Industrial applications of actinobacterial enzymes
\r\n\t
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Tiefenbacher",publishedDate:null,coverURL:"https://cdn.intechopen.com/books/images_new/10765.jpg",keywords:"Managing Urbanization, Managing Development, Managing Resource Use, Drought Management, Flood Management, Water Quality Monitoring, Air Quality Monitoring, Ecological Monitoring, Modeling Extreme Natural Events, Ecological Restoration, Restoring Environmental Flows, Environmental Management Perspectives",numberOfDownloads:null,numberOfWosCitations:0,numberOfCrossrefCitations:null,numberOfDimensionsCitations:null,numberOfTotalCitations:null,isAvailableForWebshopOrdering:!0,dateEndFirstStepPublish:"January 12th 2021",dateEndSecondStepPublish:"February 9th 2021",dateEndThirdStepPublish:"April 10th 2021",dateEndFourthStepPublish:"June 29th 2021",dateEndFifthStepPublish:"August 28th 2021",remainingDaysToSecondStep:"23 days",secondStepPassed:!1,currentStepOfPublishingProcess:2,editedByType:null,kuFlag:!1,biosketch:"A geospatial scholar working at the interface of natural and human systems, collaborating internationally on innovative studies about hazards and environmental challenges. Dr. Tiefenbacher has published more than 200 papers on a diverse array of topics that examine perception and behaviors with regards to the application of pesticides, releases of toxic chemicals, environments of the U.S.-Mexico borderlands, wildlife hazards, and the geography of wine.",coeditorOneBiosketch:null,coeditorTwoBiosketch:null,coeditorThreeBiosketch:null,coeditorFourBiosketch:null,coeditorFiveBiosketch:null,editors:[{id:"73876",title:"Dr.",name:"John P.",middleName:null,surname:"Tiefenbacher",slug:"john-p.-tiefenbacher",fullName:"John P. Tiefenbacher",profilePictureURL:"https://mts.intechopen.com/storage/users/73876/images/system/73876.jfif",biography:"Dr. John P. Tiefenbacher (Ph.D., Rutgers, 1992) is a professor of Geography at Texas State University. His research has focused on various aspects of hazards and environmental management. Dr. Tiefenbacher has published on a diverse array of topics that examine perception and behaviors with regards to the application of pesticides, releases of toxic chemicals, environments of the U.S.-Mexico borderlands, wildlife hazards, and the geography of wine. More recently his work pertains to spatial adaptation to climate change, spatial responses in wine growing regions to climate change, the geographies of viticulture and wine, artificial intelligence and machine learning to predict patterns of natural processes and hazards, historical ethnic enclaves in American cities and regions, and environmental adaptations of 19th century European immigrants to North America's landscapes.",institutionString:"Texas State University",position:null,outsideEditionCount:0,totalCites:0,totalAuthoredChapters:"1",totalChapterViews:"0",totalEditedBooks:"6",institution:{name:"Texas State University",institutionURL:null,country:{name:"United States of America"}}}],coeditorOne:null,coeditorTwo:null,coeditorThree:null,coeditorFour:null,coeditorFive:null,topics:[{id:"12",title:"Environmental Sciences",slug:"environmental-sciences"}],chapters:null,productType:{id:"1",title:"Edited Volume",chapterContentType:"chapter",authoredCaption:"Edited by"},personalPublishingAssistant:{id:"194667",firstName:"Marijana",lastName:"Francetic",middleName:null,title:"Ms.",imageUrl:"https://mts.intechopen.com/storage/users/194667/images/4752_n.jpg",email:"marijana@intechopen.com",biography:"As an Author Service Manager my responsibilities include monitoring and facilitating all publishing activities for authors and editors. 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Actinobacteria, the filamentous Gram positive bacteria are primarily saprophytes of the soils, where they contribute notably to the turnover of complex biopolymers such as lignocellulose, hemicellulose, pectin, keratin, and chitin. Undoubtedly, they are also well known as a rich source of antibiotics, enzymes, and other bioactive molecules, and are of considerable importance in pharmaceutical and other industries [1].
The ever-increasing requirement for enzymatic preparations is being met by such classical sources as animal and higher plant tissues, and that has stimulated the search for similar enzymes from the microbial world. The value of microorganisms, including actinobacteria, in the production of enzymes is enhanced by their relatively high yields, cost-efficiency, and susceptibility to genetic manipulation [2]. At present, enzymes of microbial origin are widely used in food processing, detergent manufacturing, the textile and pharmaceutical industries, medical therapy, bioorganic chemistry, and molecular biology. The wide use of enzymes reflects their characteristic specificity of action as biocatalysts. However, enzymes of identical substrate profile produced by different microorganisms may significantly vary in the optimal conditions of their productivity. For this reason, it is necessary to obtain microorganisms which produce enzymes with required substrate specificity, at particular temperature and pH ranges demanded by the production process. The biochemical heterogeneity, ecological diversity, and exceptional capacity of actinobacteria for secondary metabolites production make them an obvious target for enzymes displaying new activities and/or specificities. For many years, actinobacteria are best known as the source of majority of antibiotics. More recently, they have been found to be a promising source of a wide range of industrially important enzymes. Keeping this in mind and recognizing the significance of actinobacteria, especially Streptomyces species, as a source of novel enzymes, many studies have focused to understand the diversity of marine actinobacteria and to screen the enzyme-producing ability of actinobacteria occurring in the less explored hypersaline saltpan, mangrove, and marine sediments.
Actinobacteria are one of the ubiquitous dominant groups of Gram positive bacteria. Actinobacteria have been commercially exploited for the production of pharmaceuticals, neutraceuticals, enzymes, antitumor agents, enzyme inhibitors, and so forth [3]. These bioactive compounds are of high commercial value, and hence actinobacteria are regularly screened for the production of novel bioactive compounds. A wide array of enzymes applied in biotechnological industries and biomedical fields have been reported from various genera of actinobacteria. Since there is vital information available due to the advent of genome and protein sequencing data, actinobacteria has been continuously screened for the production of proteases, cellulases, chitinases, amylases, xylanases, and other enzymes. The industrial applications of several actinobacterial enzymes are given in Table 1.
