Enzyme Research

Guest Editors: Raffaele Porta, Ashok Pandey, and Cristina M. Rosell

Enzymes as Additives or Processing Aids in Food Biotechnology

Enzymes as Additives or Processing Aids inFood Biotechnology

Enzyme Research

Enzymes as Additives or Processing Aids inFood Biotechnology

Guest Editors: Raffaele Porta, Ashok Pandey,and Cristina M. Rosell

Copyright © 2010 SAGE-Hindawi Access to Research. All rights reserved.

This is a special issue published in volume 2010 of “Enzyme Research.” All articles are open access articles distributed under the CreativeCommons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the originalwork is properly cited.

Enzyme Research

Editorial Board

Sabbir Ahmed, UKMario Amzel, USAVasu D. Appanna, CanadaDavid Ballou, USAUlrich Baumann, SwitzerlandFabrizio Briganti, ItalyJoaquim Cabral, PortugalGerald M. Carlson, USASunney I. Chan, USAChristopher Davies, USANarasimha Rao Desirazu, IndiaJohn David Dignam, USAColin Dingwall, UKJean-Marie Dupret, FrancePaul Engel, IrelandRoberto Fernandez Lafuente, Spain

D. M. G. Freire, BrazilVilmos Fulop, UKGiovanni Gadda, USAJ. Guisan, SpainMunishwar Nath Gupta, IndiaR. S. Gupta, CanadaAlbert Jeltsch, GermanyMarilyn S. Jorns, USAMari Kaartinen, CanadaEva Nordberg Karlsson, SwedenLeszek Kleczkowski, SwedenWilliam Konigsberg, USAH. Kuhn, GermanyDavid Lambeth, USAA-Lien Lu-Chang, USAPaul Malthouse, Ireland

Michael J. McLeish, USAPeter Moody, UKWilliam David Nes, USAToshihisa Ohshima, JapanMichael Page, UKJose Miguel Palomo, SpainRobert Pike, AustraliaRaffaele Porta, ItalyAlireza R. Rezaie, USAAli Akbar Saboury, IranEngin Serpersu, USAAssia Shisheva, USAR. D. Tanner, USAJohn J. Tanner, USAGianluigi Veglia, USAQi-Zhuang Ye, USA

Contents

Enzymes as Additives or Processing Aids in Food Biotechnology, Raffaele Porta, Ashok Pandey,and Cristina M. RosellVolume 2010, Article ID 436859, 2 pages

Enzymes in Food Processing: A Condensed Overview on Strategies for Better Biocatalysts,Pedro FernandesVolume 2010, Article ID 862537, 19 pages

Some Nutritional, Technological and Environmental Advances in the Use of Enzymes in Meat Products,Anne y Castro Marques, Mario Roberto Marostica Jr., and Glaucia Maria PastoreVolume 2010, Article ID 480923, 8 pages

Enzymatic Strategies to Detoxify Gluten: Implications for Celiac Disease, Ivana Caputo, MarilenaLepretti, Stefania Martucciello, and Carla EspositoVolume 2010, Article ID 174354, 9 pages

Uses of Laccases in the Food Industry, Johann F. Osma, Jose L. Toca-Herrera,and Susana Rodrıguez-CoutoVolume 2010, Article ID 918761, 8 pages

Fungal Laccases: Production, Function, and Applications in Food Processing, Khushal Brijwani,Anne Rigdon, and Praveen V. VadlaniVolume 2010, Article ID 149748, 10 pages

Potential Applications of Immobilized β-Galactosidase in Food Processing Industries, Parmjit S. Panesar,Shweta Kumari, and Reeba PanesarVolume 2010, Article ID 473137, 16 pages

Screen-Printed Carbon Electrodes Modified by Rhodium Dioxide and Glucose Dehydrogenase,Vojtech Polan, Jan Soukup, and Karel VytrasVolume 2010, Article ID 324184, 7 pages

Preparation of Antioxidant Enzymatic Hydrolysates from Honeybee-Collected Pollen Using PlantEnzymes, Margarita D. Marinova and Bozhidar P. TchorbanovVolume 2010, Article ID 415949, 5 pages

Characterization of Activity of a Potential Food-Grade Leucine Aminopeptidase from Kiwifruit,A. A. A. Premarathne and David W. M. LeungVolume 2010, Article ID 517283, 5 pages

SAGE-Hindawi Access to ResearchEnzyme ResearchVolume 2010, Article ID 436859, 2 pagesdoi:10.4061/2010/436859

Editorial

Enzymes as Additives or Processing Aids in Food Biotechnology

Raffaele Porta,1 Ashok Pandey,2 and Cristina M. Rosell3

1 Department of Food Science, University of Naples Federico II, Portici, 80055 Napoli, Italy2 Biotechnology Division, National Institute for Interdisciplinary Science and Technology, CSIR, Trivandrum, Kerala 695019, India3 Institute of Agrochemistry and Food Technology (IATA-CSIC), 46980 Paterna, Valencia, Spain

Correspondence should be addressed to Raffaele Porta, raffaele.porta@unina.it

Received 31 December 2010; Accepted 31 December 2010

Copyright © 2010 Raffaele Porta et al. This is an open access article distributed under the Creative Commons Attribution License,which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Essential in the metabolism of all living organisms, theenzymes are increasingly used to drive chemical reactionsoutside their natural localization. In particular, the use of thebiocatalysts as food additives and in processing raw materialshas been practiced for a long time. In fact, enzymaticpreparations from the extracts of plants or animal tissueswere used well before much was known about the nature andproperties of enzymes.

Food industry is constantly seeking advanced technolo-gies to meet the demand of the consumers, and enzymes havelong been used by the industrial product makers as majortools to transform the raw materials into end-products.Their clean label (GRAS, generally recognized as safe)consideration from the legal point of view has promptedtheir extensive use in food technology. When purified andadded to food preparations, several enzymes are able toimprove their flavor, texture, digestibility, and nutritionalvalue. However, it was not until the mid of the past centurythat the rapid development in protein technology occurred,and only in the last 30 years, the use of commercial enzymeshas grown in the food industry, progressively becoming animportant aspect of the manufacturing of meat, vegetables,fruit, baked goods, milk products, and both alcoholic andnonalcoholic beverages. As a matter of fact, an increasingnumber of articles, mostly describing the enhanced productyields, have been published during the last ten years, bothin food and beverage manufacturing. Moreover, since itis desirable in different branches of food technology tochange the physical and chemical properties of protein, manypreviously unexplored enzymes are currently employed toproduce a variety of foods in which the biocatalysts replacepotentially carcinogenic or otherwise harmful chemicals.This includes also new methods in which the characteristics

of natural products are altered to fit the nutritional ortechnological needs changing.

The economic benefit of using technical enzyme prepa-rations lies in lowered process costs, in the reduction of theenvironmental impact by making use of renewable resources,and often in increasing the quality of the products. Also,preservation makes a significant impact on the quality offood as well of beverages. It is well known, for example,that modern processes convert juices into concentrates that,except for aroma, can be stored for a long time without lossin quality. Stabilizing flavor and color is also an example ofimproved preservation. Finally, the advent of biotechnologyhas also allowed significant refinements in the methodologiesoffering unpredictable solutions to many persistent problemsand opening up exciting new possibilities. Among these,enzymes are proposed as exemplary agents of “green”technology since they can also be used either to treat thebiological wastes or to prevent their formation. Currentlyused enzymes sometimes originate in animals and plants butmost come from a range of beneficial microorganisms. Thus,numerous purified enzymes are now being widely used notonly in food processing but also as food additives. In thisrespect, it is noteworthy that the enzymes, like all proteins,can cause reactions only when people have been sensitizedthrough exposure to large quantities. Therefore, since theirlevels in the food are generally very low, the enzymes arehighly unlikely to cause allergies.

This special issue of Enzyme Research is devoted tocontribute to highlight some expanding fields of enzymeapplications in food technology, mostly explaining how somedifferent biocatalysts bring advantages in some food proces-sing improvement and innovation. It comprises six reviewarticles and three research articles. The first review article

2 Enzyme Research

is a condensed and concise overview on the applications ofenzymes in food and feed processing, outlining the develop-ment of better biocatalysts through microbial screening,protein engineering, and immobilization techniques. Thesecond review article summarizes the nutritional, techno-logical, and environmental advances in meat products and,in particular, the application of the proteolytic enzymes,phytases, and transglutaminase in the meat industry. Trans-glutaminase, as well as bacterial-derived endopeptidases, arethe subject of the third review article which reports the mostrecent developments of the attempts to detoxify gluten. Thefourth and the fifth review articles describe, respectively,the uses of laccases as additives in food and beverageprocessing and the production, function, and applicationsin food industry of fungal laccases. The last review articlehas been focused on the different types of techniques usedfor the immobilization of β-galactosidase and its potentialapplications in food and dairy processing industries. Thethree research articles describe (i) a new glucose biosensorbased on a screen-printed carbon electrode modified byglucose dehydrogenase immobilized on its surface, (ii)the preparation of antioxidant hydrolysates of honeybee-collected pollen by using proteinase and aminopeptidasesof plant origin, and (iii) the characterization of a potentialfood-grade leucine aminopeptidase extracted from kiwifruit.

We sincerely hope that the present volume may re-present only the first of a special issue series in whichEnzyme Research will periodically stimulate authors topublish the highlights and original research articles reportinghow enzymes bring new advantages in food preparationimprovement and innovation.

Raffaele PortaAshok Pandey

Cristina M. Rosell

SAGE-Hindawi Access to ResearchEnzyme ResearchVolume 2010, Article ID 862537, 19 pagesdoi:10.4061/2010/862537

Review Article

Enzymes in Food Processing: A Condensed Overview onStrategies for Better Biocatalysts

Pedro Fernandes

Institute for Biotechnology and Bioengineering (IBB), Centre for Biological and Chemical Engineering, Instituto Superior Tecnico,Avenue Rovisco Pais, 1049-001 Lisboa, Portugal

Correspondence should be addressed to Pedro Fernandes, pedro.fernandes@ist.utl.pt

Received 7 July 2010; Accepted 1 September 2010

Academic Editor: Cristina M. Rosell

Copyright © 2010 Pedro Fernandes. This is an open access article distributed under the Creative Commons Attribution License,which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Food and feed is possibly the area where processing anchored in biological agents has the deepest roots. Despite this, processimprovement or design and implementation of novel approaches has been consistently performed, and more so in recent years,where significant advances in enzyme engineering and biocatalyst design have fastened the pace of such developments. This paperaims to provide an updated and succinct overview on the applications of enzymes in the food sector, and of progresses made,namely, within the scope of tapping for more efficient biocatalysts, through screening, structural modification, and immobilizationof enzymes. Targeted improvements aim at enzymes with enhanced thermal and operational stability, improved specific activity,modification of pH-activity profiles, and increased product specificity, among others. This has been mostly achieved throughprotein engineering and enzyme immobilization, along with improvements in screening. The latter has been considerablyimproved due to the implementation of high-throughput techniques, and due to developments in protein expression andmicrobial cell culture. Expanding screening to relatively unexplored environments (marine, temperature extreme environments)has also contributed to the identification and development of more efficient biocatalysts. Technological aspects are considered, buteconomic aspects are also briefly addressed.

1. Introduction

Food processing through the use of biological agents ishistorically a well-established approach. The earliest appli-cations go back to 6,000 BC or earlier, with the brewing ofbeer, bread baking, and cheese and wine making, whereasthe first purposeful microbial oxidation dates from 2,000BC, with vinegar production [1–3]. Coming to modern days,in the late XIX, century Christian Hansen reported the useof rennet (a mixture of chymosin and pepsin) for cheesemaking, and production of bacterial amylases was startedat Takamine (latter to become part of Genencor). Pectinaseswere used for juice clarification in the 1930s, and for a shortperiod during World War II, invertase was also used for theproduction of invert sugar syrup in a process that pioneeredthe use of immobilized enzymes in the sugar industry [1].Still, the large-scale application of enzymes only becamereally established in the 1960s, when the traditional acidhydrolysis of starch was replaced by an approach based in

the use of amylases and amyloglucosidases (glucoamylases), acocktail that some years latter would include glucose (xylose)isomerase [1, 2, 4, 5]. From then on, the trend for thedesign and implementation of processes and production ofgoods anchored in the use of enzymes has steadily increased.Enzymes are currently among the well established productsin biotechnology [6], from US $1.3 billion in 2002 to US $4billion in 2007; it is expected to have reached US $5.1 billionin a rough 2009 year, and is anticipated to reach $7 billionby 2013 [3, 5, 7–9]. In the overall, this pattern correspondsto a rise in global demand slightly exceeding 6% yearly[7, 9]. Part of this market is ascribed to enzymes used inlarge-scale applications, among them are those used in foodand feed applications [10]. These include enzymes used inbaking, beverages and brewing, dairy, dietary supplements,as well as fats and oils, and they have typically beendominating one, only bested by the segment assigned totechnical enzymes [11, 12]. The latter includes enzymes inthe detergent, personal care, leather, textile and pulp, and

2 Enzyme Research

paper industries [10, 13]. A recent survey on world salesof enzymes ascribes 31% for food enzymes, 6% for feedenzymes and the remaining for technical enzymes [11]. Arelatively large number of companies are involved in enzymemanufacture, but major players are located in Europe, USAand Japan. Denmark is dominating, with Novozymes (45%)and Danisco (17%), moreover after the latter taking overGenencor (USA), with DSM (The Netherlands) and BASF(Germany) lagging behind, with 5% and 4% [10, 11, 14].The pace of development in emerging markets is suggestivethat companies from India and China can join this restrictedparty in a very near future [15–17].

2. Relevant Enzymes: Tapping forImproved Biocatalysts

2.1. General Aspects and the Screening Approach. Roughlyall classes of enzymes have an application within the foodand feed area, but hydrolases are possibly the prevalent one.Representative examples of the enzymes and their role infood and feed processing are given in Table 1. The widespreaduse of enzymes for food and feed processing is easily under-standable, given their unsurpassed specificity, ability to oper-ate under mild conditions of pH, temperature and pressurewhile displaying high activity and turnover numbers, andhigh biodegradability. Enzymes are furthermore generallyconsidered a natural product [18, 19]. The whole contributesfor developing sustainable and environmentally friendlyprocesses, since there is a low amount of by-products,hence reducing the need for complex downstream processoperations, and the energy requirements are relatively low.Life-cycle assessment (LCA) has confirmed, that within therange of given practical case studies, including food and feedprocessing, the implementation of enzyme-based technologyhas a positive impact on the environment [3]. LCA is amethodology used to compare the environmental impactof alternative production technologies while providing thesame user benefits [20].

