Note: This is part 42 of the series of publications Enzyme catalyzed reactions. Part 41 is Effenberger, F. & Osswald, S. (2001) Selective hydrolysis of aliphatic dinitriles to monocarboxylic acids by a nitrilase from Arabidopsis thaliana. Synthesis 1866–1872.
F. Effenberger, Institut für Organische Chemie, Universität Stuttgart, Pfaffenwaldring 55, D-70569 Stuttgart, Germany. Fax: + 49 711685 4269, Tel.: + 49 711685 4265, E-mail: email@example.com
The nitrilase AtNIT1 from Arabidopsis thaliana was overexpressed in Escherichia coli with an N-terminal His6 tag and purified by zinc chelate affinity chromatography in a single step almost to homogeneity in a 68% yield with a specific activity of 34.1 U·mg−1. The native enzyme (≈ 450 kDa) consists of 11–13 subunits (38 kDa). The temperature optimum was determined to be 35 °C, and a pH optimum of 9 was found. Thus, recombinant AtNIT1 resembles in its properties the native enzyme and the nitrilase from Brassica napus. The stability of AtNIT1 could be significantly improved by the addition of dithiothreitol and EDTA. The substrate range of AtNIT1 differs considerably from those of bacterial nitrilases. Aliphatic nitriles are the most effective substrates, showing increasing rates of hydrolysis with increasing size of the residues, as demonstrated in the series butyronitrile, octanenitrile, phenylpropionitrile. In comparison with 3-indolylacetonitrile, the rate of hydrolysis of 3-phenylpropionitrile is increased by a factor of 330, and the Km value is reduced by a factor of 23. With the exception of fluoro, substituents in the α position to the nitrile function completely inhibit the hydrolysis.
Nitriles are found in a variety of naturally occurring compounds such as cyanolipids, cyanoglucosides, and simple aliphatic or aromatic nitriles as metabolites of micro-organisms . In nature, the hydrolysis of nitriles to the corresponding carboxylic acid and NH3 is catalyzed by nitrilases (EC 188.8.131.52) or based on the sequential action of a nitrile hydratase (EC 184.108.40.206)–amidase (EC 220.127.116.11) system [2,3]. Most nitrilases described so far have been isolated from fungi or bacteria. In recent years, however, four nitrilases (AtNIT1–AtNIT4) have been cloned from Arabidopsis thaliana, a member of the brassicaceae family [4,5]. The genes of AtNIT1–3 are clustered␣on␣chromosome␣3 and have sequence identities of more than 80% at the amino acid level, whereas AtNIT4 has a distinct chromosomal localization and is only 65% identical with AtNIT1–3 . The subdivision of the Arabidopsis nitrilases into AtNIT1–3 and AtNIT4 is also reflected by functional differences between these enzymes. Whereas AtNIT1–3 convert 3-indolylacetonitrile (IAN) into the plant hormone 3-indolylacetic acid, IAN is not a substrate for AtNIT4 [6,7]. Moreover, homologs of AtNIT1–3 have exclusively been found in Arabidopsis and other members of the brassicaceae, whereas AtNIT4 isoforms have also been reported in species from other taxonomic groups such as tobacco  and rice . In accordance with the brassicaceae-restricted occurrence of nitrilases of the AtNIT1–3 type, these enzymes seem to be involved in the degradation of nitriles released from glucosinolates, which can be found in high concentrations in various species of the brassicaceae . Recent studies have shown that AtNIT4 and two related nitrilases from tobacco are β-cyano-(l)-alanine nitrilases . As nitrilases of the AtNIT4 type have been found in taxonomically quite distinct groups, it seems likely that AtNIT4 homologs may exist in all higher plants. In␣accordance with this is the fact that the substrate of the AtNIT4-type nitrilases, β-cyano-(l)-alanine, seems to occur in all plants as the result of detoxification of cyanide, which is inevitably produced during biosynthesis of the plant hormone ethylene .
In general, nitriles are synthetically more accessible than the corresponding carboxylic acids. Chemical hydrolysis of nitriles to carboxylic acids, however, requires drastic conditions (strong mineral acids and bases and relatively high reaction temperature). Biocatalysts for the transformation of nitriles to carboxylic acids are therefore of particular interest.
