Cyanide bioremediation: the potential of engineered

Appl Microbiol Biotechnol DOI 10.1007/s00253-017-8204-x

MINI-REVIEW

Cyanide bioremediation: the potential of engineered nitrilases Jason M. Park 1 & B. Trevor Sewell 2 & Michael J. Benedik 1

Received: 16 January 2017 / Revised: 13 February 2017 / Accepted: 15 February 2017 # Springer-Verlag Berlin Heidelberg 2017

Abstract The cyanide-degrading nitrilases are of notable interest for their potential to remediate cyanide contaminated waste streams, especially as generated in the gold mining, pharmaceutical, and electroplating industries. This review provides a brief overview of cyanide remediation in general but with a particular focus on the cyanide-degrading nitrilases. These are of special interest as the hydrolysis reaction does not require secondary substrates or cofactors, making these enzymes particularly good candidates for industrial remediation processes. The genetic approaches that have been used to date for engineering improved enzymes are described; however, recent structural insights provide a promising new approach. Keywords Cyanide . Bioremediation . Nitrilase . Cyanide dihydratase . Protein engineering . Protein stability

Introduction From the time chemist Carl Wilhelm Scheele discovered prussic acid (hydrogen cyanide, H-CN) in 1762 (Cummings 2004), cyanide has retained its notoriety as a powerful poison from its use in murders, as a chemical weapon, and for terror attacks (Gracia and Shepherd 2004). It has found extensive use as a pesticide and was the active ingredient in Zyklon B, infamous for its use in concentration camps during WWII. * Michael J. Benedik [email protected] 1

Department of Biology, Texas A&M University, College Station, TX 77843-3258, USA

2

Structural Biology Research Unit, Institute for Infectious Diseases and Molecular Medicine, University of Cape Town, Cape Town 7925, South Africa

The toxicity of cyanide stems from its inhibition of cytochrome C oxidase in the electron transport chain as well as many other metalloenzymes by binding metal ions critical to the proteins’ functions (Vogel et al. 1981). The toxicity in humans can be as low as 1 mg/kg body weight; at higher doses, mortality can occur within minutes. Exposure to hydrogen cyanide gas commonly leads to convulsions, with respiratory and cardiac arrest following in rapid succession. It is the affinity of cyanide for metals that also makes it a valuable tool in mining and electroplating. Despite the associated risks, the global consumption of cyanide is over 1.5 million metric tons per year. Although most is used in the synthesis of organic chemicals (Cummings 2004), the use that poses the largest environmental threat is as a lixiviant in gold leach mining. Up to 90% of gold is extracted from ore utilizing some form of cyanide leaching. These mining practices generate large volumes of cyanide-laden effluent, which when not properly contained and remediated, have had catastrophic impacts on downstream environments and human communities (Kovac 2000; Rico et al. 2008). Conventional chemical treatments of cyanide wastewaters can be effective at reducing the toxicity from cyanide; they can be costly, require significant infrastructure, need longterm storage or transportation, and involve reagents that pose their own environmental hazards (Akcil 2003). This has sparked interest in turning to nature for biological methods for cyanide degradation (reviewed in Akcil 2003; Dash et al. 2009; Gupta et al. 2010). Cyanide is naturally produced, albeit in small quantities, by a variety of plants, animals, and microorganisms as defense molecules or during nitrogen metabolism. To deal with the toxicity of cyanide, these and competing organisms have evolved enzymes and pathways for its degradation or transformation. Gupta et al. (2010) provides a broad overview of cyanide bioremediation and the enzymes and pathways

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having potential for the degradation of industrial cyanide wastes. Among the cyanide-degrading enzymes, two related groups of enzymes from the nitrilase superfamily are particularly attractive, the microbial enzymes cyanide dihydratase (CynD) and cyanide hydratase (CHT). Recently summarized by Martínková et al. (2015), these cyanide-degrading enzymes have significant potential for biodegradation and possible utility as a biosensor for cyanide. This review discusses current methods used for degrading cyanide wastes and potential enzymatic alternatives and focuses on the microbial enzymes CynD and CHT. We discuss the potential of CynD and CHT as bioremediation tools, what is known about their distinctive structure, and the progress made in engineering these enzymes to improve their industrial applicability.

photodecomposition. Natural degradation is a lengthy process, and only 60% of the cyanide is removed in a period of 9 months (Botz 2000). The elevated localized concentration of hydrogen cyanide gas in the vicinity of the ponds adds to the safety concerns as well. A method marketed by Cyanide Destruct Systems, Inc. (Canada) uses the thermal hydrolysis of cyanide solutions. This naturally occurring reaction converting cyanide into ammonia and formic acid occurs slowly (Stuehr et al. 1963) (Eq. 1). In order to speed up the reaction, temperatures of 400–450 °F (204–230 °C) at high pressure (300 to 500 psi/ 20 to 35 bar) are used to maintain the liquid phase during the reaction. This method does not require additional reagents and is able to destroy all cyanide complexes including ironcyanide (Robey 2004).

Cyanide detoxification

HCN þ 2H2 O→NH3 þ HCOOH

Cyanide effluents generated by various industries differ in their cyanide concentration, chemical contaminants, and cyanide-metal complexes, each of which complicates remediation in different ways thereby requiring differing decontamination approaches. Cyanide compounds in effluents are classified in three categories that control the process or treatment strategy to be used. The first is the free cyanide category, which include both free cyanide ion (CN−) and hydrogen cyanide (HCN). Compounds in this category are considered extremely toxic. The second category consists of the weak acid dissociable (WAD) cyanide compounds, which are weak or moderately stable metal-cyanide complexes such as those of sodium, cadmium, copper, nickel, potassium, and zinc. These are also highly toxic. The last category consists of strong metal-cyanide complexes such as those of gold, silver, and iron. This category is considered the least toxic. These categories of cyanide discharge are important to the choice of a detoxification technique. The industrial application using cyanide significantly impacts treatment; ore leaching applications use cyanide at concentrations generally in the 5– 20 mM range, whereas metal plating industries generate more controlled volumes of waste but with cyanide concentrations often in excess of 1 M. Natural degradation or attenuation of cyanide relies on holding the wastewaters for an extended period in ponds (Ritcey 2005). This permits free cyanide (HCN or CN−) to volatilize and upon reaction with air, oxidation results in the production of ammonia and bicarbonate. The efficacy of this technique depends strongly on the cyanide species in the solution, their concentration, and the storage conditions such as temperature, aeration, depth, and area of the ponds. Depending on the pH, WAD cyanide species can be similarly dissociated at pH 4.5, yielding hydrogen cyanide that will then evaporate. Iron-cyanide complexes can also be dissociated through the action of UV radiation (sunlight), defined as