Some of the broad generalizations on the limitations ofenzymes for application as biocatalysts in commercial scale,namely, their high cost, low productivity and stability, andnarrow range of substrates, have been rebutted [21, 22].Aiming at improving the performance of biocatalysts forfood and feed applications, particular care has been given toincreasing thermal stability, enhancing the range of pH withcatalytic activity and decreasing metal ions requirements, aswell as to overcoming the susceptibility to typical inhibitorymolecules. Some examples of strategies taken to improve theperformance of relevant enzymes for food and feed are givenin Table 2. Along with these different strategies focused onthe enzyme molecule (namely, protein engineering, enzymeimmobilization), the developments in recombinant DNAtechnology that occurred in the 1980s also had a hugeimpact on the application of enzymes in food and feed. Byallowing gene cloning in microorganisms compatible withindustrial requirements, this methodology enabled cost-feasible production of enzymes that were naturally pro-duced in conditions that prevented large-scale application

(namely, enzymes from plant or animal cells, such as trans-glutaminase or even slow-growing microorganisms). Whensuccessfully implemented, the undertaken approaches allow:(a) continuous operations at relatively high temperatures;(b) eased implementation of enzyme cascade, given thereduced need for processing the reaction media (pH adjust-ments; metal ion removal/addition) throughout the interme-diate steps of a multistep biotransformation (namely, starchto high fructose syrup); and (c) the use of raw substrates,preferably as high-concentrated solutions, hence cuttingback in costs related to upstream processing and increasingproductivity [4, 23, 24]. Methodologies with a high level ofparallelization, anchored in computer-monitored microtiterplates equipped with optic fibers and temperature controlhave also been developed. These provide high-throughputcapability for a speedy and detailed characterization of theperformance of enzymes [25]. Particular focus was given tothe prediction of the long-term stability of enzymes undermoderate conditions using short-term runs (up to 3 hours).

One of the methodologies to obtain improved bio-catalyst relies on in-vitro modifications, which will beaddressed latter in this paper; another approach relies onscreening efforts, which has been consistently undertaken,as summarized recently [26–31]. Some focus is given toextremophiles, particularly thermophiles, since operationat high temperatures (roughly above 45–50◦C) minimizesthe risk of microbial contamination, a particularly deli-cate matter under continuous operation. Furthermore, theextension of some reactions in relevant food applicationsis favored at relatively high temperatures (namely, iso-merization of glucose to fructose), although care shouldbe taken to avoid an operational environment that maylead by-product formation (namely, Maillard reactions).Examples of screened enzymes include the isolation ofamylases, with some of them being calcium independent[32–38]; amylopullulanases [39]; fructosyltransferases [40];glucoamylases [41]; glucose (xylose) isomerases [42, 43];glucosidases [44, 45]; inulinases [46–49]; levansucrases [50];pullulanases [51, 52]; and xylanases [53, 54]. Other examplesof these enzymes, with some of which able to retain stabilityunder temperatures of 90◦C or higher, were reviewed byGomes and Steiner [55]. The majority of enzymes used infood and feed processing is of terrestrial microbial origin,and screening-efforts for isolation of promising enzyme-producing strains have accordingly been performed in suchbackground [3, 5, 56]. From some years now, marineenvironment has also been tapped as a source for usefulenzymes from either microbial or higher organisms origin[57–60]. This latter environment has allowed the isolationof some promising biocatalysts, such as the heat-stableinvertase/inulinase from Thermotoga neapolitana DSM 4359or inulinase from Cryptococcus aureus [61–63], amylolyticenzymes, glucosidases and proteases from severalgenera [32,44, 45, 64, 65], esterase from Vibrio fischeri [66], and glycosylhydrolases [67, 68]. Other examples of useful enzymes forfood and feed, but isolated from higher organisms [59, 69],are given in Table 3. Some of these enzymes are actuallypsychrophiles, hence performing best at low temperatures[30].

Enzyme Research 3

Operation at low temperatures is also welcome since italso reduces the risk of microbial contamination, enablessome processes to be carried out with minimal deteriorationof the raw material. These include protein processing, suchas cheese maturing and milk coagulation with proteases [59,80]; milk processing with lactase for lactose-free milk [81–83]; clarification of fruit juices with pectinases to produceclear juice [84]; or production of oligosaccharides [85].

Since extremophiles are often difficult to grow undertypical laboratory conditions if not nonculturable at all,different approaches have been developed in order to assessthe potential of enzymes from such microorganisms. Oneapproach relies on the generation and screening of targetgenes from DNA libraries, which can be obtained frommixed microbial population from environmental samples.Recombinant microorganisms can then be obtained usingmesophiles as hosts where the genes of interest fromextremophiles have been expressed [86]. In order to screenthe huge number of DNA-libraries typically generated forthe intended property, high-throughput methods have beenimplemented [87]. These methods are also widely used whenprotein engineering is carried out. This will be addressed inthe following section.

Several enzymes (namely, α-amylases; pullulanases) cur-rently used in food processing, namely, in starch hydrolysis,are actually produced by recombinant microorganisms.Despite some complexity in the implementation of theiruse in large-scale applications, partly resulting from lack ofuniformity in the US and EU legislation, quite a few enzymepreparations have been accepted for industrial use [88, 89].

3. Improving Biocatalysts:Beyond Screening

Taking advantage of the knowledge gathered on molecularbiology, high-throughput processing, and computer-assisteddesign of proteins, in-vitro improvement of biocatalystshave been consistently implemented [90–93]. Some of theresearch efforts in this area has focused on the biochemi-cal and molecular mechanisms underlying the stability ofenzymes from extremophiles [31, 94–96]. Such knowledgeis also particularly useful for protein engineering of knownenzymes, aiming at enhancing stability without compro-mising catalytic activity [97]. Enhancing the stability ofenzymes is of paramount importance when implementationof industrial processes is foreseen, since it allows for reducingthe amount of enzyme used in the process. Given thatthermostability is determined by a series of short- andlong-range interactions, it can be improved by severalsubstitutions of amino acids in a single mutant, where thecombination of each individual effect is usually roughlyadditive [98]. The targeted improvements have not beenrestricted to thermostability, but they have also addressedother features, such as broadening the range of pH where theenzyme is active, or lessening the temperature of operationwhile retaining high activity [91, 99].

Two methodologies can be used for protein engineering[97].

(i) The first is directed evolution of enzymes, throughrandom mutagenesis and recombination, where theenvironmental adaptation is reproduced in-vitro in amuch hastened timescale, towards the optimizationof the intended property. In order to control thepathway of the process, either a screening test forthe assessed feature is performed after each round ofmodification, or selective pressure is applied [100–102]. This methodology, which allows for a highthroughput, has been extensively applied, aiming formore efficient biocatalysts [103–106]. Some relevantexamples in the area of food and feed processinginclude the following.

(1) The first is the enhancement of the activity of thehyperthermostable glucose (xylose) isomerase fromThermotoga neapolitana at relatively low temperatureand pH, without decay in thermostability [107].The enzyme from the parent strain is highly activeat 97◦C, but it retains only 10% of its activity at60◦C, and requires neutral pH for optimal activity.This pattern is often reported when glucose iso-merases from hyperthermophilic strains operate inmesophilic environments. Large-scale glucose iso-merization is carried out at 55–60◦C and slightlyalkaline pH [1, 31]. This set of conditions results fromthe optimal range of pH (typically 7.0 to 9.0) andtemperature (60 to 80◦C) for glucose isomerizationdisplayed by most of the glucose isomerases used,combined with process boundary conditions. Thelatter result from by-product and color formationoccurring when the reaction is carried out at alka-line pH and high temperatures [31, 108]. There istherefore interest in selecting an enzyme able tooperate efficiently at temperatures close to thosecurrently used but at a lower pH. The mutantglucose isomerase 1F1 obtained by Sriprapundh andcoworkers displayed a roughly 5-fold higher activityat 60◦C and pH 5.5, when compared with the parentT. neapolitana isomerase, and was more thermostablethan the wild type isomerase [104, 107]. The acti-vation energy required by the triple 1F1 mutant(V185T/L282P/F186S) was roughly half of the wild-type, hence allowing for high activity at relativelylow temperatures [107]. The encouraging resultsobtained suggest the soundness of the approach toobtain a mutant glucose isomerase competitive withthose currently used, while being able to operate in aslightly acidic environment and 60◦C.

(2) The second is the enhancement of the thermostabilityof the maltogenic amylase from Thermus sp. IM6501[109], of the amylosucrase from Neisseria polysac-charea [110], of the glucoamylase from Aspergillusniger [111], of a phytase from Escherichia coli [112,113], and of a xylanase from Bacillus subtilis [114].Amylases and glucoamylases are enzymes used instarch processing, which involves temperatures typ-ically in excess of 60◦C; hence, improving thermalstability without decreasing enzyme activity is of

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Table 1: An overview of enzymes used in food and feed processing (adapted from [10, 12, 13, 68]).

Class Enzyme Role

OxidoreductasesGlucose oxidase Dough strengthening

Laccases Clarification of juices, flavor enhancer (beer)

Lipoxygenase Dough strengthening, bread whitening

Transferases

CyclodextrinCyclodextrin production

Glycosyltransferase

Fructosyltransferase Synthesis of fructose oligomers

Transglutaminase Modification of viscoelastic properties, dough processing, meat processing

Hydrolases

Amylases

Starch liquefaction and sachcarification

Increasing shelf life and improving quality by retaining moist, elastic and softnature

Bread softness and volume, flour adjustment, ensuring uniform yeastfermentation

Juice treatment, low calorie beer

Galactosidase Viscosity reduction in lupins and grain legumes used in animal feed, enhanceddigestibility

Glucanase Viscosity reduction in barley and oats used in animal feed, enhanced digestibility

Glucoamylase Saccharification

Invertase Sucrose hydrolysis, production of invert sugar syrup

Lactase Lactose hydrolysis, whey hydrolysis

Lipase Cheese flavor, in-situ emulsification for dough conditioning, support for lipiddigestion in young animals, synthesis of aromatic molecules

Proteases (namely, chymosin, papain)Protein hydrolysis, milk clotting, low-allergenic infant-food formulation,enhanced digestibility and utilization, flavor improvement in milk and cheese,meat tenderizer, prevention of chill haze formation in brewing

Pectinase Mash treatment, juice clarification

Peptidase Hydrolysis of proteins (namely, soy, gluten) for savoury flavors, cheese ripening

Phospholipase In-situ emulsification for dough conditioning

Phytases Release of phosphate from phytate, enhanced digestibility

Pullulanase Saccharification

Xylanases Viscosity reduction, enhanced digestibility, dough conditioning

Lyases Acetolactate decarboxylase Beer maturation

Isomerases Xylose (Glucose) isomerase Glucose isomerization to fructose

relevance. Starch liquefaction is performed at 105◦Cin the presence of α-amylase, upon which the effluentreaction stream has to be cooled to 60◦C, so that glu-coamylases can be used. In order to avoid, or at leastminimize, the cooling step, thermostable glucoamy-lases are aimed at. Wang and coworkers obtained amultiply-mutated enzyme (N20C, A27C, S30P, T62A,S119P, G137A, T290A, H391Y), which displayed a5.12 kJ mol−1 increase in the free energy of thermalinactivation, when compared to the wild type, thusresulting in the enhanced thermal stability of themutant. Furthermore specific activities and catalyticefficiencies remained unaltered, when mutant andwild type were compared [111]. Kim and coworkersobtained also a multiply-mutated amylase (R26Q,S169N, I333V, M375T, A398V, Q411L, P453L) whichdisplayed an optimal reaction temperature 15◦Chigher than that of the wild-type and a half-life ofroughly 170 min at 80◦C, a temperature at which

the wild-type ThMA was fully inactivated in lessthan 1 minute. However, one of the mutationsmost accountable for enhanced thermal stability,M375T, close to the active site, also led to a 23%decrease in specific activity, as compared to the wildtype [109]. The amylosucrase engineered by Emondand coworkers was a double mutant (R20C/A451T),displaying a 10-fold increase in the half-life at 50◦Ccompared to the wild-type enzyme. Actually, themutant was claimed to be the only amylosucraseusable at 50◦C. At the latter temperature, the mutantenabled the synthesis of amylose chains twice as longas those obtained by the wild-type enzyme at 30◦C,for sucrose concentrations of 600 mM. The mutantthus allowed for a process with increased yield inamylose chains (31 g L−1), lower risk of contami-nation, enhanced substrate and product solubilityand overall productivity [110]. Phytases are added toanimal feeds to improve phosphorus nutrition and

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Table 2: Some examples of strategies undertaken to improve the performance of enzymes with applications in food and feed.

Enzyme RoleTargetedimprovement

Strategy/comments Reference

α-amylaseStarch liquefaction Thermostability

Protein engineering through site-directedmutagenesis. Mutant displayed increasedhalf-life from 15 min to about 70 min (100◦C).

[70]

Starch liquefaction Activity

Directed evolution. After 3 rounds the mutantenzyme from S. cerevisiae displayed a 20-foldincrease in the specific activity when comparedto the wild-type enzyme.

[71]

Baking pH-activity profileProtein engineering through site-directedmutagenesis

[72]

l-arabinoseisomerase

Tagatose production pH-activity profile Protein engineering through directed evolution [73]

Glucoamylase Starch saccharificationSubstrate specificity,thermostability andpH optimum

Protein engineering through site-directedmutagenesis

[74]

Lactase Lactose hydrolysis Thermostability Immobilization [75]

Pullulanase Starch debranching Activity Protein engineering through directed evolution [76]

Phytase Animal feed pH-activity profileProtein engineering through site-directedmutagenesis

[77]

Xylose (glucose)isomerase

Isomerization/epimerizationof hexoses, pentoses andtetroses

pH-activity profile

Protein engineering through directedevolution. The turnover number on D-glucosein some mutants was increased by 30%–40%when compared to the wild type at pH 7.3.Enhanced activities are maintained betweenpH 6.0 and 7.5.

[78]

Substrate specificity

Protein engineering through site-directedmutagenesis. The resulting mutant displayed a3-fold increase in catalytic efficiency withL-arabinose as substrate.

[79]

Table 3: Examples of enzymes isolated from various marine higher organisms with potential of application in food and feed (adapted from[68, 69]).

Class Enzyme Source

Transferases Transglutaminase Muscles of atka mackerel (Pleurogrammus azonus), botan shrimp (Pandalus nipponensis), carp(Cyprinus carpio), rainbow trout (Oncorhynchus mykiss), scallop (Patinopecten yessoensis).

Hydrolases

Amylase

Gilt-head (sea) bream (Sparus aurata), found in Mediterranean sea and coastal North AtlanticOcean.

Turbot (Scophthalmus maximus), found mostly in Northeast Atlantic Ocean, Baltic, Black andMediterranean seas, and Southeast the Pacific Ocean

Deepwater redfish (Sebastes mentella, found in North Atlantic).

Chymotrypsin Atlantic cod (Gadus morhua), crayfish, white shrimp.

Pepsin Arctic capelin (Mallotus villosus), Atlantic cod (Gadus morhua).

Protease

Marine sponges Spheciospongia vesperia, found in Caribbean sea and South Atlantic, close toBrazil, and Geodia cydonium, found in Northeast Atlantic Ocean and Mediterranean sea.

Mangrove crab (Scylla serrata), found in estuaries and mangroves of Africa, Asia and Australia.

