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Appl Microbiol Biotechnol (2012) 93:1553–1561 DOI 10.1007/s00253-011-3525-7

ORIGINAL PAPER

Purification and characterization of heterologously expressed nitrilases from filamentous fungi Alena Petříčková & Alicja Barbara Veselá & Ondřej Kaplan & David Kubáč & Bronislava Uhnáková & Anna Malandra & Jürgen Felsberg & Anna Rinágelová & Philip Weyrauch & Vladimír Křen & Karel Bezouška & Ludmila Martínková

Received: 19 March 2011 / Revised: 21 July 2011 / Accepted: 3 August 2011 / Published online: 3 September 2011 # Springer-Verlag 2011

Abstract Nitrilases from Aspergillus niger CBS 513.88, A. niger K10, Gibberella moniliformis, Neurospora crassa OR74A, and Penicillium marneffei ATCC 18224 were expressed in Escherichia coli BL21-Gold (DE3) after IPTG Alena Petříčková and Alicja B. Veselá contributed equally to this work. Electronic supplementary material The online version of this article (doi:10.1007/s00253-011-3525-7) contains supplementary material, which is available to authorized users. A. Petříčková : A. B. Veselá : O. Kaplan : D. Kubáč : B. Uhnáková : A. Malandra : A. Rinágelová : P. Weyrauch : V. Křen : K. Bezouška : L. Martínková (*) Institute of Microbiology, Centre of Biocatalysis and Biotransformation, Vídeňská 1083, CZ-142 20 Prague, Czech Republic e-mail: [email protected] J. Felsberg Institute of Microbiology, Centre for DNA Sequencing, Academy of Sciences of the Czech Republic, Vídeňská 1083, CZ-142 20 Prague, Czech Republic

induction. N. crassa nitrilase exhibited the highest yield of 69,000 UL−1 culture. Co-expression of chaperones (GroEL/ES in G. moniliformis and P. marneffei; GroEL/ ES and trigger factor in N. crassa and A. niger CBS 513.88) enhanced the enzyme solubility. Specific activities of strains expressing the former two enzymes increased approximately fourfold upon co-expression of GroEL/ES. The enzyme from G. moniliformis (co-purified with GroEL) preferred benzonitrile as substrate (K m of 0.41 mM, Vmax of 9.7 μmol min−1 mg−1 protein). The P. marneffei enzyme (unstable in its purified state) exhibited the highest Vmax of 7.3 μmol min−1 mg−1 protein in cellfree extract, but also a high Km of 15.4 mM, for 4cyanopyridine. The purified nitrilases from A. niger CBS 513.88 and N. crassa acted preferentially on phenylacetonitrile (Km of 3.4 and 2.0 mM, respectively; Vmax of 10.6 and 17.5 μmol min−1 mg−1 protein, respectively), and hydrolyzed also (R,S)-mandelonitrile with higher Km values. Significant amounts of amides were only formed by the G. moniliformis nitrilase from phenylacetonitrile and 4-cyanopyridine.

A. Petříčková : A. B. Veselá : K. Bezouška Department of Biochemistry, Faculty of Science, Charles University in Prague, Hlavova 8, CZ-128 40 Prague, Czech Republic

Keywords Nitrilase . Chaperones . Aspergillus niger . Gibberella moniliformis . Neurospora crassa . Penicillium marneffei

A. Malandra Department of Chemistry, Chemical Engineering and Materials, University of L’Aquila, Via Campo di Pile - Zona industriale di Pile, 67100 L’Aquila, Italy

Introduction

P. Weyrauch Institute of Molecular Microbiology and Biotechnology, Westfalian Wilhelms-University Münster, Correnstrasse 3, 48149 Münster, Germany

Nitrilases are versatile enzymes in biocatalysis, mediating the hydrolysis of diverse nitriles under mild conditions. Some of their products, notably hydroxy acids, such as mandelic (Rustler et al. 2008; Sosedov et al. 2009), 3hydroxyvaleric (Wu et al. 2007), or glycolic acid (Wu et al. 2008), are of significant industrial importance. These enzymes can also be used to produce amides, which are

