Cyanide hydratase from Aspergillus niger K10: Overproduction in Escherichia coli, purification, characterization and use in continuous cyanide degradation

Process Biochemistry 49 (2014) 445–450

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Cyanide hydratase from Aspergillus niger K10: Overproduction in Escherichia coli, purification, characterization and use in continuous cyanide degradation Anna Rinágelová a,b , Ondˇrej Kaplan a,1 , Alicja B. Veselá a,c , Martin Chmátal a , Alena Kˇrenková a,c , Ondˇrej Plíhal a,2 , Fabrizia Pasquarelli a,d , Maria Cantarella d , Ludmila Martínková a,∗ a

Institute of Microbiology, Academy of Sciences of the Czech Republic, Vídenská 1083, CZ-142 20 Prague, Czech Republic ˇ Department of Biochemistry and Microbiology, Faculty of Food and Biochemical Technology, Institute of Chemical Technology, Technická 3, CZ-166 28 Prague, Czech Republic c Department of Biochemistry, Charles University in Prague, Hlavova 8, CZ-128 40 Prague, Czech Republic d Department of Industrial and Information Engineering and Economics, University of l’Aquila, via Giovanni Gronchi n.18 - Nucleo Industriale di Pile, I-67100 L’Aquila, Italy b

a r t i c l e

i n f o

Article history: Received 13 September 2013 Received in revised form 12 December 2013 Accepted 12 December 2013 Available online 21 December 2013 Keywords: Cyanide hydratase Nitrilase Aspergillus niger Continuous stirred membrane reactor

a b s t r a c t A cyanide hydratase from Aspergillus niger K10 was expressed in Escherichia coli and purified. Apart from HCN, it transformed some nitriles, preferentially 2-cyanopyridine and fumaronitrile. Vmax and Km for HCN were ca. 6.8 mmol min−1 mg−1 protein and 109 mM, respectively. Vmax for fumaronitrile and 2-cyanopyridine was two to three orders of magnitude lower than for HCN (ca. 18.8 and 10.3 ␮mol min−1 mg−1 , respectively) but Km was also lower (ca. 14.7 and 3.7 mM, respectively). Both cyanide hydratase and nitrilase activities were abolished in truncated enzyme variants missing 18–34 C-terminal aa residues. The enzyme exhibited the highest activity at 45 ◦ C and pH 8–9; it was unstable at over 35 ◦ C and at below pH 5.5. The operational stability of the whole-cell catalyst was examined in continuous stirred membrane reactors with 70-mL working volume. The catalyst exhibited a half-life of 5.6 h at 28 ◦ C. A reactor loaded with an excess of the catalyst was used to degrade 25 mM KCN. A conversion rate of over 80% was maintained for 3 days. © 2013 Elsevier Ltd. All rights reserved.

1. Introduction Nitrilases (EC 3.5.5.1) and cyanide hydratases (EC 4.2.1.66; CHTs) belong to class one of the nitrilase superfamily, which consists of enzymes acting on non-peptide C–N bonds [1]. The two

Abbreviations: aa, amino acid; CHT, cyanide hydratase; CSMR, continuous stirred membrane reactor; CynDpum , cyanide dihydratase from Bacillus pumilus; CynDstu , cyanide dihydratase from Pseudomonas stutzeri; NitAn1, cyanide hydratase/nitrilase from Aspergillus niger K10; NitAn1-C14, NitAn1-C18, NitAn1-C22, NitAn1-C26, NitAn1-C30 and NitAn1-C34, variants of NitAn1 missing 14, 18, 22, 26, 30 and 34 C-terminal aa residues, respectively; Nit(p), nitrilase from Pseudomonas fluorescens EBC 191; Nit(r), nitrilase from Rhodococcus rhodochrous J1; MWCO, molecular weight cut-off. ∗ Corresponding author. Tel.: +420 296 442 569. E-mail addresses: [email protected] (A. Rinágelová), [email protected] (L. Martínková). 1 Current address: Institute of Macromolecular Chemistry, Academy of Sciences of the Czech Republic, Heyrovského námˇestí 2, CZ-162 06 Prague, Czech Republic. 2 Current address: Centre of the Region Haná for Biotechnological and Agricultural ˇ Research, Slechtitel u˚ 813/21, CZ-783 71 Olomouc – Holice, Czech Republic. 1359-5113/$ – see front matter © 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.procbio.2013.12.008