Up until now, hydratase–amidase systems, not nitrilases, have mainly been used in practice as nitrile-hydrolyzing enzymes [3,11–14]. In this paper, we report on basic investigations of the nitrilase AtNIT1 from A. thaliana, in particular, the substrate range required for the hydrolysis of nitriles to carboxylic acids. Cloning and overexpression of AtNIT1 [4,5] (EC 18.104.22.168) will provide an interesting plant nitrilase in sufficient quantities for synthetic applications. The application of AtNIT1 to the hydrolysis of several specific substrates such as aliphatic dinitriles and 2-fluoroarylacetonitriles has been published in detail [15,16].
Materials and methods
Expression cloning of AtNIT1
AtNIT1 cDNA was cloned in the expression vector pQE10 (Qiagen), which allows isopropyl β-d-thiogalactoside- induced expression of N-terminally His-tagged recombinant protein. In brief, the coding region and part of the 3′-noncoding region of AtNIT1 cDNA were amplified from an A. thaliana cDNA library (Stratagene) with an advanced polymerase system (Clontech) using the primers AtNIT1-for (5′-GCTGCTAGATCTTATGTCAACTGT CCAAAA CGCAACTCCTTTTAACGGCGTTGCCCC ATCCACC -3′; start codon according to  in bold) and AtNIT1-rev (5′-ACAATTGATGATTCAACGCCCAAC 3′). Using the BglII sites in the 5′ overhang of AtNIT1-for and the 3′-noncoding region of the cDNA, the AtNIT1 cDNA was inserted in-frame in the BamHI site of pQE10. The resulting expression plasmid pQE10-AtNIT1 was sequenced to confirm the identity of the AtNIT1 sequence after PCR amplification. pQE10-AtNIT1 was transformed in Escherichia coli M15[pREP4] cells (Qiagen) for overexpression of AtNIT1. For induction of recombinant AtNIT1, an overnight culture was performed at 37 °C in Luria–Bertani medium supplemented with ampillicin (50 µg·mL−1) and kanamycin (20 µg·mL−1), diluted 1 : 20 with Luria–Bertani medium supplemented again with ampillicin and kanamycin, and grown at 30 °C. After 4 h, isopropyl β-d-thiogalactoside was added to a final concentration of 0.5 mm for induction of AtNIT1 expression. After an additional 6 h, cells were harvested.
Preparation of the crude extract and purification of recombinant AtNIT1
Cells were separated from the nutrient medium by centrifugation (30 min, 4 °C, 5700 g), and washed with sodium phosphate buffer A (50 mm, pH 7.8). The pellet was resuspended in buffer A (100 mL per 10 g wet weight) and sonicated (3 × 5 min, 0 °C). The homogenate was centrifuged (40 min, 4 °C, 186 000 g). The supernatant (100 mL) was degassed with argon, filtered through a membrane (70 µm) and applied to a Zn2+-charged HiTrap metal chelate affinity chromatography column (Pharmacia). The column was rinsed successively with 20 mL each of sodium phosphate buffer B (50 mm, 100 mm NaCl, pH 7.8) and buffer A until the absorbance reached the base line of column equilibration. Nonspecifically bound proteins were eluted at a flow rate of 2 mL·min−1 in a 22.5-mL linear gradient of 0–100 mm imidazole in buffer A, and successively in 5 mL of sodium phosphate buffer C (50 mm, 100 mm imidazole, pH 7.8). After additional rinsing with 11.25 mL buffer A, AtNIT1 was eluted with 11.25 mL sodium phosphate buffer (50 mm, 100 mm EDTA, pH 7.8). To the collected fractions (2.5 mL), 25 µL sodium phosphate buffer (50 mm, 100 mm dithiothreitol, pH 7.8) was added, and after measurement of enzyme activity, fractions were pooled.
Recombinant purified AtNIT1 (200 µL) was separated by size-exclusion chromatography on a Superdex 200 HR10/30 column (Pharmacia) in 50 mm sodium phosphate buffer, containing 100 mm EDTA and 1 mm dithiothreitol, pH 7.8, at a flow rate of 0.5 mL·min−1. For calibration of the column, thyroglobulin (663 kDa), apoferritin (443 kDa), alcohol dehydrogenase (150 kDa), BSA (66 kDa), carbonic anhydrase (29 kDa) and cytochrome c (12.4 kDa) (all from Sigma) were used.