ð1Þ

In addition to these non-chemical cyanide remediation approaches, there are a variety of chemical approaches that rely on the destruction of cyanide through oxidation, with each having unique advantages and disadvantages. Alkaline chlorination is the most commonly used decontamination technique. This technique is well established and requires the use of hypochlorite ion originating from chlorine bleach (sodium hypochlorite) (Eq. 2). An advantage is that cyanide is transformed into the less toxic product cyanate. The method works well for both free and WAD cyanide; however, stable metal complexes with gold, silver, copper, or nickel require longer times but can still be transformed. This method is not effective with ferrocyanide [Fe(CN)6–4] and has the disadvantage that chlorine itself is toxic and can generate toxic intermediates. CN− þ OCl− →CNO− þ Cl−

ð2Þ

These disadvantages of alkaline chlorination have led to hydrogen peroxide becoming the preferred reagent for cyanide oxidation (Knorre et al. 1984). This reaction, in alkaline medium with a catalyst such as copper, also produces the less toxic cyanate (Eq. 3a) that is subsequently hydrolyzed to carbon dioxide and ammonia (Eq. 3b). The reaction is simple to operate and is environmentally favorable since excess hydrogen peroxide decomposes to water and oxygen. Free cyanide as well as WAD cyanide can easily be transformed with this method but other cyanide complexes with nickel, silver, and iron require additional steps. Ferrocyanide [Fe(CN)6–4] is not oxidized because of the high stability of the complex; however, it can undergo precipitation using heavy metals such as copper or by photolytic degradation using UV irradiation (Kuhn and Young 2005). The major drawback of using hydrogen peroxide is the relatively high reagent cost and the continuous and accurate measurements of the reagent dose that is required (Mosher and Figueroa 1996).

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CN− þ H2 O2 →OCN− þ H2 O

ð3aÞ

CNO− þ 2H2 O→NH3 þ CO2 þ OH−

ð3bÞ

Cyanide and thiocyanate anions can also be oxidized using sulfur dioxide (SO2) in the presence of air and a soluble metal catalyst such as copper (Cu) (Devuyst et al. 1989) (Eq. 4a). This sulfur dioxide mediated destruction technique is known as INCO and marketed by INCO Ltd. The process oxidizes free and WAD cyanide and thiocyanate (Eq. 4b) but not ferrocyanides [Fe(CN)6–4] that are precipitated by the heavy metal catalyst. This approach is useful for the cyanide detoxification of slurry; however, the capital cost is high due to the complex chemical handling systems used as well as the licensing fees (Mosher and Figueroa 1996). CN− þ SO2 þ H2 O þ O2 →OCN− þ H2 SO4

ð4aÞ

SCN− þ 4SO2 þ 5H2 O þ 4O2 →OCN− þ 5H2 SO4

ð4bÞ

The chemical processes described above are successful in yielding low non-toxic cyanide concentrated effluents. The limitations arising from cost or the additional treatment of the products needed before the effluent can be released into the environment are often significant. Additionally, many of the chemical processes are not suitable for in situ remediation, necessitating either transportation of the waste or construction of a local facility. For these reasons, the bioremediation of cyanide waste is presently being considered as a simple alternative that is a safe, inexpensive, and environmentally friendly method compared to the chemical processes. Cyanide-degrading enzymes Despite being a potent poison, cyanide is commonly found in nature. Not surprisingly, therefore, most organisms produce enzymes capable of degrading cyanide either for detoxification or for assimilation as a nitrogen source. The enzymes that form the basis for biological remediation differ remarkably in their reaction conditions, co-factor requirements, or additional substrates. The three main reaction paths, reduction, oxidation, and hydrolysis, are degradative pathways: they degrade cyanide into simpler and less toxic molecules. A fourth, substitution/transfer, is commonly used for the assimilation of cyanide into an organism’s primary metabolism as a source of nitrogen. Cyanide reduction Some microorganisms such as cyanobacteria use nitrogenase to fix atmospheric nitrogen. Several have been shown to mediate cyanide degradation through the action of their nitrogenase such as Rhodopseudomonas gelatinosa (Materassi et al. 1977) and Klebsiella oxytoca (Kao et al. 2003). Nitrogenase

activity of the cyanide-degrading K. oxytoca strain is induced by the presence of potassium cyanide in the medium as a sole nitrogen source. This reaction requires anaerobic conditions because oxygen inhibits the nitrogenase enzyme mediating the pathway. Also, the enzyme requires low potential reductants such as sodium dithionite (Na2S2O4), ATP, and protons. Nitrogenase is made up of two protein complexes, an iron containing homodimer (Fe protein) and an iron-molybdenum heterotetramer (MoFe protein). The Fe protein forms a complex with MoFe protein while binding two molecules of ATP; in this complex, one electron is transferred from Fe protein to MoFe accompanied by hydrolysis of ATP into ADP. The complex is then dissociated, and the two molecules of ADP are replaced by ATP (Li et al. 1982). In 1967, Hardy and Knight showed that HCN was reduced to methane and ammonia through the action of nitrogenase (Eq. 5) (Gantzer 1990; Hardy and Knight 1967). However, using nitrogenase is problematic because it is inhibited by the CN− species which stops electron flow through the pathway and thus acts as an inhibitor (Li et al. 1982). HCN þ 6e− þ 6H þ →CH4 þ NH3

ð5Þ

The concentration of the CN− species in solution is dependent upon the pH. Its concentration increases with an increase in pH. At pH 7, HCN constitutes 99% of total cyanide (HCN + CN−) present in aqueous solution. However, at pH 9.5, 80% of total cyanide is present as CN− (Broderius 1981). Therefore, cyanide solutions are routinely kept at highly alkaline pH to inhibit the production of the volatile hydrogen cyanide. Whereas nitrogenase might be useful at neutral pH, it is ineffective in the alkaline conditions routinely found industrially because its catalysis is inhibited by the excess CN− substrate found in those conditions. Cyanide conversion by oxidation pathways Pseudomonas fluorescens NCIMB 11764 was isolated for its ability to utilize cyanide as a sole source of nitrogen in the presence of a carbon source such as glucose (Harris and Knowles 1983a). Cultures pulsed with cyanide vapor in aerobic conditions degraded the cyanide and produced stoichiometric amounts of ammonia while stimulating oxygen uptake (Harris and Knowles 1983a). However, using a P. fluorescens NCIMB 11764 cell-free extract in lieu of whole cells, cyanide degradation required the addition of NADH (Harris and Knowles 1983b). Two pathways were proposed. The first involves a cyanide monooxygenase that converts cyanide to cyanate (Eq. 6a). The latter product would be converted to ammonia and carbon dioxide through the action of a cyanase, an enzyme found in bacteria and plants such as Escherichia coli (Anderson and Little 1986) and Arabidopsis thaliana