enzymes differ in their substrates (nitriles vs. HCN, respectively) and in their major reaction products (carboxylic acids vs. amide, respectively). All characterized CHTs originate from filamentous fungi, and all the genes of putative CHTs were also found in fungal genomes. CHTs are likely to play a role in the detoxification of HCN released from cyanogenic glycosides, which occur in many plant species. CHTs could have evolved from nitrilases whose genes were acquired by fungi via horizontal gene transfer from bacteria [2]. CHTs were first reported in phytopathogenic fungi (Stemphylium loti, Leptosphaeria maculans, Gloeocercospora sorghi, genus Fusarium; for a review, see [3]). Later, CHTs from saprophytic fungi Neurospora crassa and Aspergillus nidulans and from the phytopathogenic fungi Gibberella zeae and G. sorghi were expressed in Escherichia coli and partially characterized [4]. The genus Aspergillus is rich in genes coding for putative nitrilases or CHTs, but only a few of them have been characterized. Apart from the CHT from A. nidulans, a CHT from Aspergillus niger K10 (NitAn1) was expressed in E. coli and its activities were examined in whole cells [5]. In addition, an aromatic nitrilase with a preference for 4-cyanopyridine and benzonitrile

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was partially purified from a mycelium of A. niger K10 grown on 2-cyanopyridine [6]. This enzyme was later analyzed by mass spectrometry and found to be closely related to a hypothetical nitrilase whose gene was detected in the genome sequence of Aspergillus kawachii IFO 4308 [7]. In this work, NitAn1 was purified and characterized. Previously, the enzyme was expressed in E. coli cells [8] and found to transform not only HCN, but also selected nitriles [5]. Here the kinetics of the purified enzyme were examined with HCN and its best nitrile substrates, 2-cyanopyridine and fumaronitrile. Variants of NitAn1 truncated at the C-terminus were prepared in order to assess the effect of this region on their CHT and nitrilase activities. The operational stability of the enzyme was determined using continuous stirred membrane reactors (CSMRs), which were previously used in the transformation of nitriles via the monoenzymatic (nitrilase) or bienzymatic (nitrile hydratase–amidase) pathway [9,10]. 2. Materials and methods 2.1. Database search and sequence alignment Database searches were performed using the BLASTP programme (http://blast.ncbi.nlm.nih.gov/Blast.cgi). The sequence alignment was performed using the COBALT: Constraint-based Multiple Alignment Tool (http://www.ncbi.nlm.nih.gov/tools/ cobalt/cobalt.cgi). 2.2. Nitrilase expression and purification NitAn1 was expressed without a His6 -tag in E. coli as described previously [8,11]. Briefly, the NdeI-HindIII fragment of the gene amplified from the cDNA of A. niger K10 was ligated into the corresponding sites of pET-30a(+) (Novagen) to give the pOK101 vector, which was then transformed into E. coli BL21-Gold(DE3). The cells were grown at 37 ◦ C in Luria-Bertani medium supplemented with 50 ␮g mL−1 kanamycin until OD610 reached 0.4–0.6 and gene expression was then induced by adding 0.8 mM IPTG. The cells were harvested after a further 16-h cultivation at 26 ◦ C. Genes encoding C-terminally truncated variants missing 14, 18, 22, 26, 30 or 34 aa residues (designated NitAn1-C14, NitAn1-C18, NitAn1-C22, NitAn1-C26, NitAn1-C30 and NitAn1-C34, respectively) were prepared synthetically by Generay Biotech Co., Ltd., Shanghai, China, and expressed in the same way as the wild-type enzyme. After disrupting the cells by sonication and removing cell debris by centrifugation (13,000 × g, 4 ◦ C, 30 min), the supernatant was injected into a Hi-Prep 16/10 Q FF column (Amersham Biosciences) and the proteins eluted with a linear gradient of NaCl (0.15–1 M) in Tris/HCl buffer (50 mM, pH 8.0). Active fractions were pooled, concentrated using an Amicon Ultra-4 unit (MWCO 10 kDa) and injected into a Hi-Prep 16/60 Sephacryl S-200 column. Proteins were eluted with Tris/HCl buffer (50 mM, pH 8.0, 150 mM NaCl). Active fractions were pooled, concentrated as described above and stored at −80 ◦ C.

