stabilization. J Biomol Struct Dyn 30(5):585–593. Guo F, Xu H, Xu H, Yu H. 2013. Compensation of the enantioselectivityactivity trade-off in the directed evolution of an esterase from Rhodobacter sphaeroides by site-directed saturation mutagenesis. Appl Microbiol Biotechnol 97(8):3355–3362.

1738

Biotechnology and Bioengineering, Vol. 111, No. 9, September, 2014

Havranek JJ. 2010. Specificity in computational protein design. J Biol Chem 285(41):31095–31099. Henke E, Bornscheuer UT, Schmid RD, Pleiss J. 2003. A molecular mechanism of enantiorecognition of tertiary alcohols by carboxylesterases. Chembiochem 4(6):485–493. Homann MJ, Vail R, Morgan B, Sabesan V, Levy C, Dodds David R, Zaks A. 2001. Enzymatic hydrolysis of a prochiral 3-substituted glutarate ester, an intermediate in the synthesis of an NK1/NK2 dual antagonist. Adv Synth Catal 343(6–7):744–749. Hosangadi BD, Dave RH. 1996. An efficient general method for esterification of aromatic carboxylic acids. Tetrahedron Lett 37(35):6375–6378. Kazlauskas RJ, Weissfloch ANE, Rappaport AT, Cuccia LA. 1991. A rule to predict which enantiomer of a secondary alcohol reacts faster in reactions catalyzed by cholesterol esterase, lipase from Pseudomonas cepacia, and lipase from Candida rugosa. J Org Chem 56(8):2656–2665. Knoll M, Müller M, Pleiss J, Pohl M. 2006. Factors mediating activity, selectivity, and substrate specificity for the thiamin diphosphatedependent enzymes benzaldehyde lyase and benzoylformate decarboxylase. Chembiochem 7(12):1928–1934. Krebsfänger N, Schierholz K, Bornscheuer UT. 1998. Enantioselectivity of a recombinant esterase from Pseudomonas fluorescens towards alcohols and carboxylic acids. J Biotechnol 60(1–2):105–111. Laskowski RA, MacArthur MW, Moss DS, Thornton JM. 1993. PROCHECK: A program to check the stereochemical quality of protein structures. J Appl Crystallogr 26(2):283–291. Lavecchia A, Cosconati S, Novellino E, Calleri E, Temporini C, Massolini G, Carbonara G, Fracchiolla G, Loiodice F. 2007. Exploring the molecular basis of the enantioselective binding of penicillin G acylase towards a series of 2-aryloxyalkanoic acids: A docking and molecular dynamics study. J Mol Graph Model 25(6):773–783. Ma J, Wu L, Guo F, Gu J, Tang X, Jiang L, Liu J, Zhou J, Yu H. 2013. Enhanced enantioselectivity of a carboxyl esterase from Rhodobacter sphaeroides by directed evolution. Appl Microbiol Biotechnol 97(11):4897–4906. Magnusson AO, Rotticci-Mulder JC, Santagostino A, Hult K. 2005. Creating space for large secondary alcohols by rational redesign of Candida antarctica Lipase B. Chembiochem 6(6):1051–1056. Mats H. 1998. Insights into the molecular basis for fatty acyl specificities of lipases from Geotrichum candidum and Candida rugosa. Chem Phys Lipids 93(1–2):57–65. May O, Nguyen PT, Arnold FH. 2000. Inverting enantioselectivity by directed evolution of hydantoinase for improved production of l-methionine. Nat Biotech 18(3):317–320. Mugford PF, Wagner UG, Jiang Y, Faber K, Kazlauskas RJ. 2008. Enantiocomplementary enzymes: Classification, molecular basis for their enantiopreference, and prospects for mirror-image biotransformations. Angew Chem Int Ed 47(46):8782–8793. Parida S, Dordick JS. 1993. Tailoring lipase specificity by solvent and substrate chemistries. J Org Chem 58(12):3238–3244. Park S, Forró E, Grewal H, Fülöp F, Kazlauskas RJ. 2003. Molecular basis for the enantioselective ring opening of b-lactams catalyzed byCandida antarctica lipase B. Adv Synth Catal 345(8):986–995. Perregaard J, Moltzen EK, Meier E, Sanchez C. 1995. Sigma ligands with subnanomolar affinity and preference for the sigma 2 binding site. 1.3(omega-Aminoalkyl)-1H-indoles. J Med Chem 38(11):1998–2008. Polyak I, Reetz MT, Thiel W. 2013. Quantum mechanical/molecular mechanical study on the enantioselectivity of the enzymatic Baeyer– Villiger reaction of 4-hydroxycyclohexanone. J Phys Chem B 117(17): 4993–5001. Reetz MT. 2011. Laboratory evolution of stereoselective enzymes: A prolific source of catalysts for asymmetric reactions. Angew Chem Int Ed 50(1):138–174. Rotticci D, Rotticci-Mulder JC, Denman S, Norin T, Hult K. 2001. Improved enantioselectivity of a lipase by rational protein engineering. Chembiochem 2(10):766–770. Rui L, Cao L, Chen W, Reardon KF, Wood TK. 2005. Protein engineering of epoxide hydrolase from Agrobacterium radiobacter AD1 for enhanced activity and enantioselective production of (R)-1-Phenylethane-1,2 -Diol. Appl Environ Microbiol 71(7):3995–4003.

Sakakura A, Ishihara K. 2011. Asymmetric Cu(ii) catalyses for cycloaddition reactions based on p–cation or n–cation interactions. Chem Soc Rev 40(1):163. Sambrook JR, David W. 2001. Molecular cloning: A laboratory manual. 3rd ed. New York, NY: Cold Spring Harbor Laboratory Press. Straathof AJJ, Jongejan JA. 1997. The enantiomeric ratio: Origin, determination and prediction. Enzyme Microb Technol 21(8):559–571. Tracewell CA, Arnold FH. 2009. Directed enzyme evolution: Climbing fitness peaks one amino acid at a time. Curr Opin Chem Biol 13(1):3–9.

Wang J, Wolf RM, Caldwell JW, Kollman PA, Case DA. 2004. Development and testing of a general amber force field. J Comput Chem 25(9):1157–1174. Yu MS, Lantos I, Peng Z-Q, Yu J, Cacchio T. 2000. Asymmetric synthesis of ()-paroxetine using PLE hydrolysis. Tetrahedron Lett 41(30): 5647–5651. Zhang Z-G, Parra LP, Reetz MT. 2012. Protein engineering of stereoselective Baeyer–Villiger monooxygenases. Chem Eur J 18(33):10160–10172. Zhu D, Hua L. 2010. How carbonyl reductases control stereoselectivity: Approaching the goal of rational design. Pure Appl Chem 82(1):117–128.

Guo et al.: Control of Enantioselectivity Biotechnology and Bioengineering

1739