In the first step, a library of 83 esterases from metagenomic origin was screened for activity towards the ethyl esters 1b–3b using a microtiter plate‐based method (Baumann et al., 2000) with bromothymol blue serving as pH indicator. This led to the identification of 20 enzymes, which were then subjected to analytical scale kinetic resolutions for the determination of enantioselectivity and ‐preference by gas chromatographic analysis. The best hits are shown in Fig. 2. Next, the reaction conditions were further optimized by variation of reaction temperature (from 30°C to 20°C) and the addition of the cosolvent DMSO. Thus, at least two enzymes were found for each substrate exhibiting high enantioselectivity (E‐values > 50) and complementary enantiopreference (Table 1).

Two enzymes, esterases CL1 and A3, showed high selectivity towards ethyl‐3‐phenyl butanoate 1b, whereas esterases CE and Est8 were most useful for the kinetic resolution of ethyl‐2‐phenyl propanoate 2b and the butanoate 3b. Surprisingly, in the case of esterases CE and Est8, the presence of DMSO increased the enzyme activity, without affecting significantly the enantioselectivity, probably due to the better solubility of the substrates in the reaction media. Finally, the kinetic resolutions were performed in preparative (1 g) scale (Table 2).

Esterases CE and A3 were thoroughly characterized before (Elend et al., 2006). Esterase Est8 was identified by an activity‐based screening of a metagenomic library using tributyrin as substrate. The library was set up from a soil sample collected from a vineyard near Weinheim, Germany. The protein comprises 247 amino acids giving a calculated molecular weight of 26.4 kDa. The primary structure shows a Gly–X–Ser–X–Gly pentapeptide typical of the structural motive in the immediate proximity of the active‐site serine of α/β‐hydrolases (Ollis et al., 1992). Furthermore, a Gly–Gly–Gly–Y motive was identified N‐terminal to the above‐mentioned pentapeptide, indicating that this enzyme is a member of the so‐called ‘GGG(A)X‐type’ esterases. This group of enzyme was shown to be capable of the conversion of esters of tertiary alcohols (Henke et al., 2002; Kourist et al., 2008). Est8 showed the highest identity of 65% to an esterase/lipase from an uncultured bacterium (AY833090.1).

In addition to the high enantioselectivity reported above, initial biochemical characterization of Est8 was also performed. Thus crude cell extracts of Escherichia coli expressing the gene encoding Est8 displayed the highest esterase activity at 50°C. Increasing the temperature further decreased the activity to about 10% at 70°C, 80°C and 90°C (Fig. 3).

Esterase CL1 was identified using the activity assay based on hydrolysis of tributyrin. This esterase was found in a metagenomic library isolated from biofilms in drinking water (Schmeisser et al., 2003); it comprises 318 amino acids resulting in its calculated molecular weight of 33 kDa. It reaches its maximum hydrolytic activity at pH 8, presenting the highest stability at pH 9–10. Further characterization and application of this protein are currently under investigation.

All chemicals were purchased from Sigma‐Aldrich (Steinheim, Germany). Chiral gas chromatography (GC) analyses were performed by using a Heptakis‐(2,3‐di‐O‐acetyl‐6‐O‐t‐butyldimethylsilyl)‐β‐cyclodextrin‐column (Macherey‐Nagel, Düren, Germany) on a GC‐14A gas chromatograph (Shimadzu, Tokyo, Japan) and by using a Heptakis‐(2,6‐O‐di‐O‐methyl‐3‐O‐pentyl)‐β‐cyclodextrin‐column (Macherey‐Nagel, Düren, Germany) on a GC‐2010 gas chromatograph (Shimadzu, Tokyo, Japan) equipped with an AOC‐20i Autoinjector (Shimadzu, Tokyo, Japan). Chiral HPLC analyses were performed by using the column OD‐H (Daicel Chemical Industries, Osaka, Japan) with n‐hexane and 2‐propanol as solvent on a Hitachi Elite LaChrom device (Hitachi High Technologies America, San Jose, USA). All metagenomic esterases were produced and donated by B.R.A.I.N. AG (Zwingenberg, Germany) and used as glycerol‐stabilized crude cell extracts or as lyophilisates (Brüsehaber et al., 2008). Plasmids containing the genes for the esterases A3 (Elend et al., 2006), CE (Elend et al., 2006) and CL1 (Schmeisser et al., 2003) were cloned as described.

All ethyl esters were prepared by azeotropic esterification. One equivalent (18 mmol of 1a or 3a, and 20 mmol of 2a) of the phenylalkyl carboxylic acid was stirred with 3 eq. of ethanol at 110°C in toluene as solvent with some drops of concentrated sulfuric acid as catalyst. Water formed in the esterification reaction was removed by a water trap until completion. Next, the solvent was evaporated and the product was purified by silica gel column chromatography (n‐pentane/ethyl acetate, 10:1). Using this procedure, 1b was obtained as light yellow oil (1.8 g, 9.3 mmol, 52% yield), 2b was obtained as colourless oil (2.6 g, 14.6 mmol, 73% yield) and 3b was obtained as colourless oil (2.5 g, 13 mmol, 72% yield). The structures were confirmed by NMR spectroscopy and matched literature data (Yang et al., 1999).