Fig. 1. Expression in E. coli and purification of NitAn1. Molecular weight markers (lane 1), crude extract (lane 2), pooled fractions after purification on Q-Sepharose (lane 3), pooled fractions after purification on Superdex 200 (lane 4).

The pH optimum of the purified enzyme was determined in 33 mM acetic acid/boric acid/phosphoric acid/NaOH buffer, pH 4.0–11.0, with 25 mM 2-cyanopyridine instead of HCN as the substrate, since HCN would be largely lost at a low pH. The temperature optima were determined with the same substrate in Tris/HCl buffer (50 mM, pH 8.0, 150 mM NaCl) at 25–55 ◦ C. The pH stability was determined by measuring the residual activity for 2-cyanopyridine after pre-incubation of the enzyme in 33 mM acetic acid/boric acid/phosphoric acid/NaOH buffer, pH 4.0–11.0, at 30 ◦ C for 2 h. The temperature stabilities were determined in the same way after preincubation of the enzyme in Tris/HCl buffer (50 mM, pH 8.0, 150 mM NaCl) at 25–55 ◦ C for 1 h. The substrate specificity of the purified enzyme was determined with 25 mM KCN, fumaronitrile, benzonitrile, 2-, 3- or 4-cyanopyridine or phenylacetonitrile in Tris/HCl buffer (50 mM, pH 8.0, 150 mM NaCl) at 30 ◦ C. The concentrations of nitriles and their reaction products were determined by HPLC [5]. The kinetics of the purified enzyme were determined with 2–40 mM KCN or 0.5–25 mM fumaronitrile or 2-cyanopyridine under the same conditions. 2.4. Biotransformations in CSMRs An Amicon 8050 ultrafiltration cell (Millipore, USA) with a cellulose ultrafiltration membrane (MWCO 10 kDa) was used as a CSMR with a working volume of 70 mL. The reactor was loaded with an appropriate amount of E. coli cells expressing NitAn1 and operated at a flow rate of 12 mL h−1 , 28 ◦ C and under constant stirring (250 rpm). The stock solution of 25 mM KCN in 50 mM Tris/HCl buffer was kept under an argon atmosphere. The deactivation constant, enzyme half-life and the initial reaction rate were calculated as described previously [10].

2.3. Cyanide hydratase and nitrilase assays 3. Results and discussion The CHT activity was assayed in 1.5-mL Eppendorf tubes with 0.5 mL of the reaction mixtures consisting of 25 mM KCN, Tris/HCl buffer (50 mM, pH 8.0, 150 mM NaCl) and an appropriate amount of the purified enzyme or E. coli whole cells. The reactions proceeded with shaking (Thermomixer Eppendorf Compact, 850 rpm) at 30 ◦ C and were stopped within their linear range (after 5 min) by the addition of methanol (final concentration 50%, v/v). The production of formamide was determined by HPLC analyses as described previously [5].

