- 3.1. Substrate specificity
- 3.2. Enzyme structure
- 4Cyanide hydratase
- 4.1. Substrate specificity
- 4.2. Enzyme structure
- 5Cyanide dihydratase/cyanidase
- 5.1. Substrate specificity
- 5.2. Enzyme structure
- 6Molecular genetic analysis
- 7Reaction mechanism
The enzymes nitrilase, cyanide dihydratase and cyanide hydratase are a group of closely related proteins. The proteins show significant similarities at the amino acid and protein structure level but the enzymes show many differences in catalytic capability. Nitrilases, while catalysing the hydration of nitrile to the corresponding acid, vary widely in substrate specificity. Cyanide dihydratase and cyanide hydratase use HCN as the only efficient substrate but produce acid and amide products, respectively. The similarities of all these enzymes at the amino acid level but the functional differences between them provide a rich source of material for the study of structure/function relationships in this biotechnologically important group of enzymes. This review provides an overview of current understanding of the genetics and biochemistry of this interesting group of enzymes.
Cyanide is abundant in nature and occurs both as inorganic cyanide (HCN) and as organic cyanide or nitriles (RCN). Plants synthesize a range of nitriles. 5-indole-3-acetonitrile (IAN) is an intermediate in the biosynthesis of the major plant hormone IAA (5-indole-3-acetic acid) in the Brassicaceae (Ludwig-Muller and Hilgenberg 1988; Muller and Weiler 2000). β-Cyano-l-alanine [Ala(CN)] occurs widely in plants and is produced by cyanoalanine synthase from cyanide and cysteine. This pathway may play a role in cyanide detoxification in plants (Miller and Conn 1980). The breakdown of glucosinolates (mustard oil glucosides) can also lead to the production of nitriles (Wittstock and Halkier 2002). Glucosinolates are nontoxic but breakdown of these compounds, which occurs on tissue damage, can release toxic compounds such as nitriles and isothiocyanates. Cyanogenic glycosides are produced by many plant species and several economically important plants are highly cyanogenic (Vetter 2000). Cyanogenic glycosides and glucosinolates are thought to play a part in phytopathogen resistance (Vetter 2000; Wittstock and Halkier 2002).
Cyanide and nitrile hydrolysing enzymes have been studied in a wide range of microbial species, and increasingly, in plants. The enzymatic conversion of HCN/nitrile to the corresponding acid can take place by a one-step process as exemplified by nitrilases and cyanide dihydratases (CDH) or by a two-step process with an amide intermediate as is the case with nitrile hydratases and cyanide hydratases (CH).
Cyanide hydratase, although functionally different to nitrilases and cyanide dihydratases, has been shown to be closely related to these enzymes and shows no relationship to the more functionally similar nitrile hydratase (Wang and VanEtten 1992; Cluness et al. 1993). Nitrilases, CDHs and CHs have been classified into a single enzyme family, known as the nitrilase/cyanide hydratase family, which also includes the less closely related aliphatic amidases (Novo et al. 1995). This family is part of a larger group of related proteins, which have been termed CN-hydrolases (Bork and Koonin 1994) or more recently as the nitrilase superfamily (Pace and Brenner 2001). The nitrilase superfamily has been classified into 13 branches based on sequence homology. Nine of these groups have CN hydrolase activity.
This review aims at providing a comparative analysis of the nitrilase branch of the nitrilase superfamily. This branch includes nitrilase, CDH and CH enzymes, all of which have CN hydrating activity but the enzymes differ in the product produced or in substrate specificity. CDH and CH enzymes show high specificity for HCN showing very little activity with nitriles while nitrilases in general show activity with a broad range of nitrile substrates. Nitrilases and CDH produce mainly an acid product while CH produces the amide product from HCN. The nitrilases are important for their potential application in biotransformation particularly for the production of fine chemicals for the pharmaceutical industry (reviewed by Kobayashi and Shimizu 2000; Banerjee et al. 2002) while HCN hydrating enzymes have application in the bioremediation of cyanide bearing waste (Dubey and Holmes 1995).
The enzyme nitrilase was first described by Thimann and Mahadevan (1964). The enzyme, which was isolated from barley leaves, catalysed the conversion of indoleacetonitrile (IAN) to indoleacetic acid (IAA) and was initially called indoleacetonitrilase. Substrate analysis with purified enzyme on 26 nitriles indicated that the enzyme had a broad substrate range. The rate of hydrolysis was eight times greater with 3-cyanopyridine than with IAN. The enzyme was therefore renamed nitrilase to indicate the broad substrate range of the enzyme (Thimann and Mahadevan 1964). The first bacterial nitrilase was isolated from a soil bacterium (possibly a Pseudomonas species) by selection for growth on the naturally occurring nitrile, ricinine (N-methyl-3-cyano-4-methoxy-2-pyridone), as a sole carbon source(Hook and Robinson 1964; Robinson and Hook 1964). This enzyme catalysed the hydrolysis of a range of 2-pyridones with rates between 0 and 118% relative to ricinine (100%).