Sardine Orange roughy (Hoplostethus atlanticus)

to reduce phosphorus excretion, by promoting thehydrolysis of phytate into myoinositol and inorganicphosphate. Thermal stable enzymes are needed, sincefeed pelleting is carried out at high temperature(60 to 80◦C). Phytases produced by thermophilesdo not provide a suitable approach, since they havelow activity at the physiological temperature of

animals [115]. E. coli phytases, which are appealing toindustrial application, due to the acidic pH optimum,specificity phytate, and resistance to pepsin digestion,were thus engineered in order to improve theirthermal stability, without compromising the kineticparameters. As a result, mutants were obtained,with roughly 20% increased thermostability at 80◦C

6 Enzyme Research

improved overall catalytic efficiency (kcat, turnovernumber/KM, Michaelis constant) within 50 to 150%,as compared to the wild type. No significant changesin the pH activity profile were observed, but forsome mutants, containing a K46E substitution, thatdisplayed a decrease in activity at pH 5.0 [112, 113].Xylanases catalyze the cleavage of β1,4 bonds in xylanpolymers. Accordingly, these enzymes can be usedin dough making, in baking, in brewing and inanimal feed compositions. When the latter containcereals (namely, barley, maize, rye or wheat), or cerealby-products, xylanases improve the break-down ofplant cell walls, which favors the ingestion of plantnutrients by the animals and consequently enhancesfeed consumption and growth rate. Furthermore,the use of xylanases decreases the viscosity of xylan-containing feeds [116, 117]. As referred for phytases,the formulation of commercial feed often involvessteps at high temperatures. Xylanases added to thethe formulations hence have to withstand theseconditions, while they are to display high activityat about 40◦C, which is the temperature in theintestine of animals. However, most xylanases areinactive at temperatures exceeding 60◦C, hence theneed for enhancing thermal stability [114, 117].Miyazaki and coworkers obtained a triple-mutantxylanase (Q7H, N8F, and S179C) which retained fullactivity for 2 hours at 60◦C, whereas the wild-typeenzyme was inactivated within 5 minutes under thesame conditions. The mutation also led to a 10◦Cincrease in the optimal temperature for reaction andenhanced activity at higher temperatures, albeit at thecost of decreased activity at lower temperatures, ascompared to the wild-type enzyme [114].

(3) Third is the enhancement of the activity of theamylosucrase from Neisseria polysaccharea [118].Amylosucrases can be used for the modification orsynthesis of amylose-type polymers from sucrose, buttheir industrial application is somehow thwarted bythe low catalytic efficiency on sucrose and by sidereactions leading to the formation of sucrose isomers.Van der Veen and co-works engineered mutantenzymes through error-prone PCR that displayedincreases in activity up to 5-fold and in overallcatalytic efficiency up to 2-fold, when compared tothe wild-type enzyme. Furthermore, the mutantswere able to produce amylose polymers from 10 mMsucrose on, unlike the wild-type enzyme [118]. Theirwork provides an illustrative example on the useof random mutagenesis and recombination for theenhancement of the catalytic properties of enzymeswith application on food and feed. Another examplewas provided by Tian and coworkers who engineereda phytase from Aspergillus niger 113 through geneshuffling, to obtain mutants with enhanced catalyticproperties [119]. Hence, K41E and E121F substitu-tions allowed for increases in the specific activity of2.5- and 3.1-fold, and of affinity for sodium phytate,

as expressed by decreases in KM of roughly 35%and 25%, as compared to the wild-type enzyme.Furthermore, the overall catalytic efficiency of themutants increased 1.4- and 1.6-fold as compared tothe wild type.

Other examples can be found elsewhere [120, 121].

(ii) The second methodology underlines that rationalpinpoint modifications in one or more amino acidsare made, where these changes are predicted to bringalong the envisaged improvement in the targetedenzyme function. The alterations promoted are per-formed based on the growing knowledge on thestructure and functions of enzyme. Information onthis matter mostly comes from bioinformatics, whichprovides data on amino-acid propensities and onprotein sequences. Adequate processing of the dataenable the output of generalized rules predictingthe effect of mutations on enzyme properties. Alsoused are molecular potential functions, which, onceimplemented, enable the prediction of the effectof mutations in enzyme structure [97]. Compu-tational tools used for enzyme engineering havebeen recently reviewed [122]. Enzyme engineeringthrough molecular simulations requires structuraldata from the native enzyme, which can be preferablyobtained from crystallography or NMR. Otherwisea model is built based on known enzyme structureswith homologous sequences [90]. Computationalmethods are also welcome in directed evolution,as a tool to better lead the random mutagenesis[97]. Ultimately this approach is put into practiceby producing a site-directed mutant, where selectedamino acids are replaced with those suggested fromthe outcome of modeling.

Some relevant examples of this strategy in the areaof food and feed processing are given. These mostlyaim to improve thermal stability and/or catalytic effi-ciency and/or to modify the range of pH/temperaturewhere the enzyme is active—goals that were alreadyreferred to when examples of enzyme modificationsusing random mutagenesis were addressed.

(1) The first example underlines the enhancement of thethermostability of the recombinant glucose (xylose)isomerase from Actinoplanes missouriensis [123, 124]and of glucose (xylose) isomerase from Streptomycesdiastaticus [125]; of amylases from Bacillus spp. [126,127]; and of glucoamylase from Aspergillus awamori[128]. The mutant isomerase from A. missouriensisdisplayed an enhanced thermal stability, alongsidewith improved stability at different pH, as comparedwith the original enzyme, with no changes in catalyticproperties [123, 124]. The double mutant isomerase(G138P, G247D) displayed a 2.5-fold increase inhalf-life, and additionally a 45% increase in thespecific activity, when compared to the wild type.Such features were ascribed to increased molecular

Enzyme Research 7

rigidity due to the introduction of a proline inthe turn of a random coil [125]. Multiply-mutatedamylases obtained by Declerck and coworkers dis-played considered enhanced thermal stability. Basedon the temperature at which amylase initial activityis reduced by 50% for a 10-minute incubation,this parameter went as high as 106◦C, as comparedto 83◦C for the wild-type strain. Furthermore, thethermal stabilization was not accompanied by adecrease in the catalytic activity [126]. The work byLin and coworkers on amylase mutants from Bacillussp. strain TS-23 highlighted the relevance of E219for the thermal stability of the enzyme [127]. Themutated glucoamylases engineered by Liu and Wangallowed to establish the role of several intermolecularinteractions in thermal stability of these enzymes.Thermostable enzymes were obtained through theintroduction of disulfide bonds in highly flexibleregion in the polypeptide chain of the enzyme, as wellas by the introduction of more hydrophobic residues-stabilized α-helices. Data gathered also showed thatcare had to be taken not to disrupt the hydrogen bondand salt linkage network in the catalytic center as aresult of mutagenesis, for this could lead to a decreasein the specific activity and overall catalytic efficiency[128].

(2) The second example underlines the enhancement ofthe pH-activity profile and of the thermostabilityof phytase from A. niger. This was achieved bycombining several individual mutations that allowedfor mutants that were quite active at pH 3.5. Effi-cient operation in the stomach of simple-stomachedanimals where phytate hydrolysis mostly occurs at apH around 3.5, and the wild type was ineffective, wasthus enabled. Furthermore, the hydrolytic activity ofthe mutants at pH 3.5 exceeded in roughly 1.5-foldthat of the parent one at pH 5.5, which was theoptimum of the latter. Mutants also retained higherresidual activity after incubation within 70 to 100◦C,as compared to the wild type. The work demonstratesthat cumulative improvements in pH activity andthermostability through mutation are compatible inthis phytase; see [129].

(3) The third example underlines the modification ofthe temperature- and pH activity profile of the l-arabinose isomerase from Bacillus stearothermophilusUS100 [130]. l-Arabinose isomerases catalyze theconversion of l-arabinose to l-ribulose in-vivo, butin-vitro they also isomerize d-galactose into d-tagatose [130]. The latter keto-hexose is being usedas a low-calorie bulk sweetener, since its taste andsweetness are roughly equivalent to sucrose, but thecaloric value is only 30% of that of sucrose [131,132]. Although several thermostable l-arabinose iso-merases have been isolated and characterized, most ofthese display an alkaline pH optimum. For industrialapplication this presents the same drawbacks ofby-product and color formation referred to when

the random mutation of glucose isomerases wasaddressed. Hence, again arises the need for enzymesable to isomerize l-arabinose in an acidic environ-ment and at relatively low temperature, 60 to 70◦C.Operation within the latter temperature range alsorules away the use of divalent ions, which stabilizeisomerases at high temperatures [133, 134]. Rhimiand coworkers engineered two individual mutants,harboring each N175H and Q268K mutations. Theseled to broader optimal temperature range within 50to 65◦C and to enhanced stability in acidic media,respectively, when compared to the wild type. Anengineered double mutant, harboring both modifi-cations, displayed optimal activity within a pH rangeof 6.0 to 7.0 and a temperature range within 50–65◦C. Such set of operational conditions matches thetargeted goals and again shows that the basis for pH-activity profile and thermostability in l-arabinoseisomerase are quite independent and compatible.Cumulative enhancements in both properties in thesame enzyme were thus possible [134]. A similarpattern was also observed in the previous examplededicated to a mutant phytase.

(4) The fourth example underlines the modification ofthe product profile of inulosucrase from Lactobacillusreuteri [135] and from B. subtilis [136]. Inulosucrasesare used to synthesize fructooligosaccharides or fruc-tan polymer from sucrose. The transglycosylationcatalyzed by the inulosucrase from L. reuteri leadsto a wide range of fructooligosaccharides alongsidewith minor amounts of an inulin polymer. In orderto minimize the dispersion in the products obtained,mutants R423K and W271N were obtained, whichallowed the synthesis of a significant amount ofpolymer and a lower amount of oligosaccharide,without significantly affecting the catalytic activity,when compared with the wild type. The data gatheredshowed that the −1 subsite in the inulosucrasefrom L. reuteri has a key role in the determinationof the size of the products obtained [135]. Ortiz-Soto and coworkers also showed that the productprofile of transfructosylation reactions could beadequately tuned through modification of targetresidues of an inulosucrase from B. subtilis. Theseauthors established the effect of mutations on thereaction specificity (hydrolysis/transfructosylation),molecular weight and acceptor specificity. For exam-ple, engineered mutants R360S, Y429N and R433Aonly synthesized oligosaccharides, whereas the wildtype synthesized levan, since the former are morehydrolytic. On the other hand these mutationsreduced the affinity for sucrose, and thermal stability,when compared to the wild type [136].

(5) The fifth example underlines the enhancement ofthe product profile of cyclodextrin glycosyltrans-ferases (CGTase) from differentgenera [137, 138].These enzymes promote the production of cyclodex-trins, α(1→ 4) linked oligosaccharides form starch,

8 Enzyme Research

through an intramolecular transglycosylation reac-tion. In the process, a starch oligosaccharide is cleavedand cleaved and the resulting reducing-end sugaris transferred to the non-reducing-end sugar of thesame chain [137]. The resulting cyclodextrin mayconsist of six, seven or eight, which are accord-ingly termed α, β, or γ-cyclodextrin, respectively.Given their ability to form inclusion complexes withsmall hydrophobic molecules, they are of interestfor both industrial and research applications. Wild-type CGTases typically produce a mixture of thethree cyclodextrins when incubated with starch. Thepurification of a given cyclodextrin from the reactionmixture requires several additional steps, includingselective complexation with organic solvents, whichmay prove restrictive for cyclodextrin applicationsinvolving human consumption [139, 140]. Thereis therefore a clear interest in obtaining a mutantCGTase capable of producing a particular type ofcyclodextrin in a high rate. Van der Veen and cowork-ers engineered a double-mutant (Y89D/S146P) ofCGTase from Bacillus circulans which displayed a 2-fold increase in the production of α-cyclodextrin anda marked decrease in β-cyclodextrin when comparedto the wild type. From the data gathered, theauthors suggested that hydrogen bonds (S146) andhydrophobic interactions (Y89), are likely to playa key role in to the size of cyclodextrin productsformed, and that changes in sugar-binding subsites−3 and−7 may result in mutant CGTases with alteredproduct specificity [137]. Li and coworkers were alsoable to obtain CGTase mutants from Paenibacillusmacerans strain JFB05-01 with increased specificityfor α-cyclodextrin, through mutations at subsite−3. In particular, double mutant D372K/Y89R dis-played a 1.5-fold increase in the production ofα-cyclodextrin, and a significant (roughly 45%)decrease in the production of β-cyclodextrin whencompared to the wild-type enzyme [138].

The two methods are not mutually exclusive and meth-odologies for engineering of enzymes can assemble bothstrategies [141].

Upon identification of the most adequate enzyme, thiscan be formulated adequately for better process integration.One of the most widely considered approaches for suchformulation is enzyme immobilization.

4. Immobilization

There are several issues that can be lined up to sustainenzyme immobilization. It allows for high-enzyme loadwith high activity within the bioreactor, hence leadingto high-volumetric productivities; it enables the controlof the extension of the reaction; downstream process issimplified, since biocatalyst is easily recovered and reused;the product stream is clear from biocatalyst; continuousoperation (or batch operation on a drain-and-fill basis) andprocess automation is possible; and substrate inhibition can

be minimized. Along with this, immobilization preventsdenaturation by autolysis or organic solvents, and can bringalong thermal, operational and storage stabilization, pro-vided that immobilization is adequately designed [142, 143].Immobilization has some intrinsic drawbacks, namely, masstransfer limitations, loss of activity during immobilizationprocedures, particularly due to chemical interaction orsteric blocking of the active site; the possibility of enzymeleakage during operation; risk of support deteriorationunder operational conditions, due to mechanical or chemicalstress; and a (still) relative empirical methodology, whichmay hamper scale up. Economical issues are furthermoreto be taken into consideration when commercial processesare envisaged, although immobilization can prove critical foreconomic viability if costly enzymes are used. Still, the costof the support, immobilization procedure and processing thebiocatalyst once exhausted, up- and downstream processingof the bioconversion systems, and sanitation requirementshave to be taken into consideration. In the overall, theenhanced stability allowing for consecutive reuse leads tohigh specific productivity (mass−1

product mass−1biocatalyst), which

influences biocatalyst-related production costs [1, 142].A typical example is the output of immobilized glucoseisomerase, allowing for 12,000–15,000 kg of dry-producthigh-fructose corn syrup (containing 42% fructose) perkilogram of biocatalyst, throughout the operational lifetimeof the biocatalyst [144]. Increased thermal stability, allowingfor routine reactor operation above 60◦C minimizes the risksof microbial growth, hence leading to lower risks of microbialgrowth and to less demanding sanitation requirements, sincecleaning needs of the reactor are less frequent [1, 144].A rule of thumb suggesting that the enzyme costs shouldbe a few percent of the total production costs has beenestablished [142]. The half-life of the bioreactor is also acritical issue when evaluating the economical feasibility of abioconversion process, longer half-lives favoring process eco-nomics. Examples of commercial bioreactors depict half-livesof several months to years, and the same packing can workthroughout some months to years. Among this group, areimmobilized enzyme reactors packed with glucose isomerasefor the production of high-fructose corn syrup; lactase forlactose hydrolysis, for the production of whey hydrolysatesand for the production of tagatose; aminoacylase for theproduction of amino acids; isomaltulose synthase for theproduction of isomaltulose; invertase for the productionof inverted sugar syrup; lipases for the interesterificationof edible oils, ultimately targeted at the production oftrans-free fat, of cocoa butter equivalents, and of modifiedtriacylglycerols; and β-fructofuranosidase for the productionof fructooligosaccharides [144–146]. On the other hand,despite the technical advantages of immobilization, the large-scale liquefaction of starch to dextrins by α-amylases isperformed by free enzymes, given the low cost of the enzyme[18].

