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Review

Recent Progress in the Production of Cyanide-Converting Nitrilases—Comparison with Nitrile-Hydrolyzing Enzymes

by
Ludmila Martínková
1,*,
Natalia Kulik
1,
Anastasia Sedova
2,
Barbora Křístková
1,3 and
Pavla Bojarová
1,2
1
Institute of Microbiology of the Czech Academy of Sciences, CZ-142 20 Prague, Czech Republic
2
Department of Health Care and Population Protection, Faculty of Biomedical Engineering, Czech Technical University in Prague, CZ-272 01 Kladno, Czech Republic
3
Faculty of Food and Biochemical Technology, University of Chemistry and Technology, Prague, Technická 5, CZ-166 28 Prague, Czech Republic
*
Author to whom correspondence should be addressed.
Catalysts 2023, 13(3), 500; https://doi.org/10.3390/catal13030500
Submission received: 20 December 2022 / Revised: 3 February 2023 / Accepted: 20 February 2023 / Published: 28 February 2023
(This article belongs to the Section Biocatalysis)
Browse Figures
Versions Notes

Abstract

:
Nitrilases have a high potential for application in organic chemistry, environmental technology, and analytics. However, their industrial uses require that they are produced in highly active and robust forms at a reasonable cost. Some organic syntheses catalyzed by nitrilases have already reached a high level of technological readiness. This has been enabled by the large-scale production of recombinant catalysts. Despite some promising small-scale methods being proposed, the production of cyanide-converting nitrilases (cyanide hydratase and cyanide dihydratase) is lagging in this regard. This review focuses on the prospects of cyanide(di)hydratase-based catalysts. The current knowledge of these enzymes is summarized and discussed in terms of the origin and distribution of their sequences, gene expression, structure, assays, purification, immobilization, and uses. Progresses in the production of other nitrilase catalysts are also tackled, as it may inspire the development of the preparation processes of cyanide(di)hydratases.

Graphical Abstract

1. Introduction

Nitrilases (EC 3.5.5.-) represent a branch of the nitrilase superfamily [1]. They are also probably the only branch acting on cyanides or nitriles. The known members of the other branches are generally amidases.
The substrates of nitrilases are organic cyanides (nitriles) and/or HCN, which is a form of free cyanide (fCN). The HCN/CN− ratio in fCN depends on pH. Two types of nitrilases prefer HCN as the substrate, the cyanide dihydratase (CynD) and cyanide hydratase (CynH). The latter is formerly classified as lyase (EC 4.2.1.66). Nevertheless, the classification of CynD and CynH as distinctive types of nitrilases is clearly justified by comparing their primary structures with those of nitrile-hydrolyzing enzymes (Table S1). In addition, the regions proximal to the key residues (Glu-Lys-Glu-Cys) are even more conserved, resulting in similar 3D structures (see below).
The two enzymes differ in their products, that is, formate and ammonia for CynD and formamide for CynH. Similarly, nitrilases transform nitriles into ammonia and carboxylic acid or amide or their mixture (Figure 1).
Some nitriles, typically 2-cyanopyridine (1a) and fumaronitrile (2a), are also transformed by CynHs (Figure 2).
While enzymatic hydrolysis of nitriles has been known since the 1950s, the first nitrilases that efficiently degrade HCN (CynHs) were reported about two decades later, followed by CynDs in the 1990s (see [1,2,3] for previous reviews).
The physiological role of nitrilases, including CynDs and CynHs, is not fully understood. It is generally supposed that the main functions of nitrilases are nitrogen recycling and cyanide detoxification [4]. Specifically, one of the roles of CynHs in fungi is likely to be the detoxification of HCN released from cyanogenic glycosides that serve as plant defense compounds. It is plausible that the CynH product formamide can be hydrolyzed to provide ammonia. CynDs may serve similar purposes. The bacterial genera that produce CynDs are soil and plant-associated bacteria, which is consistent with this speculation.
Nitrilases can replace conventional tools of nitrile hydrolysis, such as strong acids or bases. They usually operate at pH 7–8 and 30–40 °C. They became attractive, especially because of their stereoselectivities and regioselectivities. A few nitrilase-catalyzed processes to produce fine chemicals were brought to a high degree of technological readiness (for relevant reviews, see [5,6,7,8,9]). For example, enantioselective hydrolysis of racemic mandelonitriles 3a and 4a provided (R)-mandelic acid (3b) [10] and its o-chloro derivative 4b [11]. A regioselective hydrolysis of 1-cyanocyclohexaneacetonitrile (5a) provided 1-cyanocyclohexaneacetic acid (CCHA; 5b) (Figure 2), the precursor of gabapentin (3c) [12], a widely used anticonvulsant medicine. In addition, many studies aimed at improving nitrilase catalysts by site-directed mutagenesis or immobilization (for relevant reviews, see e.g., [5,6,13,14]).
Catalysts 13 00500 g002 550
Figure 2. Examples of nitrile conversions by (a) cyanide hydratases (CynHs) and (b) nitrilases (Nits). CynHs produce carboxylic acids and/or amides from 2-cyanopyridine (1a) and fumaronitrile (2a). The acid (1b,2b)/amide (1c,2c) ratio depends on enzyme origin [15]. Some bacterial Nits produce enantiopure R-mandelic acids 3b and 4b from (R,S)-mandelonitriles 3a and 4a [10,11], or 1-cyanocyclohexaneacetic acid 5b, the precursor of gabapentin (5c), from the corresponding dinitrile 5a [12].
Figure 2. Examples of nitrile conversions by (a) cyanide hydratases (CynHs) and (b) nitrilases (Nits). CynHs produce carboxylic acids and/or amides from 2-cyanopyridine (1a) and fumaronitrile (2a). The acid (1b,2b)/amide (1c,2c) ratio depends on enzyme origin [15]. Some bacterial Nits produce enantiopure R-mandelic acids 3b and 4b from (R,S)-mandelonitriles 3a and 4a [10,11], or 1-cyanocyclohexaneacetic acid 5b, the precursor of gabapentin (5c), from the corresponding dinitrile 5a [12].
Catalysts 13 00500 g002
In contrast, the production and application of CynHs or CynDs are less developed in terms of scale and catalyst improvement. Nevertheless, the potential of enzymes is promising for cyanide sensors and cyanide degradation in wastewaters from gold mining, metal plating, coking, chemical, and other industries. Previous comprehensive reviews on this topic summarized the literature up to about 2015 [2] and 2016 [3], with the latter focusing on the structure-activity relationships. Some general reviews on the removal of cyanide from wastewater also addressed the HCN-degrading enzymes [16,17]. Here, we summarize the main results obtained so far in this area, focusing mainly on the recent literature. We also address the major advances in the study of other nitrilases (nitrile-transforming enzymes) to the extent that they may provide new approaches for the development of HCN-degrading enzymes.