The biotechnological potential of nitrile hydrolysing enzymes has led to the isolation of a range of bacteria and fungi capable of hydrolysing nitriles. Most of these were isolated on the basis of using a particular nitrile as a carbon and/or nitriogen source. A list of bacteria, fungi and plants with well-characterized nitrilases is given in Table 1. The enzymes studied to date show a very diverse range of biochemical characteristics. In particular substrate specificity of the enzymes varies widely. Initial investigations suggested that nitrilases were specific for aromatic nitriles and nitrile hydratases for aliphatic nitriles but this distinction has to be reconsidered in the light of the growing body of information on both nitrilases and nitrile hydratases. Table 2 gives an overview of the basic biochemical characteristics available on the most studied nitrilases. While it is not possible to give a complete analysis of substrate range for each of the enzymes the most notable features for each enzyme are given and the relative rate of benzonitrile hydrolysis is given where available. None of the enzymes show identical properties. The most notable differences between the enzymes are in substrate specificity, native structure and aggregation properties, and pH optima.
|Acido vorax facilis 72W||Gavagan et al. 1999|
|Acinetobacter sp. (strain AK226)||Yamamoto and Komatsu 1991|
|Alcaligenes faecalis ATCC 8750||Yamamoto et al. 1992|
|Alcaligenes faecalis JM3||Nagasawa et al. 1990|
|Bacillus sp. strain OxB-1||Kato et al. 2000|
|Bacillus pallidus Dac521||Almatawah et al. 1999|
|Comamonas testosteroni||Levy-Schil et al. 1995|
|Klebsiella pneumoniae ssp. ozaenae||Stalker et al. 1988|
|Pseudomonas?||Hook and Robinson 1964|
|Pseudomonas sp. (SI)||Dhillon et al. 1999|
|Pseudomonas DSM 7155||Layh et al. 1998|
|Nocardia (Rhodococcus) NCIB11216||Harper 1977b|
|Nocardia (Rhodococcus) NCIB11215||Harper 1985|
|Rhodococcus rhodococcus K22||Kobayashi et al. 1990|
|Rhodococcus ATCC39484||Stevenson et al. 1992|
|Rhodococcus rhodochrous PA-34||Bhalla et al. 1992|
|Rhodococcus rhodochrous J1||Kobayashi et al. 1989|
|Fusarium oxysporum f.sp. melonis||Goldlust and Bohak 1989|
|Fusarium solani IMI196840||Harper 1977a|
|Arabidopsis thaliana||Piotrowski et al. 2001|
|Osswald et al. 2002|
|Barley||Mahadevan and Thimann 1964|
|Chinese cabbage||Rausch and Hilgenberg 1980|
|Organism||Substrate||Molecular mass (kDa)||pH||Temp. optima||Reference|
|Acinebacter sp. AK226||Ibu-CN 100% |
|pH 8||50°C||Yamamoto and Komatsu 1991|
|Alcaligenes faecalis ATCC 8750||Mandelonitrile 100% |
p-aminobenzyl cyanide 1670%
|460||32||pH 7·5||40–45°C||Yamamoto et al. 1992|
|Alcaligenes faecalis JM3||Mandelonitrile 8·62% |
p-aminobenzyl cyanide 49·1%
|260||44||pH 7·5||45°C||Nagasawa et al. 1990 |
Kobayashi et al., 1993
|Bacillus pallidus Dac521||Benzonitrile 100% |
|600||41 + 72 (groEL)||pH 6–9||65°C||Almatawah et al. 1999|
|Comamonas testosteroni||Adiponitrile 100% |
|Oligomer||38||Levy-Schil et al. 1995|
|Fusarium oxysporum f.sp. melonis||Benzonitrile 100% |
|550 (170–880)||37||pH 6–11||40°C||Goldlust and Bohak 1989|
|Fusarium solani IMI196840||Benzonitrile 100% |
|620||76||pH 7·8–9·1||Harper 1977a|
|Klebsiella pneumoniae ssp. ozaenae||Bromoxynil 100% |
|74||37 × 2||pH 9·2||35°C||Stalker et al. 1988|
|Pseudomonas sp. (SI)||Acrylonitrile 100% |
|41||41||Dhillon et al. 1999 |
Dhillon and Shivaraman 1999
|Nocardia (Rhodococcus) NCIB11216||Benzonitrile 100% |
|pH 8||30°C||Harper 1977b |
Hoyle et al. 1998
|Nocardia (Rhodococcus) NCIB11215||Benzonitrile 100% |
|560||45||pH 7–9·5||30°C||Harper 1985|
|Rhodococcus ATCC39484||Benzonitrile 100% |
|pH 7·5||30°C||Stevenson et al. 1992|
|Rhodococcus rhodococcus K22||Crotononitrile 100% |
|650||41||pH 5·5||55°C||Kobayashi et al. 1990|
|Rhodococcus rhodochrous PA-34||Benzonitrile 100% |
|45||45||pH 7·5||35°C||Bhalla et al. 1992|
|Rhodococcus rhodochrous J1||Benzonitrile 100% |
|pH 7·6||45°C||Kobayashi et al. 1989 |
Nagasawa et al. 2000
3.1. Substrate specificity