Immobilization can be performed by several methods,namely, entrapment/microencapsulation, binding to a solidcarrier, and cross-linking of enzyme aggregates, resulting incarrier-free macromolecules [142]. The latter presents analternative to carrier-bound enzymes, since these introduce

Enzyme Research 9

a large portion of noncatalytic material. This can accountto about 90% to more than 99% of the total mass ofthe biocatalysts, resulting in low space-time yields andproductivities, and often leads to the loss of more than50% native activity, which is particularly noticeable at highenzyme loadings [142]. A broad, generalized overview of theadvantages and drawbacks of the different immobilizationapproaches is given in Table 4. A typical example of thepatterns suggested by data in Table 4 was observed by Abdel-Naby when evaluating the immobilization of α-amylasethrough different methods [147]. Details on the differentmethods, as well as some illustrative examples of theirapplications, are given hereafter.

Entrapment/(micro)encapsulation, where the enzyme iscontained within a given structure. This can be: a polymernetwork of an organic polymer or a sol-gel; a membranedevice such as a hollow fiber or a microcapsule; or a (reverse)micelle. Apart from the hollow fiber, the whole processof immobilization is performed in-situ. The polymericnetwork is formed in the presence of the enzyme, leadingto supports that are often referred to as beads or capsules.Still, the latter term could preferably be used when the coreand the boundary layer(s) are made of different materials,namely, alginate and poly-l-lysine. Although direct contactwith an adverse environment is prevented, mass transferlimitations may be relevant, enzyme loading is relativelylow, and leakage, particularly of smaller enzymes fromhydrogels (namely, alginate, gelatin), may occur. This maybe minimized by previously cross-linking the enzyme withmultifunctional agent (namely, glutaraldehyde) [148, 149]or by promoting cross-linkage of the matrix after theentrapment [150]. The use of LentiKats, a polyvinyl-alcohol-based support in lens-shaped form, has been used for severalapplications in carbohydrate processing. Among these arethe synthesis of oligosaccharides with dextransucrase [149],maltodextrin hydrolysis with glucoamylase [151], lactosehydrolysis with lactase [152], and production of invert sugarsyrup with invertase [153]. In these processes the biocatalystcould be effectively reused or operated in a continuousmanner. Methodologies for large scale production of thesesupports have been implemented [154, 155]. Flavourzyme,(a fungal protease/peptidase complex) entrapped in calciumalginate [156], k-carragenan, gellan, and higher melting-fatfraction of milk fat [157], was effectively used in cheeseripening, in order to speed up the process, while avoidingthe problems associated with the use of free enzyme. Theseinclude deficient enzyme distribution, reduced yield andpoor-quality cheese, partly ascribed to excessive proteolysisand whey contamination. The enzyme complex is released ina controlled manner due to pressure applied during cheesecurd [156].

Calcium alginate beads were also used to immobilizeglucose isomerase [158] and α-amylase for starch hydrolysisto whey [159]. In the latter work, the authors observed thatincreasing the concentration of CaCl2 and of sodium alginateto 4% and 3%, respectively, enzyme leakage was minimized(a common drawback of hydrogels) while allowing for highactivity and stability. This effect was also observed in aprevious work where alginate-entrapped inulinase was used

for sucrose hydrolysis [160]. The stability of an amylaseimmobilized biocatalyst was further enhanced with theaddition of 1% silica gel to the alginate prior to gelation, asreflected by the use of the biocatalyst in 20 cycles of opera-tion, while retaining more than 90% of the initial efficiency[159]. Several enzymes, namely, chymosin, cyprosin, lactase,Neutrase, trypsin, have also been immobilized in liposomes,[161]. In a particularly favored technique immobilization ofenzymes in liposomes, known as dehydration-rehydrationvesicles (DRVs), small (diameters usually below 50 nm)unilamellar vesicles (SUVs) is prepared in distilled waterand mixed with an aqueous solution of the enzyme tobe encapsulated. The resulting vesicle suspension is thendehydrated under freeze drying or equivalent method. Uponrehydration, the resulting DRVs are multilamellar and larger(from 200 nm to a little above 1000 nm) than the originalSUVs, and can capture solute molecules [161, 162]. Recentwork in this particular application has used lactase asenzyme model and has focused on the optimization andcharacterization of the liposome-based immobilized system[163, 164]. If liposome-based biocatalysts are used in aprocess under continuous operation, biocatalyst separationhas to be integrated (namely, using an ultra-filtrationmembrane). In a different concept, based in batch mode,liposome-encapsulated lactase was incorporated in milk.After ingestion, the vesicles are disrupted in the stomachby the presence of bile salts, allowing in-situ degradation oflactose [165]. Cocktails of enzymes, namely, Flavourzyme,bacterial proteases and Palatase M (a commercial lipasepreparation), were immobilized in liposomes and success-fully used to speed up cheddar cheese ripening [166].Encapsulation in lipid vesicles has been proved a mildmethod, providing high protection against proteolysis. Thereis however some lack of consensus on the feasibility of itsapplication on large scale, as well as on the effectivenessof the methodology for controlled release of enzymes [156,157, 161, 163, 167]. Containment within an ultra-filtration(UF) membrane allows the enzyme to perform in a fullyfluid environment; hence, with little loss (if any) of catalyticactivity. However, the membrane still presents a boundaryfor overall mass transfer of substrate/products and enzymemolecules are prone to interact with the membrane material.This feature is enhanced along with the hydrophobicityof the membrane, hence immobilization in membranedevices may have some adsorptive nature, a feature thatwill be addressed in (ii). Besides, regular replacement ofthe membrane may be required. Enzyme containment by amembrane has been used for the continuous production ofgalactooligosaccharides from lactose. The reaction, with upto 80% lactose conversion out of a substrate concentrationof 250 gL−1, was carried out in a perfectly mixed reactor andenzyme was recovered in a 10 kDa nominal molecular weightcutoff. The resulting product presented some similarities tothe commercially available Vivinal prebiotic [168]. Withinthe same methodology, a hollow-fiber module was used tocontain lactase, in order to carry out lactose hydrolysis incontinuous operation. A conversion rate close to 95% in skimmilk was observed for an initial substrate concentration closeto 40 gL−1 [169].

10 Enzyme Research

Table 4: A generalized characterization of immobilization methods.

ParameterImmobilization method

Carrier bindingCLEAs, CLECs Entrapment

Covalent Ionic Adsorption

Activity High High Low Intermediate/High High

Range of application Low Intermediate Intermediate Low Intermediate/High

Immobilization efficiency Low Intermediate High Intermediate Intermediate

Cost Low Low High Intermediate Low

Preparation Easy Easy Difficult Intermediate Intermediate/Difficult

Substrate specificity Cannot be changed Cannot be changed Can be changed Cannot be changed Can be changed

Regeneration Possible Possible Impossible Impossible Impossible

Binding to a solid carrier, where enzyme-support inter-action can be of covalent, ionic, or physical nature. The lattercomprehends hydrophobic and van der Waals interactions.These are of weak nature and easily allow for enzyme leakagefrom the support, namely, after environmental shifts in pH,ionic strength, temperature or even as a result of flow rateor abrasion. On the other hand, desorption can be turnedinto an advantage if performed under a controlled manner,since it enables the expedite removal of spent enzyme andits replacement with fresh enzyme [170]. A recent paperby Gopinath andSugunan illustrates the increased trend forleakage when adsorption is compared with covalent binding,using α-amylase as model enzyme [171]. Curiously, thefirst reported application of enzyme immobilization was ofinvertase onto activated charcoal [172]. Recently invertasewas immobilized in different types of sawdust, aiming at itsapplication for sucrose hydrolysis. When wood shavings wereused as support, the immobilized invertase retained 90%of the original activity after 20 cycles of 15 minutes, eachunder consecutive batch operation; and it retained 65% ofthe original activity after 10 hours of continuous operationalregime in a column reactor [173]. Anther example is theimmobilization of pectinase in egg shell for the preparationof low-methoxyl pectin. The immobilized biocatalyst couldbe reused for 32 times at 30◦C, and it was used in afluidized-bed reactor, operated at an optimum flow rate of5 mL h−1 and 35◦C [174]. Other examples are the surfaceimmobilizations of α-amylase on alumina [175] and inzirconia [176]. Covalent binding is the strongest form ofenzyme linking to a solid support. It involves chemicallyreactive sites of the protein such as amino groups, carboxylgroups, and phenol residues of tyrosine; sulfhydryl groups;or the imidazole group of histidine. The binding can becarried out by several methods; among them are amidebond formation, alkylation and arylation, or UGI reaction.However, this often brings along loss of activity during theprocess of immobilization, due to support binding to criticalresidues for enzyme activity, and steric hindrance, amongothers. Examples include the immobilization of α-amylase[177] and of levansucrase [178] on glutaraldehyde-treatedchitosan beads, through the glutaraldehyde reaction betweenthe free amino groups of chitosan and the enzyme molecule;the immobilization of pectinase onto Amberlite IRA900 Clthrough glutaraldehyde cross-linking [179]; glucoamylase

onto dried oxidized bagasse [180], onto polyglutaraldehyde-activated gelatin [181], or onto macroporous copolymerof ethylene glycol dimethacrylate and glycidyl methacrylatethrough the carbohydrate moiety of the enzyme [182]; glu-coamylase or invertase immobilized onto montmorilloniteK-10 activated with aminopropyltriethoxysilane and glu-taraldehyde [183, 184]; and invertase immobilized on nylon-6 microbeads, previously activated with glutaraldehyde andusing PEI as spacer [185, 186]; on polyurethane treatedwith hydrochloric acid, polyethylenimine and glutaralde-hyde [187]; on poly(styrene-2-hydroxyethyl methacrylate)microbeads activated with epichlorohydrin [188]; or onpoly(hydroxyethyl methacrylate)/glycidyl methacrylate films[189]. Within this methodology for immobilization, high-light should be given to the introduction of commer-cial supports (namely, Eupergit, Sepabeads) with a highdensity of epoxide functional groups aimed at multipointattachment, typically with the ε-amino group of lysine, toconfer high rigidity to the enzyme molecule, hence enhanc-ing stabilization [190, 191]. This methodology has beenused for lactase immobilization in magnetic poly(GMA-MMA), formed from monomers of glycidylmethacrylateand ethylmethacrylate, and cross-linked with ethyleneglycoldimethacrylate [192]; for the immobilization of cyclodextringlycosyltransferases to glyoxylagarose supports for the pro-duction of cyclodextrins [193]; or for the immobilization ofdextransucrase on Eupergit C [194]. Ionic binding to a car-rier involves interaction of negatively or positively chargedgroups of the carrier with charged amino-acid residueson the enzyme molecules [195]. Ionic interaction may befavored if enzyme leakage is not an issue, since it allowsfor support regeneration, unlike immobilization by covalentbinding. Ion-exchanger resins are typical supports for ionicbinding; among them are derivatives of cross-linked polysac-charides, namely, carboxymethyl- (CM-) cellulose, CM-Sepharose, diethylaminoethyl- (DEAE-) cellulose, DEAE-Sephadex, quaternary aminoethyl anion exchange- (QAE-) cellulose, QAE-dextran, QAE-Sephadex; derivatives ofsynthetic polymers, namely, Amberlite, Diaion, Dowex,Duolite; and resins coated with ionic polymers, namely,polyethylenimine (PEI) [196]. Recent examples include theimmobilization of invertase in Dowex [197], in Duolite[198], in poly(glycidyl methacrylate-co-methyl methacry-late beads grafted with PEI [199], and in epoxy(amino)

Enzyme Research 11

Sepabeads [200]; lactase immobilization in PEI-graftedSepabeads [201]; fructosyltransferase in DEAE-cellulose forthe production of fructosyl disaccharides [202]; glucoseisomerase in DEAE-cellulose [203] or in Indion 48-R [204];glucoamylase onto SBA-15 silica [205] and in epoxy(amino)Sepabeads [200]. Ionic binding to Sepabeads-like supportshas acknowledged multipoint attachment nature. Enzymemolecules can be modified chemically or genetically mod-ified to enhance immobilization efficiency, an approach fol-lowed by Kweon and coworkers, who obtained a cyclodextringlycosyltransferase fused with 10 lysine residues to improveionic binding to SP-Sepharose [206].

Carrier-free macroparticles, where a bifunctional reagent(namely, glutaraldehyde), is used to cross-link enzyme aggre-gates (CLEAs) or crystals (CLECs), leading to a biocatalystdisplaying highly concentrated enzyme activity, high stabilityand low production costs [142, 207]. The use of CLEAsis favored given the lower complexity of the process. Thisapproach is recent, as compared with entrapment andbinding to a solid carrier, and there are still relatively fewexamples of its application to enzymes used in the area offood processing. Among those are following.

(1) First is the immobilization of Pectinex Ultra SP-L, a commercial enzyme preparation containingpectinase, xylanase, and cellulose activities [208]. TheCLEA biocatalyst displayed a slight (30%) in theVmax, maximal reaction rate/KM ratio, but a signifi-cant enhancement in thermal stability (a roughly 10-fold increase in half-life), when the pectinase activityof the immobilized biocatalyst was compared withthe free form.

(2) Second is the immobilization of lactase for thehydrolysis of lactose, where, under similar opera-tional conditions as for the free enzyme, the CLEAyielded 78% monosaccharides in 12 h as comparedto 3.9% of the free form [209].

(3) Third, CLEAs of glucoamylase, formed by eitherglutaraldehyde or diimidates, namely, dimethylmal-onimidate, dimethylsuccinimidate, and dimethylglu-tarimidate, led to biocatalysts with improved thermalstability as compared to the free form (over 2-foldincrease in half-lives) [210].

(4) Fourth, CLEAs of wild type and two mutant levan-sucrases were assayed for oligosaccharides/levan andfor fructosyl-xyloside synthesis. Although the specificactivity of the three free enzymes was 1.25- to 3-fold higher than the corresponding CLEAs, thesedisplayed a 40- to 200-fold higher specific activitythan the equivalent Eupergit-C-immobilized enzymepreparations. Furthermore, all CLEA preparationsdisplayed enhanced thermal stability when comparedwith the corresponding free enzymes [211].

(5) Fifth are CLECs of glucose isomerase, aimed at theconversion of glucose into fructose for the pro-duction of high fructose corn syrup. When placedin a packed-bed, the resulting enzyme preparationallowed for flow rates that matched or even exceeded

those processed by commercially available enzymepreparations (either free, carrier free, or carrier-bound), while achieving the same 45% yield infructose, under similar operational condition [212].

(6) Sixth, CLECs of glucose isomerase packed in acolumn were also used for the concentration/puri-fication of xylitol from dilute or impure solutions.The approach was based on the high specificityof the enzyme crystals towards xylitol, allowing itsseparation from other sugars, including the nat-ural substrates, xylose and glucose. Recovery ofthe adsorbed xylitol was achieved by elution withCaCl2 solutions, with Ca2+ being acknowledged toinactivate glucose isomerase [213].

Each method for enzyme immobilization has a uniquenature. Therefore, despite the potential of immobilization toimprove enzyme performance by enhancing activity, stabil-ity, or specificity, no specific approach tackles simultaneouslythese different features. A careful evaluation and charac-terization of the methodology addressed is thus required,which can be significantly fastened by high-throughputapproaches [214]. Again, the feasibility of its application toreactor configuration and mode of operation has also to beconsidered in the selection process of the most adequateimmobilized biocatalyst for a given bioconversion.