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their usual by-products and even major products in some cases (e.g., Osswald et al. 2002; Fernandes et al. 2006; Kaplan et al. 2006a; Sosedov et al. 2009). Most of the nitrilases studied and explored as biocatalysts have been bacterial in origin except for a few fungal and plant enzymes (for reviews see, O'Reilly and Turner 2003; Thuku et al. 2009). Besides this, nitrilases of unknown origin were obtained from metagenomic DNA (Robertson et al. 2004). According to both activity screens (Kato et al. 2000) and gene database searches (Martínková et al. 2009), filamentous fungi seem to be a rich source of nitrilases. The first two fungal nitrilases were purified in the Fusarium genus and characterized (Harper 1977; Goldlust and Bohak 1989) without determining their amino acid sequences. The other two nitrilases from the same genus were purified and characterized by ourselves and their partial amino acid sequences determined (Vejvoda et al. 2008, 2010). The nitrilase studied by us in Aspergillus niger K10 was the first fungal nitrilase to be purified from both its native (Kaplan et al. 2006a) and heterologous producer (Escherichia coli; Kaplan et al. 2011a). Gene databases proved to be an abundant source of hypothetical nitrilase sequences and three of these enzymes were recently expressed in E. coli by us, namely a nitrilase from A. niger CBS 513.88, Neurospora crassa OR74A, and Gibberella moniliformis (Kaplan et al. 2011b). Synthetic genes were prepared according to published sequences, and functional enzymes were obtained in all cases. The aim of this work was to characterize these enzymes in order to define their potential biocatalytic use. A further two enzymes were also included in this study, namely a nitrilase from Penicillium marneffei ATCC 18224 and the aforementioned enzyme from A. niger K10. Both enzymes were expressed from the corresponding synthetic genes but the latter also from the gene amplified from the cDNA of the native organism to compare the efficiency of both methods. Aiming to improve the heterologous expression of nitrilases, we also examined the effect of molecular chaperones (GroEL/ES, dnaK-dnaJ-grpE, trigger factor (TF)), which are known for their ability to facilitate protein folding and hence to improve the production of active proteins (Nishihara et al. 2000).

Materials and methods Nitrilase expression The expression of all nitrilases was performed in E. coli BL21-Gold (DE3) (Agilent Technologies—Stratagene Products, USA). The enzyme from A. niger K10 (GenBank ABX75546) was expressed using either the gene amplified from the wild-type producer (Kaplan et al. 2011a) or the synthetic gene (GeneArt, Regensburg, Germany), the codon

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frequency of which was optimized using GeneArt's own software. A strain transformed with plasmid pOK101 was used to express the former gene (Kaplan et al. 2011a). The latter gene was cloned into the Nde I and Hind III sites of plasmid pET 30a(+) to construct plasmid pOK106 and expressed in E. coli without the His-tag. The expression of nitrilases from N. crassa OR74A (GenBank CAD70472), G. moniliformis (GenBank ABF83489), and A. niger CBS 513.88 (GenBank XP_001397369) was performed using strains transformed with plasmids pOK103, pOK104, and pOK105, respectively (Kaplan et al. 2011b). The gene encoding the nitrilase in P. marneffei ATCC18224 (GenBank XP_002144951 (protein)) was synthesized with GeneArt according to the published sequence, which was obtained using the programs BLASTX and BLASTP (Altschul et al. 1997; http://blast.ncbi.nlm.nih.gov/Blast.cgi). The gene was cloned into the Nde I and Hind III sites of plasmid pET 28a (+) to construct plasmid pOK107 and expressed in E. coli with an N-terminal His-tag. The optimized gene sequences are shown in supplementary Fig. S1 (P. marneffei, GeneBank JN012233; A. niger K10, GeneBank JN243351) and in the previous work (A. niger CBS 513.88, GeneBank JN012230; G. moniliformis, GeneBank JN012231; N. crassa, GeneBank JN012232; Kaplan et al. 2011b, Electronic supplementary material). Nitrilase and chaperone co-expression E. coli BL21-Gold (DE3) (Agilent Technologies—Stratagene Products, USA) was transformed with one of the plasmids containing nit genes and one of the chaperone plasmids (pG-KJE8, pGro7, pKJE7, pG-Tf2 or pTf16; Takara Bio Inc., Japan), resulting in 30 strain harboring different nit and chaperone gene combinations. Strains lacking chaperone genes served as controls. The strains were grown in LB with kanamycin (50 μg mL−1) and chloramphenicol (20 μg mL−1). Chaperone screening was performed at 25°C in 10-mL culture volumes in shaken 50-mL Falcon tubes closed with screw caps. After OD610 reached approximately 0.5, the expression of chaperones was induced with −1 −1 L-arabinose (2 gL ) or tetracycline (5 μg L ) according to the manufacturer's instructions (Takara). At the same time, expression of the nitrilase was induced with 0.5 mM IPTG, and the cultivation continued under the same conditions for a further 17 h. Selected cultures were scaled up to a 200-mL volume using shaken 500-mL Erlenmeyer flasks closed with cotton plugs. The cultivation was carried out at 37°C for 3.5–4 h (until the culture reached an OD610 of approximately 0.5). Then, the cultivation temperature was decreased to 25°C and the expression of chaperones and nitrilase was induced as described above. The cells were harvested after a further 17 h.