Substrate range has been determined for a wide range of purified nitrilases and indicates that while most nitrilases show highest activity with aromatic nitriles some nitrilases have a preference for arylacetonitriles and others for aliphatic nitriles (Table 3). Table 3 shows the activity of the aromatic nitrilases from Rhodococcus rhodochrous ATCC 39484, R. rhodochrous J1, Nocardia (Rhodococcus) sp. NCIB11216 and NCIB 11215, Fusarium solani IMI196840, F. oxysporum f.sp. melonis and Bacillus pallidus Dac521 with a range of substrates and clearly indicates the differences in substrate range between these enzymes. When comparing data from different organisms caution has to be exercised to allow for differences in experimental protocols between laboratories. All the enzymes are more active with substituted aromatic nitriles than with benzonitrile but the best substrate is different for each enzyme. The nitrilase of Nocardia (Rhodococcus) NCIB11215 shows the greatest differences in relative activity showing 8·4 times higher activity with 3-nitrobenzonitrile than with benzonitrile, 7·2 times higher activity with 3-chlorobenzonitrile and 5·2 times higher activity with 3-bromobenzonitrile (Harper 1985). In general the enzymes prefer meta and para substitutions with poor or no activity seen with ortho-substituted substrates. The poor activity with ortho-substituted substrates is thought to be due to steric hindrance. The nitrilase from R. rhodochrous PA-34 (Bhalla et al. 1992) appears similar to this group, although substrate range information for this nitrilase is limited. The enzyme has high activity with benzonitrile and low activity with phenylacetonitrile.
|Substrate||Aromatic nitrilases||Arylacetonitrilases||Aliphatic nitirlases|
|B. pallidus Dac521||F. oxysporum f.sp. melonis||F. solani IMI196840||Nocardia sp. NCIB 11216||Nocardia sp. NCIB 11215||R. rhodochrous ATCC 39484||R. rhodochrous J1||Alcaligenes faecalis JM3||Alcaligenes faecalis ATCC 8750||R. rhodochrous K22||Acidovorax facilis 72W||Comamonas testosteroni||Pseudomonas S1||Acinebacter sp. AK226|
The enzymes from Alcaligenes faecalis strains ATCC8750 and JM3 are both arylacetonitrilases but yet show clear differences in enzymatic activity. The relative activities of these two nitrilases are shown in Table 3. JM3 shows higher activity with acrylonitrile, m-chlorobenzylcyanide, 2-thiophenacetonitrile, 3-pyridineacetonitrile, 2-phenoxypropionitrile while ATCC 8750 has higher activity with p-aminobenzylcyanide (Nagasawa et al. 1990; Yamamoto et al. 1992). The nitrilase from P. fluorescens DSM 7155 is also an arylacetonitrilase with high activity with phenylacetonitrile and similar activity to ATCC 8750 with 2-phenylpropionitrile (Layh et al. 1998). These enzymes have no activity with benzonitrile.
A number of nitrilases which show highest activity with aliphatic nitriles have been studied. Table 3 compares the substrate range of these aliphatic nitrilases. As with other nitrilases these enzymes have different activities from each other. The nitrilase of R. rhodochrous K22 is the best characterized (Kobayashi et al. 1992a). This enzyme has highest activity with the unsaturated aliphatic nitrile acrylonitrile, and the aliphatic dinitrile glutaronitrile. The nitrilases of Acidovorax facilis 72 W (Gavagan et al. 1999), Comamonas testosteroni (Levy-Schil et al. 1995) and Pseudomonas sp. S1 (Dhillon et al. 1999) are also classified as aliphatic nitrilases. The nitrilase of Acinebacter sp. AK226 is unusual in that it has very broad substrate specificity (Yamamoto and Komatsu 1991). This strain was selected for its ability to convert 2-arylpropionitriles to the corresponding acids and was shown to convert racemic 2-(4′-isobutylphenyl)-propionitrile (Ibu-CN) to S-(+)-2-(4′-isobutylphenyl) propionic acid [S-(+)-ibuprofen]. This enzyme could also be classified as an aromatic nitrilase as it has high activity with benzonitrile but as the highest activity is seen with acrylonitrile it is classified as an aliphatic nitrilase here.