\n\t\t\t\tUse\n\t\t\t | \n\t\t\t\n\t\t\t\tEnzyme\n\t\t\t | \n\t\t\t\n\t\t\t\tApplications\n\t\t\t | \n\t\t
Detergent (laundry and dish wash) | \n\t\t\tProtease | \n\t\t\tProtein stain removal | \n\t\t
Amylase | \n\t\t\tStarch stain removal | \n\t\t|
Lipase | \n\t\t\tLipid stain removal | \n\t\t|
Cellulase | \n\t\t\tCleaning, color clarification, anti-redeposition (cotton) | \n\t\t|
Mannanase | \n\t\t\tMannanan stain removal (reappearing stains) | \n\t\t|
Starch and fuel | \n\t\t\tAmylase | \n\t\t\tStarch liquefaction and saccharification | \n\t\t
Amyloglucosidase | \n\t\t\tSaccharification | \n\t\t|
Pullulanase | \n\t\t\tSaccharification | \n\t\t|
Glucose isomerase | \n\t\t\tGlucose to fructose conversion | \n\t\t|
Cyclodextrin-glycosyltransferase | \n\t\t\tCyclodextrin production | \n\t\t|
Xylanase | \n\t\t\tViscosity reduction (fuel and starch) | \n\t\t|
Food (including dairy) | \n\t\t\tProtease | \n\t\t\tMilk clotting, infant formulas (low allergenic), flavor | \n\t\t
Lipase | \n\t\t\tCheese flavor | \n\t\t|
Lactase | \n\t\t\tLactose removal (milk) | \n\t\t|
Pectin methyl esterase | \n\t\t\tFirming fruit-based products | \n\t\t|
Pectinase | \n\t\t\tFruit-based products | \n\t\t|
Transglutaminase | \n\t\t\tModify visco-elastic properties | \n\t\t|
Baking | \n\t\t\tAmylase | \n\t\t\tBread softness and volume, flour adjustment dough conditioning | \n\t\t
Xylanase | \n\t\t\tDough stability and conditioning (in situ emulsifier) | \n\t\t|
Lipase | \n\t\t\tDough stability and conditioning (in situ emulsifier) | \n\t\t|
Phospholipase | \n\t\t\tDough strengthening | \n\t\t|
Glucose oxidase | \n\t\t\tDough strengthening | \n\t\t|
Lipoxygenase | \n\t\t\tBread whitening | \n\t\t|
Protease | \n\t\t\tBiscuits, cookies | \n\t\t|
Transglutaminase | \n\t\t\tLaminated dough strengths | \n\t\t|
Animal feed | \n\t\t\tPhytase | \n\t\t\tPhytate digestibility – phosphorus release | \n\t\t
Xylanase | \n\t\t\tDigestibility | \n\t\t|
β-Glucanase | \n\t\t\tDigestibility | \n\t\t|
Beverage | \n\t\t\tPectinase | \n\t\t\tDe-pectinization, mashing | \n\t\t
Amylase | \n\t\t\tJuice treatment, low calorie beer | \n\t\t|
β-Glucanase | \n\t\t\tMashing | \n\t\t|
Acetolactate decarboxylase | \n\t\t\tMaturation (beer) | \n\t\t|
Laccase | \n\t\t\tClarification (juice), flavor (beer), cork stopper treatment | \n\t\t|
Textile | \n\t\t\tCellulase | \n\t\t\tDenim finishing, cotton softening | \n\t\t
Amylase | \n\t\t\tDe-sizing | \n\t\t|
Pectatelyase | \n\t\t\tScouring | \n\t\t|
Catalase | \n\t\t\tBleach termination | \n\t\t|
Laccase | \n\t\t\tBleaching | \n\t\t|
Peroxidase | \n\t\t\tExcess dye removal | \n\t\t|
Pulp and paper | \n\t\t\tLipase | \n\t\t\tPitch control, contaminant control | \n\t\t
Protease | \n\t\t\tBiofilm removal | \n\t\t|
Amylase | \n\t\t\tStarch-coating, de-inking, drainage improvement | \n\t\t|
Xylanase | \n\t\t\tBleach boosting | \n\t\t|
Cellulase | \n\t\t\tDe-inking, drainage improvement, fiber modification | \n\t\t|
Fats and oils | \n\t\t\tLipase | \n\t\t\tTransesterification | \n\t\t
Phospholipase | \n\t\t\tDe-gumming, lyso-lecithin production | \n\t\t|
Organic synthesis | \n\t\t\tLipase | \n\t\t\tResolution of chiral alcohols and amides | \n\t\t
Acylase | \n\t\t\tSynthesis of semisynthetic penicillin | \n\t\t|
Nitrilase | \n\t\t\tSynthesis of enantiopure carboxylic acids | \n\t\t|
Leather | \n\t\t\tProtease | \n\t\t\tUnhearing, bating | \n\t\t
Lipase | \n\t\t\tDe-pickling | \n\t\t|
Personal care | \n\t\t\tAmyloglucosidase | \n\t\t\tAntimicrobial (combined with glucose oxidase) | \n\t\t
Glucose oxidase | \n\t\t\tBleaching, antimicrobial | \n\t\t|
Peroxidase | \n\t\t\tAntimicrobial | \n\t\t|
L-Asparagine | \n\t\t\tAntitumor | \n\t\t|
Neuraminidase | \n\t\t\tAntiviral agents | \n\t\t|
Aminoacylase | \n\t\t\tRegulation of urea cycle | \n\t\t
Industrial applications of actinobacterial enzymes
Source: Goodfellow (1988).
