- Open Access
Biotechnological applications of archaeal enzymes from extreme environments
Biological Research volume 51, Article number: 37 (2018)
To date, many industrial processes are performed using chemical compounds, which are harmful to nature. An alternative to overcome this problem is biocatalysis, which uses whole cells or enzymes to carry out chemical reactions in an environmentally friendly manner. Enzymes can be used as biocatalyst in food and feed, pharmaceutical, textile, detergent and beverage industries, among others. Since industrial processes require harsh reaction conditions to be performed, these enzymes must possess several characteristics that make them suitable for this purpose. Currently the best option is to use enzymes from extremophilic microorganisms, particularly archaea because of their special characteristics, such as stability to elevated temperatures, extremes of pH, organic solvents, and high ionic strength. Extremozymes, are being used in biotechnological industry and improved through modern technologies, such as protein engineering for best performance. Despite the wide distribution of archaea, exist only few reports about these microorganisms isolated from Antarctica and very little is known about thermophilic or hyperthermophilic archaeal enzymes particularly from Antarctica. This review summarizes current knowledge of archaeal enzymes with biotechnological applications, including two extremozymes from Antarctic archaea with potential industrial use, which are being studied in our laboratory. Both enzymes have been discovered through conventional screening and genome sequencing, respectively.
Biotechnology is present everywhere and it has had a great impact on several industrial sectors, such as fine chemical and pharmaceutical industry, textile, and detergent industries, biofuel generation, bioremediation, among others. Generally, industrial processes use chemical compounds as catalysts, generating toxic byproducts . Biocatalysis has emerged as an alternative process using enzymes or cells as biocatalysts, which are more selective, efficient and environmentally friendly . Mesophilic enzymes have been used as biocatalysts but they have low stability at high temperature or extremes pH. For this reason, there is a considerable demand for more stable enzymes. One approach to overcome this need is to search for new enzymes within extremophilic microorganisms. Extremophiles are organisms that are able to thrive at extreme environmental conditions (temperature, pressure, salinity, dryness, radiation, pH or concentrations of heavy metals) (see Table 1). Most of the extremophiles belong to the Archaea domain. These microorganisms and their enzymes have unique characteristics [3, 4]. Archaea exist in a broad range of habitats, but there are a few reports of thermophilic or hyperthermophilic archaea from Antarctica . This is an extreme continent not only composed of cold environments but also of geothermal sites, such as fumaroles, hot springs, hydrothermal vents, and volcanoes. These sites are suitable for the development of these microorganisms.
Extremophilic archaea that live under extreme conditions have developed enzymes with unique structure–function properties. These enzymes, known as extremozymes, have an increased stability at high temperatures, extreme pH, in the presence of organic solvents and heavy metals and against proteolytic attack. For this reason, they are able to withstand harsh conditions during industrial processes and can be used in a diversity of biotechnological applications (see Table 2). To date, there are a variety of archaeal extremozymes, which are used as biocatalysts in different industrial sectors . In this work archaeal extremozymes with biotechnological applications and potential use will be reviewed, including an Antarctic extremozyme that has being developed in our laboratory.
Proteolytic enzymes (EC 3.4.x.x)
Proteases catalyze hydrolysis of proteins into smaller peptides or free amino acids. They are generally classified in two groups: endopeptidases (proteases that cleave peptide bonds within the protein) and exopeptidases (proteases that cleave off amino acids from the ends of the protein). Based on the mechanism of catalysis they are classified in serine proteases (EC 3.4.21.x), cysteine or thiol proteases (EC 3.4.22.x), aspartic, carboxyl or acidic proteases (EC 3.4.23.x), glutamic proteases (EC 3.4.23.x), threonine proteases (EC 3.4.25.x), and metalloproteases (EC 3.4.24.x) . Proteases are of great interest because of their versatile characteristics and different applications in industrial sectors. These enzymes represent a large percentage of the global enzyme market [12, 13]. Most proteases from extremophiles belong to the serine type and many of them come from hyperthermophilic archaea belonging to the genera Pyrococcus , Thermococcus , Desulfurococcus , Pyrobaculum , Staphylothermus , and from the thermoacidophilic archaeon Sulfolobus  (see Table 3).
In addition, there are also proteases derived from halophilic archaea belonging to the genera Haloferax , Halobacterium [21, 22], Natrinema , and Natronomonas . These enzymes are alkaline proteases, they work at elevated pH and some of them are stable at high temperatures or in organic solvents. For example, a protease from Haloferax lucentensis VKMM 007 showed maximal activity at 60 °C at pH 8 and it remains active in the presence of various polar and non-polar solvents, surfactants and reducing agents .
Biotechnological applications of proteases
In food and feed industry they are used to degrade complex proteins, predigest baby foods or soft meat. Since the latter process is carried out at 40–60 °C, thermostable proteases are mainly required for this purpose . In detergent industry they are used as additives in household laundry detergents to remove proteinaceous stains. In this industry, proteases have also been shown to resist denaturation by detergents and alkaline conditions. Thus, alkaline proteases from halophilic archaea are ideal for this purpose . In molecular biology they are used to remove proteinaceous contaminants of DNA in PCR prior to amplification. Therefore, thermostability to function in PCR is absolutely required. In peptide synthesis the process is carried out in low water/nonaqueous environments and peptides are used as precursors of sweeteners, such as aspartame. Thus, alkaline proteases resistant to organic solvents are required . Proteases can also help to reduce time during dough fermentation in bread industry and to modify mixtures containing high gluten content, through partial hydrolysis of the blend, making it soft and easy to pull and knead .
Esterases (EC 126.96.36.199) and lipases (EC 188.8.131.52)
Esterases and lipases are widely used as biocatalysts in biotechnology. Esterases (EC 184.108.40.206) hydrolyze water-soluble short acyl chain esters. On the other hand, lipases (EC 220.127.116.11) catalyze the hydrolysis of long-chain acylglycerols into glycerol and fatty acids. These enzymes display much broader substrate specificity than esterases. Esterases and lipases possess regio-, chemo-, and enantioselectivity and are stable in organic solvents. Thus, both types of enzymes are widely used in industrial processes performed in organic solvents [26, 27]. Many hyper/thermophilic esterases and lipases come from archaea belonging to the genera Pyrococcus , Pyrobaculum , Sulfolobus , Aeropyrum , and Archaeoglobus [32, 33]. These enzymes have also been reported from halophilic archaea belonging to the genera Haloarcula  and Halococcus  (see Table 4).
Biotechnological applications of esterases and lipases
Esterases and lipases are used in fine chemicals production (chemicals produced with purity higher than 90%) and pharmaceutical industry. They are used to improve the separation of numerous racemic mixtures of alcohols and acids, producing optically pure compounds, such as ibuprofen, ketoprofen and naproxen. These enzymes are used to obtain poly-unsaturated fatty acids (PUFAs) from plants and animal lipids, to produce pharmaceuticals [26, 36]. Lipases are also used as additives in detergents to remove oils and fats. Therefore, they improve washing capability of detergents and enhance removal of stringent stains, preventing scaling . In food and feed industry, lipases are used to modify the structure of some triglycerides for enhancing the flavor and physical and nutritional properties. They are also used in the ripening of cheese and in the production of human milk fat substitute and cocoa butter equivalents . Lipases are also used in pulp and paper production to remove the hydrophobic components of wood . They are also used in the synthesis of new biopolymeric materials, such as polyesters and polysaccharides, which are biodegradable and environmentally friendly . One of the current applications is in transesterification reactions of plant fats for biodiesel production . On the other hand, esterases are used to produce wine, fruit juices, beer, alcohol and flavoring and fragrance compounds present in cereals. In agrochemical industry these enzymes are used in the production of pesticides, insecticides, and nematicides . Lipases are also used in pulp and paper to remove the hydrophobic components of wood. But they are also used in the synthesis of new biopolymeric materials, such as polyesters and polysaccharides, which are biodegradable and environmentally friendly [38,39,40,41,42].
Glycosyl hydrolases (EC 3.2.1.x)
This large group of enzymes hydrolyzes glycosidic bonds between two or more carbohydrates or between carbohydrate and non-carbohydrate moieties. They degrade complex polysaccharides.
Starch degrading enzymes
The starch-degrading enzymes use as substrate starch, one of the largest renewable carbon sources in nature. Starch is a heterogeneous polysaccharide composed of the two polymers amylose (15–25%) and amylopectin (75–85%), both are high molecular weight components. Amylose and amylopectin are composed of α-d-glucose units, linked via α-1,4-glycosidic and α-1,6-glycosidic linkages respectively, forming the insoluble linear polymer amylose and the soluble branched polymer amylopectin. Because of the complex structure of starch, starch-processing requires a combination of enzymes, which depolimerize starch into oligosaccharides and smaller sugars (endoamylases and exoamylases) and enzymes to transform starch by transferring oligoglucosidic linkages and residues, creating new bonds (debranching enzymes and glycosyl-transferases) [10, 43]. Starch-degrading enzymes also represent a large percentage of the global enzyme market.