3.1. Substrate specificity of NitAn1 NitAn1 was purified to near homogeneity in two steps. The enzyme formed a major part of the total cellular proteins, and therefore the specific activity increased only 1.8-fold after purification, which proceeded with a yield of ca. 17%. SDS-PAGE analysis indicated the molecular weight of the purified protein to be ca. 40 kDa as expected (Fig. 1).

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Table 1 Substrate specificity of NitAn1. Substrate

Relative activity (%)

Amide (mol % of total product)

Vmax (␮mol min−1 mg−1 )

Km (mM)

HCN Benzonitrile 2-Cyanopyridine 3-Cyanopyridine 4-Cyanopyridine Fumaronitrile Phenylacetonitrile

100 0.062 ± 0.002 0.76 ± 0.05 0.055 ± 0.003 <0.01 0.95 ± 0.18 <0.01

100 n.d. 19.5 ± 0.5 2.22 ± 0.03 n.a. n.d. n.a.

6.8 ± 0.3 × 103 n.a. 10.3 ± 1.3 n.a. n.a. 18.8 ± 2.1 n.a.

109 ± 5 n.a. 3.7 ± 0.5 n.a. n.a. 14.7 ± 2.6 n.a.

n.a.: not assayed; n.d.: not detected. Relative activities were determined with 25 mM substrates at pH 8.0 and 30 ◦ C. The specific activity for HCN (1324 ± 48 U mg−1 ) was taken as 100%.

The relative activities of the purified enzyme were determined for HCN and selected nitriles (25 mM each; Table 1). The enzyme exhibited a high specific activity of ca. 1.3 mmol min−1 mg−1 of protein for HCN. Fumaronitrile and 2-cyanopyridine were the best of the nitrile substrates examined, being transformed at ca. 13 and 10 ␮mol min−1 mg−1 of protein, respectively. 3-Cyanopyridine and benzonitrile were transformed at less than a tenth of the rates with fumaronitrile or 2-cyanopyridine. Activities for nitrile compounds were previously found in two CHTs (Table 2). The purified CHT from Fusarium oxysporum N-10 preferentially hydrolyzed acrylonitrile, crotononitrile and methacrylonitrile [12] and E. coli cells expressing the CHT from Fusarium lateritium hydrolyzed acetonitrile, propionitrile and benzonitrile [13]. The reaction rates determined for these nitriles were less than 0.05% of the rates with HCN in both enzymes. Thus the best nitrile substrates of purified NitAn1 were transformed at higher relative rates (ca. 0.8–1% of that with HCN). Vmax for HCN was 6.8 ± 0.3 mmol min−1 mg−1 of protein, but Km was also high (109 ± 5 mM). The Vmax values for fumaronitrile and 2-cyanopyridine were two to three orders of magnitude lower than for HCN, and the Km values were also lower (ca. 7 and 30-fold, respectively). Kinetic data for HCN were comparable with those of the CHTs from G. sorghi (Vmax 4.4 ± 1.5 mmol min−1 mg−1 of protein; Km 90 ± 35) [14], F. oxysporum (Km 71 mM) [12] and F. lateritium (Km 43 mM) [15]. These data demonstrated that the CHT assays were performed at HCN concentrations far below saturating substrate conditions in this and other previous studies (e.g. [4,12]). Substrate saturation was only achieved at very high HCN concentrations, which, however, are known to inactivate CHT rapidly (at over 150 mM HCN; [15]). To the best of our knowledge, no kinetic data have been determined for substrates other than HCN with any of the CHTs studied previously. 3.2. Acid vs. amide production The only product of HCN transformation by NitAn1 was formamide. However, this enzyme mainly produced the corresponding carboxylic acids from the nitriles examined. A significant amount of amide (picolinamide) was only formed from 2cyanopyridine (almost one fifth of the total reaction product). No product of KCN or nitrile conversion was detected if the catalyst (whole cells or purified enzyme) was omitted from the reaction mixtures. Likewise, the catalyst formed no detectable product in the absence of the substrate. In contrast to NitAn1, the CHT from F. oxysporum transformed its nitrile substrates into amides according to GC analysis [12]. The formation of amides by nitrilases is affected by some electronic and steric properties of the substrates, leading to destabilization of the tetrahedral intermediate and its cleavage into amide and the free enzyme instead of acylenzyme and ammonia [16]. 2-Cyanopyridine (a sterically demanding substrate) seems to be prone to conversion into amide. Over 80% amide was produced from this substrate by another nitrilase from A. niger [6].