Aminoacylase (N-acylamino-acid amidohydrolase) catalyzes the hydrolysis of acylated D- or L-amino acids to D- or L-amino acids and an appropriate carboxylic acid: N-acetyl-(D) or (L)-amino acid> carboxylic acid+(D)-or (L)-amino acid (Figure 1). Aminoacylases are interesting and ever-increasing enzymes due to the growing demand in the pharmaceutical industry for optically active amino acids. In enzymology, an aminoacylase is an enzyme that catalyzes the following chemical reaction:
Catalytic reaction of aminoacylase
This enzyme belonged to the family of hydrolases, those acting on carbon-nitrogen bonds other than peptide bonds, specifically in linear amides. This enzyme mainly concerns D-amino acids, both natural and synthetic, such as D-phenylglycine and its derivatives which are used for the production of semisynthetic penicillins and cephalosporins. Phenylglycine obtained synthetically as a racemic mixture can be split into enantiomers by chemical or enzymatic reactions. The latter are usually applied because they are simpler and more efficient. Two methods have been proposed for the isolation of pure enantiomers of D-amino acids using enzymatic hydrolysis of racemic mixtures of their N-acetylated derivatives. In the first method, a stereospecific enzymatic hydrolysis of N-acetyl-DL-amino acids has been used to obtain a mixture of D-amino acid and unaffected N-acetyl-L-amino acid which has to be racemized before its reuse in the process, while in the second method enzymatic cleavage of N-acetyl-L-amino acid, a component of the racemic mixture, results in a mixture of L-amino acid and non-hydrolyzed N-acetyl-D-amino acid. D-amino acid is obtained from the latter as a result of chemical deacylation of the N-acetylated derivative. The second method of obtaining D-amino acids is the one applied in practice. D-aminoacylases are uncommon in microorganisms, though Szwajcer et al. [4] reported the occurrence of one such enzyme from Micrococcus agilis. Many examples of D-aminoacylases have been found in some species of actinobacteria; 427 strains of Streptomyces and 16 strains of Streptoverticillium were screened for D-aminoacylases and found only in four species of streptomycetes, namely S. olivaceus, S. roseiscleroticus, S. sparsogenes, and S. tuirus [5]. All the species produced D-aminoacylase intracellularly when inducers such as D-leucine, D-phenylglycine, D-valine, and their N-acetylated derivatives were supplemented with the production medium [6]. The D-aminoacylases obtained from S. olivaceus and S. tuirus were purified and characterized according to their substrate specificity. Both enzymes were active at pH 7.0 and both were hydrolyzed hydrophobic N-acetyl-D-amino acids rather than hydrophilic amino acids. Extracellular production of both L-aminoacylase and penicillin V amidase has been demonstrated in Mycobacterium smegmatis [7] and several strains belonging to the genus Streptoverticillium [8]. L-aminoacylase isolated from a Streptoverticillium sp. [9] displayed a high hydrolytic activity toward N-acetylated aliphatic and aromatic L-amino acids [10]. Additionally, N-acyl-L-amino acids cannot be used directly as building blocks for proteins and must first be converted to L-amino acids by aminoacylase. Again, the L-amino acid products can be used for biosynthesis or catabolized energy.
Amylase comprises a group of industrial enzymes having approximately 25% of the global enzyme market. Specifically, an extracellular amylase with the ability to digest raw starch has found important applications in bioconversion of starches and starch-based substrates. The level of alpha amylase activity in various fluids of human body is of clinical importance, e.g., in diabetes, pancreatitis, and cancer research, while plant and microbial alpha amylases are used as industrial enzymes. Starch-degrading amylolytic enzymes are of great significance in biotechnological applications ranging from food, fermentation, and textile to paper industries (Table_1). Although amylases can be derived from several sources, such as plants, animals, and microorganisms, the enzymes from microbial sources are generally used to meet industrial demands and have made significant contribution to the production of foods and beverages in the last three decades. The microbial amylases have almost completely replaced the starch hydrolyzing chemicals in starch processing industry.
Like most microorganisms, actinobacteria can also survive in both mesophilic and thermophilic conditions; they have the ability to degrade starch by hydrolysis [15]. The occurrence of amylases in actinobacteria has been a well-known phenomenon since it was established that several representatives of the genera Nocardia and Streptomyces display amylolytic activity when cultivated on media with maltose [16], although, amylolytic enzymes quite often occur in mesophilic actinobacteria. Unfortunately, only a few of them have been studied in detail, as their enzymes are similar to bacilli amylases which are relatively thermolabile, thus limiting their value in biotechnological processes. On the basis of the literature survey, the more promising amylase producers are the strains of Streptomyces\n\t\t\t\t\thygroscopicus [17], S. limosus [18], and S. praecox [19] as a result of extensive screening program from among others. To date, amylases from S. hygroscopicus and S. praecox have been used for the commercial preparation of high-maltose syrups [20]. More attention has been paid to thermostable amylases produced by the thermophilic actinobacteria namely Thermomonospora curvata [21] and T. vulgaris [22] and by a Thermoactinomyces sp. [23; 24]. The amylases from Thermomonospora species and T.\n\t\t\t\t\tvulgaris are highly active and stable at 60°-70°C and act at slightly acidic and neutral pH values (Table 2) [25].
\n\t\t\t\tEnzyme producers\n\t\t\t | \n\t\t\t\n\t\t\t\tEnzyme\n\t\t\t | \n\t\t\t\n\t\t\t\tTemperature (°C)\n\t\t\t | \n\t\t\t\n\t\t\t\tpH\n\t\t\t | \n\t\t\t\n\t\t\t\tReference\n\t\t\t | \n\t\t
\n\t\t\t\tT. fusca NTU22 | \n\t\t\tα-amylase | \n\t\t\t60 | \n\t\t\t7.0 | \n\t\t\tChao-Hsun and Wen-Hsiung, 2007 | \n\t\t
\n\t\t\t\tStreptomyces transformant T3-1 | \n\t\t\tCellulase | \n\t\t\t50 | \n\t\t\t6.5 | \n\t\t\tHung-Der and Kuo-Shu, 2003 | \n\t\t
\n\t\t\t\tThermoactinomycetes sp. HS682 | \n\t\t\tProtease | \n\t\t\t70 | \n\t\t\t11·0 | \n\t\t\tTsuchiya et al., 1991 | \n\t\t
\n\t\t\t\tStreptomyces rimosus R6-554W | \n\t\t\tLipase | \n\t\t\t50 | \n\t\t\t9.0-10.0 (4-10) | \n\t\t\tAbrami et al., 1999 | \n\t\t
\n\t\t\t\tThermomonospora fusca\n\t\t\t | \n\t\t\tXylanase | \n\t\t\t60-80 | \n\t\t\t7.0 (6-8) | \n\t\t\tMcCarthy et al., 1995 | \n\t\t
Industrially important enzymes from thermophilic actinobacteria
β-N-acetyl-D-glucosaminidase (2-acetamido-2-deoxy-β-D-glucoside) is frequently encountered in microorganisms, higher plants, and mammalian tissues. This enzyme splits hydrolytically chitobiose, N,N\'-diacetylchitobiose moieties of asparagines-linked oligosaccharides of various glycoprotein and hydrolyzes N-acetyl-β-D-galactosaminidases, yielding oligosaccharide chains from glycoproteins [30]. Thus, it has been found to be very useful for the structural determination of the carbohydrate moiety of several glycoproteins and for studying their biochemical functions and biosynthesis [31]. Generally, actinobacteria producing enzymes are synthesized at extracellular region including endo-β-N-acetyl-D-glucosaminidase H [32] and endo-β-N-acetyl-glucosaminidase L [33] isolated from Streptomyces griseus (formerly S. plicatus). The enzyme were designated as endo-β-N-acetyl-D-glucosaminidase L [34; 35] and appear to be extremely useful for structural determination of ovoalbumin and several other glycoproteins [30; 36].