There is a need today for thermostable enzymes as starch processing is performed at high temperatures. In addition, these enzymes should be independent from calcium and metallic ions for their stabilization and catalytic activity . Thus, thermostable enzymes have the advantages of lowering the cost of sugar syrup production, consuming less energy.
Endohydrolases (or endoamylases)
α-amylases (EC 18.104.22.168) randomly cleave α-1,4 linkages on the inner part of starch and related substrates, producing branched and linear α-anomeric oligo- and polysaccharides of different sizes. There are thermostable α-amylases from plants, fungi, animals and microbes . Several of these enzymes come from hyperthermophilic archaea belonging to the genera Pyrococcus [45, 46], Thermococcus [47,48,49], Desulfurococcus , Staphylothermus , Methanococcus , and Sulfolobus . In addition, there are also α-amylases from haloalkaliphilic archaea belonging to the genera Haloarcula [53,54,55], Halorubrum , Haloferax , and Natronococcus  (see Table 5). α-amylases from haloalkaliphilic archaea are active at lower temperatures and higher pH than α-amylases from hyper/thermophilic archaea. For this reason, they are not suitable for starch industry, but they can be used in detergents for medium-temperature laundering, because of their stability in detergents and organic solvents.
Exohydrolases (or exoamylases)
β-amylases (EC 22.214.171.124) attack every alternate α-1,4-glucosidic linkage of the starch, producing the dimeric sugar β-maltose. These enzymes have been found to be distributed in higher plants, fungi, bacteria and only in one archaeon. The most thermostable β-amylase and the only one from an archaeon is PF0870, which comes from Pyrococcus furiosus (see Table 5). This enzyme has an optimal temperature of 110 °C, but it does not hydrolyze starch, glycogen, pullulan, or large maltooligosaccharides .
Glucoamylases (γ-amylases; EC 126.96.36.199) are exohydrolases that cleave α-1,4-glycosidic bonds from starch or related polysaccharides, releasing single β-d-glucose units from nonreducing ends. These enzymes also hydrolyze α-1,3- and α-1,6-glycosidic bonds in high molecular weight polysaccharides. Most of the glucoamylases reported come from fungi, but there are also in bacteria and thermoacidophilic archaea belonging to the genera Picrophilus [60, 61], Sulfolobus , Thermoplasma [60, 63], and from the methanogenic archaeon Methanococccus  (see Table 5). These archaeal glucoamylases are more thermostable than those from bacteria and fungi.
Another group of exohydrolases are α-Glucosidases (EC 188.8.131.52), which break every α-1,4- glycosidic bond from the terminal nonreducing end of starch or smaller polysaccharides produced by other starch-degrading enzymes. They prefer smaller oligosaccharides, such as maltotriose, and generate glucose . These enzymes are involved in the last step of starch degradation. Most of them come from bacteria and hyperthermophilic archaea belonging to the genera Pyrococcus [65, 66] and Thermococcus [67, 68]. In addition, there are α-glucosidases from the thermoacidophilic archaea Sulfolobus  and Picrophilus , and from the acidophilic archaeon Ferroplasma acidophilum strain Y  (see Table 5).
Starch debranching enzymes
Starch-debranching enzymes are important because of their biotechnological applications. These enzymes hydrolyze α-1,6-glycosidic bonds in amylopectin and/or glycogen and related polysaccharides. Pullulanases break down pullulan, a polysaccharide produced from starch by the fungus Aureobasidium pullulans. This molecule is a linear α-glucan consisting of maltotriose units joined by α-1,6-glycosidic linkages. Pullulanases are capable of hydrolyzing α-1,6 glucosidic bonds in pullulan and other branching polysaccharides, such as starch. Since the complete hydrolysis of starch can only be achieved in the presence of debranching enzymes, pullulanases are of great interest in starch industry. Based on the substrate specificity and reaction products, these enzymes are classified into three groups: pullulanases type I, pullulanases type II, and pullulan hydrolases (type I, II, and III) .
Pullulanases type I, exclusively hydrolyze the α -1,6 glycosidic linkages of pullulan. Are produced by K. pneumoniae, Bacteroides thetaiotaomicron, Bacillus sp. KSM-1876, T. aquaticus, alkaliphilic Bacillus sp. S-l, Micrococcus sp. Y-1 .
Pullulanases type II (amylopullulanases; EC 184.108.40.206) hydrolyze α-1,6-linkages in pullulan, producing maltotriose and also hydrolyze α-1,4-linkages in linear and branched oligosaccharides, such as amylose and amylopectin. Amylopullulanases are able to convert polysaccharides, such as amylopectin, into small sugars (e.g. glucose, maltose). These enzymes are important in starch processing industry due to their specific debranching capacity. They have been reported in bacteria and hyper/thermophilic archaea belonging to the genera Pyrococcus , Thermococcus [75, 76], Desulfurococcus , Staphylothermus , and in the halophilic archaeon Halorubrum  (see Table 5). Most of amylopullulanases from hyper/thermophilic archaea are active in the absence of calcium, which is a required for their industrial use.
Pullulan hydrolase type III (EC 3.2.1.x) attacks both α-1,4 and α-1,6-glucosidic linkages in pullulan, producing a mixture of maltotriose, panose, and maltose. It also degrades starch, producing mainly maltotriose and maltose . This enzyme has been reported in hyperthermophilic archaea belonging to the genera Thermococcus [77, 82] (see Table 5).
Isoamylases (EC 220.127.116.11) are enzymes that hydrolyze α-1,6-glucosidic linkages in branched polysaccharides, such as amylopectin, glycogen, and α and β limit dextrins, producing linear malto oligosaccharides. Isoamylases are the only enzymes capable of debranching glycogen completely [79, 83]. They have been reported in plants, bacteria, and in the archaeon Sulfolobus solfataricus ATCC 35,092  (see Table 5).
Transferases are enzymes that cleave an α-1,4 glucosidic bond of the donor molecule and transfer part of this molecule to a glucosidic acceptor, forming a new glucosidic bond .
Amylomaltases (EC 18.104.22.168) catalyze the transfer of a segment of an α-1,4-d-glucan to the reducing end of an acceptor (glucose or another α-1,4-d-glucan) . These enzymes are used for producing syrups. Amylomaltases have been found in bacteria and hyperthermophilic archaea belonging to the genera Sulfolobus , Thermococccus , and Pyrobaculum  (see Table 5).
Cyclodextrin glycosyltransferases (CGTases; EC 22.214.171.124) convert starch and oligodextrins into cyclodextrins, which are six to eight α-1,4 linked glucose units with an apolar internal cavity. Most of these enzymes have been reported in bacteria and also in hyperthermophilic archaea belonging to the genera Pyrococcus , Thermococcus  as well as in Archaeoglobus fulgidus strain 7324 [89, 90], and from the haloalkaliphilic archaeon Haloferax mediterranei  (see Table 5).
Branching-enzymes (α-1,4-glucan 6-α-glycosyltransferase; EC 126.96.36.199) cleave α-1,4-glycosidic linkages of a linear oligo- or polysaccharide and transfer the branch to the same or another polysaccharide, creating a new α-1,6 glucosidic bond . These enzymes have been reported in plants, mammals, bacteria, fungi, and in the archaeon Thermococcus kodakaraensis KOD1  (see Table 5).
Biotechnological applications of starch-degrading enzymes
In food and feed industry starch-converting enzymes are used to produce valuable products (glucose, fructose, and maltose) from starch. It is possible to produce starch-based materials with gelatin-like characteristics and defined linear dextrines as texturizers, aroma stabilizers, and prebiotics . Pullulanases and amylopullulanases are used for the production of glucose, maltose, and fructose as food sweeteners. These enzymes are also used for the production of high-glucose, high-fructose, and high-maltose syrups (manufacturing of high-quality candy and ice cream). In baking industry, pullulanases are used to improve texture, volume, and flavor of bakery products . Amylomaltases can produce cycloamylose and thermoreversible starch gel, which can be used as a substitute of gelatin . They are also used to produce syrups of isomalto-oligosaccharides with low sweetness and viscosity. α-amylases, branching and debranching enzymes and β-amylases can act as anti-staling agents, preventing undesirable changes in bread . In pharmaceutical industry and human health, pullulanases can be used for the production of maltose, which can replace d-glucose in the intravenous feeding . These enzymes are also used for the production of branched cyclodextrins. Due to their apolar interior, cyclodextrins can be used as hosts for pharmaceutical important molecules (e.g. proteins) that are solubilized and stabilized. On the other hand, pullulanases debranching are used for the preparation of slowly digestible starch, which correlates with low glycemic levels . Alkaline pullulanases and α-amylases are used as additives in dishwashing and laundry detergents to remove starches under alkaline conditions . In biofuel production α-amylases, glucoamylases, pullulanases and amylopullulanases can be used for degrading starch-containing crops (e.g. wheat, corn, barley) and produce ethanol .