Enzyme activity towards 3-phenylpropionitrile was assayed using bacterial protein (0.34–135 mg) in 5 mL Tris/HCl buffer (70 mm, pH 8.5) and 50 µL 3-phenylpropionitrile in methanol (0.25 m). The reaction was carried out for 1 h at 35 °C. An aliquot of 1 mL was acidified with 50 µL HCl (5 m) and extracted with diethyl ether (5 mL). After centrifugation (5 min, 2000 g) and cooling at −30 °C for 30 min to freeze the aqueous layer, the organic layer was decanted and derivatized with ethereal diazomethane (0.2 m). After concentration, the residue was taken up in 1 mL diethyl ether and subjected to gas chromatography on a Carlo Erba Fractovap 4160 with FID and Spectra Physics minigrator using a capillary glass column (50 m) with PS086 and carrier gas 50 kPa hydrogen. Peak areas were calibrated as follows. A volume of 5 mL each of a solution of 3-phenylpropionitrile (181.5 mg) and 3-phenylpropionic acid (205.2 mg) in methanol (10 mL), and Tris/HCl buffer (990 mL, 70 mm, pH 8.5) were mixed, and 5 mL from this mixture was added to 5 mL of the 3-phenylpropionitrile solution. This procedure was repeated three times. A sample of 1 mL from each solution was treated as described above and analyzed by gas chromatography. The conversion factor was determined from the plot of ratio areas vs. ratio concentrations. One unit is defined as 1 µmol converted·min−1.
Determination of temperature and pH optimum of AtNIT1
Temperature dependence. Nitrilase activity towards 3-phenylpropionitrile was assayed as described above using purified enzyme (55.2 U·mL−1, 1.98 mg protein·mL−1) in a 1 : 5000 dilution with Tris/HCl buffer (70 mm, pH 8.5) and 50 µL 3-phenylpropionitrile in methanol (0.25 m). The reaction was initiated by the addition of substrate either directly after preliminary heating at the respective temperature for 10 min or cooling at 7 °C for 30 min and after 24 h, respectively.
PH dependence. Enzyme activity was assayed as described above using purified enzyme (56.8 U·mL−1, 1.78 mg protein·mL−1) in phosphate buffer (50 mm, pH 7.8), which was diluted (1 : 5000) at 4 °C with the respective buffer. After preliminary warming at room temperature, the reaction was initiated by the addition of 50 µL 3-phenylpropionitrile in methanol.
Purification and determination of Km values
Recombinant AtNIT1 was purified from E. coli lysates by metal chelate affinity chromatography using a Zn2+-charged HiTrap column. After a wash step with 100 mm imidazole, the tightly bound AtNIT1 was eluted with high recovery by 100 mm EDTA (Fig. 1A). This single-step purification yielded almost pure AtNIT1 (Fig. 1B) with a specific activity of 34.1 U·mg−1(Table 1) and a subunit mass of 38 kDa (Fig. 1B). Recombinant AtNIT1 was eluted during gel-filtration chromatography (Fig. 1C) in fractions corresponding to a molecular mass of ≈ 450 kDa, suggesting that native AtNIT1 occurs as a homomeric protein complex of 11–13 subunits (data not shown).
Table 1. Summary of purification of the nitrilase from A. thaliana.
Total activity (U)
Total protein (mg)
Specific activity (U·mg−1)
Recombinant AtNIT1 showed a Km value of 3.67 mm for 3-indolylacetonitrile and 0.159 mm for 3-phenylpropionitrile (Fig. 2A,B). The Km value for 3-indolylacetonitrile is in good agreement with a reported value of 5 mm.
As a crude extract of recombinant AtNIT1 had a half-time of 2 days at pH 8 and 4 °C, the influence of antioxidants and protease inhibitors on enzyme stability was investigated.
Of the applied thiols that proved to be good antioxidants (mercaptoethanol and dithiothreitol) [4,17–21], dithiothreitol had the better stabilizing effect (Table 2). The loss of enzyme activity on the addition of 2 mm dithiothreitol was 20% compared with 63% for the reference without thiol. However, increasing the dithiothreitol concentration to 5 mm did not further improve enzyme stability. The best result with protease inhibitors was achieved using EDTA at a concentration of 2 mm (Table 2). Thus, all buffers used for cell disintegration and conversions were supplemented with dithiothreitol and EDTA (2 mm each). In this way, we succeeded in significantly increasing the enzyme stability of both crude extract and purified enzyme: after 2 days at room temperature and 3 months at 4 °C, 95% and 90% enzyme activity, respectively, remained.