Endo-l, 3-α-D-glucanases (1, 3-α-D-glucan glucanohydrolase) hydrolyzes fragments of polysaccharides that contain consecutive 1, 3-linked α-D-glucosyl residues. Consequently, 1,3- α -D-glucanases are useful in detection of 1,3- α -D-linkages sequences in dextran as well as provide a route for 1,3-α-D-glucans study in fungal cell walls [37; 38]. These enzymes are produced by fungi and bacteria and are quite common among actinobacteria. Therefore, these enzymes may be useful as protective agents for odontological purposes. The presence of mutan-hydrolyzing enzymes was detected in S. chartreusis and S. werraensis [39]. Endo-1,3-β-D-glucanases (1,3-β-D-glucan glucanohydrolase) occur in bacteria, fungi, higher plants, and actinobacteria. Investigations of 1,3-β glucanases isolated from actinobacteria have been mainly carried out for their ability to degrade the cell walls of yeasts and fungi [40]. Several species of actinobacteria excrete 1, 3-β-glucanases together with chitinases. A laminarinase system consisting of three different types of l,3-β glucanases and chitinases was isolated from S. rimosus [41]. When laminarin (1,3- β glucan) was used as substrate, laminaritriose was obtained as the major product of one type of endo-1,3-β glucanase in addition to oligomeric breakdown products. The second laminarin-degrading (exo-splitting) enzyme yields predominantly laminaribiose. Another exo-l,3-β glucanase liberates glucose but not oligosaccharides from the nonreducing end of laminarin. The mycolase system produced an extracellular complex when the strain was grown on media with crab-shell chitin and fungal mycelia. δ1,3- β glucanases have also been isolated from Streptomyces (Actinomyces) cellulosae [42] and Streptomyces sp. [43]. Some of them, such as 1,3-β glucanase isolated from S. murinus, are used in wine preparation [44]. The preparations of 1,3-β glucanases have also been used to obtain some saccharides. The enzymatic preparation of laminaribiose (3-0-β gluco-pyranosyl-D-glucose) was achieved by the hydrolysis of curdlan (1,3- β- D-glucose) with the l,3-β-glucanase system from Streptomyces sp. K.27-4 [45]. The hydrolysate obtained consisted mainly of glucose and laminaribiose in an approximate ratio of 1:1 by weight. The application of the yeast Schizosaccharomyces\n\t\t\t\t\tpombe, which selectively metabolized all the glucose present in the hydrolysate, resulted in crystalline laminaribiose at 30% of yield.
Cellulose, which forms almost half of the dry weight of the earth’s biomass, is an unbranched polymer consisting of D-glucose units linked by 1,4-β glycosidic bonds. This macromolecule has a complex crystalline structure, is insoluble in water, and is quite resistant to depolymerizing enzymes and chemical reagents. Under natural conditions, cellulose is almost always combined with hemicellulose and lignin [46], which makes its degradation by microorganisms even more difficult.
Investigations on the mechanism of cellulose degradation and its possible applications have been carried out for many years [47]. Recently, the enzymatic hydrolysis of cellulose for D-glucose production has aroused an ever-increasing interest [48; 49]. Cellulose-degrading enzymes are excreted by microorganisms into the surrounding environment and as with most enzymes-degrading biopolymers they constitute a multicomponent lytic complex that acts synergistically on the cellulose. The cellulolytic system consists of three major components: 1,4-β glucan glucanohydrolase acting as endoglucanase, l,4-β-D-glucan cellobiohydrolase displaying exoglucanase activity, and β-glucosidase, which splits cellobiose. The enzymatic system of cellulases operates synergistically, i.e., endoglucanases make random scissions of the cellulose chain yielding glucose and oligosaccharides; exoglucanases attack the nonreducing end of cellulose forming cellobiose; and finally cellobiases hydrolyze cellobiose to glucose [5042]. Members of several mesophilic and thermophilic actinobacteria have been studied for their ability to degrade cellulose. Thermomonospora species have been found to be highly cellulolytic [51-55]. Cellulases produced by representatives of Thermomonospora species are multiple, extracellular exoglucanases and endoglucanases at pH of 6.0 and temperature ranging from 60° to 70°C; they also display considerable heat stability. The mesophilic actinobacteria known to produce cellulolytic complexes include Streptomyces\n\t\t\t\t\tantibioticus [56], S.\n\t\t\t\t\tflavogriseus [57-62], and S. viridosporus [63]. Mesophilic streptomycetes also produce complex cellulases at pH between 5 and 7; they show their highest activity at 40°-55°C (Table_2). Both mesophilic and thermophilic actinobacteria produced cellulolytic complexes when cultivated on media supplemented with powdered cellulose.
Proteases, generally classified into exopeptidases (cleave off peptide bonds from the ends of the protein chain) and endopeptidases (cleave peptide bonds within the protein) (Figure 2.), are the major industrial enzymes and fulfill more than 65% of the global market need [64]. These enzymes are extensively used in the food, pharmaceutical, leather, and textile industries [64; 65]. Among the extremophilic sources, thermostable proteases have been reported from certain haloalkaliphilic bacteria and actinobacteria [66; 67]. With the increasing demand of the enzymes, there will be an ever-increasing need for stable biocatalysts capable of withstanding extreme conditions of operation. Proteases generally activate a nucleophile, which will in turn attack the carbon of the peptide bond. The electrons in the carbon-oxygen double bond migrate onto the oxygen as the nucleophile attaches itself. This tetrahedral intermediate is a highly energetic intermediate, and the protease will stabilize this intermediate. The intermediate will then decompose, usually releasing the two peptide fragments.
Catalytic mechanism of protease
The ability to produce a variety of proteolytic enzyme is a well-known phenomenon in mesophilic actinobacteria; Streptomyces protease including “pronase 7M” (S. griseus) and “fradiase 7M” (S. fradiae) are commercially useful. While alkaline proteases from bacteria are extensively characterized, similar attention has not been paid to alkaliphilic actinobacteria namely S. nigellus, S. albidoflavus, and other genus Nocardiopsis, Thermomonospora, and Thermoactinomyces [68].
Chitin, a polymer occurring in crustaceans, fungi, and insects, is a fibrillar 1,4 linked 2-acetamido-2-deoxy-β-D-glucan with acetyl groups attached to nitrogen to various extents. It is found in three polymeric forms with various degrees of crystallinity. Fully deacetylated chitin is known as chitosane [69]. Enzymatic hydrolysis of chitin, liberating free N-acetyl-D-glucosamine, is caused by the chitinolytic complex which consists of chitinase (polyβ-1,4- (2-acetamido-2-deoxy)-D-glucose glycanohydrolase) and chitobiase (β-N-acetyl-D-glucosaminidase) (Figure 3.). As a result of the action of chitinase complex, chitobiose and chitotrose are released. Chitinases are specific to linear polymers of N-acetylglucosamine, but they do not split chitobiose. They hydrolyze chitin to chitobiose and to a lesser extent to chitotriose [70].
