There are thousands of areas in the U.S.A. and Europe contaminated with cyanide-containing wastes as a consequence of a large number of industrial activities such as gold mining, steel and aluminium manufacturing, electroplating and nitrile pesticides used in agriculture. Chemical treatments to remove cyanide are expensive and generate other toxic products. By contrast, cyanide biodegradation constitutes an appropriate alternative treatment. In the present review we provide an overview of how cells deal in the presence of the poison cyanide that irreversible binds to metals causing, among other things, iron-deprivation conditions outside the cell and metalloenzymes inhibition inside the cell. In this sense, several systems must be present in a cyanotrophic organism, including a siderophore-based acquisition mechanism, a cyanide-insensitive respiratory system and a cyanide degradation/assimilation pathway. The alkaliphilic autochthonous bacterium Pseudomonas pseudocaligenes CECT5344 presents all these requirements with the production of siderophores, a cyanide-insensitive bd-related cytochrome [Cio (cyanide-insensitive oxidase)] and a cyanide assimilation pathway that generates ammonium, which is further incorporated into organic nitrogen.

Chemical forms of cyanide and their toxicity

Cyanide is an ancient molecule that might be involved in the prebiotic synthesis of different nitrogenous compounds, including amino acids and nitrogenous bases [1]. Cyanides include a type of chemicals that present the cyano (−C≡N) group and they can be found in Nature in many different forms owing to the chemical properties of this group. Depending on the pH, cyanide can be found as the ion cyanide (CN) in dissolution at high pH or may evaporate as HCN (cyanhydric acid) at neutral or acid pH values (pKa 9.2). Cyanide shows a high affinity for metals, therefore complexes of cyanide and transition metals that are often very stable are frequently found in Nature. Among these metal–cyanide complexes, two types have been described: the complexes with nickel, copper or zinc are called WAD (weakly acid-dissociable), whereas the strong complexes with iron and cobalt are named SAD (strong acid-dissociable), and, in general, show dissociation constants within the range 10−17−10−52 M [2,3]. SCN (thiocyanate) results from the interaction of free cyanide and reduced sulfur forms present in ores such as pyrite and pyrrothite [4]. Nitriles (R–C≡N) are organic cyanides. Higher plants are the main producers of these compounds as cyanolipids or cyanoglucosides in their secondary metabolism [5,6]. Cyanide is highly toxic to most living organisms because it strongly binds to metalloproteins [7]. HCN is the most toxic form described for cyanide, followed by ion cyanide found in dissolution. The toxicity of metal–cyanide complexes and nitriles depend on their capacity to release free cyanide [3,8].

Cyanide is generated as a natural compound by some bacteria, algae, fungi, higher plants and even by insects, mainly for defensive or offensive purposes [6,914]. However, environmental problems caused by cyanide are due to the production of large amounts of cyanide-containing wastes by human industrial activities, including synthesis of organic nitriles, nylon, acrylic plastics, paints, dyes, drugs and chelating agents. Additionally, cyanide is used in gold mining and the metal and jewellery industries, which generate waste with high cyanide concentrations. The cyanidation process used for gold extraction in mining uses more than 20% of worldwide cyanide production [15]. During the cyanidation process, leaching of cyanide generates a great variety of metal–cyanide complexes.

Chemical treatments to remove cyanide are very expensive and they generate products that are also toxic, therefore cyanide biodegradation may be an attractive alternative treatment [1618]. In spite of cyanide toxicity, some micro-organisms are able to grow with cyanide as a nitrogen source [17,18]. A bacterial strain able to degrade cyanide under alkaline conditions was isolated from sludges of the Guadalquivir River (Córdoba, Spain). This bacterium, identified as Pseudomonas pseudoalcaligenes CECT5344, grows at an optimum pH of 9.5 and, in addition to cyanide, this strain uses cyano–metal complexes, cyanate, 3-cyanoalanine, cyanoacetamide, nitroferricyanide and a residue from the jewellery industry as a nitrogen source [1921]. The CECT5344 strain is an ideal cyanotroph because it is alkaliphilic, prevents cyanide volatilization as HCN, produces siderophores for iron acquisition [22,23], induces a cyanide-insensitive respiration chain [24] and contains enzymes suitable for cyanide degradation [20,22]. In this bacterium, cyanide also induces several responses related to iron deprivation, nitrogen starvation and oxidative stress [25] (Figure 1).