The composition of the product (acid vs. amide) is also dependent on the nitrilase itself, amide formation being typical for the enzymes from Pseudomonas fluorescens [17], Arabidopsis thaliana [18] or A. niger [6]. The major product of 2-cyanopyridine hydrolysis, picolinic acid, is an intermediate in the production of pharmaceuticals such as local anaesthetics. Nitrilases with acceptable activities for this substrate are rarely reported (for a review, see [3]). NitAn1 transformed this substrate at a satisfactory rate but its tendency to produce picolinamide was a drawback. It is possible that a screening of further CHTs could provide an enzyme which would produce picolinic acid in a higher purity. 3.3. Effect of C-terminal truncation on the CHT and nitrilase activities Some point mutations in the CHT from F. lateritium were previously found to abolish both CHT and nitrilase activities [13]. Here we examined the effect of the C-terminal region of NitAn1 on both activities. This region was proposed to participate in forming the “A surface” as one of the interfaces between the enzyme subunits [19]. The effect of deletions of this region was previously examined in cyanide dihydratases [19] and nitrilases [20,21], but not in CHTs. NitAn1 variants, in which 14–34 aa residues from the Cterminus were deleted, were examined for nitrilase activities with 2-cyanopyridine and for CHT activities. As determined by SDSPAGE, the expression levels of the variants and the wild-type NitAn1 were all similar, and the molecular weights of all the proteins were as expected (Fig. S1). C-Terminal truncation had similar effects on their CHT and nitrilase activities, which were examined using whole E. coli cells expressing the wild-type enzyme or its variants (Table 3). NitAn1-C14 exhibited CHT and nitrilase activities similar to the wild-type enzyme and also formed similar amounts of picolinamide. NitAn1-C18 exhibited a very low CHT and nitrilase activity and no activities were detected in the shorter variants. The ratio of the CHT and nitrilase activity seemed to be changed in NitAn1-C18 (in favour of the nitrilase activity) but this result may have been due to very low concentrations of the products, which were thus difficult to quantify. Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.procbio.2013.12.008. Most of the nitrilases and cyanide dihydratases from which deletion mutants were prepared were less sensitive to C-terminal deletions than NitAn1. In the nitrilase from P. fluorescens (Nit(p)), variants missing 20–75 aa residues were prepared, all but the shortest one having nitrilase activities. The activities of variants missing 47–67 aa residues were, however, significantly decreased (to 5–13%) [20]. Of the truncated variants of the nitrilase from Rhodococcus rhodochrous (Nit(r)), those missing up to 49 aa residues were active, in contrast to the variants missing 55 or 64 aa residues [21]. Comparison with the cyanide dihydratase from Pseudomonas stutzeri (CynDstu ) was not possible, as none of the tested variants of this enzyme exhibited any such activity [19]. However, the cyanide

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Table 2 Comparison of biochemical properties of cyanide hydratases produced by various fungal strains. Organism

Nitrilase activity (substrate, percentage of CHT activity)

a

Aspergillus niger Aspergillus nidulansa Gibberella zeaea Gloeocercospora sorghia Neurospora crassaa Fusarium lateritiumb Fusarium oxysporumc

Fumaronitrile, 2-cyanopyridine (ca. 0.8–1%) n.a. n.a. n.a. n.a. Benzo-, aceto-, propionitrile (ca. 0.01–0.03%) Methacrylo-, crotono-, acrylonitrile (ca. 0.03–0.04%)

Effect of temperature

Effect of pH

Reference

Optimum

Stability

Optimum

Stability

45 n.a. n.a. n.a. n.a. n.a.