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Nitrilase purification The cells were disrupted by sonication and cell debris removed by centrifugation (13,000 g, 4°C, 15 min). The nitrilases from A. niger CBS 513.88 and N. crassa were purified from cell-free extracts on a Hi-Prep 16/10 Q FF column (GE Healthcare), with a linear gradient of NaCl (0.15–1 M) in Tris/HCl buffer (50 mM, pH 8.0) at 2 mL min−1 in the first step. Active fractions were pooled, concentrated using an Amicon Ultra-4 unit (cut-off 30 kDa; Millipore) and applied on a Superdex 200 HR 10/300 GL column (GE Healthcare) using a Tris/HCl buffer (50 mM, pH 8.0, 150 mM NaCl) at 0.4 mL min−1. Alternatively, HiPrep 16/60 Sephacryl S-200 HR (GE Healthcare) was used in this step for nitrilase from G. moniliformis. Enzyme was eluted with the same buffer at 2 mL min−1. Active fractions were pooled, concentrated as described above and stored at −80°C. The nitrilase from P. marneffei was partially purified by affinity chromatography on Talon Metal Affinity Resin (Clontech). The cell-free extract was incubated with the resin at 4°C for 30 min, centrifuged (750 rpm, 5 min) and the pellet resuspended in Tris/HCl buffer (50 mM, pH 8.0). The mixture was transferred into a 25-ml column and proteins eluted with a stepwise gradient of imidazole (0–1– 150 mM) in the same buffer. Active fractions (eluted at 150 mM imidazole) were pooled and concentrated as described above. Nitrilase assay Strains were screened for optimum nitrilase–chaperone combinations using reaction mixtures (total volume 0.5 mL each) containing an appropriate amount of whole cells resuspended in Tris/HCl buffer (50 mM, pH 8.0; 150 mM NaCl) and 25 mM benzonitrile (from 500 mM stock solution in methanol). The suspensions were shaken at 30°C for 5 min, the reaction initiated by the addition of benzonitrile and terminated after a further 10 min incubation under the same conditions. Cells from cultivations on 200-mL scale were assayed in an analogous way with slight modifications; activities of G. moniliformis and P. marneffei were assayed with 25 mM benzonitrile after a 5 min reaction and those of N. crassa and A. niger with 25 mM phenylacetonitrile after a 1 min reaction. To determine the nitrilase activity in the soluble and insoluble fractions, the cells were disrupted by sonication and cell debris removed by centrifugation (13,000×g, 4°C, 30 min). The pellet was resuspended in Tris/HCl buffer (50 mM, pH 8.0; 150 mM NaCl) and it and the supernatant were used as the insoluble and soluble fraction, respectively. An appropriate amount of each fraction was taken for the above nitrilase assay. The nitrilases from G. moniliformis and