Cellulose is the most abundant polymer on earth. This polysaccharide is a structural component of the cell wall of green plants and consists of up to 15,000 glucose units linked by β-1,4-glycosidic bonds. It has a high affinity to water, but it is completely insoluble in it because of its heterogeneous structure, which consists of both amorphous and highly ordered crystalline regions. Cellulases hydrolyze β-1,4 linkages in cellulose and based on their amino acid sequences and crystal structures they have been classified into three types: endoglucanases, exoglucanases, and β-glucosidases. Due to the complex structure of cellulose, it is necessary the combination of these enzymes for the complete hydrolysis of it into glucose. Cellulose is typically embedded in a network of hemicellulose and lignin, for this reason it requires an alkaline pretreatment at high temperatures to become accessible to enzymatic hydrolysis. So, cellulose industry needs thermostable cellulases, which in addition must be active at high pH [37, 81].
The endoglucanases (Cellulases; EC 188.8.131.52) hydrolyze β-1,4 bonds of cellulose in a random manner, generating oligosaccharides, such as cellobiose and glucose. These enzymes have been reported in bacteria, fungi, plants, animals, and in the hyperthermophilic archaea belonging to the genera Pyrococcus [93,94,95,96], Ignisphaera , Metallosphaera , Thermoproteus . These enzymes are also been reported in the thermoacidophilic archaea Acidilobus saccharovorans , Sulfolobus solfataricus , and in the haloalkaliphilic archaeon Haloarcula [102,103,104] (see Table 6). Endoglucanases from Ignisphaera aggregans, Metallosphaera cuprina, Thermoproteus uzoniensis, and Acidilobus saccharovorans have been identified but they are not characterized.
β-glucosidases (Cellobiases; EC 184.108.40.206) hydrolyze soluble cellodextrins and cellobiose, releasing β-d-glucose. These enzymes have been reported in bacteria, fungi, plants, animals, and archaea belonging to the genera Pyrococcus [94, 103, 104], and the thermoacidophilic archaea Sulfolobus [104,105,106] and Thermofilum  (see Table 6).
Biotechnological applications of cellulose-degrading enzymes
In pulp and paper industry, mixtures of endoglucanases reduce the fiber coarseness. Endoglucanases decrease the pulp viscosity and cellulases enhance the bleachability of softwood kraft pulp. Cellulases and xylanases release the ink from the fiber surface, improve fiber brightness and strength properties. In food and feed industry, cellulases are used for the improvement of juice yield, pretreatment of cellulose biomass and forage crops to improve nutritional quality. These enzymes are also employed in the color extractions of juices and releasement of antioxidants from fruit pomace. β-glucosidases improve texture, flavor, aroma of fruits and vegetables, they control bitterness of citrus fruits, and are used as additives to hydrolyze nonstarch polysaccharides [10, 108]. In biofuel production cellulases are used to increase the yield of saccharification of agricultural and industrial waste for bioethanol production. These enzymes convert cellulosic materials into useful and valuable products, such as ethanol, solvents, and organic acids. They improve nutritional quality of animal feed and facilitate their digestion and absorption. Textile industry also uses these enzymes for biostoning of jeans and biopolishing of cotton. Endoglucanases improve softness and water absorbance property of fibers and provide a cleaner surface structure. Cellulases remove short fibers, create a smooth appearance, and improve color brightness. In home care industry particularly in detergents, cellulases are used as additives causing color brightening and softening of fibers and removing rough protuberances in cotton fabrics . For wine and beer industry glucanases can improve quality, fermentation, and yields of beers and wines. These enzymes together with β-glucosidases improve color extraction, maceration, clarification, filtration, stability and aroma of wines. In agricultural industry, preparations based on cellulases are used to control plant disease because they can degrade the cell wall of phytopathogens .
The starting material to produce paper is wood, which is composed of cellulose (40–45%), hemicellulose (20–30%), and lignin (15–25%). Xylan, the principal component of hemicellulose, is a heterogeneous molecule with a main chain composed of xylose residues linked by β-1,4-glycosidic bonds . Xylanases are present in bacteria, fungi, and archaea. The steps in the paper production are carried out at elevated temperatures, so this industry requires thermostable xylan-degranding enzymes .
The endo-β-1,4-xylanases (xylanase; EC 220.127.116.11) are the most predominant enzymes. They cleave β-1,4-xylosidic linkages in xylans . These enzymes have been reported in the halophilic archaeon Halorhabdus utahensis  and in the hyperthermophilic archaeon Pyrodictium abyssi  (see Table 7). On the other hand, β-1,4-Xylosidases (EC 18.104.22.168) hydrolyze β-1,4-xylans and disaccharides, such as xylobiose, generating D-xylose . These enzymes have been reported in the halophilic archaeon Halorhabdus utahensis  and in the thermoacidophilic archaeon Sulfolobus solfataricus  (see Table 7).
Biotechnological applications of xylanases
In pulp and paper industry xylanases are used in bleaching of cellulose pulp as an alternative to chlorine bleaching. The treatment with these enzymes makes the pulp more permeable to subsequent extraction of residual brown lignin from fibers, because they degrade the xylan network that traps the residual lignin. In food and feed industry xylanases in conjunction with cellulases, and amylases improve yield and clarification of fruit juices. These enzymes increase aromas, essential oils, pigments, etc. of fruits and vegetables. Xylanases are also used as ingredients during bread preparations to improve its quality. In animal feed, these enzymes along with cellulases, proteases, and lipases are used to digest raw material, reducing viscosity, which improve the digestion of nutrients [110,111,112]. In pharmaceutical industry and human health, xylanases in conjunction with proteases are used as dietary supplements or to treat poor digestion. On the other hand, hydrolytic products of xylan are used as low-calorie sweeteners .
Chitin is the second most abundant polysaccharide, after cellulose, present in fungal cell walls, insect exoesqueletons, and shells of crustacea. Chitin, a linear β-1,4 homopolymer of N-acetyl-d-glucosamine (GlcNAc) residues, is crystalline, highly hydrophobic, and insoluble in water and organic solvents. This polysaccharide is nontoxic, antimicrobial, and biodegradable polymer. It is used for the production of oligosaccharides as biologically active substances . Chitinases have been reported in bacteria, fungi, plants, insects, mammals, and archaea belonging to the genera Haloferax , Halobacterium , Pyrococcus , Sulfolobus , Thermococcus [120, 121] (see Table 7).
Biotechnological applications of chitinases
Chitinases are used for the preparation of pharmaceutical important chitooligosaccharides with anti-tumor activity and N-acetyl-d-glucosamine, which is an anti-inflammatory drug used in the treatment of osteoarthritis, ulcerative colitis and other gastrointestinal inflammation disorders. Degradation products of chitin are used in drug delivery, wound healing, anti-fungal creams and lotions, production of artificial skin, surgical stitches and dietary fiber. These chitin derivatives are non-toxic, non-allergic, biodegradable, and biocompatible. In agricultural industry chitinases are used to control fungal phytopathogens and harmful insects, degrading their chitin coats. For bioremediation they are used in the treatment of chitinous waste to fertilizer .
DNA polymerases and DNA ligases are enzymes widely used in molecular biology to perform the polymerase chain reaction (PCR) and analytical methods, respectively .
DNA polymerases (EC 22.214.171.124) are key enzymes in DNA replication in all life forms. They synthesize a new DNA strand according to the template DNA, adding a deoxyribonucleotide 5′-triphosphate onto the growing 3′-OH end of a primer strand in the presence of Mg2+ ions. Taq polymerase was the first thermostable DNA polymerase applied in PCR but it has not 3′–5′ proofreading exonuclease activity, as a result, this enzyme is unable to excise mismatches. So, when high fidelity is required to reduce the error rate, the best choice is an archaeal DNA polymerase. These DNA polymerases have 3′–5′ proofreading activity, an error rate tenfold lower than that of Taq polymerase and are more thermostable, but are slower. The most commonly used DNA polymerases are from the archaea belonging to the genera Pyrococcus (Pfu, Pwo, Deep Vent™, Platinum® Pfx) and Thermococcus (KOD1, Tli, 9°N-7) [123,124,125] (see Table 8).