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(Qian et al. 2011). Cyanase catalyzes the conversion of cyanate to ammonia and carbon dioxide utilizing bicarbonate in the process. This first pathway was supported by the presence of cyanate-degrading activity in bacterial extracts (Harris and Knowles 1983b; Kunz and Nagappan 1989). In the second pathway, a NADH-requiring cyanide dioxygenase catalyzes the conversion of cyanide into ammonia and carbon dioxide (Harris and Knowles 1983b) (Eq. 6b). Further research demonstrated that there is no co-induction between cyanide oxygenase and cyanase activities, excluding the coupled pathway of a cyanide oxygenase with cyanase (Dorr and Knowles 1989). Supporting this were additional studies showing that a cyanate-defective (ON101) mutant strain of P. fluorescens NCIMB 11764 still grew on cyanide, and vice versa, a separate mutant (JL102) while no longer capable of growth on cyanide remained able to grow on cyanate (Kunz et al. 1994). Cyanide monooxygenase HCN þ O2 þ Hþ þ NADH→HOCN þ NADþ þ H2 O ð6aÞ Cyanide dioxygenase HCN þ O2 þ Hþ þ NADH→CO2 þ NH3 þ NADþ

ð6bÞ

To demonstrate that the first step is indeed a monooxygenase, Wang et al. (1996) showed a single atom of labeled molecular oxygen was incorporated during the reaction and an intermediate X-OH formed (Eq. 7a) (Wang et al. 1996). A second atom of oxygen from water was then incorporated and the hydrolysis of the substrate into ammonia and carbon dioxide completed (Eq. 7b). HCN þ O2 þ 2NADH þ 2Hþ →½X−OH þ H2 O þ NADþ ð7aÞ

½X−OH þ H2 O→NH3 þ CO2

ð7bÞ

Studies on the purified cyanide-attacking monooxygenase showed that the enzyme is pterin-dependent (Kunz et al. 2001), requires NADH, and converts cyanide into formate and ammonia when the cyanide concentration in the medium was 10 to 50 μM (Eq. 8a) (Fernandez et al. 2004). Formate is further oxidized by formate dehydrogenase (FDH) into carbon dioxide (Eq. 8b). HCN þ O2 þ NADH þ Hþ →HNCO þ H2 O þ NADþ ð8aÞ HNCO þ NADH þ Hþ → HCONH2 þ NADþ HCONH2 þ H2 O→ HCO2 H þ NH3

HCO2 H þ NADþ →CO2 þ NADH þ Hþ

ð8bÞ

At higher cyanide concentration (1 mM), the major end product of the reaction was formamide (formamide/formate ratio, 0.6:0.3), which accumulates in the medium and is not

further degraded (Fernandez et al. 2004). Cyanide monooxygenase is proposed to consist of four different components; NADH oxidase, NADH peroxidase, cyanide dihydratase, and carbonic anhydrase (Fernandez and Kunz 2005). Cyanide monooxygenase activity is detected only when all four components are combined and NADH and reduced pterin added to the reaction yielding as products formate and ammonia. In the presence of FDH, the formate is further oxidized into CO2 (Fernandez and Kunz 2005). Cyanide hydrolysis Cyanide degradation through hydrolysis is catalyzed by members of the nitrilase superfamily (Pace and Brenner 2001) that have two different activities: cyanide hydratase and cyanide dihydratase. Cyanide hydratase (CHT) converts cyanide into formamide (Eq. 9a). It is found in numerous fungi such as Neurospora crassa, Gloeocercospora sorghi, and Fusarium lateritium (Basile et al. 2008). Cyanide dihydratase (CynD), sometimes referred to as cyanidase, converts cyanide into ammonia and formate (Eq. 9b). It has only been found in bacteria, notably Bacillus pumilus, Pseudomonas stutzeri, and Alcaligenes xylosoxidans subsp. denitrificans (Jandhyala et al. 2005). The primary substrate for cyanide hydratase and cyanide dihydratase is cyanide, and the hydrolysis reaction does not require secondary substrates or cofactors making these enzymes good candidates for industrial remediation processes. They also lend themselves for use as biosensors for rapid and accurate detection of cyanide (Keusgen et al. 2004) whereby the degradation of cyanide by cyanide dihydratase yields ammonia that is then detected by an ammonia electrode. These nitrilases will be discussed in detail in sections to follow. HCN þ H2 O→HCONH2

ð9aÞ

HCN þ 2H2 O→NH3 þ HCOOH

ð9bÞ

Substitution/transfer pathway Two primary types of cyanide assimilation reactions exist: substitution reactions and amino acid synthesis reactions. The cyanide is not degraded per se but is assimilated into primary metabolic pathways and used as a source for nitrogen for growth. Focused studies of cyanide assimilation have been done on Chromobacterium violaceum (a cyanogenic bacterium), Bacillus megaterium, and Citrobacter freundii (Michaels and Corpe 1965; Porter and Knowles 1979; Rodgers and Knowles 1978). In fungi, a few species such as Rhizopus oryzae (Padmaja and Balagopal 1985) and Rhizoctonia solani (Mundy et al. 1973) demonstrated an ability for cyanide assimilation. On the other hand, plants commonly perform cyanide assimilation.

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Amino acid synthesis reactions

COOHCOCH2 SH þ CN− →CH3 COCOOH þ SCN−

Reactions performed by the enzymes β-cyanoalanine synthase and γ-cyano-α-aminobutyric acid synthase fall into this group. β-Cyanoalanine synthase (CAS) is a mitochondrial enzyme widely distributed in higher plants (Hatzfeld et al. 2000) and insects (Ogunlabi and Agboola 2007) and also found in some bacteria (Brysk et al. 1969; Dunnill and Fowden 1965; Macadam and Knowles 1984). Even in noncyanogenic plants, β-cyanoalanine synthase was detected but with lower activity than that found in cyanogenic species (Miller and Conn 1980). This enzyme catalyzed the conversion of cyanide and cysteine to β-cyanoalanine and hydrogen sulfide (Eq. 10). Further transformation of β-cyanoalanine can be accomplished by the Pin4 nitrilase of P. fluorescens, found as a rhizosymbiont with those plants (Howden et al. 2009).