2. Origin and Distribution of the Sequences

CynDs are a relatively small group of enzymes. The two best-characterized ones are from Bacillus pumilus (CynDpum) and Pseudomonas stutzeri (CynDstut). These enzymes are about 80% identical (with a coverage of 95%) and differ mainly in their C-termini [18,19]. CynDpum and CynDstut seem strictly selective for HCN.
In addition, CynDs were recently reported in Flavobacterium indicum [20], Bacillus safensis [21], and Bacillus sp. [22]. The enzyme in F. indicum (here called CynDind) is different from the above-mentioned CynDs (its identity with CynDpum is only about 34% with a coverage of 92%). Moreover, it has a different sequence in the region containing the catalytically active cysteine residue, i.e., the ACGEN instead of the typical sequences of the well-known CynDs (MCWEH in CynDpum and CynDstut). The number of non-redundant homologous sequences with 70–99% identity and ≥90% coverage to CynDind is 151 in Flavobacteria (Table 1).
The GenBank database (https://blast.ncbi.nlm.nih.gov; accessed on 5 December 2022) contains 230 CynD sequences with ≥70% identity and ≥90% coverage to the templates (Table 1), CynDpum, CynDstut (P. stutzeri was reclassified as Stutzerimonas stutzeri), or CynDind (233 including the templates). The number of non-redundant sequences with 70–99% identity to CynDpum or CynDstut is 79 (81 including the templates). We assume that CynD activity is likely in these proteins. However, it is difficult to speculate on those that have lower levels of identity (approximately 70 sequences more if the identity threshold is set at 50%). CynD sequences are frequently found in the genus Bacillus (52), and here mainly in the species Bacillus pumilus (15) and Bacillus safensis (17). All Flavobacterium homologs were joined in one clade with a significant phylogenic distance from other CynDs (see Figure S1 for the phylogenetic tree).
All putative CynDs found were of bacterial origin with one exception, which is the hypothetical CynD of the yeast Sheffersomyces stipitis CBS 6054. The closest homologs of this protein are bacterial proteins annotated as carbon-nitrogen hydrolase family proteins in Clostridium ljungdahlii (GenBank: WP_063554590.1) with ca. 76% identity and 96% coverage, and Bacillus australimaris (GenBank: MBR8688472.1) with ca. 74% identity and 97% coverage. The closest fungal homologs are hypothetical proteins from Aureobasidium melanogenum (GenBank: KAH0440800.1) with ca. 41% identity and 88% coverage and a putative nitrilase from Hyaloraphidium curvatum (GenBank: KAI9027987.1) with ca. 36% identity and 93% coverage.
The group of CynHs is larger (see Figure S2 for the phylogenetic tree) and more sequences were confirmed to have the expected activity. The recombinantly produced CynHs originated from Fusarium lateritium [26], Aspergillus nidulans, Neurospora crassa, Gibberella zeae, Gloeocercospora sorghi [27] (reclassified as Microdochium sorghi), Aspergillus niger, Penicillium chrysogenum [28], Botryotinia fuckeliana, Pyrenophora teres [29], Stereum hirsutum [15], and Exidia glandulosa [30]. Some other CynHs were directly obtained from their natural producers (for relevant reviews, see [2,31]). Homology searches were based on some annotated CynHs. The searches for sequences with 60–99% identity to selected confirmed CynHs and more than 90% coverage provided 389 homolog sequences (including the templates), with all of them in fungi and 98.5% of them in Ascomycota (Table 2). Only six of them were identified in Basidiomycota.
In contrast to CynDs, CynHs can also transform some organic cyanides. For example, the activities for 2-cyanopyridine (1a) and fumaronitrile (2a) (Figure 2) were found in all tested CynHs [28,29,30,32]. The Vmax/KM ratio is higher for HCN than for cyanopyridine or fumaronitrile in CynH from A. niger (ca. 62, 2.8, and 1.3, respectively) [32].
Table
Table 2. Cyanide hydratases (CynHs) identified in GenBank.
Table 2. Cyanide hydratases (CynHs) identified in GenBank.
PhylumClassGenus 1Sequences Found 2Species with Confirmed CynHs (Reference)
AscomycotaDothideomycetesAureobasidium39
Alternaria17
Hortaea12
Pyrenophora4P. teres [29]
Leptosphaeria2L. maculans [33]
Stemphylium1S. loti [34]
Other46
LecanoromycetesMycoblastus2
Letharia2
Other6
LeotiomycetesBotrytis8
Monilinia4
Botryotinia1B. fuckeliana [29]
Other20
SordariomycetesFusarium43F. solani [35], F. lateritium [26], F. graminearum [27] 3
Colletotrichum21
Monosporascus8
Diaporthe6
Verticillium6
Neurospora2N. crassa [27]
Microdochium2M. sorghi [27] 4
Other39
EurotiomycetesAspergillus52A. nidulans [27], A. niger [32]
Penicillium29P. chrysogenum [28]
Other9
XylobotryomycetesCirrosporium1
SareomycetesSarea1
BasidiomycotaAgaricomycetesAuricularia3
Stereum2S. hirsutum [15]
Exidia1E. glandulosa [30]
1 The templates were CynHs from Fusarium solani (GenBank: CAC69666.1) [35] and Gloeocercospora sorghi (annotated as Microdochium sorghi in NCBI; GenBank: AAA33353.1) [27]. Identity and coverage thresholds were 60–99% and 90%, respectively. Non-redundant sequences were filtered with the tool (https://web.expasy.org/decrease_redundancy/; accessed on 5 December 2022). 2 Only the genera with the highest number of sequences or with confirmed CynHs are shown for each class. 3 Formerly Gibberella zeae. 4 Formerly Gloeocercospora sorghi.
Thus, CynDs and CynHs form relatively small fractions of nitrilases. The total number of nitrilase homologs is in the thousands according to GenBank (not shown). CynD and CynH both contain the Glu-Lys-Cys triad characteristic of the nitrilase superfamily (C164, E48, and K130 in CynDpum). Substrate positioning was studied in two amidases from the genus Pyrococcus, one of which was previously designated as a nitrilase. In these enzymes, another conserved Glu, together with Lys and the first Glu, contributes to the correct position of the substrate toward the catalytic Cys [36]. It is thus essential for activity, which is also shown by mutation studies on an amidase from Geobacillus pallidus [37]. Nitrilases, without excluding CynDs and CynHs, also contain this conserved second Glu residue (Figure 3), which corresponds to E137 in CynDpum. The sequence close to the second Glu is more conserved in CynH than in CynDs. However, we can identify a common pattern for both types of enzymes: XER/KL, where X is the hydrophobic aliphatic residue. In the selected nitrilases, only the ER motif is conserved.
CynHs or CynDs do not appear to be parts of metabolic pathways such as the “aldoxime-nitrile pathway”, although some genera of fungi (Fusarium, Aureobasidium, etc.) contain both (putative) aldoxime dehydratases (Oxds) [38] and CynHs abundantly. However, a typical substrate of nitrilases involved in the above pathway is phenylacetonitrile (a natural compound), while the activities of CynHs are negligible for this substrate [29,32].