On the other hand, DNA ligases (ATP-dependent DNA ligases, EC 126.96.36.199 and NAD+ -dependent DNA ligases, EC 188.8.131.52) are ubiquitous enzymes that ligate breaks in DNA. The first thermostable ligase was discovered in the bacterium Thermus thermophilus HB8. Most of these enzymes come from thermophilic bacteria, but there are also of them from the hyper/thermophilic archaea Pyrococcus [131,132,133], Thermococcus [133,134,135,136], Hyperthermus butylicus , Methanocaldococcus jannaschii , Methanobacterium thermoautotrophicum , Sulfophobococcus zilligii , Aeropyrum pernix K1 , Archaeoglobus fulgidus , and Sulfolobus  (see Table 9). Unlike bacterial DNA ligases, these enzymes require ATP as a cofactor.
Biotechnological applications of DNA polymerases and DNA ligases
Thermostable DNA polymerases are used in DNA amplification, sequencing or labelling. Due to the high fidelity of archaeal DNA polymerases, they are used for reducing amplification errors in PCR products. On the other hand, thermostable DNA ligases are used in the construction of sequencing primers and as LDR/LCR enzymes because of their catalytic activity for nick-joining reaction at high temperatures (90–100 °C). LDR/LCR is a technique for detecting a single-base mutation in the DNA strand and it is used for the diagnosis of genetic diseases .
Nitriles are organic compounds that contain a cyano group (−C≡N) as part of their chemical structure. They are important chemical building blocks for the synthesis of intermediates in fine chemicals and pharmaceuticals. These reactions are carried out at elevated temperatures. So, pharmaceutical industry requires thermostable nitrile-degrading enzymes (amidases and nitrilases) [1, 144].
Amidases (EC 184.108.40.206) catalyze the conversion of amides to the corresponding carboxylic acids and ammonia. These enzymes are enantioselective and have a diverse substrate spectrum. Most of them do not require metal ions to be active. There are amidases that hydrolyze aliphatic substrates (aliphatic amidases) and those that hydrolyze cyclic or aromatic amides (aromatic amidases). These enzymes have been reported in bacteria and in hyper/thermophilic archaea belonging to the genera Pyrococcus  and Sulfolobus [147,148,149] (See Table 10).
On the other hand, nitrilases (EC 220.127.116.11) hydrolyze in one step nitriles to their corresponding carboxylic acid and ammonia. They are regio-, chemo-, and enantioselective, have a wide substrate spectrum, and do not require metal ions to be active. However, most of nitrilases have a poor thermostability. Based on the substrate specificity these enzymes are classified as aliphatic nitrilases (high affinity for aliphatic nitriles), aromatic nitrilases (high affinity for aromatic and heterocyclic nitriles), and arylacetonitrilases (high affinity for arylacetonitriles). Most of these enzymes come from bacteria, but they are also present in plants, fungi and archaea. Only one recombinant nitrilase have been reported from the hyperthermophilic archaeon Pyrococcus abyssi. This enzyme is an aliphatic nitrilase with high thermostability. Nevertheless, it does not hydrolyze aromatic nitriles, which are widely used in fine chemical and pharmaceutical industries .
In our laboratory we have isolated a nitrilase from a novel Antarctic Pyrococcus sp. recently isolated from the Antarctic Peninsula, Deception Island. This microorganism was isolated from an environmental sample and it was able to grow in the presence of aromatic nitriles at temperatures above 80 °C. The gene encoding the nitrilase enzyme was identified from its genome and subsequently was cloned and overexpressed in E. coli. The recombinant nitrilase showed activity at elevated temperatures towards aromatic and aliphatic nitriles, although it hydrolyzes preferentially aromatic compounds. The specific catalytic properties of this enzyme make it a potential candidate as biocatalyst for pharmaceutical industry . Currently, the complete biochemical characterization and thermostability studies of this enzyme are taking place.
Biotechnological applications of nitrile-degrading enzymes
Amidases are used to produce optically pure compounds in pharmaceutical industry. In food industry these enzymes are used to produce glutamic acid and aspartic acid (which contribute to the tastes “umami” and “sour”) and to produce fermented condiments such as soy sauce. Another application of these enzymes is in waste water treatment [1, 150].
On the other hand, nitrilases are used for the production of active pharmaceutical ingredients (API) and drug intermediates. This includes the synthesis of common and valuable pharmaceuticals, such as non-steroidal anti-inflammatory drugs (Ibuprofen, Ketoprofen, Naproxen), antidepressants (Nialamide, Gabapentin), analgesics (Pregabalin), anticholesterol drugs (Atorvastatin, Resuvastatin), anxiolytic and anticonvulsive drugs (Pregabalin), anti-tumor agents (Taxol), antibiotics (penicillins and semisynthetic cephalosporins), anti-allergic drugs (Terfenadine), drug delivery material (polyglycolic acid), and dermatological products (glycolic acid and mandelic acid, which are used as additives in facial moisturizers). In chemical industry nitrilases are used for the production of acrylamide, which is used in molecular biology (SDS-PAGE), paper making, permanent press fabrics, and ore processing. These enzymes are used to synthesize plastics, surface coatings, adhesives, synthetic rubbers, cosmetics, perfumes, household cleaners, automotive oil additives, pesticides, and herbicides. Additionally, nitrilases are used to produce glycine, which is a raw material for synthesizing detergents. They are used for synthesizing nicotinic acid (vitamin B3), which is used in the production of feedstuff additive. Since nitrilases are able to degrade nitriles, they can also be used for the treatment of contaminated water and soil [144, 151].
This review has summarized the main industrial enzymes from archaea, which play key roles in processes carried out in food, beverages, paper, textile, detergent, and pharmaceutic industries. However, there are also archaeal enzymes with specific applications which have been patented. Some of them are summarized in Table 11.
From the archaeal enzymes currently described and used in biotechnological applications several of these enzymes belong or have been isolated from species which recently have been described and found for the first time in Antarctica. Among them Micrococcus, Thermoccocus, Pyroccocus.
The enzyme nitrilase has been recently isolated and characterized from a novel Pyroccocus specie found in Antarctica by our group and it is currently being fully identified.
Archaeal extremozymes have demonstrated to be excellent biocatalysts for industrial applications due to their stability at high temperatures, extremes of pH, organic solvents, high salt concentration, and enantio selectivity. Due to these properties, archaeal biocatalysts can be used in a wide range of biotechnological applications. They can improve the processing of starch, cellulose, chitin, and xylan and they also allow the production of enantiomerically pure drugs of common use. Global enzyme market includes many industrial sectors and for this reason it requires large amounts of different enzymes. The best option to fulfill these requirements is the use of recombinant enzymes, which are produced in large-scale and can be easily purified. Nevertheless, nowadays there is still a need for more novel enzymes that can be generated in a recombinant manner particularly from archaea.
Recent findings of interesting archaeal species in Antarctica containing novel enzymes for potential industrial applications, makes Antarctica an interesting source of new archaeal and other type of microorganisms which contain more stable and active enzymes.
The task today and for the future is to generate better molecular tools for overexpression of some of these novel enzymes which are poorly expressed in the currently available molecular tools. The search for more enzymes and efficient improvements through modern technologies, such as site-directed mutagenesis, directed evolution, rational design, and metabolic engineering in order to generate the new generation of industrial biocatalysts is still needed.
polymerase chain reaction
poly unsaturated fatty acids
ligase chain reaction
ligase detection reaction
sodium dodecyl sulfate polyacrylamide gel electrophoresis
transmissible spongiform encelopathy
Banerjee A, Sharma R, Banerjee UC. The nitrile-degrading enzymes: current status and future prospects. Appl Microbiol Biotechnol. 2003;60:33–44.
Sarrouh B, Santos TM, Miyoshi A, Dias R, Azevedo V. Up-to-date insight on industrial enzymes applications and global market. J Bioprocess Biotech. 2012;S4:002.
Koga Y, Morii H. Biosynthesis of ether-type polar lipids in archaea and evolutionary considerations. Microbiol Mol Biol Rev. 2007;71:97–120.
Cavicchioli R. Archaea-timeline of the third domain. Nat Rev Microbiol. 2011;9:51–61.
Amenábar MJ, Flores PA, Pugin B, Boehmwald FA, Blamey JM. Archaeal diversity from hydrothermal systems of Deception Island, Antarctica. Polar Biol. 2013;36:373–80.
Horikoshi K, Bull A. Prologue: definition, categories, distribution, origin and evolution, pioneering studies, and emerging fields of extremophiles. In: Horikoshi K, editor. Extrem Handb. Tokyo: Springer; 2011. p. 3–15.