4.1. Typical Bioreactors. The most common form of enzy-matic reactors for continuous operation is the packed-bedsetup, basically a cylindrical column holding a fixed bedof catalyst particles (Figure 1). These should not have sizesbelow 0.05 mm, in order to keep the pressure drop withinreasonable limits. Commercially available carriers such asEupergit C have particle sizes of roughly 0.1 mm [215].Commonly operated in down-flow mode, the range of flowrates used must be such as to provide a compromise betweenreasonable pressure drop, minimal diffusion layer and highconversion yield. Minimization of external mass-transferresistances with enhanced flow rates can be considered,leading to the fluidized-bed reactor. This is basically avariation of the packed-bed reactor, but operated in up-flow mode, where the biocatalyst particles are not in closecontact which each other; hence, pressure drop is low, andaccordingly are pumping costs. The residence time allowedby the flow rates required for fluidization may howeverresult in low conversion yields. This can be overcome byoperating a battery of reactor or by operation in recyclemode [216]. Bioconversions with free enzymes are carriedout in stirred tanks. When on their own, they are restrictedto batch mode, but when coupled to a membrane setupwith suitable cutoff, they can be integrated in a continuousprocess, since the enzymes are rejected by the membrane,which acts as an immobilization device, whereas the product(and unconverted substrate) freely permeates. Shear stressinduced by stirring creates a hazardous environment forimmobilized biocatalysts, particularly when hydrogels areconsidered, since they are prone to abrasion. In order toovercome this, a basket reactor was developed, but is seldom

12 Enzyme Research

Fluid in

Fluid out

(a)Packed-bedreactor

Fluid out

Fluid in

(b)Fluidized-bed reactor

Fluid in

Fluid out (product rich)

Ultrafiltration unitBiocatalyst

recycle

(c) Perfectly mixed reactor with recycle

Fluid in Fluid out

Free enzymeImmobilized enzyme

(d) Stirred basket reac-tor

Free enzymeImmobilized enzyme

(e) Stirred batch reactor

Figure 1: Examples of bioreactor configurations commonly used in bioconversion processed involving free or immobilized enzymes.Reactors (a) to (d) are depicted under continuous mode of operation, whereas reactor (e) is depicted.

used, possibly due to mass transfer resistances associated[18].

5. Conclusions and Future Perspectives

The integration of enzymes in food and feed processes is awell-established approach, but evidence clearly shows thatdedicated research efforts are consistently being made asto make this application of biological agents more effectiveand/or diversified. These endeavors have been anchoringin innovative approaches for the design of new/improvedbiocatalysts, more stable (to temperature and pH), lessdependent on metal ions and less susceptible to inhibitoryagents and to aggressive environmental conditions, whilemaintaining the targeted activity or evolving novel activities.This is of particular relevance for application in the foodand feed sector, for it allows enhanced performance underoperational conditions that minimize the risk of microbialcontamination. It also favors process integration, by allowingthe concerted use of enzymes that naturally have diverserequirements for effective application. Such progresses havebeen made through the ever-continuing developments inmolecular biology, the accumulated evolutionary enzymeengineering expertise, the (bio)computational tools, and theimplementation of high-throughput methodologies, withhigh level of parallelization, enabling the efficient and timelyscreening/characterization of the biocatalysts. Alongsidewith these strategies, the immobilization of enzymes has alsobeen a key supporting tool for rendering these proteins fitfor industrial application, while simultaneously enabling theimprovement of their catalytic features. Again, and despitethe developments made in this particular field, there is stillthe lack of a set of unanimously applicable rules for theselection of carrier and method of enzyme immobilization,which furthermore encompass both technical and economicrequirements. The latter can be particularly restrictive in

the food and feed sector, since most products are of relativelylow added value. Therefore, there is no universal supportand method for enzyme immobilization aimed at applicationin food and feed (let alone the overall range of possiblefields of use), and the immobilized biocatalyst fit for a givenprocess and product may be totally unsuitable for another.Given the diversity of enzyme nature and applications thispattern is unlikely to be reversed. Hence, it can be foreseenthat efforts will be towards the development of immobilizedbiocatalyst with suitable chemical, physical, and geometriccharacteristics, which can be produced under mild condi-tion, that can be used in different reactor configurations andthat comply with the economic requirements for large-scaleapplication. All these strategies either isolated or preferablysuitably integrated have been put into practice in food andfeed, to improve existing processes or to implement newones, with the latter often combined with the output of newgoods, resulting from novel enzymatic activities. Given therecent developments in this field, this trend is foreseen to befurther implemented.

Acknowledgment

Pedro Fernandes acknowledges Fundacao para a Ciencia e aTecnologia (Portugal) for financial support under programCiencia 2007.

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SAGE-Hindawi Access to ResearchEnzyme ResearchVolume 2010, Article ID 480923, 8 pagesdoi:10.4061/2010/480923

Review Article

Some Nutritional, Technological and Environmental Advances inthe Use of Enzymes in Meat Products

Anne y Castro Marques, Mario Roberto Marostica Jr., and Glaucia Maria Pastore

School of Food Engineering, University of Campinas, Monteiro Lobato st., 80, 13083-862 Campinas, SP, Brazil

Correspondence should be addressed to Mario Roberto Marostica Jr., mmarostica@gmail.com

Received 11 June 2010; Revised 31 August 2010; Accepted 14 September 2010

Academic Editor: Cristina M. Rosell

Copyright © 2010 Anne y Castro Marques et al. This is an open access article distributed under the Creative Commons AttributionLicense, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properlycited.

The growing consumer demand for healthier products has stimulated the development of nutritionally enhanced meat products.However, this can result in undesirable sensory consequences to the product, such as texture alterations in low-salt and low-phosphate meat foods. Additionally, in the meat industry, economical aspects have stimulated researchers to use all the animal partsto maximize yields of marketable products. This paper aimed to show some advances in the use of enzymes in meat processing,particularly the application of the proteolytic enzymes transglutaminase and phytases, associated with nutritional, technological,and environmental improvements.

1. Introduction

Meat products consumption (including beef, pork, mutton,goat and poultry) has increased gradually, particularlyin developing countries. Studies estimate that the worldconsumption of meat products will reach 40 kg per capita in2020 [1]. The processes involved in the conversion of muscleto meat are complex. The chemical and physical propertiesof muscle tissue and the associated connective tissue aredeterminant on meat quality [2].

The growing consumer demand for healthier productshas stimulated the development of nutritionally enhancedmeat foods. In order to achieve these nutritionally enhancedmeat foods, changes such as the use of improved raw mate-rials, reformulation of products, and technological processesare necessary [3]. These improvements, however, can bringundesirable consequences to the product, such as texturealterations in low-salt and low-phosphate meat foods [4, 5].Additionally, high costs have stimulated researchers to useall animal parts, including muscles of poorer technologicalquality, to maximize the yield of marketable products. Thishas required the development of methods to restructurelow-valued cuts and trimmings, improving appearance andtexture and increasing market value [6, 7].

Faced with new market trends, is it possible to pro-duce meat products that meet all the market requirements(healthy, with good sensory properties, low cost, and envi-ronmental friendly)? The aim of this paper is to show someadvances related to this topic, focusing on the application ofproteolytic enzymes, transglutaminase and phytases in meatproducts.

2. The Use of Proteolytic Enzymes inMeat Products

Of all the attributes of meat quality, consumers rate ten-derness as the most important. Tenderness is a charac-teristic resulting from the interaction of actomyosin effectof myofibrillar proteins, the bulk density effect of fat,and the background effect of connective tissue. There areseveral ways to tenderize meat, chemically or physically,which mainly reduce the amounts of detectable connectivetissue without causing extensive degradation of myofibrillarproteins. Treatment by proteolytic enzymes is one of themost popular methods of meat tenderization [8, 9].

Proteolytic enzymes are a multifunctional class of enz-ymes, with physiological functions that range from gener-alized protein digestion to more specific regulated processes

2 Enzyme Research

such as the activation of zymogens, blood coagulation, com-plement activation, inflammation process, and liberation ofphysiological peptides from the precursor proteins. They arefrequently used in food processing [10].

The first variation of meat tenderness is due to thecomplex endogenous calpain-calpastatin, which acts in mus-cle tissue after slaughter. Calpains are calcium-dependentproteases that degrade myofibrillar proteins (tropomyosin,troponin T, troponin I, C-protein, connectin, and desmin).Calpastastin, in turn, inactivates calpains, decreases themyofibrillar degradation, and thus reduces the tenderness.Calpastatin effect is finished after calpastatin is inactivatedby cooking. The concentration of the enzymes varies amongbreeds of species, determining the higher or lower meattenderness, due to increased or reduced proteolysis ofmyofibrillar proteins [11, 12].

Several examples of proteases application in meat prod-ucts can be found in the literature. Benito et al. [13] showedthat the fungal protease EPg222 hydrolyzed myofibrillarproteins of whole pieces of meat with 5% NaCl, favoringtenderization and improving texture of the product. Accord-ing to the authors, salt and curing agents at the level foundin dry cured meat products are powerful inhibitors of theformer endogenous enzymes. The effect of this proteasemay be of great interest to counterbalance the increase ofhardness reported in these products as a consequence ofprotein denaturation. Nalinanon et al. [14] used pepsin toobtain fish gelatin, from bigeye snapper skin, as an alternativefor porcine and bovine gelatin. Thiansilakul et al. [15]produced a protein hydrolyzate (derived from round scad)with Flavourzyme protease addition that could be used asan emulsifier and as a foaming agent with antioxidativeactivities in food systems.

The quest for valuable proteases with distinct specificityfor industrial applications is always a continuous challenge.Proteolytic enzymes from plant sources have received specialattention for being active over a wide range of temperaturesand pH [16, 17].

The name ficin (EC 3.4.22.3) refers to the endoprote-olytic enzymes from trees of the genus Ficus. These enzymeshave different properties. The most extensively studied ficinsare the cysteine proteases found in the latex of Ficus glabrataand Ficus carica. Proteases from other species are less known.In 2008, a new protease from Ficus racemosa was identified.The protein has a molecular weight of 44.500 ± 500 Da, pHoptima between pH 4.5 and 6.5 and maximum activity at60 ± 0.5◦C. These unique properties indicate this proteaseto be distinct from other known ficins. Applications of thisenzyme include its use as meat tenderizers, removal of chillhaze in beer, improvement in the processing of cereals, andplant and milk clotting enzymes for novel dairy products[16, 18].

Papain (EC 3.4.22.4) is a nonspecific thiol protease andthe major protein constituent of the latex in the tropicalplant Carica papaya. The enzyme has high thermal andpressure stability, requiring intense process conditions foradequate inactivation (to achieve 95% inactivation of papainat 900 MPa and 80◦C, 22 minutes of processing is required).Due to its proteolytic properties, it is widely used in the

food industry to tenderize meat and as an additive in flourand in beer manufacturing [18, 19]. However, papain hasa tendency to overtenderize the meat surface, making it“mushy”, which has limited its use as a commercial meattenderizer [10]. Herranz et al. [20] used papain (300 units/kgof papain) to increase the amount of free amino acids in dryfermented sausages. These precursors of volatile compounds,responsible for the ripened flavor, were tested in presence ofLactococcus lactis subsp. cremoris NCDO 763, its intracellularcell free extract (ICFE), and α-ketoglutarate. The results,however, did not show any important activity related toamino acid breakdown, and the sensory analysis showedthat neither the addition of the extract nor its use togetherwith papain or a-ketoglutarate lead to an improvement inthe sensory quality of the experimental sausages. Recently,Shimizu et al. [21] evaluated the antithrombotic activityof papain-hydrolyzate from defatted pork meat (crude andpeptides purified by cation exchange chromatography) “invivo”. The initial peptide fraction with an average molecularweight of 2500 showed antithrombotic activity after oraladministration to mice at 210 mg/kg. The fraction with anaverage molecular weight of 2517, further purified by cationexchange chromatography, showed antithrombotic activityafter oral administration at 70 mg/kg. Antithrombotic activ-ity of the last peptide fraction was equivalent to that ofaspirin at 50 mg/kg body weight.

Bromelain (or bromelin, EC 3.4.4.24) is a group of pro-teolytic enzymes present in large quantities in fruit, leaves,and stems of the Bromeliacea family, of which pineapple(Ananas comosus) is the most commonly known [22, 23].This enzyme, like other proteases, degrades myofibrillarproteins and collagen, often resulting in overtenderizationof meat [24]. Ionescu et al. [25] investigated bromelainuse in adult beef, with the best results at 10 mg/100 gmeat, with tenderization time 24 hours at 4◦C, followed bythermal treatment by increasing 1◦C/min until 70◦C (whenenzyme inactivation occurs). These conditions improvedbeef tenderness.

The ideal meat tenderizer would be a proteolytic enzymewith specificity for collagen and elastin in connective tissue,at the relatively low pH of meat, which would act eitherat the low temperature at which meat is stored or at thehigh temperature achieved during cooking [26]. Qihe et al.[8] investigated elastase from Bacillus sp. EL31410, appliedto beef tenderization, in comparison with other nonspecificproteases, such as papain, and evaluated the feasibility ofusing it for this purpose. The samples were treated for 4hours in different enzyme solutions and then were stored at4◦C for 24, 48, and 72 hours. A marked decrease in hardnesswas observed in the meat with papain and elastase and highersensory scores for tenderness were obtained from the meattreated with enzymes. However, the scores given for juicinessand taste were lower than those of the control. Rapid increaseof fragmentation of myofibrils from the enzyme-treated meatwas observed in the first 24 hours of storage, especiallyfor papain-treated meat. Meantime, elastin of myofibrillarstructure was selectively degraded by elastase when stored at4◦C for 48 hours as shown by electron microscopy. These

Enzyme Research 3

Table 1: Use of proteolytic enzymes for bioactive peptides production in meat foods.

Product Conditions Results Reference

ACE of proteinhydrolysatesfrom sardine(Sardinellaaurita)

Sardine: heads and viscera.Enzymes: alcalase, chymotrypsin,

Bacillus licheniformis NH1protease, Aspergillus clavatus ES1

protease and sardine visceraprotease.

Sardinelle proteins were digested by proteases and the ACEinhibitory activity was markedly increased. The degrees of

hydrolysis and the inhibitory activities of ACE increased withincreasing proteolysis time. The sardinelle hydrolysis with the

crude enzyme extract from sardine viscera resulted in theproduction of the hydrolysate with the highest ACE inhibitory

activity.

[32]

ACE of proteinhydrolysatesfrom shark meathydrolysate

Enzyme: Bacillus sp. SM98011protease (diluted to 4000 U/mL

with distilled water).Enzyme/substrate concentration:

1 : 5 w/v.

The hydrolysate of shark meat was rich with ACE inhibitorypeptides, and 3 novel peptides with high ACE inhibitory activity

were identified (Cys-Phe, Glu-Tyr, and Phe-Glu).

[33]

ACE of proteinhydrolysatesfrom muscle ofcuttlefish (Sepiaofficinalis)

Enzymes: trypsin, chymotrypsin,sardinelle protease, cuttlefishprotease and smooth houndsprotease. Enzyme/substrate

concentration: 3 U/mg.