3. Overexpression of Genes

The cynD and cynH genes were obtained by the amplification of genomic, complementary [27,32,39], or synthesized DNA [28,29,30]. Their expression largely followed standard protocols with Escherichia coli as the host, although it differed in methodological details such as the cultivation medium, temperature regime, IPTG concentration, etc., as shown in Table 3 and Figure 4.
The two best-characterized CynDs (CynDpum and CynDstut) were produced with an N-terminal His6-tag, which slightly affected the kinetic parameters and the temperature optimum, as shown in CynDpum [39].
Some cynH genes were produced in the autoinduction medium (glucose, glycerol, and lactose [40]). Expression levels were similar for the tagged and untagged constructs. Subsequent work was carried out with the tagged constructs to simplify purification [27].
For enzyme overproduction in E. coli, recent studies on other nitrilases have proposed methods to increase efficiency and reduce cost. One promising approach is the high-density culture (the optical density was up to 86.4 at 600 nm). In addition, the nit gene (from Pseudomonas putida) was expressed constitutively, showing a high activity per liter of culture medium (654,000 U/L for 3-cyanopyridine as the substrate) [41].
The activity and stability of nitrilase can be improved by the chaperones GroEL/GroES. This was shown for a nitrilase from Rhodococcus rhodochrous produced in E. coli or Rhodococcus ruber. From the different options tested (fusion of each chaperone with nitrilase, co-expression), the co-expression of GroEL (in R. ruber) was the most effective; it increased the specific activity and, especially, the thermostability of the nitrilase (the latter by eightfold) [42]. The GroEL/GroES co-expression was also used for the production of CynHs and other nitrilases [29] (Table 3). Earlier investigation suggested the positive effect of these chaperones on nitrilase solubility [43,44].
Table
Table 3. Overexpression of the cynD and cynH genes in Escherichia coli.
Table 3. Overexpression of the cynD and cynH genes in Escherichia coli.
EnzymeVector; His6-TagMedium; Cultivation ConditionsKinetic ParametersReference
CynDpET28a; N-terminalLB broth; (i) 37 °C → OD 0.3;
(ii) 1 mM IPTG, 30 °C, 3–4 h		Vmax 120–150 U/mg 2
KM 0.7–7.9 mM 2		[18,19,45]
CynHpET26b; C-terminalLB broth; (i) 30 or 37 °C → OD 0.5;
(ii) 1 mM IPTG, 30 °C, 3–10 h		Vmax ≈ 4400 U/mg 3
KM ≈ 90 mM 3		[39]
pET28a; N-terminal
pET26b; no tag		Autoinduction medium (g/L):
glucose 0.5, glycerol 5, lactose 2;
30 °C, overnight		n.d.[27]
pET30a; no tagLB broth; (i) 37 °C → OD 0.4–0.6;
(ii) 0.8 mM IPTG, 26 °C, 16 h		Vmax ≈ 6800 U/mg 4
KM ≈ 109 mM 4		[32]
pET22b, pGro7 (Takara) 1; C-terminalEnPresso (Biosilta); (i) 30 °C, 16 h;
(ii) 0.02 mM IPTG, 11 mM L-arabinose; 25 °C, 24 h		n.d.[29]
pET22b; C-terminal2xYT; (i) 37 °C → OD 1.0;
(ii) 0.02 mM IPTG, 20 °C, 20 h		Vmax ≈ 1335 U/mg 5
KM ≈ 22 mM 5		[30]
1 Encoding GroEL/ES chaperone (expression induced by L-arabinose). 2 Bacillus pumilus CynD and mutants. 3 Gloeocercospora sorghi (reclassified: Microdochium sorghi). 4 Aspergillus niger. 5 Exidia glandulosa. n.d., no data.
Catalysts 13 00500 g004 550
Figure 4. Summary of the methods used for the production, purification, and activity assays used for (a) cyanide hydratases (CynDs) and (b) cyanide hydratases (CynHs). See Table 3 and Table 4 for the details of gene expression and activity assays, respectively.
Figure 4. Summary of the methods used for the production, purification, and activity assays used for (a) cyanide hydratases (CynDs) and (b) cyanide hydratases (CynHs). See Table 3 and Table 4 for the details of gene expression and activity assays, respectively.
Catalysts 13 00500 g004
Table
Table 4. Cyanide (di)hydratase activity assays.
Table 4. Cyanide (di)hydratase activity assays.
Enzyme/Reaction ConditionsCompound DeterminedReagentMethodReference
CynD/4 mM KCN, pH 7.7, r.t.Free cyanidePicric acid, Na2CO3Spectrophotometric[18,19,45]
CynD/4 mM KCN, pH 7.7, r.t.AmmoniaNessler (K2HgI4)Spectrophotometric[18]
CynH/10 mM KCN, pH 7.4, r.t.Free cyanidePicric acid, Na2CO3Spectrophotometric[27]
CynH/25 mM KCN, pH 8.0, 30 °CFormamide-HPLC (ion-exchange column)[32]
CynH/25 mM KCN, pH 8.0, 30 °CFormamide-HPLC (silica gel column)[15]
CynH/25 mM KCN, pH 9.0, 30 °CFormamide(i) Hydroxylamine/NaOH, (ii) HCl/FeCl3Spectrophotometric[30]
r.t., room temperature.
The above cultivations were carried out on small scale. However, the feasibility of scaling up the production of a related enzyme (nitrilase acting on mandelonitrile) was demonstrated for E. coli harboring the nit gene from A. faecalis [10]. The whole-cell catalyst produced R-MA from 0.5 M (R,S)-mandelonitrile. The cells were prepared in 420 L, 4200 L, and 12,000 L of an autoinduction medium.
There are only very few examples of the use of Pichia pastoris (reclassified as Komagataella phaffii) as the host for nit gene expression. However, this host organism has some significant advantages in post-translational modification, and restricted leakage and solubility of the recombinantly produced protein [46]. The associated approach has been demonstrated with nitrile-hydrolyzing enzymes [46,47,48].
The nitrilase production in K. phaffii was intracellular [46,47] or extracellular [48]. Thus, this host was used to co-express the nit gene of Pseudomonas fluorescens and the oxynitrilase gene from Manihot esculenta, and the whole-cell catalyst was used for the biocatalytic synthesis of (S)-mandelic acid from benzaldehyde [47]. An efficient and robust nitrilase for the production of CCHA was also prepared in K. phaffii. The strain carried multiple copies of the nit gene from Acidovorax facilis. In addition, the production was optimized by selecting a suitable (alcohol oxidase 1) promoter and co-expressing a chaperone (ER oxidoreductin 1). The resulting whole-cell catalyst was quite stable for one day at 55 °C and could be used for at least 20 times [46]. Another strain of this organism produced the nitrilase from Alcaligenes faecalis for the synthesis of R-MA from mandelonitrile; the enzyme was produced constitutively and secreted into the culture medium [48].