Polymer of β- (1-4)-N-acetyl-D-glucosamine units
Chitinolytic complexes commonly occur in bacteria, fungi, and especially in actinobacteria. The chitinase have been isolated from culture filtrates of S. griseus [71], S. antibioticus [70], Amycolatopsis (Streptomyces) orientalis [72], and several strains of Streptomyces spp. [73-75]. Chitinases are produced in abundance where strains are cultivated on chitin-supplemented media. Purified chitinases are more active at pH 5.0 but they are not heat-stable. Their properties have been considered from the view of fungal cell wall degradation [76] and utilization of chitin wastes [77].
Cholesterol esterase, which converts cholesterol esters into free cholesterol, is used in clinical tests for determining the cholesterol level in blood serum [78]. Until now, little is known about the properties of cholesterol esterases. These enzymes differ not only in their optimum pH for production but also in their substrate specificity. For example, cholesterol esterase isolated from S. lavendulae had lipolytic activity [79]. A lipase of the arylesterase group able to hydrolyze specifically phthalate esters to a free phthalic acid and simple n-alcohols was isolated from a Rhodococcus (Nocardia)\n\t\t\t\t\terythropolis [80]. The enzyme, which was most active at pH 8.6 and at 42°C, hydrolyzed several phthalate esters and to a lesser extent olive oil and tributyrin. The lipase production by several Streptomyces strains was reported by Chakrabarti et al. [81] but details were not given on substrate specificity. There is an increasing interest in lipases, especially those which display high stereo-specificity and may be useful for resolution of racemic acids and alcohols applied as chiral substrates in organic synthesis [82]. More attention has been paid to lipases from actinobacterial origin since these microorganisms are known for their ability to produce various secondary metabolites and hence provide a potential source of enzymes with substrate specificity.
Phospholipases, the enzymes capable of selective cleavage of ester bonds in glycerophosphatides, occur widely in both animal and plant kingdoms. Because of their high specificity, they are used for the analysis of phospholipid components of biological membranes as well as for clinical diagnostic tests. Phospholipases are classified into four groups, A, B, C, and D (Table 3). Serum choline phospholipids are hydrolyzed by phospholipase-D and the amount of liberated choline can be estimated quantitatively. Phospholipase-D from streptomycetes has been found useful for the determination of serum choline-phospholipids and in clinical diagnostic tests [83; 84].
\n\t\t\t\tEnzyme\n\t\t\t | \n\t\t\t\n\t\t\t\tName of the producer\n\t\t\t | \n\t\t\t\n\t\t\t\tOptimum pH\n\t\t\t | \n\t\t\t\n\t\t\t\tLeading references\n\t\t\t | \n\t\t
Phospholipase - A2\n\t\t\t | \n\t\t\t\n\t\t\t\tStreptomyces cinnamomeus\n\t\t\t | \n\t\t\t7.0 | \n\t\t\tOkawa and Yamaguchi (1976a) | \n\t\t
Phospholipase - B | \n\t\t\t\n\t\t\t\tS. hiroshimensis\n\t\t\t | \n\t\t\t9.0 | \n\t\t\tWalker and Walker (1975) | \n\t\t
Phospholipase - C | \n\t\t\t\n\t\t\t\tS. griseus\n\t\t\t | \n\t\t\t7.5 | \n\t\t\tVerma and Khuller (1983) | \n\t\t
Phospholipase - D | \n\t\t\t\n\t\t\t\tS. chromofuscus\n\t\t\t | \n\t\t\t8.0 | \n\t\t\tImamura and Horiuti (1979) | \n\t\t
Practical significance of phospholipases isolated from actinobacteria
Xylan, a hemicellulose, is composed of l,4-β-linked D-xylose units that form a linear backbone to which 4-O-methyl-D-glucuronic acid and L-arabinose are attached as side chains (Figure 4). This polymer, which occurs together with cellulose, is degraded by xylanases. Xylanases can be found in large amounts in both microorganisms as well as several invertebrates [85]. Together with other carbohydrases, xylanases play an important role in the degradation of terrestrial biomass [86]. Like cellulases they occur in microorganisms in the form of extracellular complexes, which consist of endo and exoxylanases that differ in substrate specificity. Xylanases produced by mesophilic actinobacteria belong to the endotype (l,4-β-D-xylan xylanohydrolase). They have been isolated and purified from several species of streptomycetes, such as S. flavogriseus [67], S. lividans [87], and Streptomyces sp. [88]. These enzymes were produced by microorganisms grown on media with xylan or its hydrolysates as a carbon source and/or in the presence of nonmetabolizable inducer [89]. The isolated xylanases exhibited their higher activity at pH 5.0-7.0 and at 40-60°C. Little attention has been given to xylanases produced by thermophilic actinobacteria [90; 68]. Thermostable xylanases isolated from Thermomonospora strains are heat-stable and most active at 60-70°C and at pH from 5.0 to 8.0. Apart from attempts to apply them to biodegradation of hemicelluloses and xylanases, they have also been used in the food industry for the production of D-xylose. Since this is not assimilated by mammalian organisms, it is used as an artificial sweetener in dietetic preparations [86]. An original method of obtaining xylobiose was developed by Kusakabe et al. [91] who prepared a pure xylobiose using a xylan hydrolysate from corncobs and rice straw and xylanase produced by Streptomyces sp. E-86. D-xylose, formed during the hydrolysis, was eliminated by the yeast Candida parapsilosis, which utilized xylose as a carbon source.
Structure of xylan and digestion of xylose by using xylanase
N-acetylmuramidase, an enzyme resembling lysozyme in action, cleaves the N-acetyl-muramyl-β,4-N-acetylglucosamine bonds of the polysaccharide chain of peptidoglycan, liberating free-reducing groups of N-acetylmuramic acid. N-acetylmuramidase (mucopeptide N-acetylmuramoyl-hydrolases) belongs to the group of bacteriostatic enzymes comprising glycosidases that hydrolyze peptidoglycan (murein), which is basic component of the bacterial cell wall. Murein, composed of glycan strands consisting of alternating acetylated amino sugars, N-acetylglucosamine and N-acetylmuramic acid linked by β (1-4) glycosidic bonds mutually cross-linked by peptide chains, forms a mono- or multilayer net covered with lipopolysaccharides, phospholipids, and lipoproteins. The peptide moiety of murein is composed of short chains of unbranched aliphatic amino acids and/or amino acids that form stem peptides linked to the carboxyl group of N-acetylmuramic acid and cross-linked by interpeptide bridges [92]. Peptidoglycans, especially available high in Gram-positive bacteria, are highly diversified. The determination of the primary structure of peptidoglycans has revealed differences between the bacteria and it provides significant taxonomic tools [93]. The enzyme was isolated from Streptomyces globisporus [94; 95], and is also found in other streptomycetes, including Streptomyces sp. [96] S. griseus [97], S. erythraeus [98], S. (Actinomyces) levoris [99], and S. rutgersensi [100].