The crystal structure of 3-mercaptopyruvate sulfurtransferase from Leishmania major (Alphey et al. 2003) showed that the enzyme forms a persulfurated cysteine in its active site. The enzyme then interacts with cyanide to form thiocyanate. This enzyme is involved in cysteine metabolism via the mercaptopyruvate pathway. Cysteine is metabolized to form mercaptopyruvate by cysteine transaminase. 3Mercaptopyruvate is then converted to pyruvate by 3Mercaptopyruvate sulfurtransferase. This enzyme is widely distributed in prokaryotes and eukaryotes and is evolutionary related to rhodanese. In mammalian cells, the enzyme is present in the mitochondria as well as in the cytoplasm. In the past, efforts were employed in finding a cyanideantidote administered to mass-exposed cyanide victims. Such an antidote is advantageous in the case of chemical disaster or terrorist attack. As mentioned earlier, current thiosulfate antidotes rely on rhodanese, which is concentrated in the mitochondria of the liver and kidneys but not in the other major organs such as the heart and the central nervous system. 3-Mercaptopyruvate sulfurtransferase seems to be a better candidate for cyanide detoxification. Sulfanegen sodium, a novel prodrug of 3-mercaptopyruvate, alleviated cyanide toxicity when tested in animals (Belani et al. 2012; Brenner et al. 2010).

HCN þ HSCH2 CHNH2 CO2 H→NCCH2 CHNH2 CO2 H ð10Þ

þ H2 S

Substitution reactions Another enzyme type carrying out cyanide substitution reactions is sulfurtransferase. In 1933, K. Lang discovered rhodanese, which catalyzes the reaction between cyanide and thiosulfate to produce thiocyanate and sulfite (Eq. 11). S2 O3

2−

−

þ CN →SO3

2−

−

þ SCN

ð11Þ

Rhodanese, also known as thiosulfate/cyanide sulfurtransferase, is found in animals with high levels primarily in the mitochondrial fraction of the liver and kidneys (Cipollone et al. 2007). It also occurs in plants like the leaf of the cassava plant Manihot utilissima and microorganisms such as Bacillus subtilis and Pseudomonas aeruginosa (Cipollone et al. 2008). This mitochondrial enzyme is the key enzyme used by animals for the detoxification of cyanide. It is suggested that a low-protein diet is responsible for the chronic disease linked to consumption of cyanogenic foods such as cassava. With a constant intake of non-lethal dose of cyanogens, the body increases the synthesis of rhodanese and the demand for sulfur containing amino acids used as co-factor. Chronic consumption of cyanogenic food needs to be combined with a rich protein diet to avoid cyanide linked diseases such as konzo, a permanent paraparesis. Rhodanese deficiency is also linked to the Leber’s hereditary optic neuropathy, a rare mitochondrially inherited loss of vision (Poole and Kind 1986). 3-Mercaptopyruvate sulfurtransferase is another member of the transferase family. The enzyme has two substrates, 3mercaptopyruvate and cyanide, and catalyzes their conversion to pyruvate and thiocyanate (Eq. 12).

ð12Þ

Nitrilase superfamily Nitrilase enzymes carry out hydrolysis of nitrile groups to carboxylic acid or amide products. A cyanide-hydrolyzing enzyme that emerged to be a member of this enzyme group was first reported in sorghum tissue infected with G. sorghi, a fungus that causes zonate leaf spot in Sorghum species (Fry and Munch 1975). These enzymes are dependent on a cysteine for their catalytic activity (Kobayashi et al. 1992). The extent of the enzyme family was first recognized by bioinformatics studies (Bork and Koonin 1994). Extension of these studies revealed that nitrilases comprise one branch in a 13-branch superfamily of enzymes; members of the other 12 branches all perform hydrolysis of carbon-nitrogen bonds by amidase, carbamoylase, or amide condensation activity (Pace and Brenner 2001). Sequence alignment and structural studies identified a conserved cysteine-glutamate-lysine catalytic triad (Knorre et al. 1984), which was later extended to a catalytic tetrad by the inclusion of a second glutamate based on functional and structural studies (Kimani et al. 2007; Soriano-Maldonado et al. 2011; Weber et al. 2013). All members of the superfamily have a characteristic αββα monomer fold, which in almost all cases pairs to form αββα-αββα dimers (Kimani et al. 2007; Pace and Brenner 2001). A characteristic that distinguishes the nitrilase branch is the large spiral-shaped oligomeric structure found among the microbial nitrilases (Thuku et al. 2009).

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Nitrilases The nitrilases were initially discovered in the 1960s upon their isolation from barley leaves (Thimann and Mahadevan 1964). These nitrilases were found to hydrolyze carbon-nitrogen triple bonds (nitrile or cyano groups) producing ammonia and the corresponding carboxylic acid or amide product (Basile et al. 2008; Kiziak and Stolz 2009; Piotrowski et al. 2001; Woodward 2011). In the plant, they may also play a role in detoxifying cyanide, which is a by-product of ethylene synthesis (Jenrich et al. 2007). Since their discovery, nitrilases have been identified in a several bacteria, fungi, yeast, and plants with activities against a wide range of nitriles (Gong et al. 2012). Nitrilase-related proteins have also been discovered in a variety of other contexts leading to the categorization of the nitrilase superfamily of enzymes (Pace and Brenner 2001). Cyanide-degrading nitrilases CynD is a nitrilase that is able to degrade cyanide into the much less toxic ammonia and formate (Meyers et al. 1993; Watanabe et al. 1998). The two most studied CynD enzymes are those of P. stutzeri (CynDstut) and B. pumilus (CynDpum), both isolated from cyanide contaminated environments. B. pumilus C1 was isolated using enrichment cultures started with soil from a cyanide wastewater dam in the Transvaal, South Africa (Meyers et al. 1991), and P. stutzeri AK61 was isolated from cyanide-laden wastewater of a metal plating plant (Watanabe et al. 1998). A third CynD protein was also identified in the bacterium A. xylosoxidans subsp. denitrificans and has since been patented (Igvorsen et al. 1991); little work has been published on it. However, the sequence from a presumably similar enzyme from Alcaligenes strain DN25 is present in the GenBank database (KT965153). The second type of cyanide-degrading nitrilase is the fungal cyanide hydratase (CHT) (not to be confused with metal containing nitrile hydratases). These have been identified either through their cyanide-degrading activity or by genome mining in a variety of fungi including N. crassa, G. sorghi, and F. lateritium and numerous others (Basile et al. 2008). CHT enzymes differ from CynD in that they degrade cyanide to produce formamide instead of ammonia and formate. It is clear from sequence conservation that the active sites in CynD and CHT enzymes are very similar. The CHT enzymes are characterized by an insertion of about 14 amino acids immediately adjacent to the active site (189–203 in the enzyme from N. crassa). A reaction mechanism that accounts for these two different activities in a totally satisfactory manner has never been proposed, and therefore, it is worth examining the mechanism in more detail. A proposal that the active site cysteine attacks the nitrile carbon (Hook and Robinson 1964; Mahadevan and Thimann 1964) leading to the formation of a thioimidate has found wide acceptance (O’Reilly and Turner 2003; Jandhyala et al. 2005). Zhang et al. (2014) has suggested that