4. Activity Assays

The assays of cyanide-transforming enzymes are based on different principles (Table 4 and Figure 4), and the use of different methods can make it difficult to compare the results. The activities of CynDs or CynHs were often determined based on the rate of substrate (fCN) consumption. However, the fCN concentrations or pH were different in different studies. Alternatively, the rate of product (ammonia and formamide) formation was used. In CynHs, the concentration of substrate was often lower than KM. The KM values of CynHs are very high (up to ~100 mM, Table 3) [2]. Therefore, the specific activities may be underestimated. In addition, the temperature and pH can affect the stability of fCN, which evaporates significantly at a pH of 8.5 or lower [49].
For example, CynDs were tested with 4 mM fCN at pH = 7.7, and the consumption of fCN was determined with the picric acid reagent [18,19,45]. Alternatively, the product ammonia was determined using the Nessler reagent under the same conditions [18] or at a pH ranging from 5.5 to 9.5 [19].
The activities of the first recombinant CynHs were determined using 10 mM KCN at pH = 4.5–11. In stability assays, the residual activities of the enzymes were determined at pH = 7.4. The concentrations of the remaining fCN were monitored using the picric acid reagent [27]. The activity of a new CynH from E. glandulosa was determined with 25 mM KCN at pH = 9–10.5, and the residual fCN was determined in the same way. The formation of formamide was monitored spectrophotometrically using the hydroxylamine reagent according to a previous method [50]. At pH = 9.0, the 22 mM formamide was obtained from 25 mM KCN after 1 h. The rest of the substrate was probably lost by evaporation. However, the activities calculated from the consumption of fCN and the formation of formamide were comparable after 1 to 2 min of reactions at pH = 9.0 and 30 °C [30]. Alternatively, the concentrations of the formed formamide were determined by HPLC/UV using an ion exchange column [28]. The formamide can also be analyzed by HPLC/UV using a silica gel column [51] and this method was used with the CynH from S. hirsutum [15]. However, the sensitivity of the methods is low due to the low UV absorbance of formamide.