The nitrilase of K. pneumoniae ssp. ozaenae was isolated by selection on the herbicide bromoxynil (3,5-dibromo-4-hydroxybenzonitrile) as nitrogen source (McBride et al. 1986). The purified enzyme has no activity with benzonitrile, very low activity with 4-hydroxybenzonitrile but good activity with the 3′-5-dihalogenated 4-hydroxybenzonitrile compounds bromoxynil, chloroxynil and ioxynil (Stalker et al. 1988). The only other nitrilase shown to have activity with bromoxynil and ioxynil is the aromatic nitrilase of Nocardia sp. NCIMB 11215 but the pattern of activity of the two enzymes is very different as the Nocardia enzyme has high activity with benzonitrile (Table 3) and the bromoxynil activity is <0·5% of the benzonitrile activity. The K. pneumoniae ssp. ozaenae appears to be more substrate specific than other nitrilases with a strong preference for halogenated 4-hydroxybenzonitriles.
The nitrilases of Arabidopsis thaliana have been studied in some detail in recent years. Arabidopsis thaliana has four nitrilases, identified initially from genome sequencing (Bartling et al. 1992, 1994; Bartel and Fink 1994). AtNIT1, 2 and 3 are isoenzymes, which are found only in the Brassicaceae. The predicted protein sequences of the three enzymes are >80% identical. The genes for these enzymes are clustered in a 13·8-kb region on chromosome 3 (Hillebrand et al. 1998). The fourth nitrilase of A. thaliana, AtNIT4, is not linked to these genes and the predicted protein sequence is only 65% identical with AtNIT1–3 (Bartel and Fink 1994). AtNIT4 homologs are found in many plant species such as tobacco (Tsunoda and Yamaguchi 1995) and rice (Piotrowski et al. 2001). Enzymatic characterization of AtNIT1–3 indicates a preference for aliphatic substrates with the most efficient substrate tested being phenylpropionitrile (PPN). IAN is a relatively poor substrate (2% activity relative to PPN) for the enzymes but it is postulated that the enzymes may still play a role in IAA production in particular situations (Vorwerk et al. 2001). The major physiological role for AtNIT1–3 appears to be in the metabolism of nitriles released by the breakdown of glucosinolates (Vorwerk et al. 2001). The AtNIT4 nitrilase of A. thaliana is a β-cyano-l-alanine hydratase and has very little activity with a wide range of other nitriles tested (Piotrowski et al. 2001).
3.2. Enzyme structure
The subunit molecular mass and native structure has been determined for many nitrilases. Most nitrilases consist of a single polypeptide with a molecular mass of ca 40 kDa (32–45 kDa), which aggregates to form the active enzyme as indicated in Table 2. The preferred form of the enzyme seems to be a large aggregate of 6–26 subunits. The nitrilase of F. oxysporum f.sp. melonis shows many bands with nondenaturing PAGE with sizes from 170 to 880 kDa. The bands increase in size by ca 70 kDa (2 × 34 kDa subunits). Activity assays have shown all these bands to be active. This indicates that the enzyme can be active with 4–26 subunits with addition of subunits always taking place in pairs (Goldlust and Bohak 1989). The enzyme of R. rhodochrous PA-34 is active as a monomer with a molecular mass of 45 kDa (Bhalla et al. 1992). The bromoxynil-specific nitrilase of Klebsiella pneumoniae ssp. ozaenae is active as a dimer (Stalker et al. 1988). The nitrilase of Fusarium solani IMI196840 is unusual in that it has a subunit molecular mass of 72 kDa. Other characteristics of this enzyme are similar to other nitrilases and in particular the enzyme aggregates to an active form of 620 kDa (Harper 1977a).
Some of these enzymes aggregate as a consequence of substrate activation (Table 2). This phenomenon was first observed with the enzyme from Nocardia (Rhodococcus) NCIB 11216 (Harper 1977b). When purified in the absence of benzonitrile this enzyme had an elution volume, on gel filtration, corresponding to a molecular mass of 47 kDa while enzyme purified in the presence of benzonitrile the molecular mass was determined to be 560 kDa. The association of the subunits was shown to be pH, temperature and enzyme concentration dependent with a pH optimum of 7·3 and very slow association below 20°C. The nitrilases of Rhodococcus ATCC 39484, Alcaligenes faecalis ATCC8750 and R. rhodochrous J1 also show substrate-dependent activation (Stevenson et al. 1992; Yamamoto et al. 1992; Nagasawa et al. 2000). Nagasawa et al. (2000) demonstrated that subunit association of the nitrilase of R. rhodochrous J1 took place in the presence of benzonitrile but not in the presence of acrylonitrile although acrylonitrile is a good substrate for the activated enzyme.