deprotonation of the sulfhydryl by the adjacent glutamate activates the cysteine residue for the attack. In the light of this acceptance of the thioimidate, it is surprising that its existence has never been conclusively shown (Stevenson et al. 1992) as mass spectroscopy at the time of the attempt to do so was unable to distinguish between the potential thioimidate and thioester that were found in the case of a slowly reacting substrate. Addition of water to the to the putative thioimidate leads to a tetrahedral intermediate involving the cysteine that would lead to a thioester in either the case of CynD or the release of the amide in the case of CHT (Jandhyala et al. 2005). It is also not known what active site differences are necessary to distinguish nitrilase and amidase activities. Indeed, the relative locations of the primary active site residues as visualized in crystal structures of enzymes having the two different activities are extremely similar, within the limits of resolution, for the amidase from Geobacillus pallidus and the nitrilase from Synechocystis species PCC6803 (Fig. 1). Some related nitrilases can release both amide and acid products in ratios that depend on the substrate (Fernandes et al. 2006; Piotrowski et al. 2001). This suggests the possibility of a two-step process in which the amide is the output of the first step and the input of the second step—however, Williamson et al. (2010) argued against this on the basis of the constancy of the acid/amide ratio for a particular substrate as a function of time and the failure of the G. pallidus nitrilase to accept an amide substrate. However, at least one nitrilase, that from Rhodococcus rhodochrous J1 (Kobayashi et al. 1999), has been demonstrated to hydrolyze an amide substrate, albeit slowly, confirming

Fig. 1 Superposition of the active site residues of the amidase from Geobacillus pallidus and the nitrilase from Synechocystis species PCC6803

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at least some functional similarity of the active site of these two branches of the superfamily. An acceptable explanation for these observations should emerge from a detailed functional and structural analysis of the determinants of amidase and nitrilase activity, but this has yet to be accomplished. Sequence analysis and point mutagenesis of the conserved active site residues in CynDpum (the CynD from B. pumilus) demonstrated the enzyme’s relationship to other nitrilases (Watanabe et al. 1998). Nitrilase structure The crystal structures of multiple representatives of members of the nitrilase superfamily demonstrate the conservation of the core structure. The cyanide hydrolyzing enzymes CynD and CHT have never been visualized at crystallographic resolution, but the quaternary structure has been shown by negative stain electron microscopy to be in the form of either short spirals (Jandhyala et al. 2003; Meyers et al. 1993) or extended helices. Several have been reconstructed in three dimensions (Dent et al. 2009; Sewell et al. 2003; Woodward et al. 2008). The oligomerization of the core structure into extended spirals that is seen primarily in the nitrilase branch of the superfamily (Thuku et al. 2009) distinguishes these enzymes from the majority of the amidases. Nitrilase αββα-αββα dimer Despite great sequence divergence within the nitrilase superfamily, its members show significant secondary and tertiary structural homology (Thuku et al. 2009). The monomer structure was first revealed to be a αββα sandwich based on the crystal structure of the Nit domain from the NitFhit protein (Pace et al. 2000) and the Dcarbamoylase (Nakai et al. 2000). In this and other crystal structures, the αββα monomer is paired to form a αββα-αββα dimer, a defining characteristic of the superfamily (Gordon et al. 2013; Kumaran et al. 2003; Nakai et al. 2000; Pace and Brenner 2001; Pace et al. 2000). The interface pairing the monomer subunits into this αββα-αββα dimer is commonly referred to as the Asurface (Sewell et al. 2003). Earlier secondary structure predictions and circular dichroism data (Mueller et al. 2006; Sewell et al. 2003; Thuku et al. 2007) and more recent crystal structure determination (Zhang et al. 2014) from nitrilase proteins indicate that the αββα-αββα dimer structure is conserved in the nitrilase branch. Models of the dimers for some cyanide-degrading nitrilases based on the dimer structures from more distant members of the superfamily have been built (Dent et al. 2009; Sewell et al. 2003, 2005; Thuku et al. 2007; Williamson et al. 2010). The recent structure of the nitrilase from Synechocystis sp. PCC6803 (Nit6803) (Zhang et al. 2014) has been used as a basis for a model to enable a better

interpretation of the effects of point mutations on the spiral structure (Park et al. 2016) (Fig. 2). Spiral oligomer The basic dimer structure can associate further to form a variety of oligomeric complexes that include tetramers (Nakai et al. 2000; Pace et al. 2000), hexamers (Kimani et al. 2007), and C-shaped octamers (Lundgren et al. 2008). While all structures share the common dimer interface, or A-surface, the way these dimers associate into larger quaternary structures varies among the different superfamily branches. Most nitrilases including CynD, CHT, and others were commonly found to have native molecular masses in excess of 400 kDa and for some over 1 MDa (Meyers et al. 1993; Nagasawa et al. 2000; Thuku et al. 2009; Watanabe et al. 1998). Upon examination by electron microscopy, these nitrilases appear as large spiral oligomers (Dent et al. 2009; Jandhyala et al. 2003; Meyers et al. 1993; Sewell et al. 2003; Thuku et al. 2007; Vejvoda et al. 2008; Woodward 2011; Woodward et al. 2008). Three-dimensional reconstructions from negatively stained protein have revealed these large oligomers to be left handed corkscrew-shaped spirals (Dent et al. 2009; Jandhyala et al. 2003; Sewell et al. 2003; Thuku et al. 2007; Woodward et al. 2008) (Fig. 3). The crystal packing of

Fig. 2 A model of three monomers of CynDpum based on the structure of nitrilase from Synechocystis sp. PCC6803 (Nit6803). The monomers are colored orange, purple, and brown, respectively. The predicted A-surface between the orange and purple monomers is highlighted in yellow and corresponds to residues 165–174 and 193–201 in CynD. Two insertions in the nitrilase sequences, relative to other members of the superfamily, are candidates for the interacting regions of the C-surface. Region 1, colored red, corresponds to residues 55–72 and interacts in Nit6803. Region 2, colored green, corresponds to residues 222–235. This region does not interact in Nit6803 but does interact in Drosophila melanogaster β-alanine synthase. The C-terminal tails were not present in the structure of Nit6803 and are not modeled. The N- and truncated C-termini are indicated in the orange monomer (color figure online)