≤35 ≤37 ≤43 ≤37 ≤43 n.a.

8–9 5–6.5 6–8 7 6–7 n.a.

5.5–10 n.a. n.a. n.a. n.a. n.a.

This work [4] [4] [4] [4] [13]

30

≤50

7.5

7–9

[12]

n.a.: not assayed. a Enzyme purified from E. coli. b Whole E. coli cells. c Enzyme purified from the wild-type strain.

Table 3 Conversion of KCN and 2-cyanopyridine by E. coli whole cells expressing NitAn1 and its variants with C-terminal deletions. Variant

Chain length (aa residues)

Aa residues deleted

NitAn1 (wild-type) NitAn1-C14 NitAn1-C18 NitAn1-C22, NitAn1-C26, NitAn1-C30, NitAn1-C34

356 342 338 322–334b

– 14 18 22–34b

Specific activity (␮mol min−1 mg−1 dry cell weight) ± SDa KCN

2-Cyanopyridine

385 ± 48 367 ± 14 0.31 ± 0.05 n.d.

3.47 ± 0.06 3.39 ± 0.03 0.079 ± 0.002 n.d.

n.d.: not detected. a At 30 ◦ C and pH 8.0. b In 4 aa residue increments.

dihydratase in Bacillus pumilus (CynDpum ) withstood more extensive deletions (up to 38 aa residues) than CynDstu before losing its activity [19]. Thus, compared to Nit(p), Nit(r) and CynDpum , NitAn1 activity was dramatically affected by deletions of less (18) aa residues. Except for Nit(p) variants, in all other enzymes including NitAn1, the deletions were positioned outside the conserved motifs (towards the C-terminus), as indicated by the sequence alignment (Fig. 2).

3.4. Temperature and pH optima and stabilities The temperature optimum of NitAn1 was 45 ◦ C. The enzyme was still active at 50 ◦ C with ca. 36% of its maximum activity, but nearly inactive at 55 ◦ C. It was fairly stable at temperatures up to ca. 35 ◦ C. Its operational range was between pH 5.5 and 10.5, and it was also acceptably stable between pH 5.5 and 10.0 (Fig. S2). Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.procbio.2013.12.008.

Although CHTs exhibit relatively high aa sequence identities (mostly over 60%), enzymes with differing catalytic properties were found among them [4]. Of these CHTs, the enzyme from A. nidulans was the most similar to NitAn1 with ca. 84% aa sequence identity, while the enzymes from N. crassa, G. zeae and G. sorghi exhibited lower aa sequence identities (around 60%) to NitAn1. With some caution – due to the use of different assays based either on determination of KCN [4] or formamide (this work) – we compared the properties of NitAn1 with those of the previously characterized CHTs (Table 2). The pH profiles of NitAn1 and the previously characterized CHTs were different. The activity range of the CHT from A. nidulans was shifted towards lower values (4.5–10) compared to NitAn1 (pH 5.5–10.5). The CHTs from G. sorghi and G. zeae only exhibited considerable activities between pH 6 and 10, while the CHT from N. crassa was active over the broadest pH range (4.5–10.5). pH optima were between 8 and 9 in NitAn1 but lower in the previously characterized CHTs (Table 2). NitAn1 and the CHT from A. nidulans were more sensitive to elevated temperatures than the other CHTs, being quickly deactived at temperatures over

ABX75546 AAW79573 Q03217 AAN77003 BAA11653

279 273 274 262 261

GLLFVDIDLDECHLSKSLADFGGHYMRPDLIRLLVDTNRKDLV GILYADIDLGVIGVAKAAYDPVGHYSRPDVLRLLVNREPMTRV GILYADIDLSAITLAKQAADPVGHYSRPDVLSLNFNQRHTTPV GIAYADIDVERVIDYKYYIDPAGHYSNQS-LSMNFNQQPTPVV GIAYAEIDIEKIIDFKYYIDPVGHYSNQS-LSMNFNQSPNPVV