The most active hydrolysate was obtained with the crude proteaseextract from the hepatopancreas of cuttlefish (64.47 ± 1.0% at

2 mg of dry weight/mL) with a degree of hydrolysis of 8%. Threenovel peptides with high ACE-inhibitory activity were formed:

Val-Tyr-Ala-Pro, Val-Ile-Ile-Phe, and Met-Ala-Trp.

[34]

findings suggest that Bacillus elastase could be a promisingsubstitute for papain as a favorable meat tenderizer.

Sullivan et al. [17] studied the tenderization extent(Warner-Bratzler shear and sensory evaluation) and modeof action (myofibrillar or collagen degradation) of sevenenzyme randomized treatments (papain, ficin, bromelain,homogenized fresh ginger, Bacillus subtilis protease, andtwo Aspergillus oryzae proteases) in Triceps brachii andSupraspinatus. Except for ginger treatment, all steaks treatedwith enzymes showed improvement in both sensory andinstrumental tenderness analysis. If this enzyme could bepurified further, applications in meat would be promising.Among the results presented, papain was the enzyme thatcaused the greatest tenderness in meat, but juiciness andtextural changes were negatively affected. The authors alsoconcluded that all enzyme treatments resulted in increasedtenderness with no difference between high- and low-connective tissue muscles.

Kiwifruit has also been studied as a source of actinidin,an important proteolytic enzyme. Han et al. [10] investigatedthe ability of prerigor infusion of kiwifruit juice (10% bodyweight) to improve the tenderness of lamb. The enhancedproteolytic activity in lamb carcass was associated withsignificant degradation of the myofibrillar proteins, resultingin new peptides and activation of m-calpain during post-mortem ageing. Thus, kiwifruit juice is a powerful and easilyprepared meat tenderizer, which could contribute efficientlyand effectively to the meat tenderization process. However,studies have show kiwifruit to cause allergic reactions, andactinidin to be one of most important allergens, both inchildren and adults [27–29]. Thus, caution should be takenwhen considering tenderizing meat using kiwifruit juice.

Currently, besides the extensive use in meat processes,proteases are being investigated with the aim of transformingthe byproducts of these processes. For example, keratinases,serine proteases which are capable of degrading hard and

insoluble keratin proteins, can be applied in the conversionof large amounts of chicken feather waste generated frompoultry into highly digestible animal feed [30, 31].

Proteases are also being used for other purposes, suchas the production of bioactive peptides against hypertension[32–34] and reduction of the power of allergenic meat foods[35]. The angiotensin I-converting enzyme (ACE), a trans-membrane dipeptidyl peptidase which degrades bradykinin,shows the potential of cleaving any peptide, includingvasoactive peptides such as angiotensin-I. The nutritionaltherapy approach and the use of nutraceuticals from meatis a good way of continual healthcare for patients withhypertension [36]. These studies are more detailed in Table 1.

3. The Use of Transglutaminase inMeat Products

Transglutaminase (TGase; protein-glutamine γ-glutamyltra-nsferase, EC 2.3.2.13) is an enzyme with the ability toimprove the functional characteristics of protein such astexture, flavor, and shelf life. TGase initially attracted interestbecause of its capacity to reconstitute small pieces of meatinto a steak. It can adhere to the bonding surfaces of foodsuch as meat, fish, eggs, and vegetables as a thin layer, andit exhibits strong adhesion in small amounts. The enzymecatalyses acyl transfer reactions between the γ-carboxyamidegroup of peptide bound glutamine residues and a varietyof primary amines, including the ε-amino group of lysineresidues, resulting in the formation of high molecularweight polymers. In the presence of primary amines, TGasecan cross-link the amines to the glutamines of a protein(acyl-transfer reaction). In the absence of lysine residuesor other primary amines, water will react as a nucleophile,resulting in deamidation of glutamines. All three of theseTGase reactions can modify the functional properties offood proteins [4, 37, 38].

4 Enzyme Research

Table 2: Studies using microbial tranglutaminase (MTGase) in meat food.

Product Conditions Results Reference

Chicken andbeef sausages

Proportion of MTGase to MHC∗ =1 : 500. Heat treatment: 40◦C/30 minutes

using a thermo-minder; 80◦C/30minutes, using a water bath shaker.

∗MHC: myosin heavy chain.

MTGase affected the breaking strength score in both meattypes, especially for beef cooked at 80◦C. The functional

properties of MTGase make it a good protein-binding agent,positively helping the functionality of proteins to improvethe texture and gelation of sausages. Some variation in gel

improvement level between chicken and beef sausages wereobserved, in response to MTGase, as well as to the original

glutamyl and lysine contents.

[2]

Dry-cured ham

Treatments: no treatment (control);immersion in a saline (NaCl with

200 ppm of KNO3 and 100 ppm ofNaNO2) aqueous solution (3%, w/v) for10 minutes at 4◦C; and even distributionof a mixture of salts (NaCl with 200 ppmof KNO3 and 100 ppm of NaNO2) on the

surfaces for 1 minute and after 10minutes of setting time. Bindingtemperature: 0◦C, 7◦C and 24◦C.

MTGase: powder and liquid (MTGase at0.1% in solution of NaCl 3%).

MTGase provided enough stable cross-links in the course ofthe salting and drying processes. The highest binding forceand rate were obtained by treating the meat surface with amixture of salts (NaCl including KNO3 and NaNO2) then

adding MTGase.

[42]

Fish (Trachurusspp., horsemackerel)

High pressure treatment (300 MPa, 25◦C,15 minutes), combined with a prior or asubsequent setting step (25◦C, 2 hours),1.5% chitosan and/or 0.02% MTGase.

MTGase led to an increase in hardness and a considerabledecrease in elasticity and breaking deformation. MTGase

activity was greater when setting was applied beforepressurization than after; moreover, there was no synergismderived from the addition of chitosan and MTGase together.

[45]

Ground beef

Preparation A: MTG (1 g/100 g) andmaltodextrin (99 g/100 g); Preparation B:

MTG (0.5 g/100 g), SC (60 g/100 g) andmaltodextrin (39.5 g/100 g). MTG:

product weight/meat weight.

MTG with sodium caseinate (SC) led to a slight increase inpeak temperature (Tmax) values of myosin. MTGase

treatment caused a slight decrease in Tmax values of myosin[51]

Restructuredcooked porkshoulder

Phosphate-free product. Salt levels: 2%and 1%; MTG: 0%, 0.075% and 0.15%.

Processing conditions: 72◦C/65′ minutesand 78◦C/65′ minutes.

MTG affected consistency and overall acceptability of theproduct. MTG had no effect on firmness, juiciness, color,

odor, taste and saltiness. MTG can be used at a level of 0.15%with reduced salt level (1%) and processing at 72◦C/65

minutes to produce phosphate-free restructured cooked porkshoulder with acceptable sensory attributes.

[37]

TGase is widely distributed among mammals, plants,invertebrates, amphibians, fish, birds, and microorganisms,but the extremely high cost of transglutaminase from animalorigin has hampered its wider application and has initiatedefforts to find an enzyme of microbial origin. The industrialproduction of transglutaminase is done mainly from avariant of Streptoverticillium mobaraense (namely MTGase).The pH optimum of MTGase is around 5 to 8. However,even at pH 4 or 9, MTGase still expresses some enzymaticactivity. The optimum temperature for enzymatic activityis 50◦C, and MTGase shows activity even during chillingtemperatures (under 4◦C); this property is used to bind rawpieces of meat under refrigeration to produce restructuredmeat products [4, 5, 38–40].

TGase has been widely applied in meat products such aschicken and beef sausages [2], ham [41, 42], doner kebab[43], frankfurters [44], fish [45], and so forth. Some studieswith TGase in meat food are detailed in Table 2.

An important functional property of transglutaminase isthe ability to induce gelation in meat foods. The gelation

resulted from protein aggregation in food is highly relatedto the enzymes reactions as well as the biological activities ofsome additives [2]. The TGase catalyses the interconnectionsof myofibrils, improves the gel elasticity of meat protein,and forms a protein-rotein network. Gel strength is furtherenhanced by heat treatment subsequent to the action ofTGase [46].

Herrero et al. [47] determined the effect of addingdifferent levels of MTGase to meat systems (meat emulsionat 0.0%, 0.05%, and 0.10%). This addition produced asignificant increase in hardness, springiness, and cohesive-ness. Data revealed secondary structural changes in meatproteins due to MTGase action; significant correlations werefound between these secondary structural changes in meatproteins and the textural properties of meat systems. Fortet al. [48] studied the heat-induced gelling properties, atacid pH, of porcine plasma previously treated with MTGaseunder high pressure (HP), when kept under refrigerationconditions for different times. The results indicated thatalthough the cross-linking activity of MTGase was enhanced

Enzyme Research 5

under pressure, consequently improving the thermal geltexture, the most significant effects, particularly on gelhardness, were obtained by keeping the treated plasmasolutions under refrigeration for at least 2 hours beforegelation. Literature also shows a species-specific variation inthe ability of MTGase to catalyze the cross-linking of muscleproteins. Proteins in chicken, beef, and pork respond differ-ently to MTGase, generating different products (polymers)and, consequently, differ in terms of both rheological andphysiochemical properties [39, 49].

The fact that MTGase reacts differently to myofibrilsof different species may be because of the variation inmuscle physiology and morphogenesis, the identity of freeamino acids, especially those with the ability to react withMTGase, the amount and distance between transferableamino acids, and the amount of MTGase inhibitors. It isnecessary to understand the protein reactions induced byMTGase binding in meat proteins because of the importanteconomic benefits of using it to improve the textural qualityof meat products [46].

TGase application in low-salt and phosphate-free meatproducts has been extensively investigated. Dry-cured meatand restructured meat products are traditionally preparedusing high salt and phosphate contents, which, with the aidof mechanical action, promote the extraction of myofibrillarproteins; upon cooking, these form a stable protein matrixwith a beneficial effect on product characteristics, such ascohesion and cook yield. The exclusion of salt and phosphateled to products with poor physicochemical properties.Addition of transglutaminase has been proposed as a meansof inducing gelation, reducing or eliminating the need to addNaCl and phosphate products. Furthermore, combinationsof TGase with suitable nonmeat ingredients are also neededto overcome the problems in NaCl-free meat products [4–6, 41, 44, 50]. Askin et al. [43] indicated that the MTGasewith sodium caseinate (SC) or nonfat dry milk could be usedto produce salt-free low-fat turkey doner kebab (a MiddleEast product); the results were more significant when theenzyme was used with SC. Trespalacios et al. [3] showed thatthe simultaneous application of MTGase and high pressure(700 and 900 MPa) on chicken batters with the addition ofegg proteins, low salt and no phosphates resulted in increasedcutting force, hardness, and chewiness of gels.

TGase has been tested with other ingredients in meatproducts. Aktas et al. [51], for example, showed that the com-bination of MTGase with SC could form more cross-linkingbonds between meat proteins in ground beef than when usedseparately; therefore, usage of MTGase with SC may be moresuitable in restructuring meat products. Carballo et al. [50]analyzed the effect of microbial transglutaminase/sodiumcaseinate (MTGase/SC-1.5 g/100 g) systems on meat battercharacteristics (water-binding and textural properties of rawand cooked products) in the presence of NaCl (1.5 g/100 g)and sodium tripolyphosphate (0.5 g/100 g) for pork, chicken,and lamb. Products combining salts and MTGase/SC hadhigher hardness and chewiness, and the efficiency of theMTGase/SC system as a texture conditioner of cookedproducts varied with the meat source. They concludedthat transglutaminase with caseinate form a viscous sol

which could act as a glue to bind restructured meat piecestogether.

Colmenero et al. [44] observed that the combination ofTGase with caseinate, KCl, or fibre (caseinate > KCl > fibre)led to harder, springier, and chewier frankfurters with betterwater- and fat-binding properties (emulsion stability andcooking loss) than those made with TGase only. Accordingto the authors, caseinate has proven to be a good substratefor TGase, facilitating cross-linking and promoting theformation of a much more stable gel matrix during heating.Some previous studies reported several problems related tomoisture loss of meat products induced by TGase. Hong etal. [52], however, suggested that the combination of TGasewith sodium alginate can improve water-binding ability andproduce cold-set myofibrillar protein gelation at an evenlower salt level than TGase alone. In the meat processingindustry, cold-set meat binding is a useful technique formaking raw meat products [53].

Because of the many promising applications of MTGasecatalyzed modification of food proteins, attention should befocused on the nutritional value of resultant cross-linkedproteins. It is obvious that modified MTGase and nativeproteins differ only with respect to ε(γ-glutamyl) lysinebonds, and the rest is totally the same. The cross-linkedproteins can be readily absorbed in the body [4]. A studyconducted by our research group showed that MTGase didnot interfere on the protein quality of soy protein isolate ingrowing Wistar rats (unpublished data).

4. Phytase: Environment Approach and Use asFeed Additive in Meat Animal Production

Phytase is not an ingredient largely used in meat productsformulation; however, some environmental approaches andits use as additive in meat animal production should bementioned.

Nowadays, producers and consumers require more thana good sensory and nutritional product: they are alsoconcerned about the impact of the food chain on theenvironment. So, not only should the appearance or shelflife of meat foods be taken into consideration, but alsothe resources used for the production and the conse-quent damages to the environment. Phytase (myo-inositolhexaphosphate phosphohydrolase) has been used in order toreduce costs of meat food production, as well as to reduceenvironmental contamination by excrement generated dur-ing the production of animals, including swines, poultry, andfish. Phytase is the enzyme used to hydrolyse the phytatemolecule and release phosphorus [54–56]. Microbial phytasehas the ability of hydrolyzing dietary phytate, the salt ofphytic acid (myo-inositol hexaphosphate; IP6), to liberate sixphosphorus and inositol in the gastrointestinal tract [57].

Typically, swine and poultry diets contain around10 g kg−1 phytate-bound phosphorus (phytate-P), but it isonly partially used by the animals because they do notgenerate sufficient endogenous phytase activity. The phytasesupplementation can enhance P absorption and reduce Pexcretion, which are both nutritionally and ecologically

6 Enzyme Research

beneficial [54]. Brenes et al. [58] conducted an experimentto study the effect of microbial phytase supplementation(0, 200, 400, and 600 U/kg) in chicks fed different levelsof available phosphorus. The bone status is very critical inpoultry production, because phosphorus deficiency resultsin breakage or bone defect during processing. The treatmentof poultry feed with phytase increased weight gain; feedconsumption; Ca, P, and Zn retention; tibia ash, tibia Ca, P,and Zn contents; tibia weight; plasma Ca, P, Mg, Zn, and totalprotein content; and serum aspartate aminotransferase, ala-nine aminotransferase, and lactate dehydrogenase activities.Phytase supplementation reduced linearly serum alkalinephosphatase activity. In conclusion, the results indicate thatthe addition of phytase to maize and soybean low-availablephosphorus meals improves the performance and increasesCa, P, and Zn utilization in chicks.