Van den Burg B. Extremophiles as a source for novel enzymes. Curr Opin Microbiol. 2003;6:213–8.
Wȩgrzyn A, Zukrowski K. Biotechnological applications of archaeal extremozymes. CHEMIK. 2014;68:710–22.
Reed CJ, Lewis H, Trejo E, Winston V, Evilia C. Protein adaptations in archaeal extremophiles. Archaea. 2013;2013:373275.
Satyanarayana T, Littlechild J, Kawarabayasi Y. Thermophilic microbes in environmental and industrial biotechnology: biotechnology of thermophiles. 2nd ed. Dordrecht: Springer; 2013.
Joshi S, Satyanarayana T. Biotechnology of cold-active proteases. Biology (Basel). 2013;2:755–83.
de Miguel Bouzas T, Barros-Velázquez J, Villa TG. Industrial applications of hyperthermophilic enzymes: a review. Protein Pept Lett. 2006;13:645–51.
Eichler J. Biotechnological uses of archaeal extremozymes. Biotechnol Adv. 2001;19:261–78.
Zhan D, Bai A, Yu L, Han W, Feng Y. Characterization of the PH1704 protease from Pyrococcus horikoshii OT3 and the critical functions of Tyr120. PLoS ONE. 2014;9:e103902.
Foophow T, Foophow T, Tanaka S, Koga Y, Takano K, Kanaya S. Subtilisin-like serine protease from hyperthermophilic archaeon Thermococcus kodakaraensis with N- and C-terminal propeptides. Protein Eng Des Sel. 2010;23:347–55.
Cowan DA, Smolenski KA, Daniel RM, Morgan HW. An extremely thermostable extracellular proteinase from a strain of the archaebacterium Desulfurococcus growing at 88 degrees C. Biochem J. 1987;247:121–33.
Völkl P, Markiewicz P, Stetter KO, Miller JH. The sequence of a subtilisin-type protease (aerolysin) from the hyperthermophilic archaeum Pyrobaculum aerophilum reveals sites important to thermostability. Protein Sci. 1994;3:1329–40.
Mayr J, Lupas A, Kellermann J, Eckerskorn C, Baumeister W, Peters J. A hyperthermostable protease of the subtilisin family bound to the surface layer of the archaeon Staphylothermus marinus. Curr Biol. 1996;6:739–49.
Gogliettino M, Riccio A, Cocca E, Rossi M, Palmieri G, Balestrieri M. A new pepstatin-insensitive thermopsin-like protease overproduced in peptide-rich cultures of Sulfolobus solfataricus. Int J Mol Sci. 2014;15:3204–19.
Manikandan M, Pašić L, Kannan V. Purification and biological characterization of a halophilic thermostable protease from Haloferax lucentensis VKMM 007. World J Microbiol Biotechnol. 2009;25:2247–56.
Izotova LS, Strongin AY, Chekulaeva LN, Sterkin VE, Ostoslavskaya VI, Lyublinskaya LA, et al. Purification and properties of serine protease from Halobacterium halobium. J Bacteriol. 1983;155:826–30.
Ryu K, Kim J, Dordick JS. Catalytic properties and potential of an extracellular protease from an extreme halophile. Enzyme Microb Technol. 1994;16:266–75.
Shi W, Tang XF, Huang Y, Gan F, Tang B, Shen P. An extracellular halophilic protease SptA from a halophilic archaeon Natrinema sp. J7: gene cloning, expression and characterization. Extremophiles. 2006;10:599–606.
Stan-Lotter H, Doppler E, Jarosch M, Radax C, Gruber C, Inatomi KI. Isolation of a chymotrypsinogen B-like enzyme from the archaeon Natronomonas pharaonis and other halobacteria. Extremophiles. 1999;3:153–61.
Antranikian G, Vorgias CE, Bertoldo C. Extreme environments as a resource for microorganisms and novel biocatalysts. Adv Biochem Eng Biotechnol. 2005;96:219–62.
Jaeger KE, Dijkstra BW, Reetz MT. Bacterial biocatalysts: molecular biology, three-dimensional structures, and biotechnological applications of lipases. Annu Rev Microbiol. 1999;53:315–51.
Fojan P, Jonson PH, Petersen MTN, Petersen SB. What distinguishes an esterase from a lipase: a novel structural approach. Biochimie. 2000;82:1033–41.
Alquéres SMC, Branco RV, Freire DMG, Alves TLM, Martins OB, Almeida RV. Characterization of the recombinant thermostable lipase (Pf2001) from Pyrococcus furiosus: effects of thioredoxin fusion tag and triton X-100. Enzyme Res. 2011;2011:316939.
Shao H, Xu L, Yan Y. Biochemical characterization of a carboxylesterase from the archaeon Pyrobaculum sp. 1860 and a rational explanation of its substrate specificity and thermostability. Int J Mol Sci. 2014;15:16885–910.
Suzuki Y, Miyamoto K, Ohta H. A novel thermostable esterase from the thermoacidophilic archaeon Sulfolobus tokodaii strain 7. FEMS Microbiol Lett. 2004;236:97–102.
Gao R, Feng Y, Ishikawa K, Ishida H, Ando S, Kosugi Y, et al. Cloning, purification and properties of a hyperthermophilic esterase from archaeon Aeropyrum pernix K1. J Mol Catal B Enzym. 2003;24–25:1–8.
Kim S, Lee W, Ryu Y. Cloning and characterization of thermostable esterase from Archaeoglobus fulgidus. J Microbiol. 2008;46:100–7.
Chen CKM, Lee GC, Ko TP, Guo RT, Huang LM, Liu HJ, et al. Structure of the alkalohyperthermophilic Archaeoglobus fulgidus lipase contains a unique C-terminal domain essential for long-chain substrate binding. J Mol Biol. 2009;390:672–85.
Camacho RM, Mateos JC, González-Reynoso O, Prado LA, Córdova J. Production and characterization of esterase and lipase from Haloarcula marismortui. J Ind Microbiol Biotechnol. 2009;36:901–9.
Legat A, Denner EBM, Dornmayr-Pfaffenhuemer M, Pfeiffer P, Knopf B, Claus H, et al. Properties of Halococcus salifodinae, an isolate from permian rock salt deposits, compared with Halococci from surface waters. Life. 2013;3:244–59.
Panda T, Gowrishankar BS. Production and applications of esterases. Appl Microbiol Biotechnol. 2005;67:160–9.
Robb F, Antranikian G, Grogan D, Driessen A. Thermophiles: biology and technology at high temperatures. 1st ed. Boca Raton: CRC Press; 2008.
Aidemir S, Akin N, Kocak C. Effect of lipase enzyme on the ripening of white pickled cheese. J Food Lipids. 2007;8:205–13.
Fischer K, Messner K. Adsorption of lipase on pulp fibers during biological pitch control in paper industry. Enzyme Microb Technol. 1992;14:470–3.
Sandoval G, Rivera I, Barrera-Rivera KA, Martinez-Richa A. Biopolymer synthesis catalyzed by tailored lipases. Micromol Sym. 2010;289:135–9.
Fukuda H, Kondo A, Noda H. Biodiesel fuel production by transesterification of oils. J Biosci Bioeng. 2001;92:405–16.
Adrio JL, Demain AL. Microbial enzymes: tools for biotechnological processes. Biomolecules. 2014;16:117–39.
Sivazankar B. Sugar, sweeteners, honey and confectionery. Food Process Preserv. 6th ed. New Delhi: PHI Learning; 2009. p. 329–39.
Prakash O, Jaiswal N. α-Amylase: an ideal representative of thermostable enzymes. Appl Biochem Biotechnol. 2010;160:2401–14.
Grzybowska B, Szweda P, Synowiecki J. Cloning of the thermostable α-amylase gene from Pyrococcus woesei in Escherichia coli: isolation and some properties of the enzyme. Mol Biotechnol. 2004;26:101–10.
Jung JH, Seo DH, Holden JF, Park CS. Maltose-forming α-amylase from the hyperthermophilic archaeon Pyrococcus sp. ST04. Appl Microbiol Biotechnol. 2014;98:2121–31.
Wang S, Lu Z, Lu M, Qin S, Liu H, Deng X, et al. Identification of archaeon-producing hyperthermophilic α-amylase and characterization of the α-amylase. Appl Microbiol Biotechnol. 2008;80:605–14.
Ying Q, Zhang C, Guo F, Wang S, Bie X, Lu F, et al. Secreted expression of a hyperthermophilic α-amylase gene from Thermococcus sp. HJ21 in Bacillus subtilis. J Mol Microbiol Biotechnol. 2013;22:392–8.