The nitrilases of P. fluorescens DSM 7155 and Bacillus pallidus Dac521 proteins copurify with chaperonin proteins (Layh et al. 1998; Almatawah et al. 1999). The purified enzyme from DSM 7155 has a native molecular mass of 130 kDa, which denatures into three bands, of 57, 40 and 38 kDa, on SDS-PAGE. N-terminal sequencing indicates that the 40 and 38 kDa proteins are slightly different from each other and are related to known nitrilases. The 57 kDa protein is almost totally identical (97%) to chaperonin CPN60 (GroEL). Layh et al. (1998) suggest that CPN60 may play a role in assembly of the basic heterodimer into a higher molecular weight complex. The nitrilase of Dac521 was purified with a native molecular mass of 600 kDa, which gave two bands of 41 and 72 kDa on SDS-PAGE (Almatawah et al. 1999). The N-terminal sequence of the 41 kDa band showed good homology with other nitrilases while the 72-kDa band showed almost complete identity to the GroEL protein from B. subtilis Marburg (Tozawa et al. 1995). The GroEL protein was very tightly associated with the nitrilase protein and may play a role in stabilizing the complex in this strain. Interestingly, the nitrilase gene of C. testosteroni has been cloned and overexpressed in E. coli (Levy-Schil et al. 1995). The protein was expressed at a high level of 30% total cellular protein but only 10% of this was soluble. It was found that the solubility of the protein could be greatly enhanced by overexpression of GroEL chaperonin in the same cell and the enzyme activity increased fivefold. However, the significance of this is unclear, as overexpression of GroEL increases the solubility of many foreign proteins in E. coli, as seen with the nitrile hydratase of C. testosteroni (Stevens et al. 2003) and pig heart mitochondrial citrate synthase (Haslbeck et al. 2003).
4. Cyanide hydratase
The enzyme, cyanide hydratase (formamide hydrolyase EC4·2·1·66), was first identified, by Fry and Millar (1972), in the fungus Stemphylium loti. This fungus is a pathogen of the cyanogenic plant, birdsfoot trefoil (Lotus corniculatus). Fry and Millar (1972) found that cyanide tolerance in S. loti was due to an enzyme, induced by exposure to cyanide, which converts HCN to formamide. Fry and Millar noted the similarity between the broad pH range of this enzyme and the nitrilase from barley (Thimann and Mahadevan 1964) and the ricinine nitrilase (Hook and Robinson 1964). The native molecular mass of the S. loti enzyme was not determined but as it eluted in the void volume of a Sephadex G-200 column it was thought to be very large. Since this initial identification the enzyme has been found in many fungal species but not in bacteria or plants (Miller and Conn 1980; Rust et al. 1980; Wang et al. 1999).
4.1. Substrate specificity
The enzymes are much more substrate specific than nitrilases with HCN being the most effective substrate for all the enzymes. The enzymes from F. solani and F. oxysporum N-10 can hydrolyse a metal–cyano complex, tetracyanonickelate (II) (TCN) (Barclay et al. 1998; Yanase et al. 2000) but the activity, at least in F. oxysporum N-10, is very low (ca 0·05% of the HCN activity) (Yanase et al. 2000). Fusarium oxysporum N-10 cyanide hydratase also has some activity (ca 0·05% HCN activity) with a number of aliphatic nitriles showing best activity with acrylonitrile, methacrylonitrile and crotononitrile (Yanase et al. 2000). The product of nitrile hydrolysis is the corresponding amide, which indicates that the enzyme is catalysing a monohydrolysis of the substrate and therefore the reaction is a nitrile hydratase reaction rather than a nitrilase reaction. Nolan et al. (2003) have recently shown that the cyanide hydratase of F. lateritium has low nitrilase activity (0·02–0·4% HCN activity) with benzonitrile, propionitrile and acetonitrile.
4.2. Enzyme structure
Detailed biochemical analysis has been carried out on the enzyme from Gloeoecercospora sorghi (Fry and Munch 1975; Wang et al. 1992; Wang and VanEtten 1992), F. lateritium (Cluness et al. 1993), F. solani (Barclay et al. 1998), F. oxysporum N-10, Leptosphaeria maculans (Sexton and Howlett 2000) and F. solani IMI 196840 (Nolan et al. 2003). The cyanide hydratases analysed to date show many similarities to each other and represent a much more closely related group of enzymes than the nitrilases discussed above. The subunit molecular mass for all the enzymes is ca 40 kDa. The enzymes from G. sorghi, F. lateritium and F. solani have native molecular masses >300 kDa while that of L. maculans is 160 kDa.