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P. marneffei were assayed with 25 mM benzonitrile and the nitrilases from N. crassa and A. niger CBS 513.88 with 25 mM phenylacetonitrile under the above conditions. Km and Vmax values were determined with 1–25 mM of substrates. Temperature and pH optima were assayed at 15–60°C and pH 4.0–12.0 (using 100 mM acetic acid/boric acid/ phosphoric acid/NaOH buffers). The residual activities after pre-incubation of enzymes at 25–60°C or pH 4.5–12.0 were assayed at optimum temperature of each enzyme (43°C and 30°C for nitrilases from G. moniliformis and P. marneffei, respectively; 38°C for other enzymes) and pH 8.0. Analytical HPLC Benzonitrile, its analogues, phenylacetonitrile, (R,S)-2phenylpropionitrile, (R,S)-mandelonitrile, and their reaction products (acids, amides) were analyzed using a Chromolith Flash RP-18 (Merck, 25×4.6 mm) with a mobile phase consisting of CH3CN (10–25%) and H3PO4 (0.1%) in water at 2 mL min−1 and 35°C. 4-Cyanopyridine and its reaction products were analyzed as described previously (Malandra et al. 2009); 2- and 3-cyanopyridine and their reaction products were analyzed in the same way. Protein assay Protein concentration was determined according to Bradford (1976) using Bradford Protein Assay (Bio-Rad Laboratories, USA) with bovine serum albumin as the standard. SDS-PAGE Proteins were analyzed by 12% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) (Laemmli 1970) followed by Coomassie Brilliant Blue R-250 staining.

Results Variability of fungal nitrilases Putative fungal nitrilases/cyanide hydratases available in the GenBank can be classified into a number of sequence clades (see supplementary Fig. S2). With the aim to exploit the diversity of these enzymes, we selected four of them, which shared a relatively low homology degree. All of them were only distantly related (with less than 40% amino acid identities) to the nitrilase from A. niger K10, for which both its amino acid sequence and its biochemical properties have been already known (Kaplan et al. 2011a). The nitrilases from A. niger CBS 513.88, N. crassa OR74A,

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and G. moniliformis exhibited less than 50% amino acid identities to each other (see supplementary Fig. S3 for multiple sequence aligment). The P. marneffei enzyme shared a higher identity to G. moniliformis than the other enzymes (56%). The identity degree of all these fungal nitrilases to well-known bacterial nitrilases was approximately 40%. The nitrilase from N. crassa is “related to aliphatic nitrilase” according to GenBank, and the enzymes from G. moniliformis and P. marneffei were postulated to act on aromatic nitriles due to their similarity to characterized nitrilases from genus Fusarium (Vejvoda et al. 2010). Preliminary whole-cell experiments confirmed the expected substrate specificities, indicating that the substrate preference of the enzymes differed between the nitrilase from G. moniliformis on one side and nitrilases from N. crassa and A. niger on the other (Kaplan et al. 2011b). The latter enzymes acted preferentially on phenylacetonitrile unlike that from G. moniliformis and previously characterized fungal nitrilases in general (Martínková et al. 2009), which exhibited their maximum activities for benzonitrile or 4-cyanopyridine. High relative activities of E. coli cells expressing the nitrilase from P. marneffei for benzonitrile and 4-cyanopyridine suggested that this enzyme was also an aromatic nitrilase (data not shown). The enzyme studied previously by us in A. niger K10 (Kaplan et al. 2011a) was more closely related to cyanide hydratases (with approximately 60–85% amino acid identities) than to other fungal nitrilases studied here (maximum amino acid identity of approximately 38% with P. marneffei nitrilase). In compliance with this finding, it produced a high ratio of by-product amide, which is the sole product of cyanide hydratase (i.e., formamide from HCN; O'Reilly and Turner 2003). These enzymes served as a representative set of diverse nitrilases from filamentous fungi for comparing their heterologous expression in the presence and absence of molecular chaperones and for investigating their catalytic properties. Nitrilase in Aspergillus niger K10 It was hypothesized that the enzyme from A. niger K10 was not correctly folded in E. coli, as its specific activity for benzonitrile decreased significantly and its substrate specificity changed compared to the enzyme isolated from the native organism (Kaplan et al. 2011a). MALDI-TOF analysis showed that the recombinant enzyme retained a C-terminal 46-amino acid peptide which was cleaved in the native organism (ibid.). It is possible that enzyme misfolding led to this missing posttranslational modification. A similar posttranslational modification in Rhodococcus rhodochrous J1 was probably due to autocatalytic cleavage