Fish meal is becoming an increasingly expensive resourceas the world demand is rising. Much of the current researchin commercial fish feed formulation is therefore focusing onhow to replace fish meal by cheaper and more readily avail-able protein sources of plant origin, and good availability ofphosphorus in feed for aquatic animals is also important.The effect of a supplemental fungal phytase (0 or 1400 U kg−1

feed−1) on performance and phosphorus availability onjuvenile rainbow trout fed diets with a high inclusion ofplant based protein and on the magnitude and compositionof the waste phosphorus production was tested. Growthand feed conversion ratios were not significantly affectedby the increased dietary phosphorus level or supplementalfungal phytase, but this last one improved the availability ofphytate-phosphorus from an average of 6 to 64%. The fishretained 53%–79% of the ingested phosphorus, while 24%–44% was recovered in the feces. This study demonstrated thatphytase supplementation will be advantageous to the fish andthe environment if supplemented to low-phosphorus dietscontaining a large share of plant-derived protein [59].

Phytate, which cannot be digested by shrimp due to lackof phytase, becomes a pollutant in the aquatic environment.A study with tiger shrimp (P. monodon juveniles) fed withsoybean meal showed that although phytase supplementa-tion has no effect on shrimp growth, there is significantlylower total phosphorus excretion, and this result is usefulfor low-pollution shrimp feed [60]. This has been drivenby recognition of the ecological need to reduce P levels ineffluents and an increasing scientific and practical apprecia-tion of the roles of phytate and phytase in animal nutrition.Moreover, the recent proliferation of phytases in the market-place has generated price reduction and facilitated theirinclusion in pig and poultry diets [57].

5. Conclusion

The new demands of meat products by consumers make thefood industry search continuously for higher quality, betterprices, and less environmental damage. Enzyme applicationin the manufacturing of meat products and some possibilitiesto treat their waste have become important alternatives tomeet these needs. Further studies to apply enzymes in meat

technology are important to optimize existing processes, aswell as to develop new methods of application.

Acknowledgment

The authors thank CNPq (Conselho Nacional de Desenvolvi-mento Cientıfico e Tecnologico) for financial support.

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8 Enzyme Research

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SAGE-Hindawi Access to ResearchEnzyme ResearchVolume 2010, Article ID 174354, 9 pagesdoi:10.4061/2010/174354

Review Article

Enzymatic Strategies to Detoxify Gluten:Implications for Celiac Disease

Ivana Caputo,1, 2 Marilena Lepretti,1 Stefania Martucciello,1 and Carla Esposito1, 2

1 Department of Chemistry, University of Salerno, 84084 Salerno, Italy2 European Laboratory for the Investigation of Food-Induced Diseases, University Federico II, 80131 Naples, Italy

Correspondence should be addressed to Carla Esposito, cesposito@unisa.it

Received 4 July 2010; Accepted 14 September 2010

Academic Editor: Raffaele Porta

Copyright © 2010 Ivana Caputo et al. This is an open access article distributed under the Creative Commons Attribution License,which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Celiac disease is a permanent intolerance to the gliadin fraction of wheat gluten and to similar barley and rye proteins that occursin genetically susceptible subjects. After ingestion, degraded gluten proteins reach the small intestine and trigger an inappropriateT cell-mediated immune response, which can result in intestinal mucosal inflammation and extraintestinal manifestations. Todate, no pharmacological treatment is available to gluten-intolerant patients, and a strict, life-long gluten-free diet is the onlysafe and efficient treatment available. Inevitably, this may produce considerable psychological, emotional, and economic stress.Therefore, the scientific community is very interested in establishing alternative or adjunctive treatments. Attractive and novelforms of therapy include strategies to eliminate detrimental gluten peptides from the celiac diet so that the immunogenic effectof the gluten epitopes can be neutralized, as well as strategies to block the gluten-induced inflammatory response. In the presentpaper, we review recent developments in the use of enzymes as additives or as processing aids in the food biotechnology industryto detoxify gluten.

1. Celiac Disease

Celiac Disease (CD) is a condition affecting 1:70–1:200individuals worldwide that may be diagnosed at any age[1, 2]. In a population-based study, increasing prevalenceand high incidence of CD (1:47) in elderly people (older than52 years of age) have been remarked [3]. CD is a permanentfood intolerance to the ingested gliadin fraction of wheatgluten and similar alcohol-soluble proteins of barley andrye in genetically susceptible subjects [4, 5]. Most patientstolerate oats without any signs of intestinal inflammationprobably because oat avenins are phylogenetically moredistant from the analogous proteins in wheat, rye, andbarley [6]. Nonetheless, a few individuals with clinical oatintolerance have avenin-reactive mucosal T cells that cancause mucosal inflammation [7]. In children prone to CD,exposure to wheat, barley, and rye in the first three monthsof life significantly increases the risk of developing CDcompared with exposure between 4 and 6 months [8]

whereas breastfeeding exerts a protective effect and the riskof CD is reduced if children are still being breast-fed whendietary gluten is introduced [9].

The clinical features of CD vary considerably [2]. Intesti-nal symptoms are frequent in children diagnosed withinthe first two years of life. However, asymptomatic patientscan be found: failure to thrive, chronic diarrhoea, vomiting,abdominal distension, muscle wasting, anorexia, and generalirritability are present in most cases. The wider use ofserological screening tests is making it easier to recognizeextra-intestinal manifestations such as short stature, anaemiaunresponsive to iron therapy, osteoporosis, ataxia, periph-eral neuropathies, hypertransaminasemia, and unexplainedinfertility [10]. It is also noteworthy that CD is associatedwith a high prevalence of concomitant autoimmune diseases(approximately 5–10 times greater than in the general pop-ulation), such as endocrine autoimmune diseases, thyroiddiseases, and selective IgA deficiency, as well as of geneticdisorders, such as Down and Turner’s syndromes [11].

2 Enzyme Research

Because symptoms may improve with a gluten-free diet,it is thought that gluten plays a key role in the pathogenesisof this disease.

2. Gluten

The grain protein content of wheat varies between 8 and 17percent, depending on genetic make-up and external factorsassociated with the crop. A unique property of wheat flouris that, when in contact with water, the insoluble proteinfraction forms a viscoelastic protein mass known as gluten.Gluten, which comprises roughly 78 to 85 percent of the totalwheat endosperm protein, is a very large complex mainlycomposed of polymeric (multiple polypeptide chains linkedby disulphide (SS) bonds) and monomeric (single chainpolypeptides) proteins known as glutenins and gliadins,respectively. Gliadin consists of proteins containing α/β-, γ-,and ω-gliadins. In contrast to α/β- and γ-gliadins, whichform three and four intramolecular (SS) bonds, respectively,ω-gliadins lack cysteine residues. Glutenin is a heterogeneousmixture of SS-linked polymers with a largely unknownpolymer structure. A glutenin polymer consists of gluteninsubunits of high or low molecular weight that are connectedby intermolecular SS bonds. Glutenins confer elasticity, whilegliadins mainly confer viscous flow and extensibility to thegluten complex. Thus, gluten is responsible for most of theviscoelastic properties of wheat flour doughs, and it is themain factor dictating the use of a wheat variety in bread andpasta making. Gluten viscoelasticity, for end-use purposes, iscommonly known as flour or dough strength [12, 13].

Gliadins and glutenins have a unique amino acid com-position with a high content of proline (15%), hydrophobicamino acids (19%), and glutamine (35%), hence they arenamed prolamins. Moreover, they contain domains withnumerous repetitive sequences rich in those amino acids.Because of this glutamine- and proline-rich structure, glutenproteins are resistant to complete digestion by pancreatic andbrush border proteases [14, 15].

3. Posttranslational Modification of Gluten byTissue Transglutaminase

CD is triggered by an inappropriate T cell-mediated immuneresponse to dietary gluten proteins. Consequently, CDpatients display various degrees of intestinal inflammation,ranging from mild intraepithelial lymphocytosis to severesubepithelial mononuclear cell infiltration that results intotal villous atrophy coupled with crypt hyperplasia. Themost evident expression of autoimmunity in CD is thepresence of serum antibodies to tissue transglutaminase(tTG), the main autoantigen of endomysial antibodies [16].

tTG is a member of a Ca2+-dependent enzyme familyinvolved in post-translational modifications of proteins.tTG prevalently catalyzes the formation of stable isopeptidebonds between the γ-carboxamide group of the protein-bound glutamine residue and an appropriate amino group,either the ε-amino group of a protein-bound lysine residueor a small biogenic amine molecule such as putrescine,

spermine, spermidine, and histamine. However, the absenceof suitable nucleophilic amines and a low pH favours tTGdeamidation of protein-bound glutamine residues [17].

The presence of tTG-specific autoantibodies only inpatients who have gluten in their diet suggests that thegeneration of such antibodies in CD requires gluten asan exogenous trigger. The proposed mechanism by whichautoimmunity develops in CD is that the enzyme tTG gen-erates additional antigenic epitopes by cross-linking gliadinpeptides to itself and/or to other protein substrates, and thisstimulates mucosal T cells to produce autoantibodies againsttTG and gliadin [18] (Figure 1). Since the existence of tTG-specific T cells in the intestinal mucosa of untreated patientsis not proven, it is hypothesized that the production of anti-tTG antibodies is driven completely by intestinal gliadin-specific T cells. The observation that anti-tTG antibodytiters fall and can become undetectable during a gluten-free diet suggests that B cell activity depends on persistentantigen presentation. In a pioneering study in 1990, Portaet al. demonstrated that wheat glutelins and gliadins, aswell as purified A-gliadin, act as acyl donor substrates fortTG [19]. In particular, by performing incubations in vitroboth in the presence of radiolabeled polyamines and in theirabsence, Porta et al. showed that these proteins were ableto produce not only γ(glutamyl)polyamine adducts but alsopolymeric complexes, probably through intermolecular ε(γ-glutamyl)lysine crosslinks. In the case of A-gliadin, the singlelysil residue occurring in the amino acid sequence (K-186)is assumed to act as an acyl acceptor site. It is worth notingthat the increase of both tTG activity in situ and tTG proteinhas been detected at critical sites of celiac mucosae, such asthe intestinal brush border and subepithelial compartments[20].

The involvement of tTG in the pathogenesis of CDcould be also due to another distinct but interdependentpathway via a gliadin-derived peptide deamidation reaction(Figure 1). Gluten peptides are specifically recognized byhuman leukocyte antigen (HLA)-DQ2/DQ8, a class II majorhistocompatibility complex [21]. Indeed, CD is stronglyassociated with the genes encoding HLA-DQ2, and gluten-specific CD4+ intestinal T cells can be isolated from intestinalbiopsies of CD patients but not from healthy controls [21].By contrast, there is no evidence of T cell-mediated reactivityagainst dietary gliadin in the nonceliac mucosa. Moreover,gliadin-specific T lymphocytes from CD intestinal mucosaare mainly of the Th1/Th0 phenotype, which after gliadinrecognition, release prevalently proinflammatory cytokinesdominated by interferon (IFN)-γ [22] and interleukin (IL)-10 [23]. It has been hypothesized that tTG might be responsi-ble for the deamidation of specific glutamine residues withinnaturally digested gluten peptides, especially at low pH.Such tTG-catalyzed posttranslational modification generatesnegatively charged amino acid residues that bind with anincreased affinity to the HLA-DQ2 or HLA-DQ8 molecules,thus potentiating T cell activation [24, 25]. Recognition ofthe T cell epitope has been particularly difficult since gliadin’speculiar amino acid composition and its high glutaminecontent make it an excellent tTG substrate. A 33 mer peptide,containing three of the most immunogenic epitopes, was

Enzyme Research 3

Gliadin peptides

NH2

NH2

NH2

tTG

OH

C C

C

C

O

H2O

Deamidation

(CH2)4

(CH2)4

(CH2)2

(CH2)2

(CH2)2

(CH2)2

Ca2++NH3

+NH3

Q

K

N

H

GliadinDeamidated gliadinTransamidated gliadin

Gliadin-tTG complex

DQ2/DQ8T cell receptorCD4Anti-tTG antibodyCytokines

O

O

O

X PE

K

tTG

Q X P

APC

T T

B

Transamidation

Plasma cell

+

Macrophage

Q

Th1

Figure 1: Tissue transglutaminase (tTG)-mediated post-translational modifications in celiac disease. Gliadin peptides reach the subepithelialregion of the intestinal mucosa. Here, tTG deamidation of specific glutamines of gliadin peptides generates potent immunostimulatoryepitopes that are presented via HLA-DQ2/DQ8 on antigen-presenting cells (APC) to CD4+ T cells. Activated gliadin-specific CD4+ T cellsproduce high levels of pro-inflammatory cytokines, thus inducing a Th1 response that results in mucosal remodelling and villous atrophy.In addition, tTG transamidation activity generates tTG-gliadin complexes that bind to tTG-specific B cells, are endocytosed and processed.Gliadin-DQ2/DQ8 complexes are then presented by the tTG-specific B cells to gliadin-specific T cells, a process that leads to the productionof anti-tTG antibodies.

identified as one of the main stimulators of the inflammatoryresponse to gluten, resistant to intestinal proteases [26, 27].However, Camarca et al. recently demonstrated that therepertoire of gluten peptides recognized by adult celiacpatients is larger than had been previously thought, and itdiffers from one individual to another. Indeed, they foundseveral active gluten peptides with a large heterogeneity ofresponses [28].

Although the role of gluten in activating gluten-specificT lymphocytes in the lamina propria is well established, ithas been demonstrated that gluten contains peptides that canstimulate cells of the innate immune system. The prototypeof innate peptides is peptide 31–43/49, which has beenshown to be toxic for CD patients both in vitro and in vivo[29, 30]. Peptide 31–43/49 can reorganize intracellular actin

filaments [31], induce maturation of bone-marrow-deriveddendritic cells [32], and, by affecting epithelial growth factor-receptor decay, induce epithelial cell proliferation [33].The peptide also stimulates the synthesis and release ofthe proinflammatory cytokine IL-15 that can promote anadaptive immune response [34] involving CD4+ T cells thatrecognize various deamidated gliadin peptides [25]. Most ofthe events related to innate immune activation were inhibitedby antibodies neutralizing IL-15, thus confirming that thiscytokine mediates intestinal mucosal damage induced byingestion of gliadin. In particular, Barone et al. investigatedthe molecular mechanisms of the gliadin-induced IL15increase and discovered that gliadin peptide 31–43 increasesthe levels of IL-15 on the cell surface of CaCo-2 cells probablyby interfering with its intracellular trafficking [35].

4 Enzyme Research

4. Treatment of Celiac Disease

To date, no pharmacological treatment is available to gluten-intolerant patients. A strict, life-long, gluten-free diet isthe only safe and efficient treatment available, althoughit results in a social burden. Adhering to a gluten-freediet can have a significant negative impact on perceivedquality of life and may produce considerable psychological,emotional, and economic stress. Moreover, this requirementfor dietary compliance is made more difficult by theexclusion of wheat, rye, and barley from the diet, whichare important sources of iron, dietary fibre, and vitamin B,especially for adolescents and adults who need continuousmonitoring by dieticians [36]. A lifelong gluten-free dietcan be extremely difficult since gluten may be presentin nonstarchy foods such as soy sauce and beer, as wellas in nonfood items including some medications, postagestamp glue, and cosmetics (e.g., lipstick). CD patients can,therefore, be exposed inadvertently to gluten. Moreover, evenafter many years of gluten avoidance, CD patients neveracquire tolerance to gliadin, and re-exposure to the antigenreactivates the disease. Finally, it is worth noting that a smallgroup of patients with CD (2%–5%) fail to improve clinicallyand histologically upon elimination of dietary gluten. Thiscomplication is referred to as refractory CD, and it imposesa serious risk for developing lethal enteropathy-associated T-cell lymphoma.