Ps. pseudoalcaligenes CECT5344 responds to cyanide by the induction of a complex enzyme machinery

Figure 1
Ps. pseudoalcaligenes CECT5344 responds to cyanide by the induction of a complex enzyme machinery

Cyanide induces several mechanisms of resistance that are based on the Cio, which is related to the cytochrome bd oxidase from enterobacteria, and enzymes involved in protection against reactive oxygen species (ROS) such as an alkyl hydroperoxide reductase (AHP), a ferritin-like protein (FLP) and a heat-shock protein (HSP). Cells respond to iron-limitation in cyanide-containing medium by releasing siderophores (SDF) to this medium. In the strain CECT5344, enzymes that could be related with cyanide degradation are also induced such as the cyanase (CynS) and a nitrilase. Additionally, cyanide causes nitrogen starvation as suggested by the induction of the regulatory protein PII-2. NDH, NADH:ubiquinone oxidoreductase (complex I); SDH, succinate dehydrogenase; UQ, ubiquinol/ubiquinone pool.

Figure 1
Ps. pseudoalcaligenes CECT5344 responds to cyanide by the induction of a complex enzyme machinery

Cyanide induces several mechanisms of resistance that are based on the Cio, which is related to the cytochrome bd oxidase from enterobacteria, and enzymes involved in protection against reactive oxygen species (ROS) such as an alkyl hydroperoxide reductase (AHP), a ferritin-like protein (FLP) and a heat-shock protein (HSP). Cells respond to iron-limitation in cyanide-containing medium by releasing siderophores (SDF) to this medium. In the strain CECT5344, enzymes that could be related with cyanide degradation are also induced such as the cyanase (CynS) and a nitrilase. Additionally, cyanide causes nitrogen starvation as suggested by the induction of the regulatory protein PII-2. NDH, NADH:ubiquinone oxidoreductase (complex I); SDH, succinate dehydrogenase; UQ, ubiquinol/ubiquinone pool.

Siderophores are required for growth with cyanide

Iron is required in many biological systems for redox reactions and structural functions. However, iron solubility at neutral pH in aerobic environments is very scarce, and, consequently, micro-organisms produce siderophores to survive. Siderophores are molecules, usually with a molecular mass of 500–1000 Da, which are specialized in Fe3+-uptake mechanisms [26]. To assimilate iron from the environment many micro-organisms, including bacteria, yeast and fungi, solubilize iron from insoluble precipitates that are a part of hydroxide polymers. Pathogenic organisms have to remove iron from iron-containing proteins such as the blood haem proteins transferrin and lactoferrin. Siderophores are also produced by a great variety of higher plants [27]. More than 500 types of siderophores have been identified in bacteria, and some of them are able to produce more than one type of siderophore. For example, Escherichia coli is able to produce the 669 Da catecholate-type siderophore enterobactin and the 740 Da hydroxamate-type siderophore ferrichrome.

Many siderophores have been studied extensively and their biosynthetic pathways described. Thus the enterobactin isolated from culture supernatants of Salmonella enterica serovar Typhimurium and E. coli is synthesized from 2,3-dihydroxybenzoic acid and L-serine. Vibrio cholerae can use iron in the haem group and haemoglobin. This bacterium produces vibriobactin, a siderophore synthesized from 2,3-dihydroxybenzoate derived from chorismate and L-threonine [26]. Pyochelin and pyoverdin are siderophores found in the opportunistic pathogen Pseudomonas aeruginosa. Pyochelin is generated from salicylic acid, cysteine and S-adenosylmethionine with the consumption of ATP and NADPH. The chromophore from pyoverdin seems to have phenylalanine and/or tyrosine as precursors [28]. As indicated above, the biosynthetic pathways of siderophores are diverse, although all siderophores can be divided into two groups: the catecholate-type siderophores are fluorescent, whereas the hydroxamate-type siderophores are non-fluorescent [22]. In addition, there are many other compounds which combine these types of siderophores or present other chelating compounds such as hydroxy acids [28]. The synthesis of some siderophores is performed by enzymes known as NRPSs (non-ribosomal peptide synthetases), which are multimodular and produce peptide products of a specific sequence without an RNA template [26].