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(Thuku et al. 2007) and the misfolded enzyme from A. niger may have lost this function. However, molecular chaperones did not improve this enzyme's activity significantly (Kaplan et al. 2011a; this work—supplementary Table S4). In this work, the enzyme from A. niger K10 was produced from both the gene amplified from the native organism and the synthetic gene optimized according to E. coli codon bias. However, the latter approach was less efficient with an active nitrilase production an order of magnitude lower (about 20 vs. 200 UL−1 of culture). It is possible that using the optimized gene led to a translation speed that was too high, which hampered the folding of the recombinant product and could finally result in its degradation or aggregation. Regions that are comprised of rare codons can delay peptide elongation and thus provide time for correct folding of the nascent protein. Indeed, the optimization of codon frequency in heterologously expressed genes was reported to lead to an enhanced proportion of misfolded and aggregated protein (for a review, see Sabate et al. 2010). Nitrilases in Aspergillus niger CBS 513.88 and Neurospora crassa OR74A A screening of molecular chaperone effects on total nitrilase production (supplementary Table S4) did not indicate any significant effect of chaperone co-expression in strains expressing the A. niger CBS 513.88 or N. crassa enzyme. Only in strains simultaneously expressing GroEL/ES chaperones and trigger factor was a slight increase in nitrilase activity per liter of culture observed (by a factor of 1.25 times). Activity measurement in the soluble and insoluble cell fractions showed that the percentage of nitrilase activity in the soluble fraction was also increased in strains co-expressing these chaperones compared to the controls (from 53% to 69% and 65% to 77% in strains expressing nitrilases from N. crassa and A. niger, respectively). Therefore, these strains were used to purify the enzymes. A more significant increase in nitrilase production was, however, achieved by modifying the cultivation method in terms of the culture volume (200 vs. 10 mL), type of cultivation vessels (flasks vs. Falcon tubes), and initial cultivation temperature (37°C vs. 25°C). Thus, the cells expressing nitrilases from N. crassa and A. niger yielded up to 69,000 and 23,000 UL−1 of culture (6,700 and 3,650 UL−1 of cell suspension of OD610 =1), respectively, for phenylacetonitrile. Purification of enzymes from N. crassa and A. niger to near homogeneity (Fig. 1) was accomplished in two steps with a 2.4 and threefold increase in specific activity, respectively, and approximately 50% yields. SDS-PAGE suggested that traces of GroEL could co-purify with

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optimum (7.0–9.5) than the enzyme from N. crassa (8.0–8.5) (data not shown). The former enzyme was also more stable at alkaline pH values (Fig. 2a). Both enzymes were relatively stable at temperatures below 35°C and still retained 40–60% activity after 1-h preincubation at 40°C (Fig. 2b). Nitrilases in Gibberella moniliformis and Penicillium marneffei

Fig. 1 Expression in E. coli and purification of nitrilases from Neurospora crassa OR74A, Aspergillus niger CBS 513.88, Gibberella moniliformis, and Penicillium marneffei ATCC 18224. Lane 1 marker. Lanes 2, 4, 6, 8 cell-free extracts from strains expressing nitrilases from N. crassa, A. niger, G. moniliformis, and P. marneffei, respectively. Lanes 3, 5, 7, 9 (partially) purified nitrilases from N. crassa, A. niger, G. moniliformis, and P. marneffei, respectively. Nitrilase from P. marneffei was purified by affinity chromatography. Other enzymes were purified in two steps (Q-Sepharose, Superdex 200). Upper bands in lanes 5–9 indicate co-purification with GroEL

nitrilase from A. niger. According to purification results (data not shown), the nitrilases in N. crassa and A. niger formed approximately 42% and 34% of the total soluble protein, respectively. An examination of the substrate specificity of the purified nitrilases confirmed the findings made with whole cells (Kaplan et al. 2011b); the enzymes from N. crassa and A. niger CBS 513.88 acted preferentially on phenylacetonitrile. The purified enzyme from N. crassa exhibited an approximately 1.6 times higher Vmax value for phenylacetonitrile than that from A. niger (Table 1). (R,S)Mandelonitrile was also hydrolyzed at significant rates, but, compared to phenylacetonitrile, the Km values for this substrate were higher, especially in the enzyme from A. niger (more than three times; Table 1). The Km values determined with 2-phenylpropionitrile were the lowest, but the Vmax values were 30–175 times lower than with phenylacetonitrile (ibid.). Benzonitrile and cyanopyridines were transformed at approximately 100 times lower rates than phenylacetonitrile by both enzymes (data now shown). The enzymes formed almost no amide from the examined substrates. The temperature optima of both enzymes were approximately 38°C and the pH optima at slightly alkaline values. The nitrilase from A. niger exhibited a broader pH Table 1 Substrate specificity of arylaliphatic nitrilases