Neuraminidase (acylneuraminyl hydrolase) splits 2,3-, 2,6-, and 2,8- and 2,9-glucosidic linkages which join terminal nonreducing N- or O-acetylated neuraminyl residues present in oligosaccharides and glycoprotein. Neuraminidases or sialidases occur widely in bacteria, viruses, animal tissues, and biological fluids [101]. These enzymes, isolated from various sources and differing in their substrate specificity, are applied in a wide area of biological and immunological research, particularly in cell surface and clinical studies [23; 102]. In actinobacteria, neuraminidases have been found in representatives of the genera Corynebacterium, Mycobacterium and Nocardia [101], Streptomyces griseus [103; 104], Actinomyces naeslundii, and A. viscosus [105]. Neuraminidases isolated from Streptomyces strains showed optimum activity at pH 3.5-5.0 and at 50°-60°C. They differed notably in their properties, including substrate specificity. Neuraminidase from Clostridium perfringens and Vibrio cholerae are able to split all types of sialic acid linkages.
Proteolytic enzymes of microbial origin were classified by Morihara [106], on the basis of their catalytic mechanism, into serine, thiol, metallo, and acid proteases according to the general systematic scheme introduced by Hartley [107]. The ability to produce a variety of proteolytic enzymes is a well-known phenomenon in mesophilic actinobacteria [106]. There is also an increasing interest in proteases derived from thermophilic actinobacteria including members of the genera Thermoactinomyces (Micromonospora), Thermomonospora, and Streptomyces. These actinobacteria are still not fully exploited as a potential source of thermostable enzymes acting not only over a wide range of pH but also great number of proteases with wide spectrum of substrate specificity. An increasing interest is observed in the application of actinobacterial proteases in bioorganic chemistry. For commercial purposes, they are routinely obtained as by-products formed during biosynthesis of antibiotics in the logarithmic phase of growth [107], from the fermentation broths of Streptomyces fradiae [109], S. griseus [110], and S. rimosus [108]. The preparations obtained are enzymatic complexes that contain a mixture of endo- and exopeptidases; as commercial preparations they are known as pronase (S. griseus) or fradiase (S. fradiae). Actinobacterial proteolytic complexes provide an excellent source of protease in various substrates specificity. Of the actinobacterial protease complexes available, the most attention has been given to pronase obtained from S. griseus.
Pronase like enzymes are produced not only by S. griseus but also by members of several other species of streptomycetes. A trypsin-like serine protease was isolated from S. erythraeus [111] and S. fradiae [112]; carboxypeptidase T, which has a mixed specificity compared with pancreatic carboxypeptidase A and B, is produced extracellular by a Thermoactinomyces strain [113]. Aminopeptidases have been isolated from culture filtrates of S. rimosus grown under conditions conducive to the industrial biosynthesis of oxytetracycline [114] and from other Streptomyces sp., including S. mauvecolor [34], S. peptidofaciens [115], and S. sapporonensis [116].
The important application of the L-asparaginase enzyme is in the treatment of acute lymphoblastic leukemia, Hodgkin’s disease, acute myelocytic leukemia, acute myelomonocytic leukemia, acute and chronic lymphocytic leukemia, lymphosarcoma treatment, reticulosarbom, and melanosarcoma [117]. L-asparaginase broadly distribute among the plants, animals, and microorganisms. The microbes are a better source of L-asparaginase, because they can be cultured easily and the extraction and purification of enzyme from them is also convenient, facilitating large-scale production [118]. L-asparaginase has been arousing considerable interest as it displays an antineoplastic activity against a variety of murine neoplasms. As an antineoplastic agent, L-asparaginase from Escherichia coli has been the most widely available [119], but the enzyme also occurs in actinobacteria. It has been isolated from several actinobacteria including Mycobactserium bovis [120] and M. tuberculosis [121]. These enzymes were active at pH 8-9. L-asparaginase obtained from Streptomyces karnatakensis [122], like others, was also intracellular and showed stereospecificity not only toward the L-isomer but was also able to hydrolyze D-asparagine to a smaller extent. The L-asparaginase production was reported from various actinobacteria namely Psedonocardiae endophytica VUK-10 [123]; Streptomyces sp. WS3/1 [124]; Streptomyces sp. (SS7) [125]; S. halstedii [126]; Streptomyces acrimycini NGP [127].
Penicillin amidase (penicillin amidohydrolase) is an enzyme that hydrolyzes penicillins to 6-aminopenicillanic acid (6-APA) and carboxylic acid. The cleavage of penicillin into 6-APA and side chain is a reaction in which the penicillin nucleus, the basis for the production of semisynthetic penicillins, is obtained. Penicillin amidases in the form of immobilized preparations are applied in the production of 6-APA on an industrial scale [128]. Penicillin acylases are classified into three groups on the basis of their substrate specificity: the first group includes the enzymes that hydrolyze phenoxymethyl-penicillin (penicillin V); the second, those that act on benzylpenicillin (penicillin G); and the enzymes of the third group display specificity with respect to D-a-amino-benzylpenicillin, ampicillin [129]. The penicillin hydrolysis reaction procedes in an alkaline medium and at lower pH values and is reversible. This property was exploited to synthesize semisynthetic penicillins and cephalosporins by the application of penicillin amidase preparations [128] occur in bacteria and actinobacteria and to a minor extent in yeasts and moulds [129-131]. The majority of those found in actinobacteria such as Mycobacterium, Nocardia, and Streptomyces [131; 132] are able to hydrolyze phenoxymethyl-penicillin [133]. These enzymes are mainly intracellular and they display optimum penicillin hydrolysis at pH 7.0 to 8.0. Similarly to acylases isolated from the other sources, they catalyze penicillin synthesis in an acidic medium at pH from 4.0 to 5.5 [131]. So far they have not been reported to be of commercial significance.