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truncated at this auto-cleavage site, results in spontaneous and active extended oligomers, removing the need for activation by benzonitrile (Thuku et al. 2007). However, this substrateinduced auto-catalytic cleavage, assembly and activation is unique (to date) for the J1 nitrilase. CynD from B. pumilus normally forms 18 subunit spirals at neutral pH. When the pH is lowered to 5.4, the oligomer becomes extended into long non-terminating spirals; this structural change was also associated with a slight increase in cyanide-degrading activity. It was proposed that this increase in activity results from activation of the terminal subunits of the 18 mers when they associate into the longer spirals (Jandhyala et al. 2003, 2005); in other words, there is a smaller percentage of inactive terminal monomers when the protein is assembled into longer spirals rather than shorter ones. Supporting this suggestion, mutations in CynD isolated for enhanced tolerance to high pH were also found to form long spirals at neutral pH (Wang et al. 2012). Recently, we have identified a mutation R67C that disrupts complete oligomerization in CynDpum; these incomplete CynDpumR67C oligomers are unable to catalyze the degradation of cyanide (Park et al. 2016), further supporting the hypothesis that formation of complete oligomers is necessary for activity. Oligomeric surfaces Fig. 3 A three-dimensional reconstruction of a CynDpum created by cryo-electron microscopy (Sewell and Watermeyer, unpublished). The reconstruction has been contoured at a level that encloses the atoms of a model based on the structure of Nit6803 and segmented to show how the monomers fit together to form the fiber. The diameter of the helical fiber is 13 nm as indicated by the two vertical lines

the recently crystallized Nit6803 nitrilase (Zhang et al. 2014) reflects the spiral oligomer seen in all nitrilases. Further analysis and higher resolution structures will increase our understanding of this distinct nitrilase spiral and enable rational engineering of these enzymes. Spiral formation and nitrilase function In all nitrilases that have been examined, the formation of the spiral oligomer is obligatory for activity. The positioning of a catalytic glutamate in the active site upon oligomerization has been proposed as the activating event (Williamson et al. 2010). The nitrilase from R. rhodochrous J1 normally purifies as a dimer; however, addition of the substrate benzonitrile triggers the enzyme to both oligomerize into C-shaped or variable length spirals and become active. Following activation by benzonitrile, the substrate specificity of the J1 nitrilase includes additional substrates that themselves cannot trigger activation (Nagasawa et al. 2000). The C-shaped oligomers of the J1 nitrilase also associate into spiral structures following a spontaneous autocleavage of their C-terminus. Expression of the J1 nitrilase,

Since active nitrilases form spiral structures, it is clearly important to understand the nature of the interactions that lead to the formation of these spirals. Three-dimensional reconstructions of nitrilases from electron microscopy have been of sufficient clarity to enable the docking of dimeric models based on the crystal structures of homologs (Dent et al. 2009; Sewell et al. 2003; Thuku et al. 2007; Williamson et al. 2010). Figure 4 shows how, in the case of the cyanide dihydratase from P. stutzeri, the elongation of the spiral is the result of interactions in two major regions that were, respectively, named the A- and C-surfaces (Thuku et al. 2009). The Asurface is a conserved intermonomer association in almost all structures of the nitrilase superfamily members described to date, and thus, the spirals were thought of as being made by the association of such dimeric building blocks. The primary interface between adjacent dimers is the C-surface (Sewell et al. 2003), but the details of this interaction remain obscure because there are no high-resolution structures of spiral nitrilases. Other interactions across the groove between adjacent turns in the spiral have also been identified from the reconstructions. Details of these interactions vary between different nitrilases, and they have been named the D-, E-, and F-surfaces. The D- and/or F-surfaces are symmetric interactions seen in 3D reconstructions of both CynD and CHT, as well as related nitrilases (Dent et al. 2009; Sewell et al. 2003, 2005;

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the nitrilase CynD from B. pumilus and P. stutzeri, mutations in the predicted A-surface helices disrupted activity (Sewell et al. 2005), possibly through this interaction preventing proper placement of the active site cysteine. Another region thought to participate in the A-surface is the protein C-terminus. In some structures of nitrilase superfamily members, the C-terminus interacts with and strengthens the Asurface (Kimani et al. 2007; Lundgren et al. 2008). Surprisingly, nitrilase C-termini are highly divergent and are longer than many other superfamily members; additionally, the sequence and secondary structure characteristics of Cterminal regions vary greatly even among closely related nitrilases (Sewell et al. 2005). Further evidence pointing to the role of intramolecular interactions involving the Cterminal region has also been presented by Crum et al. (2015a).

Fig. 4 A model of CynDstut comprising 14 monomers fitting in the threedimensional reconstruction based on electron microscopy in negative stain. Each monomer is differently colored. The C-terminal tail and the two C-surface regions are not included in this model (Sewell et al. 2003). The A-surface that interfaces the two central monomers and an adjacent C-surface interface of adjacent dimers are indicated

Thuku et al. 2007; Woodward et al. 2008). They are thought to stabilize the spiral and are attractive targets for manipulations that lead to enhanced stability. In the nitrilase CynD from B. pumilus and P. stutzeri, an asymmetric interaction is seen between the dimer at the terminal end of the spiral and the previous turn. This interaction is termed the E-surface and is thought to be responsible for terminating the spiral by altering the structure in such a way that additional dimers can no longer be added (Sewell et al. 2003, 2005). A-surface The A-surface is formed by two α-helices on each monomer which interact to form the αββα-αββα pair (Fig. 2). The sequence and pattern of interactions across this interface varies, but the A-surface structure is seen in all superfamily crystal structures as the basis for dimer formation (Gordon et al. 2013; Kimani et al. 2007; Kumaran et al. 2003; Wang et al. 2001). The conservation of dimer formation at the A-surface may stem from its possible role in positioning the active site cysteine, as seen in the crystal structure of the putative CN hydrolase from yeast (Kumaran et al. 2003) thereby explaining the role of oligomerization in enzyme activation. In