The test for hydrolytic activity towards the target substrates was based on a pH assay (Baumann et al., 2000). Wells were filled with 20 µl of a substrate stock solution (200 mM in DMSO), 5 µl of a solution of the pH indicator bromothymol blue (BTB, 16 mM in 16% ethanol and 84% water), 30 µl of a glycerol stock solution of the different esterases and 145 µl of sodium phosphate buffer (5 mM, pH 7.4) to get a final volume of 200 µl. Both positive and negative controls were present in each MTP. Positive controls contained 5 µl of BTB solution, 175 µl of sodium phosphate buffer (5 mM, pH 7.4) and 20 µl of DMSO containing the correspondent acids 1a–3a at final concentrations of 1–20 mM. Negative controls were identical, but without substrates. Each MTP was then sealed with adhesive foil, incubated at 37°C for 72 h and stirred at 400 r.p.m. in a thermoshaker (Eppendorf, Hamburg, Germany). Active esterases were then identified by visible colour changes and used in analytical scale experiments.

To a stirred solution of the ethyl esters 1b–3b (0.076 mmol of 1b or 2b, or 0.15 mmol of 3b) in 1.8 ml of sodium phosphate buffer (50 mM, pH 7.4) and 10% DMSO if necessary, 200 µl of esterase solution was added to a total volume of 2 ml. The reaction mixture was stirred at 1000 r.p.m. in a thermoshaker at 20°C or 30°C. Samples (400 µl) were taken after 2, 7 and 24 h. These samples were acidified with 20 µl of 1 N HCl and subsequently extracted three times with 400 µl diethyl ether. The combined organic layers were dried over anhydrous sodium sulfate and afterwards the organic solvent was evaporated in a Speed Vac (Thermo Fisher Scientific, Langenselbold, Germany). The mixture was then treated with TMS diazomethane (Hashimoto et al., 1981) in order to convert the carboxylic acid (hydrolysis product) in its methyl ester to allow analysis of substrate and product concentration and optical purity in one run by GC. Conversion and E values were then calculated from the enantiomeric excesses from substrate and product following the equations developed by Chen and colleagues (1982).

To a 40 mM solution of 1b–3b (∼1 g) in sodium phosphate buffer (50 mM, pH 7.4) containing 10% (v/v) DMSO, esterase (lyophilisate dissolved in buffer, units as given in Table 2) was added and hydrolysis was performed at 20°C or 30°C. After TLC analysis indicated 50% conversion, the reaction was stopped by addition volume of 1 N HCl, and substrate and product were extracted three times with diethyl ether. After drying the organic phase over anhydrous Na₂SO₄, the solvent was removed by evaporation under reduced pressure and the product was purified by column chromatography (n‐pentane/ethyl acetate). The purity of the biocatalysis products was confirmed by NMR spectroscopy and was in all cases > 99%.

Using this protocol, (R)(−)‐1b was obtained as a light yellow oil (411 mg, 21 mmol, 41.4% yield). Enantiomeric excess was determined after hydrolysis to 1a as 59%ee. (S)(+)‐1a was obtained as colourless oil (230 mg, 14 mmol, 27.7% yield, 72%ee); (S)(+)‐1b was obtained as light yellow oil (420 mg, 2.2 mmol, 42% yield). Enantiomeric excess was determined after hydrolysis to 1a as 87%ee; (R)(−)‐1a was obtained as light yellow oil (330 mg, 2 mmol, 38.5% yield, 98%ee); (R)(−)‐2b was obtained as colourless oil (164 mg, 9.2 mmol, 16.4% yield). Enantiomeric excess was determined after hydrolysis to 2a as 82%ee; (S)(+)‐2a was obtained as a white powder (181 mg, 12 mmol, 21.5% yield, 97%ee); (S)(+)‐2b was obtained as a colourless oil (218 mg, 12.2 mmol, 28.3% yield). Enantiomeric excess was determined after hydrolysis to 2a as 92%ee; (R)(−)‐2a was obtained as a white powder (180 mg, 12 mmol, 21.3%, 56%ee); (R)(−)‐3b was obtained as colourless oil (380 mg, 1.99 mmol, 38% yield). Enantiomeric excess was determined after hydrolysis to 3a as 69%ee; (S)(+)‐3a was obtained as yellow oil (360 mg, 2.18 mmol, 41.9% yield, > 99%ee); (S)(+)‐3b was obtained as colourless oil (150 mg, 0.78 mmol, 15% yield). Enantiomeric excess was determined after hydrolysis to 3a as 97%ee; (R)(−)‐3a was obtained after column chromatography (n‐pentane/ethyl acetate) as yellow oil (330 mg, 2 mmol, 39% yield, 95%ee).