Table 2. Effect of antioxidants and protease inhibitors on enzyme activity. Enzyme activity was determined after incubation of 5 mL crude enzyme extract in Tris/HCl buffer (70 mm, pH 8.0) with the respective antioxidant (neat) or a stock solution of protease inhibitors (50-fold concentration; Protease-Inhibitor-Set, Boehringer-Mannheim). The reaction was carried out for 48 h at room temperature with vigorous sti2;ring. The initial activity of 112 U·L−1 is 100% in the case of antioxidants and 97 U·L−1 in the case of inhibitors.
The nitrilases investigated so far generally show highest activity in the temperature range 35–40 °C, no matter what the enzyme source [18,21–23]. However, as little is known about their stability at higher temperatures, which is a decisive factor in their application as biocatalysts in chemical reactions, the effect of temperature on AtNIT1 stability was investigated. Recombinant AtNIT1 shows a sharp temperature optimum at 35 °C, determined after 1 h of incubation, with a gentle slope at < 35 °C and a steeper slope at > 35 °C (Fig. 3). Enzyme stability at different temperatures was determined after 24 h of incubation. At 25 °C and 35 °C, only a slight decrease in activity was found. At 35 °C, the relative enzyme activity amounts to ≈ 80%, whereas the enzyme was almost completely deactivated at 40 °C. The highest absolute enzyme activity, however, was found at 35 °C, and, moreover, the stability at this temperature is sufficient for applications in longer lasting biotransformations.
PH dependence of AtNIT1
The pH dependence of AtNIT1 was investigated with different buffer systems in order to guarantee sufficient buffering capacity in the range pH 6–10 (Fig. 4).
As can be seen from Fig. 4, the choice of the buffer system affects the enzyme activity slightly, changing from Tris/HCl to glycine/NaOH. With both buffer systems, however, an activity optimum of pH 9.0 was found, with 97% of the maximum activity being measured at pH 8.5. The decrease in enzyme activity at pH values > 9 is not an irreversible process: acidifying an enzyme solution with pH 10 back to pH 9 resulted in > 80% recovery of activity. The pH optimum measured in this way is in slight contrast with the value of pH 7.5 reported for the nitrilase from A. thaliana, possibly arising from the deviant structure at the N-terminus. However, several bacterial nitrilases also clearly have a basic pH optimum [23–27].
Substrate range of recombinant AtNIT1
The substrate range of recombinant AtNIT1 was investigated using structurally varied aromatic and aliphatic nitriles (Table 3). The activities given in Table 3 are referred to the specific nitrilase activity towards butyronitrile. As can be seen, aliphatic nitriles are the most effective substrates, showing increased rates of hydrolysis with increasing size of␣the hydrophobic residue, in the order butyronitrile, octanenitrile, phenylpropionitrile. In contrast with 3-phenylpropionitrile, arylacetonitriles such as benzyl cyanide were converted 20 times more slowly. Aromatic nitriles, such as benzonitrile, were converted even more slowly (270 times) than phenylpropionitrile. The assumed natural substrate of AtNIT1, 3-indolylacetonitrile , was found to be a poor substrate (Table 3). Also the hydrolytic rate of cinnamonitrile, an α,β-unsaturated system, is significantly diminished compared with the corresponding saturated phenylpropionitrile. A double bond in the β,γ-position, however, has almost no effect on enzyme activity, as can be seen if 4-phenyl-3-butenenitrile is compared with 4-phenylbutyronitrile. Suitability as a substrate is strongly influenced by the substituents in the 2-position. All substituents other than fluoro inhibit enzymatic hydrolysis almost completely (Table 3). Nitriles with substituents in the 3-position, for example 3-methylbutyronitrile, are also poor substrates, but the decrease in the hydrolytic rate is less pronounced. Interestingly, benzoylglycine nitrile is a much better substrate for AtNIT1 than glycine nitrile itself.
Table 3. Relative activities of recombinant AtNIT1-catalyzed hydrolysis of nitriles. The reactions were performed in Tris/HCl buffer (70 mm, with dithiothreitol and EDTA, 2 mm each, pH 8) at room temperature. At a concentration of 1.25 mm, all substrates were completely soluble; the enzyme concentration was varied so that the reaction time for all substrates was in the range 2–4 h for conversion of 15–40%. Relative activities are referred to the specific nitrilase activity towards butyronitrile [1.393 µmol·min−1·(mg protein)−1= 100%].
Relative activity (%)
Relative activity (%)
a See also literature data . b24 h reaction time. cHydrolysis at pH 7.0.