C-surface The C-surface is the primary interaction between dimers in the formation of the larger spiral structure of nitrilases. This surface can be seen as the interaction between adjacent dimers in the fitted dimer models (Dent et al. 2009; Sewell et al. 2003; Thuku et al. 2007) (Fig. 4) showing that the C-surface lies perpendicular to the A-surface. Together, these two surfaces establish the shape of the spiral structure. The angle across the C-surface varies among nitrilases and determines the degree of twist in the spiral. The C-surface involves the participation of two regions in the nitrilases, C-surface regions 1 and 2. When sequences of nitrilases are aligned with homologs of known structure, these two regions stand out as insertions that are missing in sequences of proteins outside of the nitrilase branch of the superfamily (Park et al. 2016; Sewell et al. 2005). The C-surface has been visualized in two crystal structures, the Synechocystis sp. PCC6803 Nit6803 nitrilase and the Drosophila melanogaster β-alanine synthase (βaS) from the larger nitrilase superfamily. In the structure of Nit6803, the two relative insertion regions 1 and 2 participate heavily, along with a third region, in the C-surface interaction (Zhang et al. 2014). The βaS resembles the C-shaped oligomers of the J1 and G. pallidus nitrilases and was the first in which a C-surface interaction could be viewed at atomic resolution. The βaS structure contains only one of the relative insertion regions, which is similar to the nitrilase C-surface region 2 (Lundgren et al. 2008). D- and F-surfaces Symmetric cross-spiral interactions vary between CynD and CHT but are seen in 3D reconstructions as contacts between consecutive turns of the nitrilase spiral; these contacts comprise the D- and F-surfaces. The CynD spirals from both B. pumilus and P. stutzeri are more tightly wound (Sewell et al. 2005; Thuku et al. 2007) than the CHTs from G. sorghi and N. crassa (Dent et al. 2009; Woodward et al. 2008). A mixture of charged residues on the D-surface in

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CynD and the J1 nitrilase suggests that salt bridging across this surface could help stabilize the spiral (Sewell et al. 2005; Thuku et al. 2007). However, mutations removing predicted salt bridging did not noticeably alter activity in CynD (Sewell et al. 2005), although the mutation Q86R, which is predicted to add a salt bridge at the D-surface, increased CynD’s thermal stability and tolerance of higher pH (Wang et al. 2012). E-surface A curious feature of CynD from B. pumilus and P. stutzeri is the spirals that are of defined (but different) length at neutral pH, implying that termination of oligomerization limits the molecular size. In these enzymes, an asymmetric, Bacross the groove^ interaction is seen between the terminal dimer and the previous turn of the spiral. This interaction at the E-surface was proposed to be responsible to the selftermination of CynD. Variations in E-surface interactions are seen between CynD species, CynDpum terminates at 18 subunits (Jandhyala et al. 2003) and CynDstut at 14 subunits (Sewell et al. 2003). As mentioned before, changes in pH or mutations in CynDpum can disrupt this E-surface and selftermination (Jandhyala et al. 2005; Wang et al. 2012). These changes are also associated with improvements in activity and stability. Manipulating the E-surface interactions may enable us to modulate the enzymes size, activity, and thermostability. C-terminal tail The highly variable C-terminus in the nitrilases has been of particular interest for its role in oligomerization and stability. The oligomeric nitrilases have a Cterminus that is 40–100 amino acids longer relative to many of the nitrilases with known structures. Furthermore, among cyanide-degrading nitrilases, the sequence of this region is highly variable even between closely related enzymes (Thuku et al. 2009) such as two different alleles of B. pumilus (Jandhyala et al. 2003). The C-terminus has been predicted to participate in oligomerization, but its role is likely to vary given its heterogeneity. In the 3D reconstruction of CynDstut, the C-terminus is predicted to lie in the center of the spiral where it could potentially make multiple contacts (Sewell et al. 2003), but this has not been directly visualized at sufficiently high resolution to enable an interpretation. C-terminal extensions in other superfamily members suggest possible roles for this region in nitrilase oligomerization. The crystal structure of the amidase from G. pallidus has the C-terminus interlocking and strengthening the A-surface in the dimer (Kimani et al. 2007; Lundgren et al. 2008). Modeling of the fungal cyanide hydratase from N. crassa, based on the amidase structure, places part of the C-terminal region at both the A- and C-surfaces (Dent et al. 2009). In βaS from D. melanogaster, the tail folds in the center of the Cshaped spiral where it contributes to both the A- and Csurfaces (Lundgren et al. 2008). The tail region was missing completely in the crystal structure of Nit6803, so the details of its role in the true nitrilases remain to be determined.

Modifications to the C-terminal tail have a variety of effects on spiral length. In the R. rhodochrous J1 nitrilase, the extended C-terminus prevents oligomerization. An auto-lytic cleavage removes the last 39 amino acids of the C-terminus. This allows the protein to oligomerize into the active spiral structure. Expression of this protein with shorter or longer truncations resulted in short, poorly formed oligomers, leading to the hypothesis that the position of the C-terminus in the center of the spiral allows it to influence the shape of the spiral and the alignment of the cross-spiral D-surface (Thuku et al. 2007). Consistent with this is the pH dependent change in spiral size of CynDpum. When the pH is lowered to 5.4, CynDpum shifts from 18 subunit spirals terminating at the E-surface to long variable length spirals. This behavior is thought to be driven by one or more histidines that are located at an interface, possibly the C-surface or in the C-terminal tail, gaining a charge. It is postulated that this changes the conformation of this region and alters the shape of the spiral, preventing the Esurface from aligning (Jandhyala et al. 2005). Another CynDpum variant (8A3) that lacks these histidines in the tail does not show this pH transition (Jandhyala et al. 2003). Truncations of the C-terminus are tolerated to varying degree. C-terminal truncations in CynD show different levels of tolerance between the B. pumilus and P. stutzeri enzymes. CynDpum was able to tolerate large deletions up to 38 residues and retain activity. CynDstut, by comparison, was inactive with nearly every truncation. CHT from N. crassa was intermediate and able to tolerate truncations of 29 amino acids and retain partial activity. Engineering of CynD The use of nitrilases has significant potential to be less costly or environmentally hazardous than conventional chemical strategies; nonetheless, improvements to the enzyme would make them easier to adapt for use in industry. Their tolerance to industrial conditions, substrate specificity, specific activity, and overall stability are often less than ideal (Gong et al. 2012). Protein engineering therefore could lead to improved nitrilase properties. Before the recent crystallization of Nit6803, no crystal structure was available for any member of the nitrilase branch of the superfamily. The gap in structural understanding has been a significant obstacle to rational engineering of these enzymes. This hurdle has been partially circumvented by modeling which was further informed by the results of random mutagenesis and high throughput screens (Abou Nader 2012; Crum et al. 2015a, b, 2016; Wang et al. 2012). Random mutagenesis The largest obstacle to using CynD and CHT to degrade cyanide wastes has been an inability to tolerate the pH at which cyanide-contaminated solutions are generally found. Cyanide is kept at very alkaline conditions to