Enzymes are considered as a potential biocatalyst for many biological reactions. Particularly, the microbial enzymes have extensive uses in industries and medicines. The microbial enzymes are also more active and stable than plant and animal enzymes. In addition, the microorganisms, particularly actinobacteria, represent an alternative efficient source of enzymes because they can be cultured in large quantities by fermentation and owing to their biochemical diversity and susceptibility to gene manipulation. Industries are looking for new microbial strains in order to produce different enzymes to fulfill the current enzyme requirements. Hence, the actinobacteria as biofactory of potential enzyme as well as secondary metabolites production, fulfill the requirements of several industrial enzymes. In a world with a rapid increasing of population and approaching exhaustion of many natural resources, enzyme technology offers a great potential for many industries to help meet the challenges they will face in the years to come.
Moore’s law along with the Dennard scaling has been the key driving factor that has enabled steady progress in the semiconductor industry over the last three decades. However, continuous scaling of Complementary Metal Oxide Semiconductor (CMOS) transistor into the sub-nanometer regime faces severe challenges due to short channel effects, exponentially rising leakage currents and increased variations in manufacturing process. To combat these challenges, numerous approaches have been explored. One of the promising approaches is to look for alternatives transistor structures that could potentially overcome these challenges. In this regard, Quantum-dot Cellular Automata (QCA), Carbon Nano-Tube Field Effect Transistor (CNTFET) and Single Electron Transistor (SET) are proposed to replace or supplement CMOS technology [1, 2].
\nFrom circuits and systems design perspective, several approaches are explored to reduce die area and lower energy consumption. One of the approaches is to use Multi-Valued Logic (MVL) design. MVL circuit design has been explored in CMOS technology for last few decades using circuit styles like voltage-mode CMOS logic (VMCL), I2L logic and current-mode CMOS logic (CMCL) [3, 4]. Binary logic packs two logic values between available voltage levels, whereas MVL packs more than two logic values between the available voltage levels. Ternary logic is simplest form of MVL. Due to reduced noise margins, MVL logic design is less reliable and more prone to defects. Significant efforts have been directed towards defect and fault tolerance for binary logic and less effort has been directed towards defect and fault tolerant techniques for MVL.
\nIn digital design, transistor is used as switch that transitions between logic states. The switch that controls this transition is the threshold voltage of the transistor. Threshold voltage is a manufacturing process parameter that is set to a unique value during manufacturing. To realize CMOS MVL circuits, multi-threshold transistors are deployed. In CMOS, multi-threshold transistors are realized by applying body bias voltages that exploit body effect to alter threshold voltages [5]. However, CNTFETs are fundamentally different that they allow realization of multi-threshold transistors by tuning a few process parameters. This property has been effectively exploited to realize various forms of ternary logic circuits using CNTFET [6, 7].
\nDespite several advantages, there are significant issues with the reliable realization of CNTFET circuits. There are fundamental limitations that are specific to Carbon Nanotubes (CNT) that pose major challenges [8]. CNTs are graphene sheets rolled into tubes [9]. Multiple CNTs are deployed in the channel region to provide the required drive currents needed for reliable operation [10]. These CNTs can be either metallic or semiconducting depending on the arrangement of the carbon atoms in the tube. Metallic tubes can result in circuit malfunctioning due to source-drain shorts [8]. It is also not possible to guarantee perfect positioning and alignment of these CNTs in large CNTFET circuits [8]. To harness the potential benefits of CNTFET technology, variation aware defect and fault tolerant techniques are needed for reliable operation of CNTFET MVL circuits.
\nIn this chapter, we present MVL realization of CNTFET circuits and discuss techniques for defect and fault tolerance in MVL CNTFET circuits. The rest of this chapter is organized as follows: Section 2 discusses basic logic primitives that are needed for understanding MVL. Section 3 describes the CNTFET transistor and its operation. Section 4 provides a detail description of various circuit styles that have been proposed to realize MVL CNTFET circuits. Section 5 describes variation in CNTFET devices. Section 6 describes a technique for fault tolerance in MVL CNTFET circuits.
\nRaychowdhury et al. detail and discuss the logic primitives that are needed for understanding MVL [6]. Let us consider an \n
The mapping of \n
Consider \n
Input \n | \nOutput \n | \n
---|---|
\n\n | \n\n\n | \n
\n\n | \n\n\n | \n
\n\n | \n\n\n | \n
Truth table for complement operator [9].
Ternary nand operator can be defined as
\nHere is an example of min operator: \n
Ternary nor operator can be defined as:
\nwhere belongs to the set \n
Input a | \nInput b | \nOutput \n | \nOutput \n | \n
---|---|---|---|
0 | \n0 | \n2 | \n2 | \n
1 | \n0 | \n2 | \n1 | \n
2 | \n0 | \n2 | \n0 | \n
0 | \n1 | \n2 | \n1 | \n
1 | \n1 | \n1 | \n1 | \n
2 | \n1 | \n1 | \n0 | \n
0 | \n2 | \n2 | \n0 | \n
1 | \n2 | \n1 | \n0 | \n
2 | \n2 | \n0 | \n0 | \n
Ternary gates truth table [7].
\nFigure 1 shows the cross-section of a CNTFET transistor. Similar to MOSFETs, CNTFETs have four terminals: drain, gate, source and substrate [11]. CNTFET transistors are constructed by replacing the silicon channel in CMOS with carbon nanotubes (CNT). CNTs are sheets of Graphene rolled into tubes. Depending on the direction in which the sheets are rolled in the channel, CNTs can be either metallic or semi-conducting. This property of CNT being metallic or non-metallic depending on their rolled direction is termed as chirality [7]. In CMOS transistors, drive current depends on the channel width [5]. However, in the case of CNTFET the drive currents during conduction state depends on the number of CNTs in the channel along with gate length, chirality and pitch distance [12]. The \n
Side view of a CNTFET transistor.
where \n
where \n
This property has been effectively exploited to design MVL circuits using CNTFET. In practice, multi-diameter CNTFET is realized by CNT synthesis techniques that can fabricate CNTs with desired chirality [14]. Also, post-processing techniques for adjusting the threshold voltage of multiple tube CNTFET have also been demonstrated [15].
\nThis section discusses two circuit implementations of CNTFET ternary inverters along with a circuit implementation of a CNTFET ternary Muller C element. First circuit is a resistor based ternary inverter that was proposed by Raychowdhury et al. [9]. Second circuit is a ternary inverter using static complementary circuit style that was proposed by Lin et al. [7]. We will also discuss the circuit implementation and operation of a ternary Muller C element proposed by Sundararajan et al. [16].
\n\nFigure 2 shows the circuit diagram of a resistive load ternary inverter. The circuit consists of two N-channel transistors and two resistors. CNTFET with two different diameters are deployed in this circuit. Transistor \n
Resistive CNTFET ternary inverter [6]. Value of \n\n\nR\n1\n\n\n is \n\n100\n\n\n\n\nK\nΩ\n\n.