321 315 316 303 302

ABX75546 AAW79573 Q03217 AAN77003 BAA11653

322 316 317 304 303

V---------REDRVNGGVEYTRTVDRVGLSTPLD[5]DSEN HYVQPQSLPETSVLAFGAGA-DAIRSEENPEEQ-GDK N-TAISTIHATHTLVPQSGALDGVRELNGADEQRA[5]SDE[8] K-----QLNDNKNEVL---TYEAIQYQNGMLEEKV R-----KIGERDSTVF---TYDDLNLSVSDEEPVV ---[5]

356 350 366 330 334

Fig. 2. Multiple alignment of C-terminal sequences of NitAn1 (ABX75546), nitrilases from Rhodococcus rhodochrous J1 (Q03217) [21] and Pseudomonas fluorescens EBC191 (AAW79573) [20] and cyanide dihydratases from Bacillus pumilus (AAN77003) and Pseudomonas stutzeri (BAA11653) [19]. C-Terminal aa residues are highlighted with a grey background for inactive mutants, enclosed in rectangles in partially active mutants and underlined in fully active mutants.

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15–20% of the cyanide was lost due to its instability. These losses occurred either in the reactor or at the outflow, as the concentration of KCN in the stock solution was stable. CHTs were found to be promising for the elimination of KCN from wastewater due to their high specific activity and to their resistance to moderately alkaline conditions and metal ions [4]. The use of NitAn1 in continuous KCN degradation supported this opinion. CSMRs seem to be suitable for optimizing the performance of CHTs in model and real wastewater samples and for the design of the corresponding processes at laboratory scale.

100 80

Conversion (%)

A

449

60 40 20 0 0

10

20

30

40

50

60

70

4. Conclusion

Time (h) (U mg dry cell weight)

100,00 y = 68,986e

10,00

-0,1236x

1,00

-1

Reacon rate

B

0,10 0,01 0

10

20

30

40

Time (h) Fig. 3. Degradation of 25 mM KCN by NitAn1 in CSMRs with a 70-mL working volume. E. coli whole cells expressing NitAn1 were loaded into the reactor at 0.2 mg (triangles) or 9 mg of dry weight (diamonds). (A) Conversion (%); (B) instantaneous reaction rate (not calculated for the higher load). Flow rate 12 mL h−1 , temperature 28 ◦ C, stirring (250 rpm). The conversion and reaction rates were calculated from the concentrations of formamide in the outflow (see Section 2.4. for details).

35–37 ◦ C. In contrast, the enzymes from N. crassa and G. zeae were acceptably stable at up to 43 ◦ C [4]. The enzyme from F. oxysporum was stable at up to 50 ◦ C despite its lower temperature optimum (30 ◦ C) [12]. 3.5. Continuous KCN conversion Whole cells expressing NitAn1 were examined for their operational stabilities in CSMRs. To this end, the reactor was loaded with a low amount of E. coli cells (0.2 mg of dry cell weight) expressing NitAn1 and the continuous reaction was carried out for over 40 h at 28 ◦ C. The conversion achieved more than 80% after 2 h but then began to decline (Fig. 3A). For the linear part of the semilogarithmic plot of the reaction rate vs. time (Fig. 3B), the operational deactivation constant, kd , was calculated to be 0.124 h−1 , corresponding to an enzyme half-life of 5.6 h, and the initial reaction rate was calculated to be 69 ␮mol min−1 mg−1 dry cell weight, i.e. ca. 138 ␮mol min−1 mg−1 protein. This was much less than in the batch experiments with whole cells (385 ␮mol min−1 mg−1 dry cell weight; Table 3) or with the purified enzyme (ca. 1.3 mmol min−1 mg−1 protein). This difference was probably mainly due to lower operational substrate concentrations in the reactor but also to different reaction conditions (temperature, mixing). Using the whole-cell catalyst in excess (9 mg of dry cell weight) made it possible to achieve a high and stable level of substrate conversion, which was maintained at 82.7 ± 2.3% for 3 days, as calculated from the concentrations of formamide in the outflow (Fig. 3A). The apparent reaction rate in this reactor was only ca. 0.47 ␮mol min−1 mg−1 dry cell weight since, due to the high enzyme excess, only a small fraction of the enzyme could act as the catalyst. Residual KCN was hardly detectable, indicating that ca.