5. Structural and Computational Studies

The first CynD structure was resolved for a CynDpum mutant (Q86R, H305K, H308K, H323K) recently (pdb code 8c5i, deposition date 18 January 2023). Most previous studies were based on homology models and were optionally combined with electron microscopy. These approaches were used for mutation analyses and predictions of molecular contact surfaces (interfaces) or C-termini (see below).
In their active form, nitrilases are largely oligomers of 4 to 22 subunits. In addition, some produce extended spirals of different lengths [1]. The active form of CynDpum is a helical oligomer comprised of 18 monomer units [52]. The number of subunits in the similar structure of CynDstut is 14 [53]. Active nitrilase monomers have been rarely reported. One of them is the recently described CynD from Flavobacterium indicum [20].
The contact interfaces between the monomers in a helical oligomer are designated A, C, D, and E, whereas in nitrilase Nit4 from Arabidopsis thaliana, the F interface has also been found, which depends on oligomer termination [54]. The positions of A, C, and D surfaces for Cynpum are shown in Figure 5a (the E interface is formed by the rotation of the terminal dimers in the helix).
The C interface contains residues 55–72 and 222–235 in CynDpum (54–71 and 221–234 in CynDstut) [24]. The length of the first C-interface loop is similar for all the identified homologs, with the conserved motif YP[W/Y] at the beginning of the loop. The importance of these regions for oligomerization and activity has been demonstrated by mutation studies [23,24,55] (Section 6).
Catalysts 13 00500 g005 550
Figure 5. (a) Contact interfaces in the helical form of CynDpum. The monomers in one of the dimers are shown in magenta and blue and the rest of the helix is shown by surface representation. Contact interfaces are highlighted in yellow and labeled. The A interface is formed between monomers in a dimer, the C interface is formed between two dimers in the helix, and the D interface is formed between helical turns. (b) Adjacent dimers (colored differently). The C-terminal part of the encircled monomer is highlighted in blue to visualize its 3-D orientation. (c) Multiple sequence alignment of the C-terminal regions of CynDpum mutant (pdb 8c5i), Nit4 from Arabidopsis thaliana (pdb 6i00) [54], nitrilase from Pseudomonas fluorescens EBC191 (pdb 6zby), nitrilase from Synechocystis sp. PCC6803 (pdb 3wuy) [56], nitrilase-superfamily enzyme from Pyrococcus abyssi (pdb 3ivz) re-classified from nitrilase to amidase [36], CynH from Microdochium sorghi (AAA33353.1) [27], CynH from Fusarium solani (CAC69666.1) [35], CynH from Stereum hirsutum (XP_007307917.1) [15], CynD from Flavobacterium indicum (CCG52320.1) [20], CynD from Stutzerimonas stutzeri (BAA11653.1), and CynD from Bacillus pumilus (AAN77004.1) [24,55].
Figure 5. (a) Contact interfaces in the helical form of CynDpum. The monomers in one of the dimers are shown in magenta and blue and the rest of the helix is shown by surface representation. Contact interfaces are highlighted in yellow and labeled. The A interface is formed between monomers in a dimer, the C interface is formed between two dimers in the helix, and the D interface is formed between helical turns. (b) Adjacent dimers (colored differently). The C-terminal part of the encircled monomer is highlighted in blue to visualize its 3-D orientation. (c) Multiple sequence alignment of the C-terminal regions of CynDpum mutant (pdb 8c5i), Nit4 from Arabidopsis thaliana (pdb 6i00) [54], nitrilase from Pseudomonas fluorescens EBC191 (pdb 6zby), nitrilase from Synechocystis sp. PCC6803 (pdb 3wuy) [56], nitrilase-superfamily enzyme from Pyrococcus abyssi (pdb 3ivz) re-classified from nitrilase to amidase [36], CynH from Microdochium sorghi (AAA33353.1) [27], CynH from Fusarium solani (CAC69666.1) [35], CynH from Stereum hirsutum (XP_007307917.1) [15], CynD from Flavobacterium indicum (CCG52320.1) [20], CynD from Stutzerimonas stutzeri (BAA11653.1), and CynD from Bacillus pumilus (AAN77004.1) [24,55].
Catalysts 13 00500 g005
The C-termini, which are part of the A interface, also maintain interactions required for the formation of the quaternary structure in nitrilases. They span at least two adjacent dimers in Nit4 from Arabidopsis thaliana (pdb 6i00) [54] and the nitrilase from Pseudomonas fluorescens EBC191 (pdb 6zby). The orientation of the C-terminus in the dimer of the recently reported structure of CynDpum is shown in Figure 5b. This C-terminus is longer than that in Nit4 (see the multiple alignment in Figure 5c) and spans three adjacent dimers (Figure 5b).
A conserved motif D[P/F]XGHY [1] was identified for C-termini in bacterial, fungal, and plant nitrilases (Figure 5c). In contrast to CynDstut, CynDpum and their homologs (in which this motif is conserved) the CynD of F. indicum and its homologs (with only one outlier sequence) have a DILGHY motif and a shorter C-terminus (Figure 5c).
CynHs are also multimers [1]. The structure of CynH has not yet been elucidated, and thus, most studies rely on homology models reported for, e.g., the CynHs from N. crassa [1] and Stereum hirsutum [15]. The lengths of the sequences that form the C interface are similar for CynHs and CynDs. A conserved motif YP[W/Y] is present in most of the identified CynH homologs, except for three sequences. The presence of a WPV motif in the active site of CynH limits the size of substrates that can be accepted by CynHs [15]. The corresponding sequence in CynDpum is W188PG190. The C-terminus in CynH homologs has a conserved motif (DXXGHY) similar to that of CynDs (Figure 5c).

6. Engineered Enzymes

The potential of enzyme improvement by mutations was described for CynDs. Although the methodology was based on random mutagenesis in some cases, it was largely based on site-directed mutagenesis of the contact interfaces (mainly A and C) and the C-terminus. These mutations mainly altered the pH range, thermostability, and oligomerization/activity of the enzymes (see Section 6.1., Section 6.2. and Section 6.3., respectively). However, similar studies are still largely lacking for CynHs. In contrast, mutations in the contact interfaces or C-termini were also reported for nitrilases [54,57].
Several nitrilase mutations may also be relevant to CynDs or CynHs. For example, the mutants that alter the reaction product toward amide were studied in detail for the nitrilase from Pseudomonas fluorescens [13]. A similar change in the product was achieved in the nitrilase from Synechocystis sp. by replacing F193 with Asn [58]. In contrast, mutations of several residues in the active site environment (e.g., mutation of the second key glutamate E142 to Ala) in this enzyme resulted in a loss of catalytic activity tested toward fumaronitrile, 2-butenenitrile, benzonitrile, 2-cyanopyridine, and 3-cyanopyridine [56].

6.1. Improvement of the pH Range

A hybrid (called CynDpum-stut) was created with the C-terminal part of CynDpum being replaced by that of CynDstut [19]. It exhibited a different pH profile compared to the wild-type CynDs. The mutant was highly active at pH = 9.0 and fairly active at pH = 9.5. Moreover, unlike the wild-type enzymes, it retained some activity at pH = 10.0. Other mutants with improved activity at alkaline pH were produced by point mutations [59], although they were outperformed by the above hybrid (Table 5).
In contrast, some wild-type CynHs are quite active at pH = 9.0 or higher [27,30]. Therefore, their improvement is not so urgent in terms of pH ranges.

6.2. Stability Enhancement

The stability of wild-type CynDs and their mutants was examined at 42 °C. In general, the purified protein (10 μg/mL) was incubated at pH = 7.7 for 1 or 2 days. The hybrid CynDpum-stut retained 50% of the initial activity after 17 h at 42 °C, whereas the wild-type CynDs retained 50% of activity after 1 h [19].
The GERDST motif in the C-terminus of CynDpum had a positive effect on the thermostability of CynDpum [19]. The significance of the individual amino acid residues in the GERDST motif in CynDpum (not conserved, Figure 5c) was investigated using alanine scanning. All mutants retained activity, but three of them were much less active than the wild-type CynDstut [18].
The thermostability of the hybrid was further improved by a point mutation when K39 was replaced by R [19]. The K39R mutant of CynDpum was selected among those produced by the error-prone PCR. Two other mutants (E327K and D172N) also showed improved thermostability, whereas the K39R/D172N/E327K triple mutant was still better [45].
Some wild-type CynHs are quite thermostable. The CynH from N. crassa showed the highest thermostability of the four CynHs tested, with 40% of residual activity after 2-day incubation at 43 °C [27].