Transistor | \nChirality Vector (n,m) | \nDiameter(nm) | \n\n\n | \n
---|---|---|---|
\n\n | \n(19,0) | \n1.487 | \n0.29 | \n
\n\n | \n(10,0) | \n0.783 | \n0.56 | \n
CNTFET chirality, \n
The problem with the resistive load ternary inverter is that resistors are large, bulky and are prone to noise. Also, static resistors draw leakage currents from the supply and are not suitable for implementation in large scale CNTFET circuits. As an alternative to resistive load ternary inverter, Lin et al. proposed a static complementary version of ternary inverter that employs P-channel and N-channel CNTFETs as shown in Figure 3. The resistor in pull up network of the circuit in Figure 2 is replaced with the two P-Channel CNTFET with varying diameters and the voltage divider resistor is replaced with two diode connected complementary CNTFET. The diode connected transistors have a nominal threshold voltage (\n
Static complementary CNTFET ternary inverter [7].
Transistor | \nChirality Vector (n,m) | \nDiameter(nm) | \n\n\n | \n
---|---|---|---|
\n\n | \n(13,0) | \n1.018 | \n0.43 | \n
\n\n | \n(19,0) | \n1.487 | \n0.29 | \n
\n\n | \n(10,0) | \n0.783 | \n0.56 | \n
CNTFET chirality, \n
In this section, we will review the circuit implementation of ternary Muller C element described by Sundararajan et al. [16]. Muller C element is a common logic gate that is deployed in asynchronous logic [17, 18]. Figure 4 shows a circuit schematic and a logic representation of the Muller C element [19]. The basic operation of the C element can be described as follows: When the logic values of inputs \n
Binary CMOS Muller C element: Circuit schematic and its logic representation. It has two inputs a, b and an output c. the C element consists of “C-not” and “S” gates [16].
where \n
Input a | \nInput b | \nOutput c | \n
---|---|---|
0 | \n0 | \n0 | \n
0 | \n1 | \n\n\n | \n
0 | \n2 | \n\n\n | \n
1 | \n0 | \n\n\n | \n
1 | \n1 | \n1 | \n
1 | \n2 | \n\n\n | \n
2 | \n0 | \n\n\n | \n
2 | \n1 | \n\n\n | \n
2 | \n2 | \n2 | \n
Ternary Muller C-element truth table. \n
Schematic for all inputs at logic \n\n0\n\n.
Schematic for all inputs at logic \n\n1\n\n.
Schematic for all inputs at logic \n\n2\n\n.
Transient results of static complementary Muller C [16].
This section describes the process variation in CNTFET technology. Process variation (PV), an artifact of aggressive technology scaling, causes uncertainty in integrated circuit characteristics. Imperfect lithography, doping fluctuations and imperfect planarization are some of the causes of variation that strongly affect the channel length, oxide thickness, width and eventually the threshold voltage of CMOS transistors. Such variation leads to unpredictable circuit behavior. Due to the random nature of manufacturing process, various effects such as ion implantation, diffusion and thermal annealing have induced significant fluctuations of the electrical characteristics in nanoscale CMOS [24]. CNTFETs are also affected by manufacturing variation caused by imperfect fabrication. CNTFETs not only suffer from traditional process variations, which are in common with the CMOS technologies but they also have their unique source of variations. Paul et al. showed that the CNTFET devices are significantly less sensitive to stochastic variations such as process-induced variations due to their inherent device structures and geometric properties [25]. In addition, CNTFETs are subject to CNT specific variation sources including: CNT type variations, CNT density variations, CNT diameter variations, CNT alignment variations and CNT doping variations [23]. Figure 9 shows the contributions of each of the aforementioned sources of variations to the on-state current \n
Contributions of various CNT specific variation sources [23].
As discussed in Section 5, CNT density is the critical parameter that needs to be accounted for variation aware design. CNT density variation manifests itself as varying amounts of CNT under the gate of every CNTFET transistor. CNT density variation follows a normal distribution with mean \n
Percentage of \n\n\nt\npd\n\n\n variation for the ternary Muller C element.
The C element was driving a capacitance of 25 \n
This section will review a method for fault tolerance in CNTFET Multi-valued logic that was proposed by Sundararajan et al. [16]. The method called Restorative FeedBack (RFB), provides fault resilience against Single Event Upset (SEU) [19]. Triple Modular Redundancy (TMR) is one of the mostly commonly used method for fault tolerance in computer systems [30]. In TMR, a logic circuit is replicated three times and a majority vote is taken on the combined outputs. Figure 11 shows the TMR method, where three copies of a logic circuit are made and a majority vote is taken on those combined copies to obtain a fault free logic value. TMR provides resilience against single error faults. The main drawback of TMR is that logic overhead is high due to replication and and a fault in majority voter can render the technique inefficient. Also, simultaneous error in two copies of the logic value can result in propagation of the error value in the downstream logic. From a MVL perspective, TMR cannot be applied to MVL as it is ambiguous for MVL. As an alternative to TMR, Winstead et al. proposed RFB method, which is an improved version of TMR [19]. RFB method can correct single error faults and CMOS implementations were shown for both binary and ternary logic [19]. RFB method replaces the majority gate in TMR with Muller C elements. The idea of RFB method is based on the inherent fault correcting capabilities of the Muller C element. RFB method is based on two key ideas related to the Muller C element:
If the C element output is connected back to one of C element’s input, then noise in other C element’s input will be suppressed every time the C element evaluates.
Muller C element has inherent fault correcting properties. C element only changes its output state when all inputs have the same logic value. This property can be exploited for fault masking.
Triple modular redundancy (TMR).
\nFigure 12 shows an example of how a noisy input signal will be rendered less noisy, each time the signal passes through the C element. Suppose \n
Muller C element with feedback.
The magnitude of \n
From Eq. (12), we notice that as \n
The RFB circuit based on Muller C-element gates. The \n\nS\n\n gate is a storage element [21].
RFB setup phase.
RFB restoration phase.
Transient results of ternary CNTFET RFB method. The input sequence includes the six possible single error patterns. Signals \n\n\nx\n1\n\n\n and \n\n\nx\n2\n\n\n are correct, while \n\n\nx\n3\n\n\n is an error. The error is transferred to \n\n\ny\n2\n\n\n in the set-up phase and is corrected in the restoration phase [16].
This chapter discussed a technique for fault tolerance in MVL CNTFET logic. The described technique leverages the error suppression capability of Muller C element to correct single bit error in CNFTET MVL. To realize the technique, a ternary Muller C element is needed. This chapter also discussed the basics of MVL, provided an overview of CNTFET and also discussed the process variation in CNTFET.
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