Lévêque E, Haye B, Belarbi A. Cloning and expression of an α-amylase encoding gene from the hyperthermophilic archaebacterium Thermococcus hydrothermalis and biochemical characterisation of the recombinant enzyme. FEMS Microbiol Lett. 2000;186:67–71.
Canganella F, Andrade CM, Antranikian G. Characterization of amylolytic and pullulytic enzymes from thermophilic archaea and from a new Fervidobacterium species. Appl Microbiol Biotechnol. 1994;42:239–45.
Kim JW, Flowers LO, Whiteley M, Peeples TL. Biochemical confirmation and characterization of the family-57-like α-amylase of Methanococcus jannaschii. Folia Microbiol (Praha). 2001;46:467–73.
Haseltine C, Rolfsmeier M, Blum P. The glucose effect and regulation of α-amylase synthesis in the hyperthermophilic archaeon Sulfolobus solfataricus. J Bacteriol. 1996;178:945–50.
Fukushima T, Mizuki T, Echigo A, Inoue A, Usami R. Organic solvent tolerance of halophilic α-amylase from a haloarchaeon, Haloarcula sp. strain S-1. Extremophiles. 2005;9:85–9.
Hutcheon GW, Vasisht N, Bolhuis A. Characterisation of a highly stable α-amylase from the halophilic archaeon Haloarcula hispanica. Extremophiles. 2005;9:487–95.
Onodera M, Yatsunami R, Tsukimura W, Fukui T, Nakasone K, Takashina T, et al. Gene analysis, expression, and characterization of an intracellular α-amylase from the extremely halophilicarchaeon Haloarcula japonica. Biosci Biotechnol Biochem. 2013;77:281–8.
Moshfegh M, Shahverdi AR, Zarrini G, Faramarzi MA. Biochemical characterization of an extracellular polyextremophilic α-amylase from the halophilic archaeon Halorubrum xinjiangense. Extremophiles. 2013;17:677–87.
Pérez-Pomares F, Bautista V, Ferrer J, Pire C, Marhuenda-Egea FC, Bonete MJ. α-Amylase activity from the halophilic archaeon Haloferax mediterranei. Extremophiles. 2003;7:299–306.
Kobayashi T, Kanai H, Aono R, Horikoshi K, Kudo T. Cloning, expression, and nucleotide sequence of the α-amylase gene from the haloalkaliphilic archaeon Natronococcus sp. strain Ah-36. J Bacteriol. 1994;176:5131–4.
Comfort DA, Chou CJ, Conners SB, VanFossen AL, Kelly RM. Functional-genomics-based identification and characterization of open reading frames encoding α-glucoside-processing enzymes in the hyperthermophilic archaeon Pyrococcus furiosus. Appl Environ Microbiol. 2008;74:1281–3.
Serour E, Antranikian G. Novel thermoactive glucoamylases from thermoacidophilic archaea Thermoplasma acidophilum, Picrophilus torridus and Picrophilus oshimae. Antonie van Leeuwenhoek, Int J Gen Mol Microbiol. 2002;81:73–83.
Schepers B, Thiemann V, Antranikian G. Characterization of a novel glucoamylase from the thermoacidophilic archaeon Picrophilus torridus heterologously expressed in E. coli. Eng Life Sci. 2006;6:311–7.
Kim MS, Park JT, Kim YW, Lee HS, Nyawira R, Shin HS, et al. Properties of a novel thermostable glucoamylase from the hyperthermophilic archaeon Sulfolobus solfataricus in relation to starch processing. Appl Environ Microbiol. 2004;70:3933–40.
Dock C, Hess M, Antranikian G. A thermoactive glucoamylase with biotechnological relevance from the thermoacidophilic euryarchaeon Thermoplasma acidophilum. Appl Microbiol Biotechnol. 2008;78:105–14.
Uotsu-Tomita R, Tonozuka T, Sakai H, Sakano Y. Novel glucoamylase-type enzymes from Thermoactinomyces vulgaris and Methanococcus jannaschii whose genes are found in the flanking region of the α-amylase genes. Appl Microbiol Biotechnol. 2001;56:465–73.
Chang ST, Parker KN, Bauer MW, Kelly RM. α-Glucosidase from Pyrococcus furiosus. Methods Enzymol. 2001;330:260–9.
Costantino HR, Brown SH, Kelly RM. Purification and characterization of an α-glucosidase from a hyperthermophilic archaebacterium, Pyrococcus furiosus, exhibiting a temperature optimum of 105 to degrees 115C. J Bacteriol. 1990;172:3654–60.
Galichet A, Belarbi A. Cloning of an α-glucosidase gene from Thermococcus hydrothermalis by functional complementation of a Saccharomyces cerevisiae mal11 mutant strain. FEBS Lett. 1999;458:188–92.
Piller K, Daniel RM, Petach HH. Properties and stabilization of an extracellular α-glucosidase from the extremely thermophilic archaebacteria Thermococcus strain AN1: enzyme activity at 130 degrees C. Biochim Biophys Acta. 1996;1292:197–205.
Park JE, Park SH, Woo JY, Hwang HS, Cha J, Lee H. Enzymatic properties of a thermostable α-glucosidase from acidothermophilic crenarchaeon Sulfolobus tokodaii strain 7. J Microbiol Biotechnol. 2013;23:56–63.
Angelov A, Putyrski M, Liebl W. Molecular and biochemical characterization of α-glucosidase and α-mannosidase and their clustered genes from the thermo acidophilic archaeon Picrophilus torridus. J Bacteriol. 2006;188:7123–31.
Ferrer M, Golyshina OV, Plou FJ, Timmis KN, Golyshin PN. A novel α-glucosidase from the acidophilic archaeon Ferroplasma acidiphilum strain Y with high transglycosylation activity and an unusual catalytic nucleophile. Biochem J. 2005;391:269–76.
Hii SL, Tan JS, Ling TC, Ariff A. Pullulanase: role in starch hydrolysis and potential industrial applications. Enzyme Res. 2012;2012:921362.
Ho Kim C, Nashiru O, Ko JH. Purification and biochemical characterization of pullulanase type I from Thermus caldophilus GK-24. FEMS Microbiol Lett. 1996;138:147–52.
Schwerdtfeger RM, Chiaraluce R, Consalvi V, Scandurra R, Antranikian G. Stability, refolding and Ca2+ binding of pullulanase from the hyperthermophilic archaeon Pyrococcus woesei. Eur J Biochem. 1999;264:479–87.
Han T, Zeng F, Li Z, Liu L, Wei M, Guan Q, et al. Biochemical characterization of a recombinant pullulanase from Thermococcus kodakarensis KOD1. Lett Appl Microbiol. 2013;57:336–43.
Erra-Pujada M, Debeire P, Duchiron F, O’Donohue MJ. The type II pullulanase of Thermococcus hydrothermalis: molecular characterization of the gene and expression of the catalytic domain. J Bacteriol. 1999;181:3284–7.
Jiao YL, Wang SJ, Lv MS, Xu JL, Fang YW, Liu S. A GH57 family amylopullulanase from deep-sea Thermococcus siculi: expression of the gene and characterization of the recombinant enzyme. Curr Microbiol. 2011;62:222–8.
Duffner F, Bertoldo C, Andersen JT, Wagner K, Antranikian G. A new thermoactive pullulanase from Desulfurococcus mucosus: cloning, sequencing, purification, and characterization of the recombinant enzyme after expression in Bacillus subtilis. J Bacteriol. 2000;182:6331–8.
Li X, Li D, Park KH. An extremely thermostable amylopullulanase from Staphylothermus marinus displays both pullulan- and cyclodextrin-degrading activities. Appl Microbiol Biotechnol. 2013;97:5359–69.
Siroosi M, Amoozegar MA, Khajeh K, Fazeli M, Habibi Rezaei M. Purification and characterization of a novel extracellular halophilic and organic solvent-tolerant amylopullulanase from the haloarchaeon, Halorubrum sp. strain Ha25. Extremophiles. 2014;18:25–33.
Ahmad N, Rashid N, Haider MS, Akram M, Akhtar M. Novel maltotriose-hydrolyzing thermoacidophilic type III pullulan hydrolase from Thermococcus kodakarensis. Appl Environ Microbiol. 2014;80:1108–15.
Niehaus F, Peters A, Groudieva T, Antranikian G. Cloning, expression and biochemical characterisation of a unique thermostable pullulan-hydrolysing enzyme from the hyperthermophilic archaeon Thermococcus aggregans. FEMS Microbiol Lett. 2000;190:223–9.
Ray R. Microbial Isoamylases: an overview. Am J Food Technol. 2011;6:1–18.
Park H-S, Park J-T, Kang H-K, Cha H, Kim D-S, Kim J-W, et al. TreX from Sulfolobus solfataricus ATCC 35092 displays isoamylase and 4-α-glucanotransferase activities. Biosci Biotechnol Biochem. 2007;71:1348–52.