5. Cyanide dihydratase/cyanidase
Ingvorsen et al. (1991) found that Alcaligenes xylosoxidans ssp. denitrificans strain DF3 catalysed the hydrolysis of HCN to formate without forming formamide as a free intermediate. The enzyme was named cyanidase. Enzymes catalysing the direct hydrolysis of HCN to formate have also been identified in Bacillus pumilus C1 (Meyers et al. 1993), P. fluorescens NCIMB 11764 (Kunz et al. 1994) and P. stutzeri AK61 (Watanabe et al. 1998). The enzyme activity has been named cyanidase, cyanide nitrilase and cyanide dihydratase by different authors but in this review the name cyanide dihydratase (CDH) will be used.
5.1. Substrate specificity
The CDH enzymes are cyanide specific. None of the CDH enzymes studied to date produce any formamide from HCN and therefore do not show any cyanide hydratase activity. Pseudomonas fluorescens NCIMB 11764 (Kunz et al. 1994) metabolizes HCN by a number of different routes. The major activity under aerobic conditions is an NADH-dependent cyanide oxygenase, which produces CO2 and NH3. This enzyme is unrelated to the nitrilase/cyanide hydratase family of enzymes. The strain also produces formamide and formic acid. The relative amounts of formamide and formic acid produced, depends on growth conditions. Kunz et al. (1994) propose that the strain contains both cyanide hydratase and cyanide dihydratase activities but as the enzymes have not been purified it is also possible that the strain contains a single enzyme with both activities or that formic acid is a product of formamidase activity.
5.2. Enzyme structure
The native enzyme isolated from Alcaligenes xylosoxidans ssp denitrificans strain DF3 has a molecular mass of >300 kDa. SDS-PAGE indicated the enzyme consisted of two subunits of 39 and 40 kDa with identical NH2–terminal sequences. The possibility that the slightly different molecular masses are due to post-translational modification of the protein or proteolytic degradation was not excluded in this study (Ingvorsen et al. 1991). Interestingly, the enzyme isolated from Bacillus pumilus C1 shows three bands (41·2, 44·6 and 45·6 kDa) on SDS-PAGE and a native molecular mass of 417 kDa. The N-terminal sequences of the three polypeptides are also identical. It is proposed that the different subunits may be due to gene duplication or post-translational modification (Meyers et al. 1993) The only other purified CDH, from P. stutzeri AK61, consists of a single subunit of 38 kDa aggregated to a native molecular mass of >100kDa (Watanabe et al. 1998).
6. Molecular genetic analysis
The genes encoding the nitrilases of Klebsiella pneumoniae ssp. ozaenae (Stalker et al. 1988), R. rhodochrous strains K22 and J1 (Kobayashi et al. 1992b; Kobayashi et al. 1992a), A. faecalis JM3 (Kobayashi et al. 1993) and C. testosteroni (Levy-Schil et al. 1995) have been sequenced. A nitrilase gene has also been sequenced from Bacillus sp. strain OxB-1 (Kato et al. 2000). The nitrilase gene of OxB-1 is linked to the gene for the aldoxime dehydratase which catalyses the first step in aldoxime degradation, which is the dehydration of the aldoxime to the nitrile. Nitrilase genes have been identified in a number of plant species and are classified into Nit4 type nitrilases and Nit1/2/3 type nitrilases. For the purposes of this discussion the A. thaliana Nit1 and Nit4 nitrilases will be used as representative plant nitrilases. Cyanide hydratase genes have been cloned and sequenced from G. sorghi (Wang and VanEtten 1992), F. lateritium (Cluness et al. 1993), L. maculans (Sexton and Howlett 2000) and F. solani (Barclay et al. 2002). The only cyanide dihydratase sequence available is from P. stutzeri AK61 (Watanabe et al. 1998). A comparative analysis, of the available amino acid sequences, is shown in Fig. 1. Analysis indicates that the CHases are very closely related to each other with a percentage similarity of 74·1–88·7% but as all the CHases sequenced to date are fungal in origin this probably reflects lack of evolutionary divergence rather than any functional significance. The plant nitrilases, AtNIT1 and AtNIT4, show 63·6% similarity. This high level of homology again probably reflects lack of evolutionary divergence because, as discussed above, the enzymes have been shown to be functionally different (Piotrowski et al. 2001; Vorwerk et al. 2001; Osswald et al. 2002). The bacterial nitrilases show much less conservation