Substrate

Phenylacetonitrile (R,S)-Mandelonitrile 2-Phenylpropionitrile

The enzymes from G. moniliformis (Kaplan et al. 2011b) and P. marneffei were both expressed at lower levels compared to the above arylaliphatic nitrilases (Fig. 1). A screening of strains harboring various chaperone plasmids indicated that the activities of strains with the pGro7 plasmid exhibited a nitrilase activity per liter of culture that was six to seven times higher than that of the controls (supplementary Table S4). Other plasmids had no positive effect on the nitrilase activities. A more detailed examination of this effect in larger culture volumes (200 vs. 10 mL; Table 2) gave a similar result in G. moniliformis, total activity increasing by a factor of approximately 6.8 in presence of GroEL/ES. The strain expressing the enzyme from P. marneffei exhibited an approximately 2.9-fold increase in nitrilase activity on co-expression of GroEL/ ES under the same conditions. This effect was mainly due to increased specific activities of the cultures. In G. moniliformis, the specific activities of the culture broth and cell-free extract were a factor of 4.4 and 2.3 times higher than the controls, respectively. There was also a significant increase in the proportion of activity in the soluble fractions. Similar effects were observed in strains producing the nitrilase from P. marneffei; the coexpression of GroEL/ES increased the specific activity of the culture broth and cell-free extract approximately 4.3 and 2.9 times, respectively. The increase in nitrilase solubility was even more significant than in the previous enzyme. In both cases, the nitrilases were purified from the strains with chaperones. Both nitrilases co-purified with GroEL as indicated by SDS-PAGE (Fig. 1). The two-step purification of the enzyme from G. moniliformis resulted in an approximately ninefold increase in the specific activity. Replacement of Sephacryl S-200 by Superdex 200 in the second step did not improve the purity of the enzyme significantly as concluded from SDS-PAGE (data not shown), while yield

Aspergillus niger

Neurospora crassa

Km (mM)

Vmax (U mgprotein−1)

Km (mM)

Vmax (U mg−1

3.4 11.4 0.80

10.6 12.4 0.35

2.0 3.4 1.3

17.5 9.9 0.10

protein)

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Fig. 2 pH (a) and temperature (b) stabilities of purified nitrilases from Neurospora crassa OR74A (filled squares), Aspergillus niger CBS 513.88 (filled diamonds), and Gibberella moniliformis (filled triangles) and cell-free extract of cells producing nitrilase from Penicillium marneffei ATCC 18224 (filled circles) (0.2, 0.2, 0.3 and 1.6 mg protein per milliliter, respectively). a Enzymes were preincubated at pH 4.5–12.0 and room temperature for 2 h. b Enzymes were pre-incubated at 25–60°C and pH 8.0 for 1 h. Specific activites, which were determined under standard conditions prior to preincubation of the enzymes, were taken as 100%

was higher with Sephacryl S-200 (58%) than Superdex 200 (36%). The nitrilase from P. marneffei was purified approximately fourfold with 27.5% yield in one step. The reason for the relatively low yield of the enzyme may have been its instability, which also caused a significant activity loss during storage at −80°C or 4°C. Therefore, effects of temperature and pH on enzyme stability as well as the substrate specificity of this enzyme were examined with cell-free extracts (Fig. 2; Table 3).

The nitrilase from G. moniliformis exhibited the highest Vmax and lowest Km values for benzonitrile, but phenylacetonitrile was transformed with much lower Vmax and higher Km values (Table 3). 3- and 4-Cyanopyridine were good substrates of the enzyme but 2-cyanopyridine was not transformed at all probably due to steric hindrance. The nitrilase from G. moniliformis was highly active over a broader pH range (7–10; data not shown) than the arylaliphatic nitrilases, and a similar observation was made regarding pH stability (Fig. 2a). It also exhibited the highest temperature optimum (approximately 43°C; data not shown) and thermostability (Fig. 2b) of the enzymes examined. In contrast, the temperature optimum of the nitrilase from P. marneffei was the lowest (30–35°C; data not shown) and the enzyme stability decreased more sharply at temperatures over 35°C compared to the other enzymes (Fig. 2b). The enzyme was relatively stable over a pH region of 6.5–8.5 (Fig. 2a). Its substrate specificity was broad (Table 3), comparable Vmax and Km values being determined for benzonitrile and phenylacetonitrile. The enzyme was also active with cyanopyridines including the sterically hindered 2-cyanopyridine but the high Km values of these compounds indicated their inefficient binding by the enzyme. 4-Cyanopyridine was transformed by this enzyme with the highest Vmax of all substrates tested. The enzyme from P. marneffei did not produce any significant amount of amides. The production of amide from benzonitrile and chlorobenzonitriles by nitrilase from G. moniliformis was low, not exceeding 6% in total product, but this enzyme produced 42% and 54% of amide from phenylacetonitrile and 4-cyanopyridine, respectively.