Egorova K, Antranikian G. Biotechnology. 1st ed. Garrett R, Klenk H, eds. Archaea Evol Physiol Mol Biol. Malden: Blackwell Publishing; 2007.
Xavier KB, Peist R, Kossmann M, Boos W, Santos H. Maltose metabolism in the hyperthermophilic archaeon Thermococcus litoralis: purification and characterization of key enzymes. J Bacteriol. 1999;181:3358–67.
Kaper T, Talik B, Ettema TJ, Bos H, Van Der Maarel MJEC, Dijkhuizen L. Amylomaltase of Pyrobaculum aerophilum IM2 produces thermoreversible starch gels. Appl Environ Microbiol. 2005;71:5098–106.
Lee MH, Yang SJ, Kim JW, Lee HS, Kim JW, Park KH. Characterization of a thermostable cyclodextrin glucanotransferase from Pyrococcus furiosus DSM3638. Extremophiles. 2007;11:537–41.
Rashid N, Cornista J, Ezaki S, Fukui T, Atomi H, Imanaka T. Characterization of an archaeal cyclodextrin glucanotransferase with a novel C-terminal domain. J Bacteriol. 2002;184:777–84.
Labes A, Schönheit P. Unusual starch degradation pathway via cyclodextrins in the hyperthermophilic sulfate-reducing archaeon Archaeoglobus fulgidus strain 7324. J Bacteriol. 2007;189:8901–13.
Bautista V, Esclapez J, Pérez-Pomares F, Martínez-Espinosa RM, Camacho M, Bonete MJ. Cyclodextrin glycosyltransferase: a key enzyme in the assimilation of starch by the halophilic archaeon Haloferax mediterranei. Extremophiles. 2012;16:147–59.
Murakami T, Kanai T, Takata H, Kuriki T, Imanaka T. A novel branching enzyme of the GH-57 family in the hyperthermophilic archaeon Thermococcus kodakaraensis KOD1. J Bacteriol. 2006;188:5915–24.
Ando S, Ishida H, Kosugi Y, Ishikawa K. Hyperthermostable endoglucanase from Pyrococcus horikoshii. Appl Environ Microbiol. 2002;68:430–3.
Kataoka M, Ishikawa K. Complete saccharification of β-glucan using hyperthermophilic endocellulase and β-glucosidase from Pyrococcus furiosus. Biosci Biotechnol Biochem. 2014;78:1537–41.
Kishishita S, Fujii T, Ishikawa K. Heterologous expression of hyperthermophilic cellulases of archaea Pyrococcus sp. by fungus Talaromyces cellulolyticus. J Ind Microbiol Biotechnol Biotechnol. 2015;42:137–41.
Nakahira Y, Ishikawa K, Tanaka K, Tozawa Y, Shiina T. Overproduction of hyperthermostable β-1,4-endoglucanase from the archaeon Pyrococcus horikoshii by tobacco chloroplast engineering. Biosci Biotechnol Biochem. 2013;77:2140–3.
Göker M, Held B, Lapidus A, Nolan M, Spring S, Yasawong M, et al. Complete genome sequence of Ignisphaera aggregans type strain (AQ1.S1T). Stand Genomic Sci. 2010;3:66–75.
Liu LJ, You XY, Zheng H, Wang S, Jiang CY, Liu SJ. Complete genome sequence of Metallosphaera cuprina, a metal sulfide-oxidizing archaeon from a hot spring. J Bacteriol. 2011;193:3387–8.
Mardanov AV, Gumerov VM, Beletsky AV, Prokofeva MI, Bonch-Osmolovskaya EA, Ravin NV, et al. Complete genome sequence of the thermoacidophilic crenarchaeon Thermoproteus uzoniensis 768-20. J Bacteriol. 2011;193:3156–7.
Mardanov AV, Svetlitchnyi VA, Beletsky AV, Prokofeva MI, Bonch-Osmolovskaya EA, Ravin NV, et al. The genome sequence of the crenarchaeon Acidilobus saccharovorans supports a new order, Acidilobales, and suggests an important ecological role in terrestrial acidic hot springs. Appl Environ Microbiol. 2010;76:5652–7.
Huang Y, Krauss G, Cottaz S, Driguez H, Lipps G. A highly acid-stable and thermostable endo-β-glucanase from the thermoacidophilic archaeon Sulfolobus solfataricus. Biochem J. 2005;588:581–8.
Li X, Yu HY. Halostable cellulase with organic solvent tolerance from Haloarcula sp. LLSG7 and its application in bioethanol fermentation using agricultural wastes. J Ind Microbiol Biotechnol. 2013;40:1357–65.
Li X, Yu HY. Characterization of a halostable endoglucanase with organic solvent-tolerant property from Haloarcula sp. G10. Int J Biol Macromol. 2013;62:101–6.
Shin KC, Nam HK, Oh DK. Hydrolysis of flavanone glycosides by β-glucosidase from Pyrococcus furiosus and its application to the production of flavanone aglycones from citrus extracts. J Agric Food Chem. 2013;61:11532–40.
Kim BN, Yeom SJ, Kim YS, Oh DK. Characterization of a β-glucosidase from Sulfolobus solfataricus for isoflavone glycosides. Biotechnol Lett. 2012;34:125–9.
Cobucci-Ponzano B, Aurilia V, Riccio G, Henrissat B, Coutinho PM, Strazzulli A, et al. A new archaeal β-glycosidase from Sulfolobus solfataricus: seeding a novel retaining β-glycan-specific glycoside hydrolase family along with the human non-lysosomal glucosylceramidase GBA. J Biol Chem. 2010;285:20691–703.
Li D, Li X, Dang W, Tran PL, Park SH, Oh BC, et al. Characterization and application of an acidophilic and thermostable β-glucosidase from Thermofilum pendens. J Biosci Bioeng. 2013;115:490–6.
Kuhad RC, Gupta Ri, Singh A. Microbial cellulases and their industrial applications. Enzyme Res. 2011;2011:280696.
Vieille C, Zeikus GJ. Hyperthermophilic enzymes: sources, uses, and molecular mechanisms for thermostability. Microbiol Mol Biol Rev. 2001;65:1–43.
Wainø M, Ingvorsen K. Production of β-xylanase and β-xylosidase by the extremely halophilic archaeon Halorhabdus utahensis. Extremophiles. 2003;7:87–93.
Andrade CM, Aguiar WB, Antranikian G. Physiological aspects involved in production of xylanolytic enzymes by deep-sea hyperthermophilic archaeon Pyrodictium abyssi. Appl Biochem Biotechnol. 2001;91–93:655–69.
Cannio R, Di Prizito N, Rossi M, Morana A. A xylan-degrading strain of Sulfolobus solfataricus: isolation and characterization of the xylanase activity. Extremophiles. 2004;8:117–24.
Polizeli MLTM, Rizzatti ACS, Monti R, Terenzi HF, Jorge JA, Amorim DS. Xylanases from fungi: properties and industrial applications. Appl Microbiol Biotechnol. 2005;67:577–91.
Juturu V, Wu JC. Microbial xylanases: engineering, production and industrial applications. Biotechnol Adv. 2012;30:1219–27.
Sarma PVSRN, Prakash JM, Das SN, Kaur M, Purushotham P, Podile AR. Microbial chitinases: natural sources, mutagenesis, and directed evolution to obtain thermophilic counterparts. In: Satyanarayana T, Littlechild J, Kawarabayasi Y, editors. Thermophilic Microbes Environ Ind Biotechnol Biotechnol Thermophiles. 2nd ed. Dordrecht: Springer Netherlands; 2013. p. 649–69.
Hou J, Han J, Cai L, Zhou J, Lü Y, Jin C, et al. Characterization of genes for chitin catabolism in Haloferax mediterranei. Appl Microbiol Biotechnol. 2014;98:1185–94.
García-Fraga B, Da Silva AF, López-Seijas J, Sieiro C. Functional expression and characterization of a chitinase from the marine archaeon Halobacterium salinarum CECT 395 in Escherichia coli. Appl Microbiol Biotechnol. 2014;98:2133–43.
Mine S, Ikegami T, Kawasaki K, Nakamura T, Uegaki K. Expression, refolding, and purification of active diacetylchitobiose deacetylase from Pyrococcus horikoshii. Protein Expr Purif. 2012;84:265–9.
Staufenberger T, Imhoff JF, Labes A. First crenarchaeal chitinase found in Sulfolobus tokodaii. Microbiol Res. 2012;167:262–9.