within the group. The highest degree of homology, at 53%, is seen between the aliphatic nitrilase of C. testosteroni (Levy-Schil et al. 1995) and the nitrilase of Bacillus sp. strain OxB-1which is associated with aldoxime degradation (Asano and Kato 1998). Interestingly these two nitrilases show a high level of homology to the cyanide dihydratase from P. stutzeri. The two nitrilases from R. rhodochrous J1 and K22 are only 48·2% homologous which reflects their very different substrate specificities (see Table 3). A phylogenetic tree of the available sequences is shown in Fig. 2. The tree supports the assignment of the three enzymatic activities, nitrilase, cyanide hydratase and cyanide dihydratase, to a single protein family as nitrilase proteins are found in all branches of the tree. The highly specific bromoxynil nitrilase of Klebsiella pneumoniae ssp. ozaenae (Stalker et al. 1988) is closely related to the cyanide hydratases which are also very substrate specific. An alignment of the sequences is shown in Fig. 3. Only 25 residues are conserved in all the proteins while there are six residues which are conserved differently in CH and nitrilases/CDH. The importance of some of these residues has been determined by site-directed mutagenesis (Table 4). Residue numbers refer to Leptosphaeria maculans (Sexton and Howlett 2000). The cysteine 163 residue has been shown to be essential for activity of nitrilases (Kobayashi et al. 1992a,b, 1993; Piotrowski et al. 2001; Vorwerk et al. 2001), cyanide dihydratase (Watanabe et al. 1998) and cyanide hydratase (Brown et al. 1995). This residue is proposed to be part of a catalytic triad of cys163, glu46 and lys129, which has been identified from the crystal structures of NitFhit and N-carbamyl-d-amino acid amidohydrolases (Nakai et al. 2000; Pace et al. 2000). These proteins are members of the nitrilase superfamily of carbon–nitrogen hydrolases (Pace and Brenner 2001). Mutation of the lys129 residue of cyanide dihydratase resulted in no enzyme activity and no CDH protein could be purified from the cells (Watanabe et al. 1998). Watanabe et al. (1998) identified a number of positions where mutation led to an increase of activity (E180Q and D264N) or Km (Y52F). Most of the residues tested appear to play a role in correct folding of the enzymes as mutation alters protein stability. Interestingly the phenylalanine residue at position 170 in CH, one of the residues conserved differently in CH and nitrilases/CDH, appears to form part of the active site of this enzyme as mutation of this residue to the nitrilase conserved residue leads to complete loss of activity but normal protein levels (Nolan et al. 2003). The nitrilase activity of CH is also lost in this mutant indicating that the active site is the same for both activities.
|Mutation||Enzyme||Effect||Relative enzyme activity||Reference|
|T12Q||F. lateritum CH||Reduced activity||52–90||Nolan et al. (2003)|
|T12P||F. lateritum CH||Reduced stability||0||Nolan et al. (2003)|
|S13A||F. lateritum CH||Reduced activity||57–81%||Nolan et al. (2003)|
|E46Q||P. stutzeri CDH||Reduced stability||0||Watanabe et al. (1998)|
|Y52F||P. stutzeri CDH||Reduced activity||Increased Km (×4), 68%||Watanabe et al. (1998)|
|S102A||P. stutzeri CDH||Reduced stability||0||Watanabe et al. (1998)|
|E103Q||P. stutzeri CDH||Reduced activity||57%||Watanabe et al. (1998)|
|K136R||F. lateritum CH||Reduced activity||48%||Nolan et al. (2003)|
|C163A||F. lateritum CH||Reduced activity||0||Brown et al. (1995)|
|A. faecalis Nit||Kobayashi et al. (1993)|
|R. rhodochrous K22 Nit||Kobayashi et al. (1992a)|
|R. rhodochrous J1 Nit||Kobayashi et al. (1992b)|
|A. thaliana Nit 1/2/3||Vorwerk et al. (2001)|
|A. thaliana Nit4||Piotrowski et al. (2001)|
|C163S||P. stutzeri CDH||Reduced activity||0||Watanabe et al. (1998)|
|R. rhodochrous K22 Nit||Kobayashi et al. (1992a)|
|R. rhodochrous J1 Nit||Kobayashi et al. (1992b)|
|E165Q||P. stutzeri CDH||Reduced stability||0||Watanabe et al. (1998)|
|F170L||F. lateritum CH||Reduced activity||0||Nolan et al. (2003)|
|E180Q||P. stutzeri CDH||Increased activity||120%||Watanabe et al. (1998)|
|H183N||P. stutzeri CDH||Reduced stability||0||Watanabe et al. (1998)|
|D264N||P. stutzeri CDH||Increased activity||170%||Watanabe et al. (1998)|
|D275E||F. lateritum CH||Reduced activity||71–100%||Nolan et al. (2003)|
|V281A||F. lateritum CH||Reduced activity||92||Nolan et al. (2003)|