Discussion The role of molecular chaperones consists of assisting newly synthetized polypeptide folding (Sabate et al. 2010). The bacterial chaperone network involves the Hsp70 (DnaK, DnaJ, GrpE) and GroEL/ES systems and TF (Gupta

Table 2 Effect of GroEL/ES chaperone co-expression on nitrilase production and solubility in Escherichia coli strains harboring nit genes from Gibberella moniliformis and Penicillium marneffei ATCC 18224 Gene source

Gibberella moniliformis Penicillium marneffei

Chaperone

– GroEL/ES – GroEL/ES

Total activity [U L−1 culture]

233±13 1584±145 48±3 141±22

Data are means of three independent experiments

Specific activity [U L−1 culture of OD610 =1]

76±7 335±87 7±0.2 30±2

Specific activity [U mg−1 cell-free extract]

0.28±0.12 0.64±0.03 0.024±0.004 0.070±0.014

Percentage (%) of activity in soluble fraction

insoluble fraction

57±3 70±4 31±3 52±8

43±3 30±4 69±3 48±8

Appl Microbiol Biotechnol (2012) 93:1553–1561 Table 3 Substrate specificity of aromatic nitrilases

a

Partially purified enzyme

b

Cell-free extract

c

No activity

Substrate

Benzonitrile 3-Chlorobenzonitrile 4-Chlorobenzonitrile 2-Cyanopyridine 3-Cyanopyridine 4-Cyanopyridine Phenylacetonitrile

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Gibberella moniliformisa

Penicillium marneffeib

Km (mM)

Vmax (U mgprotein−1)

Km (mM)

Vmax (U mgprotein−1)

0.41 0.93 1.3 –c 3.2 2.2 0.75

9.7 6.6 3.9 –c 4.2 7.3 1.2

7.9 11.1 2.6 11.7 34.4 15.4 9.1

0.16 1.9 0.13 0.46 0.21 7.3 0.10

et al. 2010). Though co-expression of these chaperones was often used as a means to facilitate overexpression of eukaryotic proteins in E. coli, the outcome of this method is hard to predict (ibid.). Plasmids containing genes coding for components of the above chaperone network (individually or in combination) were designed (Nishihara et al. 2000) and became commercially available. The importance of chaperones in nitrilase folding was indicated by experiments with nitrilases, where they copurified with some of these molecules. The enzymes from wild-type producing strains belonging to Pseudomonas fluorescens and Bacillus pallidus co-purified with proteins closely related to GroEL (Layh et al. 1998; Almatawah et al. 1999). The recently described nitrilase from A. niger copurified with hsp60, the eukaryotic homologue of GroEL (Kaplan et al. 2006a). When co-expressing GroEL/ES in E. coli with an aliphatic nitrilase from Comamonas testosteroni, the specific activity of the cells increased fivefold and the soluble enzyme fraction from 10% to 58% (Lévy-Schil et al. 1995). The same chaperonin system also exhibited a positive effect on the functional expression in E. coli of another enzyme of the nitrilase superfamily, D-carbamoylase (Chao et al. 2000; Sareen et al. 2001). As suggested in a recent study of ten protein kinases (Haacke et al. 2009), the effect of chaperones may be case-specific. Similar conclusions were made in a study of endostatin, ORP150 protein and human lysozyme (Nishihara et al. 2000). Therefore, we screened for chaperone effects using a large array of chaperone– nitrilase combinations. The chaperone effects were actually different in strains expressing different nitrilases. Significant benefits of GroEL/ES, which binds to unfolded proteins and creates a protected environment for their folding (Lund 2009), were observed in aromatic nitrilases from Gibberella and Penicillium but not in arylacetonitrilases. This effect could be also dependent on the nitrilase expression levels, which were higher in the latter enzymes. Chaperone-assisted overexpression facilitated purification of the enzymes by increasing the specific activity of