Ferric siderophores must be transported inside the cells through outer membrane receptors, which can be classified into two general types depending on the structure of the siderophore molecule, the catecholate and the hydroxamate types [29]. Usually, these transporters are of the ATP-dependent ABC (ATP-binding cassette) type [30]. The uptake of Fe(III)-ferrichrome transport is an example of ferri-siderophore transport in Gram-negative bacteria. After ferrichrome binds to the receptor, FhuA interacts with the periplasmic domain of TonB, facilitating the force proton motive-driven active transport of ferrichrome to the periplasm. The ferri-siderophore binds to the periplasmic protein FhuD, which binds a wide range of hydroxamates, before interacting with the cytoplasmic permease FhuBC. In the cytoplasm, the complex is reduced and the ferrous ion is removed and chelated by an acceptor molecule [30].

Under iron-rich conditions, synthesis and transport of siderophores are repressed. Transcription of the iron-controlled genes is negatively regulated through the interaction of the Fe2+-binding repressor, the Fur protein, with the operator sequences within the promoter regions of the operons [3133].

Micro-organisms capable of growing with cyanide require the production of siderophores to chelate iron because cyanide bonds very strongly to this transition metal [22]. As mentioned above, the alkaliphilic bacterium Ps. pseudoalcaligenes CECT5344 is able to grow with either free cyanide or metal–cyanide complexes as the sole nitrogen source under alkaline conditions [19,20]. The production of siderophores by the CECT5344 strain (Figure 1) was previously demonstrated by using CAS (chrome azurol S) agar plates [19,20].

Bacterial cyanide-insensitive respiration

E. coli shows two terminal UQO, (ubiquinol oxidases) in the aerobic electron transfer chain, the bo-type UQO, which is expressed under high oxygen tension, and the bd-type UQO, which, by contrast, predominates under low oxygen conditions [34]. The E. coli bd-type UQO consists of two polypeptide chains, the 58 kDa subunit I and the 43 kDa subunit II. The cytochrome bd-type is encoded by the cydA and cydB genes. This enzyme presents three types of cytochrome haem species based on their optical spectra, the cytochromes d, b558 and b595. The cytochrome d has a chlorine chromophore with a maximum absorbance at 628 nm in its reduced form. Also the dioxygen molecule forms a very stable adduct with ferrous cytochrome d showing a band at 568 nm. The cytochrome b558, the ubiquinol-binding site, shows in its reduced form α and β bands at 562 and 532 nm respectively. The cytochromes d and b595 constitute a haem–haem binuclear centre different from that of the haem–copper terminal oxidase. Cyanide is able to interact with this haem–haem binuclear centre in a particular way that could explain why the bd-type UQO is more resistant to cyanide (up to 1 mM cyanide) than bo-type UQO [34]. In Ps. aeruginosa, several terminal oxidases have been described [35,36]. The cytochrome co oxidase is very sensitive to low cyanide concentrations, whereas the cytochrome bd-related oxidase named Cio (cyanide-insensitive oxidase), which is encoded by the cioA and cioB genes, is cyanide-insensitive [24]. A third type of oxidase is the cytochrome baa3, which has been described in a strain of Ps. aeruginosa, although it is not found in all strains of Ps. aeruginosa. It is a quinol oxidase more resistant to cyanide than the cytochrome co oxidase but it is still sensitive to this compound in the micromolar range [35]. As in E. coli, the Cio terminal oxidase is expressed in Ps. aeruginosa under low oxygen tension because this UQO has more affinity for oxygen than the cytochrome co oxidase. Additionally, its cyanide insensitivity could lead to bacterial strains able to survive in the presence of this toxic chemical. In fact, there are several Pseudomonas strains are capable of synthesizing hydrogen cyanide as a metabolic product at a concentration large enough to inhibit the cytochrome co oxidase present in different bacterial strains [35]. Alteration of the cyanide-insensitive terminal oxidase in Ps. aeruginosa has been reported to lead to cell division defects and multiple antibiotic sensitivity [36]. The transcriptional activator RoxR binds to the promoter region of the cioAB genes and seems to participate with the sensor kinase RoxS in the regulation of the transcription of loci that play a role in energy generation in Ps. aeruginosa [37]. Alcaligenes eutrophus H16 shows a cyanide-insensitive cytochrome bo that is required for energy transduction. Thus 80% of the proton electrochemical gradient and ATP are formed in membranes during H2 oxidation in autotrophically grown cells derived from the cytochrome bo in association with a membrane-bound hydrogenase [38]. In Achromobacter, two terminal oxidases have been described, the cyanide-sensitive cytochrome o and the cytochrome a2 that has low affinity for cyanide. In this case, both terminal oxidases coexist under low oxygen conditions [39]. The facultative alkaliphilic Bacillus YN-2000 is able to grow in the presence of cyanide and presents a respiratory oxygen-reducing system composed of a non-proteinaceous component with a molecular mass of 662 Da and a catalase [40]. The cyanotroph Ps. pseudoalcaligenes CECT5344 is able to grow with 2 mM cyanide but tolerates up to 30 mM cyanide [19] and in the presence of this compound the bacterium induces the cio genes involved in the cyanide-insensitive respiration (Figure 1). Mutation of the cioA gene leads to an intolerance to cyanide [24].