Acid amides as byproducts of AtNIT1-catalyzed nitrile hydrolysis
Acid amide was first detected as a major product of AtNIT1-catalyzed nitrile hydrolysis with fumaronitrile as substrate. In this reaction, which was followed by gas chromatography, less than 10% of the expected amount of 3-cyanoacrylic acid, estimated from the calibration, was formed. A product mixture of an unidentified product and 3-cyanoacrylic acid in the ratio 93 : 7 (Table 4) was found by HPLC. As demonstrated by co-injection, fumaric acid was not formed in the reaction. After extractive separation from 3-cyanoacrylic acid and subsequent recrystallization from chloroform, the unknown product was isolated in 68% yield and unambiguously characterized as 3-cyanoacrylamide by elemental analysis and NMR spectroscopy. The amide to acid ratio was independent of conversion. The isolated amide was not hydrolyzed to 3-cyanoacrylic acid under these reaction conditions. Moreover, the hydrolytic rate and ratio of acid to amide did not depend on enzyme purity, giving similar results with both crude extract and highly purified enzyme. From blank experiments, it could also be excluded that it was an impurity of nitrile hydratase.
As 3-cyanoacrylamide had been identified as a byproduct of fumaronitrile hydrolysis, the AtNIT1-catalyzed hydrolysis of other substrates with donor and acceptor substituents was investigated with respect to amide formation (Table 4). Amides have also been found as major products in the AtNIT1-catalyzed hydrolysis of α-fluoroarylacetonitriles . An α-fluoro substituent, however, does not conclusively result in amide formation as can be seen in the hydrolysis of α-fluorobutyronitrile, yielding 95% of the corresponding acid (Table 4). Nevertheless, both a fluoro substituent in the α position and a second nitrile group conjugated to the nitrile (fumaronitrile) seem to play a decisive role in amide formation.
The assumption that electron-withdrawing substituents favor the formation of amides was supported by the hydrolysis of differently 3-substituted acrylonitriles (Table 4). Whereas 3-nitroacrylonitrile was hydrolyzed to 3-nitroacrylamide as sole product, in the case of the donor-substituted 3-methoxyacrylonitrile and crotononitrile, the corresponding acids were formed almost quantitatively (Table 4). As 3-nitroacrylonitrile tends to decompose under basic conditions, the reaction was performed at pH 6 (Table 4), where the amide was formed only by enzyme-catalyzed hydrolysis and not by chemical reaction, as confirmed by a blank experiment. Table 4 reveals that the relative activity is almost completely independent of the kind of substituent.
Analysis of the substrate range with a variety of structurally different aromatic and aliphatic nitriles revealed that aliphatic nitriles are hydrolyzed more efficiently than the natural substrate IAN or structurally related aromatic nitriles. With a relative activity of only 2.2%, compared with butyronitrile, 3-indolylacetonitrile is a poorer substrate for AtNIT1 than benzyl cyanide (31% relative activity). This finding is in agreement with literature data, showing that IAN is one of the weakest substrates . The order of the relative AtNIT1 activity towards the substrates 3-phenylpropionitrile, 4-phenylbutyronitrile and benzyl cyanide (Table 3) also corresponds to that just recently reported . 2-Substituted substrates such as 2-methylbutyronitrile and 2-phenylpropionitrile, however, were almost completely unacceptable for AtNIT1, indicating that substituents in the 2-position, other than fluoro, inhibit the hydrolysis. The broad substrate range observed for AtNIT1 in this study is in good agreement with reports showing that AtNIT1 acts on a variety of aliphatic and aromatic substrates [4,9] and is in contrast with the high specificity of AtNIT4 for β-cyano-(l)-alanine . Its broad substrate range, recombinant accessibility and reasonable stability make AtNIT1 a promising candidate for applications in organic chemistry, in particular the synthesis of optically active 2-fluorocarboxylic acids, which are very useful as analogs of pheromones and antirheumatics, for example . Also the monohydrolysis of aliphatic dinitriles to monocarboxylic acids is of great industrial interest because selective chemical hydrolysis is virtually impossible .
The formation of amides as byproducts of nitrilase-catalyzed reactions was first reported as early as 1964 [28,29]. Furthermore, in subsequent publications [19,30–32], small amounts of amides (< 15%) could be detected besides the carboxylic acids during nitrilase catalysis. In all cases, the amide to acid ratio was independent of reaction conditions (temperature and pH) and the applied enzyme concentrations. In their basic work on the four A. thaliana nitrilases NIT1–4, Bartel & Fink  described the conversion of IAN into 3-indolylacetic acid and indole-3-acetamide and found that the latter is not a substrate for these enzymes. For the hydrolysis of β-cyano- (l)-alanine, catalyzed by NIT4, Piotrowski et al.  reported the simultaneous formation of asparagine and aspartic acid in a ratio of 1.5 : 1, independent of reaction conditions. A dependence of the amide to acid ratio on the substituents, however, has not been reported in the literature so far.