|K292Q||P. stutzeri CDH||Reduced stability||0||Watanabe et al. (1998)|
|D296N||P. stutzeri CDH||Reduced stability||0||Watanabe et al. (1998)|
|H300N||P. stutzeri CDH||Reduced stability||0||Watanabe et al. (1998)|
|Y301F||P. stutzeri CDH||Reduced stability||0||Watanabe et al. (1998)|
|M302S||F. lateritum CH||Reduced activity||78%||Nolan et al. (2003)|
Nitrilases, CHs and CDHs appear to be in all cases inducible enzymes. The level of induction can be very high. For e.g. the isovaleronitrile induced nitrilase of R. rhodochrous J1 corresponds to 35% of the soluble protein of the cell (Komeda et al. 1996) while the cyanide induced CH of F. lateritium represents ca 25% of the soluble protein (Cluness et al. 1993). Little is known about the genetic regulation of these enzymes. Komeda et al. (1996) identified a gene, nitR, which is found downstream of the R. rhodochrous J1 nitrilase gene. This gene encodes a protein with significant homology to AraC type transcriptional regulators (Martin and Rosner 2001) and NitR appears to be a positive regulator of nitA nitrilase gene expression (Komeda et al. 1996). Transcription analysis of nitR indicates that it is probably expressed by read-through from the nitA promoter unlike most AraC type regulators whose genes are transcribed divergently from the genes they are regulating (Komeda et al. 1996). A putative nitrilase regulator has also been found in Bacillus sp. strain OxB-1 (Kato et al. 2000). The nitrilase gene of OxB-1 is found 65 bp upstream of the phenylacetaldoxime dehydratase gene (oxd) and the putative regultor gene (orf2) is 314 bp upstream of the nitrilase gene and is expressed divergently. The product of orf2 is ca 27% identical to NitR of R. rhodochrous J1 and other AraC type regulators.
7. Reaction mechanism
For most nitrilases the only products produced are the corresponding acid and ammonia. However, it has been shown that the purified nitrilases of F. oxysporum f.sp. melonis (Goldlust and Bohak 1989), R. rhodochrous ATCC39484 (Stevenson et al. 1992), Pseudomonas DSM7155 (Layh et al. 1998), and the ricinine nitrilase of Pseuodomonas sp. (Hook and Robinson 1964) produce a small amount of amide product and therefore have nitrile hydratase activity. In these cases the amide product is usually <5% of the total reaction products. The AtNIT4 enzyme from A. thaliana has high NHase activity producing 1·5 times more asparagine than aspartic acid from the β-cyano-l-alanine substrate (Piotrowski et al. 2001). Recently it has been shown that the AtNIT1 enzyme of A. thaliana also has high NHase activity with some substrates (Osswald et al. 2002). For e.g. with fumaronitrile as substrate the ratio of amide to acid products was 93 : 7 while with crotononitrile the ratio was 1 : 99. The nitrilase of R. rhodochrous J1 has been shown to use benzamide as a substrate, albeit at a very low rate (0·00022%) compared with benzonitrile (Kobayashi et al. 1998), and therefore has amidase activity. The cyanide hydratase enzyme from F. lateritium has a small amount of nitrilase activity (Nolan et al. 2003). Kobayashi et al. (1998) have proposed a mechanism for nitrilase/cyanide hydratase activity which accounts for all the activities seen with these enzymes. This mechanism is shown in Fig. 4. It is an adaptation of the earlier mechanisms proposed by Hook and Robinson (1964) and Mahadevan and Thimann (1964). The mechanism proposes nucleophilic attack on the nitrile carbon atom by the sulphydryl group of the nitrilase which leads to a tetrahedral intermediate, via an enzyme-thioimidate route ( in Fig. 4). This intermediate can be broken down to the acid, as happens most often with nitrilases and cyanide dihydratase, or can release the amide as happens with cyanide hydratase. The nitrile hydratase and amidase activities of nitrilases and the nitrilase activity of cyanide hydratase support this mechanism.
The nitrilase family of enzymes are a group of closely related enzymes with diverse biochemical capabilities. Analysis shows that some of these enzymes have broad substrate specificity while others are very substrate specific. Continued molecular analysis and structure/function studies should enhance our understanding of these enzymes. The isolation of new organisms with new specificities will also increase our understanding of these enzymes and increase the biotechnological capacity for nitrile biocatalysis and cyanide bioremediation.