Cyanide degradation pathways: enzymes and bacterial genomes distribution

Different cyanide degradation pathways based on hydrolytic, oxidative and substitution/addition reactions have been described in some micro-organisms [17,18,41]. In Pseudomonas fluorescens NCIMB 11764, it has been described that the hydrolysis of cyanide produces formic acid and ammonium [42]. The oxidative degradation of cyanide occurs either in a single step to produce CO2 and NH4+ or in two steps with the formation of cyanate as intermediate, which is further converted into CO2 and ammonium by the cyanase [18,43]. However, no link between cyanide and cyanate degradation pathways has been demonstrated. The substitution/addition reactions produce either thiocyanate from cyanide and S2O32–, catalysed by the rhodanese [17] or 3-cyanoalanine from cyanide and serine or cysteine, catalysed by a 3-cyanoalanine synthase [4446]. The 3-cyanoalanine can be further degraded via a nitrilase activity to produce aspartate and ammonia or through a nitrile hydratase to produce asparagine [47]. Cyanohydrins (2-hydroxynitriles) are putative substrates for nitrilases or nitrile hydratases [48,49]. Degradation of these nitriles can be performed either in a single step by a nitrilase to produce the acid form and ammonia or in two steps catalysed by a nitrile hydratase that generates the corresponding amide that can be converted further into the acid form and ammonium by an amidase [4850]. Two types of nitrile hydratases have been described based on their metal cofactor, which may be either iron or cobalt [51]. Cyanide has been described as an effective inhibitor of nitrile hydratases because it binds to their metal cofactors, although several nitrile hydratases from pseudomonads are cyanide-resistant [48]. Several loci involved in cyanide degradation are shown (Figure 2).

Schematic organization of bacterial gene clusters involved in cyanide degradation

Figure 2
Schematic organization of bacterial gene clusters involved in cyanide degradation