Tanaka T, Fukui T, Atomi H, Imanaka T. Characterization of an exo-β-d-glucosaminidase involved in a novel chitinolytic pathway from the hyperthermophilic archaeon Thermococcus kodakaraensis KOD1. J Bacteriol. 2003;185:5175–81.
Andronopoulou E, Vorgias CE. Purification and characterization of a new hyperthermostable, allosamidin-insensitive and denaturation-resistant chitinase from the hyperthermophilic archaeon Thermococcus chitonophagus. Extremophiles. 2003;7:43–53.
Hamid R, Khan MA, Ahmad M, Ahmad MM, Abdin MZ, Musarrat J, et al. Chitinases: an update. J Pharm BioAllied Sci. 2013;5:21–9.
Ishino S, Ishino Y. DNA Polymerases and DNA Ligases. In: Satyanarayana T, Littlechild J, Kawarabayasi Y, editors. Thermophilic microbes Environ Ind Biotechnol Biotechnol thermophiles. 2nd ed. Dordrecht: Springer Netherlands; 2013. p. 429–58.
Terpe K. Overview of thermostable DNA polymerases for classical PCR applications: from molecular and biochemical fundamentals to commercial systems. Appl Microbiol Biotechnol. 2013;97:10243–54.
Cline J, Braman J, Hogrefe H. PCR fidelity of pfu DNA polymerase and other thermostable DNA polymerases. Nucl Acids Res. 1996;24:3546–51.
Dąbrowski S, Kur J. Cloning and expression in Escherichia coli of the recombinant His-tagged DNA polymerases from Pyrococcus furiosus and Pyrococcus woesei. Protein Expr Purif. 1998;14:131–8.
Dietrich J, Schmitt P, Zieger M, Preve B, Rolland JL, Chaabihi H, et al. PCR performance of the highly thermostable proof-reading B-type DNA polymerase from Pyrococcus abyssi. FEMS Microbiol Lett. 2002;217:89–94.
Takagi M, Nishioka M, Kakihara H, Kitabayashi M, Inoue H, Kawakami B, et al. Characterization of DNA polymerase from Pyrococcus sp. strain KOD1 and its application to PCR. Appl Environ Microbiol. 1997;63:4504–10.
Cambon-Bonavita MA, Schmitt P, Zieger M, Flaman JM, Lesongeur F, Raguénès G, et al. Cloning, expression, and characterization of DNA polymerase I from the hyperthermophilic archaea Thermococcus fumicolans. Extremophiles. 2000;4:215–25.
Kim YJ, Lee HS, Bae SS, Jeon JH, Lim JK, Cho Y, et al. Cloning, purification, and characterization of a new DNA polymerase from a hyperthermophilic archaeon, Thermococcus sp. NA1. J Microbiol Biotechnol. 2007;17:1090–7.
Keppetipola N, Shuman S. Characterization of a thermophilic ATP-dependent DNA ligase from the euryarchaeon Pyrococcus horikoshii. J Bacteriol. 2005;187:6902–8.
Zakabunin AI, Kamynina TP, Khodyreva SN, Pyshnaya IA, Pyshnyi DV, Khrapov EA, et al. Gene cloning, purification, and characterization of recombinant DNA ligases of the thermophilic archaea Pyrococcus abyssi and Methanobacterium thermoautotrophicum. Mol Biol. 2011;45:229–36.
Petrova TE, Bezsudnova EY, Dorokhov BD, Slutskaya ES, Polyakov KM, Dorovatovskiy PV, et al. Expression, purification, crystallization and preliminary crystallographic analysis of a thermostable DNA ligase from the archaeon Thermococcus sibiricus. Acta Crystallogr Sect F. 2012;68:163–5.
Nakatani M, Ezaki S, Atomi H, Imanaka T. A DNA ligase from a hyperthermophilic archaeon with unique cofactor specificity. J Bacteriol. 2000;182:6424–33.
Kim YJ, Lee HS, Bae SS, Jeon JH, Yang SH, Lim JK, et al. Cloning, expression, and characterization of a DNA ligase from a hyperthermophilic archaeon Thermococcus sp. Biotechnol Lett. 2006;28:401–7.
Rolland JL, Gueguen Y, Persillon C, Masson JM, Dietrich J. Characterization of a thermophilic DNA ligase from the archaeon Thermococcus fumicolans. FEMS Microbiol Lett. 2004;236:267–73.
Kim JH, Lee KK, Sun Y, Seo GJ, Cho SS, Kwon SH, et al. Broad nucleotide cofactor specificity of DNA ligase from the hyperthermophilic crenarchaeon Hyperthermus butylicus and its evolutionary significance. Extremophiles. 2013;17:515–22.
Wang Y, Xie J-J, Han Z, Liu J-H, Liu X-P. Expression, purification and biochemical characterization of Methanocaldococcus jannaschii DNA ligase. Protein Expr Purif. 2013;87:79–86.
Sriskanda V. Characterization of an ATP-dependent DNA ligase from the thermophilic archaeon Methanobacterium thermoautotrophicum. Nucleic Acids Res. 2000;28:2221–8.
Supangat S, An YJ, Sun Y, Kwon S-T, Cha S-S. Purification, crystallization and preliminary crystallographic analysis of a multiple cofactor-dependent DNA ligase from Sulfophobococcus zilligii. Acta Crystallogr Sect F. 2010;66:1583–5.
Jeon SJ, Ishikawa K. A novel ADP-dependent DNA ligase from Aeropyrum pernix K1. FEBS Lett. 2003;550:69–73.
Kim DJ, Kim O, Kim HW, Kim HS, Lee SJ, Suh SW. ATP-dependent DNA ligase from Archaeoglobus fulgidus displays a tightly closed conformation. Acta Crystallogr Sect F. 2009;65:544–50.
Lai X, Shao H, Hao F, Huang L. Biochemical characterization of an ATP-dependent DNA ligase from the hyperthermophilic crenarchaeon Sulfolobus shibatae. Extremophiles. 2002;6:469–77.
Gong JS, Lu ZM, Li H, Shi JS, Zhou ZM, Xu ZH. Nitrilases in nitrile biocatalysis: recent progress and forthcoming research. Microb Cell Fact. 2012;11:142.
Cabrera MA, Blamey JM. Cloning, overexpression, and characterization of a thermostable nitrilase from an Antarctic Pyrococcus sp. Extremophiles. 2017;5:861–9.
Fu L, Li X, Xiao X, Xu J. Purification and characterization of a thermostable aliphatic amidase from the hyperthermophilic archaeon Pyrococcus yayanosii CH1. Extremophiles. 2014;18:429–40.
Suzuki Y, Ohta H. Identification of a thermostable and enantioselective amidase from the thermoacidophilic archaeon Sulfolobus tokodaii strain 7. Protein Expr Purif. 2006;45:368–73.
Scotto D’Abusco A, Ammendola S, Scandurra R, Politi L. Molecular and biochemical characterization of the recombinant amidase from hyperthermophilic archaeon Sulfolobus solfataricus. Extremophiles. 2001;5:183–92.
Mueller P, Egorova K, Vorgias CE, Boutou E, Trauthwein H, Verseck S, et al. Cloning, overexpression, and characterization of a thermoactive nitrilase from the hyperthermophilic archaeon Pyrococcus abyssi. Protein Expr Purif. 2006;47:672–81.
El-Ghonemy D. Microbial amidases and their industrial applications: a review. Med Microbiol Diagnosis. 2014;4:1.
DeSantis G, DiCosimo R. Applications of nitrile hydratases and nitrilases. In: Tao J, Lin G, Liese A, editors. Biocatal Pharm Ind Discov Dev Manuf. 1st ed. Singapore: Wiley; 2009. p. 153–81.
Sutton JM, Raven NDH. Biological indicator. US09102976; 2015.
Oh DK YM. Thermostable beta-glycosidase and a method for preparing compound K and aglycon protopanaxadiol using the same. KR1020120034169; 2012.
Bourn W, Faurholm B FJ. Modified DNA polymerases. US8481685B2; 2013.
Yang X, Xu B, Wang W ZC. Construction method of recombinant Escherichia coli for expressing endoglucanase. CrN104561071; 2015.
Bylina EJ, Swanson RV, Mathur EJ LD. Glycosidase enzymes. US20140323356; 2014.
Both authors contribute equally to the development of this work. Both authors read and approved the final manuscript.
We would like to acknowledge the financial support of Becas CONICYT and Instituto Antártico Chileno (INACH).
The authors declare that they have no competing interests.
Availability of data and materials
All data generated or analyzed during this study are included in this published article.
Consent for publication
Ethics approval and consent to participate
Funding support was provided by National Commission for Scientific and Technological Research of Chile (CONICYT) and Chilean Antarctic Institute (INACH),
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.