NitAn1 is the first CHT to be purified and characterized from an A. niger strain. The enzyme has both CHT and nitrilase activity, but the latter is much lower. CHT kinetics for HCN and nitriles were compared for the first time. Nitrilase activities, although lower than in this work, were also previously found in two other CHTs. It is possible that minor nitrilase activities are common in CHTs but have largely gone unnoticed. The effect of C-terminal deletions on CHT activity was determined for the first time. Both CHT and nitrilase activities in NitAn1 were affected by these mutations in the same way. In this and other studies, the C-terminal region was shown to have a high impact on the catalytic properties of nitrilases, cyanide dihydratases and CHT, but the tolerance to the deletions in this region was different in different enzymes. The high catalytic efficiency and acceptable stability makes NitAn1 promising for KCN elimination from wastewater. Its resistance to coke plant wastewater is currently being examined by ourselves. Continuous reactors, which proved to be suitable for the examination of CHT stability, will also be useful for this purpose. NitAn1 also has a good activity for 2-cyanopyridine (a precursor of picolinic acid – a pharmaceutical intermediate) which is an unusual substrate for nitrilases. Acknowledgements Czech Science Foundation (P504/11/0394), Technology Agency of the Czech Republic (TA01021368), Institute of Microbiology of the Academy of Sciences of the Czech Republic, v.v.i. (RVO61388971). References [1] Brenner C. Catalysis in the nitrilase superfamily. Curr Opin Struct Biol 2002;12:775–82. [2] Podar M, Eads JR, Richardson TH. Evolution of a microbial nitrilase gene family: a comparative and environmental genomics study. BMC Evol Biol 2005;5: 42. [3] O’Reilly C, Turner PD. The nitrilase family of CN hydrolysing enzymes—a comparative study. J Appl Microbiol 2003;95:1161–74. [4] Basile LJ, Willson RC, Sewell BT, Benedik MJ. Genome mining of cyanide degrading nitrilases from filamentous fungi. Appl Microbiol Biotechnol 2008;80:427–35. [5] Kaplan O, Veselá AB, Petˇríˇcková A, Pasquarelli F, Piˇcmanová M, Rinágelová A, et al. A comparative study of nitrilases identified by genome minig. Mol Biotechnol 2013;54:996–1003. [6] Kaplan O, Vejvoda V, Plíhal O, Pompach P, Kavan D, Bojarová P, et al. Purification and characterization of a nitrilase from Aspergillus niger K10. Appl Microbiol Biotechnol 2006;73:567–75. [7] Kaplan O, Vejvoda V, Plíhal O, Pompach P, Kavan D, Bojarová P, et al. Erratum to: Purification and characterization of a nitrilase from Aspergillus niger K10 (vol 73, pg 567, 2006). Appl Microbiol Biotechnol 2013;97:3745–6. [8] Petˇríˇcková A, Veselá AB, Kaplan O, Kubáˇc D, Uhnáková B, Malandra A, et al. Purification and characterization of heterologously expressed nitrilases from filamentous fungi. Appl Microbiol Biotechnol 2012;93:1553–61. [9] Cantarella M, Cantarella L, Gallifuoco A, Spera A, Martínková L. Nicotinic acid bio-production by Microbacterium imperiale CBS 489-74: effect of 3-cyanopyridine and temperature on amidase activity. Proc Biochem 2012;47:1192–6.

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