6.3. Mutants with Altered Oligomerization and Activity

Mutations in the regions that form the A- and C-surfaces (Section 5) have been studied, and some of them have been shown to affect activities. Complete deletion of residues 220–234 (the C surface) in CynDstut (219–233 in CynDpum) resulted in inactive forms. Mutation of residues 55–72 (another part of the C surface) in CynDpum resulted in up to >90% activity loss. Some of these mutants, such as R67C, formed smaller oligomers including a dimer [24]. In contrast, the Y70C mutant showed a normal oligomer size (18 subunits) despite no activity. This suggested another mechanism of action that disabled CynD from reaching the desired conformation [23].
The CynDpum mutants with shortened C-terminal parts (the A surface) exhibited decreased activities. Mutants terminated at residues 303 and 293 retained 87% and 27% activity of the wild-type enzyme (330 residues), respectively. The mutant with only 279 residues was inactive. Similar studies were performed for deletion mutants of CynDstut (334 residues) in which residues after 330, 310, and 302 were removed. The mutants had residual activities of 97%, 13%, and 0%, respectively [18].
Similar trends were observed for CynH from N. crassa (351 residues; deletion of residues after 339, 323, and 307). The mutants showed residual activities of 46%, 22%, and 0%, respectively [18]. Mutants of CynH of A. niger (356 residues) showed similar behaviors. Deletion of 14 residues of the C-terminus had no significant effect on activity, but deletion of 18 or more residues (after residue 338, 334, or 322) had nearly abrogated activity [32]. The exact position of the C-terminus in CynDpum has been determined for the recently resolved structure (8c5i). The role of the C-terminal part is likely to be similar to that reported for the Nit4 nitrilase (pdb 6i00) (see Section 5).
The effect of the A and C surfaces on the nitrilase conformation was also discussed in a previous study on plant nitrilases. Two plant nitrilases were manipulated by combining (“swapping”) their fragments with additional modifications of the above surfaces. This not only improved the catalytic properties but also increased the solubility of the resulting mutant [60]. Thus, these findings could have broader applications in nitrilases.

7. Catalyst Forms and Uses

The recombinantly produced CynDs and CynHs were largely purified by affinity chromatography [18,19,27,29,30,45] (Figure 4). The purification yields were rarely reported. The yield was reported to be 14.4% for the CynH from E. glandulosa using cobalt affinity chromatography [30]. In contrast, the CynH from A. niger did not contain a tag and was purified in two steps (ion exchange chromatography and size exclusion chromatography). The purification yield was 17% [32].
The purified CynDpum was maintained at −80 °C [19]. In contrast, these conditions were not suitable for the purified CynHs due to the precipitation of the enzyme [27]. Therefore, the CynHs were stored at 4 °C [27,30]. Thus, the CynH from E. glandulosa retained over 4/5 of its activity after more than three months under these conditions [30].
The purified CynHs were tested for the detoxification of fCN in samples of real industrial wastewater or model samples (Table 6). For example, some of them were shown to be resistant to typical pollutants occurring together with free cyanide in coking or petrochemical wastewater (sulfide, thiocyanate, ammonia, and phenol) [30] or real coking wastewater [61] (Figure 6). Some were also active in the solutions of Ag or Cu salts mimicking diluted electroplating wastewaters [30] or diluted real wastewaters of this type [27] (Figure 7).
Studies on the immobilization of CynDs or CynDs are very few (Table 6). The whole cells of E. coli producing CynDpum were entrapped in agar or polyacrylamide and tested on a real mining effluent diluted to 17.6 mM fCN (Figure 8). Both catalysts almost completely degraded 5 mM fCN after 2 h. Moreover, the agar-immobilized catalyst could be reused many times (with 91% and 60% fCN conversion at cycles 22 and 30, respectively). In contrast, the cells immobilized with polyacrylamide showed lower operational stability (the conversion decreased rapidly after about five runs). Both were much more active than the free whole cells at pH = 9.0 and also retained almost full activity at pH = 10.0, where the free whole cells were virtually inactive [62]. This suggests that adopting immobilization to protect the enzyme from the bulk medium is a useful alternative to the much more laborious mutations.
Immobilized CynDs were also proposed to be used in cyanide biosensors as summarized previously [2]. In addition, a new biosensor was proposed based on the CynD from F. indicum [63] (Figure 9).
The CynH from A. niger was used in the form of a whole-cell catalyst held in place by a semipermeable cellulose membrane. The reaction was carried out in a bench-scale flow reactor fed with a solution of 25 mM KCN. The conversion was maintained above 80% for 3 days [32].
The CynHs also act on organic cyanides, and therefore, their uses for synthetic purposes may be possible. For example, a few CynHs exhibited significant activities for 2-CP, which is not well accepted by most of other nitrilases [28]. The major product of this reaction is picolinic acid (fine chemical for synthesis, L-Trp metabolite, biomarker in neurological, and other diseases [64]).
Recent studies on the different types of nitrilases have shown other immobilization methods, such as encapsulation of purified nitrilase in biosilica [65], a modification of an earlier method [66], or cross-linked enzyme aggregates [67], a type of immobilized catalyst first proposed by the Sheldon group [68,69].

8. Conclusions

In general, CynDs and CynHs represent small fractions of the large group of nitrilases. Both of them exhibit a significant sequence conservation except for their C-terminal parts. However, there is a fine line between CynDs and the nitrilases with other substrate specificities. A clear classification of the enzymes would help search for additional CynD candidates. CynDs and CynHs have several potential applications in environmental technology and analytics. The wild-type enzymes are highly active for fCN, although CynDs are relatively unstable. The enzymes can be substantially improved by creating artificial variants and by immobilization. Both approaches have been found successful with CynDs. Similar progress has not been achieved with CynHs. However, some CynHs with promising properties (activity at pH = 9–10, thermostability, and shelf-life) have been obtained directly by retrieving appropriate sequences from databases. Further engineering of the CynDs could be based on the recently deposited structure of a CynD. The next issue to be addressed is to achieve low-cost expression of the genes (e.g., by autoinduction as demonstrated for some CynHs) and its scale-up. The CynHs may also have applications in organic synthesis. The improvement and overexpression of CynDs and CynHs and the scale-up of their production could be inspired by the achievements reported for other nitrilases.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/catal13030500/s1, Table S1. Percentage of amino acid sequence identities between cyanide dihydratases, cyanide hydratases, and nitrile-converting nitrilases; Figure S1. Complete phylogenetic tree of cyanide dihydratase homologs; Figure S2. Complete phylogenetic tree of cyanide hydratase homologs [15,70,71,72,73,74].