the cells and the enzyme ratio in the soluble fractions. The latter effect was also significant in strains expressing arylaliphatic nitrilases from N. crassa and A. niger with a combination of GroEL/ES and TF chaperones. Therefore, the corresponding strains were used for enzyme purification. As a result, four new nitrilases (two aromatic and two arylaliphatic) were purified or partially purified in this study. The nitrilases from G. moniliformis and P. marneffei were not obtained as homogeneous proteins, as they copurified with GroEL as did a number of the wild-type nitrilases. This co-purification could indicate incomplete enzyme folding; a similar phenomenon was observed with protein kinases and the GroEL and DnaJ chaperones (Haacke et al. 2009). Aromatic nitrilases occur frequently in bacteria, primarily rhodococci (for a review, see O'Reilly and Turner 2003). All of the fungal nitrilases purified previously also belonged to this group of enzymes (Harper 1977; Goldlust and Bohak 1989; Vejvoda et al. 2008, 2010). Comparison of the kinetic data of the nitrilase from G. moniliformis for (hetero) aromatic nitriles and phenylacetonitrile suggested this enzyme could be also classified as an aromatic nitrilase. Nevertheless, the Vmax value of this enzyme for benzonitrile was more than 10 times lower than the specific activities of most of the nitrilases from the related taxon Fusarium (Goldlust and Bohak 1989; Vejvoda et al. 2008, 2010). This could be at least partly caused by the co-purification of GroEL, but could be also due to partial misfolding. On the other hand, the recombinant enzyme and the wild-type enzymes from related species exhibited similar substrate specificities and optimum reaction conditions. A nitrilase activity was recently reported in Penicillium multicolor by ourselves (Kaplan et al. 2006b), but the enzyme was not purified. No other nitrilases were described in the Penicillium genus as far as we know. The advantage of the new enzyme from P. marneffei is its relatively wide substrate specificity. Similarly to the nitrilase in P. multicolor it exhibited, however, a lower temperature stability than most other nitrilases.

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The substrate specificity of the enzymes from N. crassa and A. niger indicated that these enzymes were arylacetonitrilases. This category of nitrilases has significant industrial potential in the production of enantiopure carboxylic acids and amides (e.g., Fernandes et al. 2006; Rustler et al. 2008; Banerjee et al. 2009; Sosedov et al. 2009). They have so far been reported in the bacteria of the Alcaligenes (for reviews, see O'Reilly and Turner 2003; Thuku et al. 2009), Pseudomonas (Kiziak et al. 2005), or Halomonas (Chmura et al. 2008) genera. The nitrilase from P. fluorescens is one of the best described arylacetonitrilases (Kiziak et al. 2005, 2007; Kiziak and Stolz 2009). The enzymes from Alcaligenes faecalis or P. fluorescens (GenBank AAW79573) only shared an approximately 40% amino acid sequence similarity with the new arylacetonitrilases from fungi. However, both of these enzymes showed similar substrate specificities as the bacterial arylacetonitrilases. In conclusion, four new nitrilases were purified and characterized in heterologous producers expressing fungal nit genes. Two of the enzymes (from A. niger and N. crassa) were expressed at high levels, yielding 23,000 and 69,000 units per liter of culture, respectively. The high production and straightforward purification of these enzymes enabled tens of milligrams of them to be obtained from 1 L of E. coli cultures. These new arylacetonitrilases can be useful in the production of mandelic acid from mandelonitrile, or other carboxylic acids substituted at the α-position. Further studies should demonstrate the substrate range, enantioselectivity, chemoselectivity, and possible application areas of these enzymes. Acknowledgments The authors wish to thank Hynek Mrazek MSc. for his technical help with nitrilase purification. Financial support via projects P504/11/0394, 305/09/H008 (Czech Science Foundation), IAA500200708 (Grant Agency of the Academy of Sciences of the Czech Republic), LC06010, OC09046 (Ministry of Education of the Czech Republic), COST/ESF CM0701 (STSM fellowships COSTSTSM-CM0701-4765 and −4766 to A. Malandra) and Institutional Research Concept AV0Z50200510 (Institute of Microbiology) is gratefully acknowledged.

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