Gene annotation: rhdA and moeZ (rhodanese), motA (flagellar motor protein), PA4955 (HDOD domain-containing protein), psd (phosphatidylserine decarboxylase), Avin07470 (exonuclease), Avin07460 (ribosome-associated GTPase), lysS (putative lysyl-tRNA synthetase protein), NE2354 (hypothetical protein), NE2352 (Mov34 family protein), CV3563 (conserved hypothetical protein), CV3562 (probable ion transporter), cysK (cysteine synthase), CV3560 (conserved hypothetical protein), CV3559 (conserved hypothetical protein), CV3558 (probable acetylpolyamine aminohydrolase), coaX (pantothenate kinase), hslO (chaperonin), pabB (p-aminobenzoate synthetase component I), pabA (p-aminobenzoate synthetase component II), nitA and nit (nitrilase), NHα and NHβ (subunits of nitrile hydratase), hyp (hypothetical protein), yqjI (6-phosphogluconate dehydrogenase), polY1 (DNA polymerase IV), oxaA2 (OxaA-like protein precursor), moaR (transcriptional activator), mdlA (mandelate racemase), mdlB (S-mandelate dehydrogenase), mdlC (benzoylformate decarboxylase), araC (transcriptional regulator), oxdA (aldoxime dehydratase), ami (amidase), PFLU3211 (cobalamine biosynthesis protein), SAM (radical S-adenosylmethionine superfamily member), GNAT (GCN5-related acetyltransferase), AIRs (AIR synthase), GT (glycosyltransferase) and FLAVO (flavoprotein).

Figure 2
Schematic organization of bacterial gene clusters involved in cyanide degradation

Gene annotation: rhdA and moeZ (rhodanese), motA (flagellar motor protein), PA4955 (HDOD domain-containing protein), psd (phosphatidylserine decarboxylase), Avin07470 (exonuclease), Avin07460 (ribosome-associated GTPase), lysS (putative lysyl-tRNA synthetase protein), NE2354 (hypothetical protein), NE2352 (Mov34 family protein), CV3563 (conserved hypothetical protein), CV3562 (probable ion transporter), cysK (cysteine synthase), CV3560 (conserved hypothetical protein), CV3559 (conserved hypothetical protein), CV3558 (probable acetylpolyamine aminohydrolase), coaX (pantothenate kinase), hslO (chaperonin), pabB (p-aminobenzoate synthetase component I), pabA (p-aminobenzoate synthetase component II), nitA and nit (nitrilase), NHα and NHβ (subunits of nitrile hydratase), hyp (hypothetical protein), yqjI (6-phosphogluconate dehydrogenase), polY1 (DNA polymerase IV), oxaA2 (OxaA-like protein precursor), moaR (transcriptional activator), mdlA (mandelate racemase), mdlB (S-mandelate dehydrogenase), mdlC (benzoylformate decarboxylase), araC (transcriptional regulator), oxdA (aldoxime dehydratase), ami (amidase), PFLU3211 (cobalamine biosynthesis protein), SAM (radical S-adenosylmethionine superfamily member), GNAT (GCN5-related acetyltransferase), AIRs (AIR synthase), GT (glycosyltransferase) and FLAVO (flavoprotein).

The cyanide degradation pathway in Ps. pseudoalcaligenes CECT5344 involves the specific formation of ammonium that is further incorporated into amino acids by the glutamine synthase [29]. The genome of the CECT5344 strain contains several nitrilases that could be involved in cyanide degradation and also a cyanase-encoded gene, cynS, but cyanate and cyanide in this bacterium are metabolized by independent degradation pathways [21].

Enzymology and Ecology of the Nitrogen Cycle: A Biochemical Society Focused Meeting held at University of Birmingham, U.K., 15–17 September 2010. Organized and Edited by Jeff Cole (University of Birmingham, U.K.), Rosa María Martínez-Espinosa (University of Alicante, Spain), David Richardson (University of East Anglia, Norwich, U.K.) and Nick Watmough (University of East Anglia, Norwich, U.K.).

Abbreviations

     
  • Cio

    cyanide-insensitive oxidase

  •  
  • UQO

    ubiquinol oxidase

We thank Gemasur, Saveco and Avenir for fruitful collaborations.

Funding

This work was funded by the Ministerio de Ciencia e Innovación [grant numbers BIO2008-04542-C02-01 and BIO2008-04542-C02-02] and PET_0048, Junta de Andalucía [grant number CVI1728], Junta de Extremadura [grant number PRI07A097 and Ayuda a grupos BioMic] and Fondo Europeo de Desarrollo (FEDER) from the European Union 2007–2013. V.L.-A. is recipient of a postdoctoral fellowship from the Ministerio de Ciencia e Innovación, Spain.

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