Until now the reaction mechanism of nitrilase-catalyzed hydrolysis has not been confirmed experimentally. The mechanism postulated [19,28,29,33] involves the donation of a cysteine from the enzyme to the nitrile group to yield a thioimidate, which subsequently forms a tetrahedral intermediate A by addition of water. Generally, NH3 is eliminated from this intermediate A to give a thioester, which reacts with a further water molecule to give the carboxylic acid (Fig. 5). Therefore, the formation of the acid amide from A logically arises from the elimination of cysteine.
It has been shown for the chemical hydrolysis of thioimidate esters [34,35] that the formation of thiol ester is favored in acidic medium (pH < 2.7), whereas at higher pH values (pH > 2.7) the formation of amide dominates. This result was explained by a facilitated elimination of NH3 caused by protonation of the amino group in the tetrahedral intermediate.
Although, as mentioned, some papers have dealt with the mechanism of the nitrilase-catalyzed hydrolysis of nitriles, a relationship between the chemical structure of the substrate and the amount of acid amide formation has not so far been described. For AtNIT1-catalyzed nitrile hydrolysis, we could demonstrate for the first time such a structural relationship, because the amide formation clearly depends on the kind of substituent. The preference for acid amide formation by α-fluoro substituents or by acceptor groups (CN, NO2) in π-conjugated nitriles is clear evidence of an electronically preferred formation and stabilization of the tetrahedral intermediate A in the enzyme–substrate complex. Because the crystal structure of the active site of AtNIT1 is not known, how the stabilization of the tetrahedral intermediate assists the elimination of cysteine to yield the acid amide cannot be explained.
Chemical hydrolysis of many nitriles with labile substituents catalyzed by acid or base is virtually impossible because of the drastic reaction conditions required. Therefore, over the last few years, biocatalysts capable of hydrolyzing nitriles to carboxylic acids have been intensively investigated . In␣most cases, however, nitrile hydratase–amidase systems have been described, although not exclusively . The nitrilase AtNIT1 from A. thaliana is the first plant nitrilase to be investigated with respect to its synthetic potential. Because of optimized expression, the enzyme is now accessible in sufficient quantities. Clear optimization of enzyme stability under the reaction conditions, which is important for practical application, could be achieved by addition of the protease inhibitor EDTA (Table 2). Therefore, slowly reacting nitriles can also be hydrolyzed without problem.
The most important criteria for practical applications, however, are the substrate range and selectivity of an enzyme. In contrast with other nitrile-hydrolyzing enzymes, the nitrilase AtNIT1 stands out as having a very broad substrate range (Table 3). Although longer-chain aliphatic nitriles are the most effective substrates, hydrolysis of aromatic nitriles is also catalyzed. Because of the clearly improved enzyme stability, AtNIT1-catalyzed hydrolysis is also applicable to aromatic nitriles. Moreover, AtNIT1 shows a very interesting stereoselectivity and chemoselectivity. The influence of substituents in the α position to the nitrile function has already been mentioned. Because the enzyme does not accept any substituents at the α position except fluoro, compounds bearing several different cyano groups can be selectively hydrolyzed. Hydrolysis of racemic 2-fluoroarylacetonitriles proceeds enantioselectively . In dinitriles with chemically comparable cyano groups (e.g. adiponitrile), only one cyano group is hydrolyzed exclusively to give the corresponding cyanocarboxylic acids [16,31,37], opening up interesting possibilities for organic synthesis, for example the preparation of certain lactams . Furthermore, AtNIT1 exhibits cis/trans selectivity with α,β-unsaturated nitriles , as also reported for other enzymes [19,39,40].
Because of its broad substrate range on the one hand and unusual regioselectivities and stereoselectivities, the nitrilase AtNIT1 from A. thaliana is a very interesting biocatalyst in organic synthesis.
This work was generously supported by the Fonds der Chemischen Industrie. We acknowledge Dr K. Trummler for assistance in enzyme purification, Dr S. Förster for fermentation, and Dr A. Baro for preparing the manuscript.