Author Contributions

Conceptualization, L.M., N.K. and P.B.; methodology, N.K.; data curation, N.K.; writing—original draft preparation, L.M., N.K. and A.S.; writing—review and editing, L.M., N.K., B.K. and P.B.; supervision, L.M. and P.B.; project administration, L.M. and P.B.; funding acquisition, L.M., P.B. and A.S. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by Czech Science Foundation, grant number GA22-06785S, Czech Technical University in Prague—Faculty of Biomedical Engineering, grant number SGS21/181/OHK4/3T/17, and Czech Academy of Sciences, grant number RVO61388971.

Data Availability Statement

The complete phylogenetic trees of cyanide dihydratases and cyanide hydratases are available at https://itol.embl.de/tree/147231254143208591671005621 (accessed on 20 December 2022) and https://itol.embl.de/tree/17825516828117701671187763 (accessed on 20 December 2022) respectively.

Conflicts of Interest

The authors declare no conflict of interest.

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Catalysts 13 00500 g001 550
Figure 1. Conversion of hydrogen cyanide (HCN) by cyanide dihydratase (CynD) or cyanide hydratase (CynH), and conversion of nitriles (RCN) by nitrilase (Nit). CynHs also convert some nitriles at lower rates than those of HCN [2]. Unlike CynDs or CynHs, Nits include several subgroups with different substrate specificities [1].
Figure 1. Conversion of hydrogen cyanide (HCN) by cyanide dihydratase (CynD) or cyanide hydratase (CynH), and conversion of nitriles (RCN) by nitrilase (Nit). CynHs also convert some nitriles at lower rates than those of HCN [2]. Unlike CynDs or CynHs, Nits include several subgroups with different substrate specificities [1].
Catalysts 13 00500 g001
Catalysts 13 00500 g003 550
Figure 3. Sequence conservation diagrams of selected regions in cyanide dihydratases (CynDs) from Flavobacterium indicum (CCG52320.1), Stutzerimonas stutzeri (BAA11653.1) and Bacillus pumilus (AAN77004.1), cyanide hydratases (CynHs) from Microdochium sorghi (AAA33353.1) and Fusarium solani (CAC69666.1), and nitrilases (not utilizing HCN as substrate) from Trametes versicolor (XP_008032838.1), Synechocystis sp. (3wuy), and Arabidopsis thaliana (pdb 6i00). Red dots mark active site residues. Blue dots mark the second Glu of the catalytic tetrad. The alignment was constructed using CLUSTAL Omega. The diagrams were constructed with WebLogo (http://weblogo.berkeley.edu/logo.cgi). Active site amino acids were identified based on the previous study [1].
Figure 3. Sequence conservation diagrams of selected regions in cyanide dihydratases (CynDs) from Flavobacterium indicum (CCG52320.1), Stutzerimonas stutzeri (BAA11653.1) and Bacillus pumilus (AAN77004.1), cyanide hydratases (CynHs) from Microdochium sorghi (AAA33353.1) and Fusarium solani (CAC69666.1), and nitrilases (not utilizing HCN as substrate) from Trametes versicolor (XP_008032838.1), Synechocystis sp. (3wuy), and Arabidopsis thaliana (pdb 6i00). Red dots mark active site residues. Blue dots mark the second Glu of the catalytic tetrad. The alignment was constructed using CLUSTAL Omega. The diagrams were constructed with WebLogo (http://weblogo.berkeley.edu/logo.cgi). Active site amino acids were identified based on the previous study [1].
Catalysts 13 00500 g003
Catalysts 13 00500 g006 550
Figure 6. Removal of free cyanide (fCN) and phenol compounds (phenol, m-cresol, and p-cresol) from coking wastewater by cyanide hydratase (CynH) and tyrosinase, respectively. The reactions cannot proceed simultaneously due to the inhibition of tyrosinase (a copper enzyme) by fCN. Therefore, in the first step, fCN was converted to formamide by CynH overproduced in Escherichia coli and purified. In the second step, phenol compounds were degraded by the tyrosinase extracted from the fruiting bodies of Agaricus bisporus. Formamide, unlike fCN, was not inhibitory to tyrosinase [61].
Figure 6. Removal of free cyanide (fCN) and phenol compounds (phenol, m-cresol, and p-cresol) from coking wastewater by cyanide hydratase (CynH) and tyrosinase, respectively. The reactions cannot proceed simultaneously due to the inhibition of tyrosinase (a copper enzyme) by fCN. Therefore, in the first step, fCN was converted to formamide by CynH overproduced in Escherichia coli and purified. In the second step, phenol compounds were degraded by the tyrosinase extracted from the fruiting bodies of Agaricus bisporus. Formamide, unlike fCN, was not inhibitory to tyrosinase [61].
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Catalysts 13 00500 g007 550
Figure 7. Removal of free cyanide (depicted as HCN) in model or diluted real electroplating wastewaters by cyanide hydratases (CynHs) overproduced in Escherichia coli. (a) The model wastewater was 100 mM glycine/NaOH buffer at pH = 9.0, supplemented with 100 mM KCN and 1 mM of AgNO3 or 1 mM CuSO4. Conversion of HCN was catalyzed by the purified CynH from Exidia glandulosa [30]. (b) An electroplating bath wastewater with ca. 1 M fCN was 10 times diluted with 1 M MOPS buffer at pH = 8.0. Conversion of HCN was catalyzed by the purified CynH from Neurospora crassa [27]. The Cu or Ag content was not specified. r.t., room temperature.
Figure 7. Removal of free cyanide (depicted as HCN) in model or diluted real electroplating wastewaters by cyanide hydratases (CynHs) overproduced in Escherichia coli. (a) The model wastewater was 100 mM glycine/NaOH buffer at pH = 9.0, supplemented with 100 mM KCN and 1 mM of AgNO3 or 1 mM CuSO4. Conversion of HCN was catalyzed by the purified CynH from Exidia glandulosa [30]. (b) An electroplating bath wastewater with ca. 1 M fCN was 10 times diluted with 1 M MOPS buffer at pH = 8.0. Conversion of HCN was catalyzed by the purified CynH from Neurospora crassa [27]. The Cu or Ag content was not specified. r.t., room temperature.