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prevent the formation of hydrogen cyanide gas. Directed evolution using error-prone-PCR together with high throughput screening under alkaline conditions yielded mutants with improved alkaline tolerance in CynD from B. pumilus C1 (Wang et al. 2012). From a library of over 2000 highly mutated CynDpum clones expressed in E. coli, two alleles, C5 (Q86R, E96G, D254E) and H7 (E35K, Q322R, E327G), were identified that degrade cyanide at elevated pH. In C5, all three mutated residues were found to contribute to the pH tolerance of CynD, both in vivo and in vitro, with Q86R having the largest impact. H7 on the other hand gained its pH tolerance from E327G alone (Wang et al. 2012). These mutants were also found to have enhanced thermal stability. The C5 and the C5 + E327G mutants retained 60% activity after 8 h at 42 °C, while the wild-type enzyme only retained 20% activity. When the purified mutant enzymes were examined by electron microscopy, both C5 and E327G were seen to preserve oligomer formation at pH 9, a condition where the wild type only forms short incomplete oligomers. Q86R was also found to prevent self-termination, resulting in long extended oligomers similar to those seen in CynDpum at pH 5.6. The Q86 position is modeled to participate in the D-surface interaction (J. Watermeyer, unpublished results). The Q86R mutation may introduce additional interactions, which could explain the stabilizing phenotype. The E327G mutation on the other hand lies in the Cterminus, which could be influencing multiple oligomeric surfaces (Wang et al. 2012). Another characteristic of CynD that has been targeted for improvement has been the catalytic rate. Screening a different library generated by error-prone PCR for mutants was able to degrade cyanide samples within shortened reaction times. Three mutant alleles CD12 (E327K), DD3(K93R), and 7G8(D172N, A202T) were found to have higher catalytic activity. This improvement primarily stemmed from increased stability, similar to the pH tolerance mutants. The single mutants E327K, K93R, and D172N all had increased thermal stability at 42 °C with D172N having the most dramatic effect. Combining these three stabilizing mutations yielded further improvements in stability, the K93R-D172N-E327K triple mutant retained close to 60% activity after 8 h at 42 °C. In contrast to the alkaline tolerant mutants, neither K93R, D172N, nor E327K had increased activity at elevated pH. In fact, D172N actually lowered the pH optimum of the enzyme (Crum et al. 2016). These mutations are located on the D-surface and C-terminus. K93 is predicted to lie on the either the D- or E-surfaces where it could stabilize the spiral structure or alter the oligomer length; however, this has not yet been tested. The residue E327 in the C-terminus is clearly a critical position, two independent mutants E327G and E327K were found having an effect on stability. While both E327G and E327K improve thermal stability, they behave differently under alkaline conditions with only E327G improving activity (Wang et al. 2012).

D172N is modeled to be at the A-surface interaction and may participate in dimer formation. A highly destabilizing mutation, A202T, is similarly location to the A-surface, suggesting that the A-surface could be a productive target for further modification. C-terminal hybrids As described earlier, truncations in the C-terminus have highly variable phenotypes. C-terminal truncations in CynD from B. pumilus are tolerated whereas those in P. stutzeri are not. Genetic exchange that switched the Cterminal tails between CynDpum and CynDstut was also tolerated in the CynDpum enzyme body (CynDpum-stut), but the reverse hybrid (CynDstut-pum) was inactive (Sewell et al. 2005). CHT from N. crassa is able to tolerate truncations of 29 amino acids and retain partial activity; however, substitution for either of the CynD C-termini abolishes activity. CynDpum is able to tolerate the CHT C-terminus, but with reduced activity (Crum et al. 2015b). Further emphasizing this one sided compatibility, the CynDpum-stut hybrid was found to be significantly more thermostable and have greater tolerance to alkaline conditions than either parent CynD or any mutant CynD identified to date (Crum et al. 2015b). The region partly responsible for the incompatibility of the reverse CynDstut-pum was narrowed to six residues from the CynDpum C-terminus. When these residues were changed back to the native P. stutzeri sequence in the CynDstut-pum hybrid (CynDstut-pumGERDST), partial activity was restored. Furthermore, restoring only the glycine and glutamate was sufficient to restore about 10% of wild-type activity (Crum et al. 2015a). Because the C-terminus is seen to interact with both the Aand C-surfaces in crystal structures from nitrilase homologs, the inactive CynDstut-pum hybrid was examined in combination with substitutions in the two A-surface and the two C-surface regions for the CynDpum. Among these double hybrids, Asurface region 2 (195–206) was restored partial activity to CynDstut-pum that was otherwise inactive but did not remove the need for the C-terminus. Furthermore, the CynDpum Asurface region 2 slightly reduced activity when put in CynDstut alone indicating that the effect was not only increased stability. Together, these results indicate a specific interaction between the C-terminus and the A-surface of CynDpum (Crum et al. 2015a).

Conclusions Cyanide-degrading nitrilases CynD and CHT remain the most attractive candidates for enzymatic remediation of cyanideladen wastewaters. They are distinguished from other enzymatic alternatives by their simple reaction mechanism, requiring no supplemented substrates or co-factors, and by their

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independence from metabolically active cells. The evidence suggests a correlation between enzyme activity and the formation of the spiral oligomeric form, to the extent that stabilizing the spiral leads to improvement in the enzyme characteristics. The primary drawback of the cyanide dihydratases, namely, low tolerance to the alkaline conditions normally found in cyanide contaminated aqueous sites, has been notably improved in mutants and hybrid enzymes and could almost certainly be improved further using a strategy based on more detailed knowledge of the structure of the enzyme. The hybrid CynDpum carrying the heterologous C-terminus from P. stutzeri (CynDpum-stut) (Crum et al. 2015b) could well serve as a baseline for optimization of this enzyme. The residues forming the oligomeric surfaces are attractive targets for future manipulation because of the apparent link among nitrilase oligomerization, stability, and catalysis. Apart from engineering these proteins genetically, strategies such as chemical crosslinking, immobilization, or protection within alkaline bacteria may prove useful to achieving a product ready for direct application. Compliance with ethical standards Funding This work was supported by the Welch Foundation (A1310), the Texas Hazardous Waste Research Center (513TAM0032H), and the National Research Foundation of South Africa. Conflict of interest The authors declare that they have no conflicts of interest. Ethical approval This article does not contain any studies with human participants or animals performed by any of the authors.

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