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Catalysts 13 00500 g008 550
Figure 8. Biodegradation of free cyanide (depicted as HCN) in diluted gold-mining wastewater (“barren solution”, i.e., the solution from which no more gold can be obtained) by immobilized whole-cell catalyst based on the cyanide dihydratase (CynD) from Bacillus pumilus. The enzyme was overproduced in Escherichia coli and used as whole cells immobilized in agar or polyacrylamide beads [62].
Figure 8. Biodegradation of free cyanide (depicted as HCN) in diluted gold-mining wastewater (“barren solution”, i.e., the solution from which no more gold can be obtained) by immobilized whole-cell catalyst based on the cyanide dihydratase (CynD) from Bacillus pumilus. The enzyme was overproduced in Escherichia coli and used as whole cells immobilized in agar or polyacrylamide beads [62].
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Catalysts 13 00500 g009 550
Figure 9. Potentiometric determination of free cyanide (fCN, depicted as HCN) using whole cells of Flavobacterium indicum with cyanide dihydratase (CynD) activity. The cells were entrapped in agar and the resulting catalyst was adsorbed on a nylon membrane. The sensor was used to determine fCN concentrations between 1.5 × 10−9 and 2.5 × 10−3 M [63].
Figure 9. Potentiometric determination of free cyanide (fCN, depicted as HCN) using whole cells of Flavobacterium indicum with cyanide dihydratase (CynD) activity. The cells were entrapped in agar and the resulting catalyst was adsorbed on a nylon membrane. The sensor was used to determine fCN concentrations between 1.5 × 10−9 and 2.5 × 10−3 M [63].
Catalysts 13 00500 g009
Table
Table 1. Cyanide dihydratases (CynDs) identified in GenBank.
Table 1. Cyanide dihydratases (CynDs) identified in GenBank.
PhylumClassGenusSequences Found 1Species with Confirmed CynD Activity (Reference)
FirmicutesBacilliBacillus52 2B. pumilus [23,24,25]
Paenibacillus12 3
Other4
ClostridiaClostridium6
Lacrimispora1
ProteobacteriaGammaproteobacteriaAcinetobacter1
Stutzerimonas1S. stutzeri [18]
BetaproteobacteriaBurkholderia2
ActinobacteriaActinomycetiaBrevibacterium1
BacteroidotaFlavobacteriaFlavobacterium147F. indicum [20]
Other5
AscomycotaSaccharomycetesScheffersomyces1
1 The templates were CynDs from Bacillus pumilus (CynDpum, GenBank: AAN77004.1), Stutzerimonas stutzeri (CynDstut, GenBank: BAA11653.1) and Flavobacterium indicum (CynDind, Genbank: CCG52320.1). Identity and coverage thresholds were 70–99% and 90%, respectively. 2 Bacillus safensis—17, Bacillus pumilus—15, Bacillus thuringiensis—5, and Bacillus cereus—4. 3 Paenibacillus alvei—8.
Table
Table 5. Improved variants of cyanide dihydratase from Bacillus pumilus (CynDpum).
Table 5. Improved variants of cyanide dihydratase from Bacillus pumilus (CynDpum).
VariantIncreased
Thermostability		Activity at pH ≥ 9.0Reference
K39R+(+)[45]
Q86R+++[59]
D172N(+)-[45]
E327K(+)(+)[45]
E327G-++[59]
E35/Q322R/E327G-++[59]
Q86R/E96G/D254E+++[59]
Q86R/E96G/D254E/E327G+++[59]
K39R/D172N/E327K++-[45]
307NHQKNE312 replaced with GERDST 1+(+)[19]
CynDpum-stut hybrid 2++++[19]
CynDpum-stut K39R+++++[19]
1 Corresponding sequence (306–311) in CynDstut. 2 The CynDpum-stut contained 286 residues of CynDpum and the C-terminus of CynDstut (from residue 287 to end).
Table
Table 6. Forms of catalysts for the degradation of free cyanide and their uses.
Table 6. Forms of catalysts for the degradation of free cyanide and their uses.
EnzymeCatalyst FormPropertiesApplicationCyanide Concentration; RemovalReference
CynDWhole cells immobilized in agarRecyclability (>20-fold); Activity at pH 9–10Elimination of cyanide from mining waste17.6 mM, 98% 1[62]
Whole cells immobilized in polyacrylamideRecyclability (5-fold); Activity at pH 9–1017.6 mM, 98% 1; 528 mM, 43% 2
CynDWhole cells immobilized in agarOptimal activity at pH 8.0, 35 °C Half-life 4 h at 40 °CCyanide sensorn.a.[63]
CynHWhole cells in a flow reactorOperational stability (3 days)Elimination of cyanide 125 mM, >80% 3[32]
CynHPurifiedActivity at up to pH 10; Prolonged shelf-life (>3 months)Elimination of cyanide0.6–100 mM; 96–100% 3[30]
PurifiedActivity at up to pH 10Elimination of cyanide from electroplating waste100 mM, 30–100% 4[27]
1 Diluted wastewater. 2 Undiluted wastewater. 3 Model solution. 4 Lower conversion in the presence of copper than silver.
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Martínková, L.; Kulik, N.; Sedova, A.; Křístková, B.; Bojarová, P. Recent Progress in the Production of Cyanide-Converting Nitrilases—Comparison with Nitrile-Hydrolyzing Enzymes. Catalysts 2023, 13, 500. https://doi.org/10.3390/catal13030500

AMA Style

Martínková L, Kulik N, Sedova A, Křístková B, Bojarová P. Recent Progress in the Production of Cyanide-Converting Nitrilases—Comparison with Nitrile-Hydrolyzing Enzymes. Catalysts. 2023; 13(3):500. https://doi.org/10.3390/catal13030500

Chicago/Turabian Style

Martínková, Ludmila, Natalia Kulik, Anastasia Sedova, Barbora Křístková, and Pavla Bojarová. 2023. "Recent Progress in the Production of Cyanide-Converting Nitrilases—Comparison with Nitrile-Hydrolyzing Enzymes" Catalysts 13, no. 3: 500. https://doi.org/10.3390/catal13030500

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