In 2000, the Codex Alimentarius Commission of theWorld Health Organization and the FAO described gluten-free foods consisting of, or made only from, ingredientswhich do not contain any prolamines from wheat or anyTriticum species, such as spelt, kamut or durum wheat, rye,barley, oats, or their crossbred varieties with a gluten level notexceeding 20 ppm [37, 38]. At present, gluten-free productsare not widely available; they are usually expensive, and theyhave poor sensory and shelf life properties. Research anddevelopment are currently focused on improving mouth-feel, flavour, and rheology of gluten-free products. Gluten isresponsible for most of the viscoelastic properties of wheatflour doughs, and its absence can result in a baked breadwith a crumbling texture, poor colour and other postbakingquality defects [37]. For all these reasons the search forsafe and effective therapeutic alternatives to a gluten-freediet, which are compatible with a normal social lifestyle,is of great importance. Advances in our understandingof the complex mechanisms involved in CD pathogenesishave opened several promising avenues for therapeuticintervention aimed at targeting each factor involved in thedisease onset, some of which are being tested in early clinicaltrials [39]. The identification of T-cell stimulatory gliadinsequences (33 mer peptide) is important so that peptideanalogues of gliadin epitope(s) can be engineered to generatepeptides that exert antagonistic effects. Of course, thechances of success of using peptide analogues to modulatespecific immune responses could be hampered by the wideheterogeneity of the gliadin T-cell epitopes. Elucidation ofthe hierarchy of pathogenic gliadin epitopes and their coreregion is required before a peptide-based therapy can bedesigned [40]. Antibodies to IL-15 have also been proposed

as a treatment strategy, particularly in cases of refractorysprue because of intraepithelial lymphocyte activation inthis condition [41], as well as antibodies to IFN-γ [42].Other promising treatment strategies are aimed at preventinggliadin presentation to T cells by blocking HLA-binding sitesand using IL-10 as a tool for promoting tolerance [23]. Toreverse the toxic effects induced by gliadin in human intesti-nal cells and gliadin-sensitive HCD4-DQ8 mice, Pinier et al.proposed a completely different strategy based on the use ofsynthetic sequestering polymeric binders that can complexand neutralize gliadin in situ. Coadministration of syntheticpolymeric binders and gliadin to HLA-HCD4/DQ8 miceattenuated gliadin-induced changes in the intestinal barrierand reduced intraepithelial lymphocyte and macrophage cellcounts [43]. Recent new therapeutic approaches includecorrection of the intestinal barrier defect against gluten entry.An intestinal permeability blocker (AT1001), which is aninhibitor of the zonulin pathway that acts to prevent gliadinfrom inducing increased intestinal permeability, is currentlyin a phase IIb clinical study [44]. Finally, a vaccine that coulddesensitize or induce tolerance in individuals with CD hasbeen proposed [45, 46].

Besides therapeutic treatments, transgenic technologyand breeding ancient varieties have been tried with the goalof developing grains that have a low or zero content ofimmunotoxic sequences, but with reasonable baking quality.However, these approaches are difficult due to the numberand the repetition of sequence homologies in the cerealprotein family, and because cereals like wheat are hexaploid[47, 48].

5. Enzyme Therapy

Enzyme supplement therapies are focused on inactivatingimmunogenic gluten epitopes (Table 1).

5.1. Oral Administration of Bacterial Endopeptidases. Afteringestion, degraded gluten proteins reach the small intestine.However, because of their unusually high proline andglutamine content, especially in immunodominant gliadinpeptides like the 33 mer, gluten is poorly degraded by theenzymes present in the gastrointestinal tract. Hence, oralenzyme therapy has been suggested as an alternative to thegluten-free diet. Promising enzymes (expressed in variousmicroorganisms) tested are the prolyl oligopeptidases fromFlavobacterium meningosepticum, Sphingomonas capsulate,and Myxococcus xanthus. These enzymes are capable ofdegrading proline-containing peptides that are otherwiseresistant to degradation by proteases in the gastrointestinaltract in vitro [49–51]. However, most of these enzymes areirreversibly inactivated in the stomach by pepsin and acidicpH, thus failing to degrade gluten before it reaches the smallintestine [49]. Encapsulation of these prolyl oligopeptidaseswas proposed in order to protect them from gastric juices[51]. However, in a recent ex vivo study, using biopsy-derived intestinal tissue mounted in Ussing chambers, it wasobserved that only high doses of prolyl oligopeptidase werecapable of eliminating the accumulation of immunogenic

Enzyme Research 5

peptides in the serosal compartment [50]. This indicates that,even if the enzyme were encapsulated, it is too inefficientto degrade gluten before it reaches the proximal part ofthe duodenum, the site where gluten triggers inflammatoryT-cell responses [50]. Mitea et al. recently investigated anew prolyl endoprotease from Aspergillus niger. This enzymewas found to degrade gluten peptides and intact glutenproteins efficiently in the stomach, to such an extent thathardly any traces of gluten reached the duodenal com-partment [52]. Moreover, the optimum pH of this enzymeis compatible with that found in the stomach and theenzyme is resistant to degradation by pepsin. Finally, prolylendoprotease from Aspergillus niger is derived from the foodgrade microorganism and is available on an industrial scale.These results indicate that this enzyme might be suitablefor oral supplementation to degrade gluten proteins in foodbefore they reach the small intestine [52]. Recently, Gasset al. evaluated a new combination therapy, consisting oftwo gastrically active enzymes that detoxify gluten before itsrelease in the small intestine. They used a glutamine-specificendoprotease (EP-B2; a cysteine endoprotease from ger-minating barley seeds) and a prolyl-specific endopeptidasefrom Sphingomonas capsulata, for its ability to digest glutenunder gastric conditions. Endoprotease EP-B2 extensivelyhydrolyzes the gluten network in bread into relativelyshort (but still inflammatory) oligopeptides, whereas prolyl-specific endopeptidase from Sphingomonas capsulata rapidlydetoxifies oligopeptides after primary proteolysis at internalproline residue level to yield nontoxic metabolites [53].A practical advantage of this combination product is thatboth enzymes are active and stable in the stomach and cantherefore be administered as lyophilized powders or simplecapsules or tablets.

5.2. Pretreatment of Whole Gluten with Bacterial-DerivedPeptidase. An alternative approach to detoxify gluten isrepresented by the digestion of wheat gluten peptideswith bacterial-derived peptidase during food processing andbefore administration to patients. Traditional methods toprepare cereal foods, including long fermentation times byselected sourdough lactic acid bacteria, have mostly beensubstituted by the indiscriminate use of chemical and/orbaker’s yeast leavening agents. Under these circumstances,cereal components (e.g., proteins) are subjected to verymild or absent degradation during manufacture, resulting,probably, in reduced digestibility compared to traditionaland ancient sourdough baked goods [54]. Di Cagno et al.selected four sourdough lactobacilli (L. alimentarius 15M,L. brevis 14G, L. sanfranciscensis 7A, and L. hilgardii 51B)that showed considerable hydrolysis of albumin, globulin,and gliadin fractions during wheat sourdough fermentation.These lactobacilli had the capacity to hydrolyze the 31–43 fragment of A-gliadin in vitro and, after hydrolysis,greatly reduced the agglutination of K 562(S) subclonecells of human myelogenous leukemia origin by a toxicpeptic-tryptic digest of gliadins [55]. On the basis of theseresults, and with the goal of decreasing gluten intolerancein humans, the authors investigated a novel bread making

method that used selected lactobacilli to hydrolyze variousPro-rich peptides, including the 33 mer peptide, for theproduction of sourdoughs made from a mixture of wheatand nontoxic oat, buckwheat, and millet flours [56]. After24 hours of fermentation, wheat gliadins and low-molecular-mass, alcohol-soluble polypeptides were almost completelyhydrolyzed. Proteins extracted from sourdough were usedfor in vitro agglutination tests on K 562(S) subclone cells ofhuman origin and to produce two types of bread, containingca. 2 g of gluten. The latter were used in an in vivo double-blind acute challenge of CD patients. Agglutination testingof K 562(S) cells and the acute in vivo challenge showedimproved tolerance of breads containing 30% wheat flour[56]. The reported data suggest that long-time fermentationin the presence of a mixture of selected lactic acid bacteriaseemed to be indispensable to reduce toxicity. In actual fact,different probiotic bacterial strains have their characteristicset of peptidases, which may diverge considerably fromeach other and have variable substrate specificities. Inline with this concept, different probiotic bacterial strainshave been tested. Among probiotic preparations, VSL#3,a highly concentrated mixture of lactic acid and bifido-bacteria, was able to hydrolyze completely the α2-gliadin-derived epitopes 62–75 and 33 mer (750 ppm) [57]. It isinteresting to underline that probiotics, defined as theviable microorganisms that exhibit a beneficial effect onthe health of the host by improving its intestinal microbialbalance, could directly modulate the function of epithelialcells. It has been reported that different probiotic strains,including the VSL3# preparation, increase epithelial barrierfunction by stabilizing tight junctions and inducing mucinsecretion in epithelial cells [57, 58]. Furthermore, severalprobiotic bacterial strains are able to protect the epithe-lium, presumably by the aforementioned mechanisms, fromvarious insults, including pathogenic bacteria [59, 60] andinflammatory cytokines [61, 62]. More recently, Rizzello etal. showed that fermentation with a complex formula ofsourdough lactobacilli decreased the concentration of glutento below 10 ppm [63]. Specifically, they used a mix of tensourdough lactobacilli that were selected for their peptidasesystems capability to hydrolyze Pro-rich peptides, includingthe 33 mer peptide, together with fungal proteases, that areroutinely used as improvers in the baking industry. In thisway, wheat and rye breads or pasta, if supplemented withgluten-free flour-based structuring agents, may be toleratedby CD patients. Agglutination testing of K 562(S) cells andan acute in vivo challenge showed improved tolerance ofbreads containing 30% wheat flour. Moreover, prolongedin vivo challenge of CD patients confirms reduced toxicityof gluten fermented with selected lactobacilli and fungalproteases. In fact, CD patients (age >12 years old) ona gluten-free diet for at least five years were challengedwith a daily intake of 10 g of hydrolysed gluten (<20 ppmof gluten) for 2 months. Intestinal functional tests, aswell as CD serum antibodies (anti-tTG, anti-endomysium),and duodenal histology and immunohistochemistry atthe beginning and after 60 days of challenge showedthat all parameters were normal, i.e., no villous atrophy)[64].

6 Enzyme Research

Table 1: Potential enzyme therapies for celiac disease.

Target fordetoxification

Detoxifying agent Mechanism of actionStatus ofresearch

Ingested gliadinpeptides

Prolyl endopeptidases from:

Hydrolysis ofproline-rich peptides ofgliadin

PreclinicalS. capsulate [49]

F. meningosepticum [50]

M. xanthus [51]

Prolyl endopeptidases from: A. niger [52] Clinical trial

Prolyl endopeptidases from: S. capsulate incombination with a glutamine-specificendoprotease (EP)-B2 from germinating barley[53]

Clinical trial

FlourSourdough lactobacilli-derived peptidases [56] Hydrolysis of

proline-rich peptides ofgliadin

Clinical trial

Sourdough lactobacilli-derived peptidases incombination with fungal proteases [64]

Clinical trial

Flour Transglutaminase enzymes [67]Transamidation ofgliadin peptides withlysine methyl ester

Preclinical

Mucosal tTGIrreversible thiol-reactive reagents, competitivepeptidic, and nonpeptidic substrates [69]

unspecific or specifictTG inhibition

Preclinical

The use of proteases from germinating wheat seeds hasalso been proposed to create safe cereal products for CDpatients [65, 66].

5.3. Transamidation of Gliadin. tTG-catalyzed deamidationof specific glutamine residues within naturally digestedgluten peptides generates negatively charged amino acidresidues that bind with an increased affinity to the HLA-DQ2/DQ8 molecules, thus potentiating T cell activation.Based on this assumption, Gianfrani et al. proposed anenzyme strategy to inactivate immunogenic peptide epitopesand, at the same time, to preserve the integrity of theprotein structure via the transamidation of wheat flourwith a food-grade enzyme and an appropriate amine donor[67]. Interestingly, a recent study showed that the formationof the DQ2-α-II epitope was blocked by 5-biotinamidopentylamine and by monodansylcadaverine, reagents knownto cross-link glutamine residues [68]. To this end, the authorstreated wheat flour with tTG and lysine methyl ester; thelysine-modified gliadin peptides lost almost completely theiraffinity to bind to HLA-DQ2. Moreover, lysine-modifiedgliadin peptides caused a drastic reduction in gliadin-specificIFN-γ production in intestinal T-cell lines derived from CDpatients where the mucosal lesion was mainly induced by theproduction of IFN-γ from these gluten-specific T cells. Thisresult suggests that transamidation neutralized the immunereactivity of a large repertoire of epitopes. Similar resultswere obtained by using microbial TG, which is different fromtTG since it is calcium independent and is a low-molecular-weight protein that exhibits advantages in food industrialapplications. This enzyme is commercially available as adough improver that adds stability and elasticity to thedough. Additionally, bread volume and crumb texture arepositively influenced by the addition of microbial TG,

especially for flours with low gluten content and poor bakingperformance.

5.4. Transglutaminase Inhibitors. tTG plays an importantrole in CD pathology as it catalyzes deamidation andcross-linking of specific gluten peptides and converts theminto potent epitopes recognized by intestinal T-cells. Inorder to restrain the T cell-mediated immune response todietary gluten, a different approach could be to considertTG as a potential therapeutic target [69]. The inhibitionof the tTG-catalyzed deamidation of specific glutamineresidues within naturally digested gluten peptides mightnot generate negatively charged amino acid residues andtherefore might not increase the binding to the HLA-DQ2/DQ8 molecules (thus potentiating T cell activation).Several inhibitors acting with different mechanisms thattarget the TG cross-linking activity have been developed andtested, mainly in vitro [69, 70]. Among the tTG inhibitorstested we can find several nonspecific irreversible thiol-reactive reagents, also named suicide TG inhibitors, suchas cystamine, able to inhibit tTG via the formation ofan enzyme-inhibitor complex. Furthermore, we can findcompetitive nonpeptidic tTG amino donor substrates, suchas 1,4-diaminobutane (Fibrostat), which is used topicallyin a clinical trial to treat abnormal wound healing [71],and competitive peptidic tTG amino donor. Finally, we canfind amine acceptor pseudosubstrates able to inhibit tTGactivity by diverting it from the natural protein substrate.Recently, Hoffmann et al. used a blocking peptide approachto reduce the processing of gliadin by tTG. The authorsshowed that these peptides have a potential for glutendetoxification and could be evaluated as an alternative fordesigning new food products for gluten-intolerant patients[72]. Several factors must be taken into consideration when

Enzyme Research 7

designing a tTG inhibitor: inhibitory potency, optimal sizeof the compound, resistance towards intestinal proteolyticactivities (to this end, amino acid residues will be replacedby peptidomimetics), and selectivity towards tTG. In fact,the lack of specificity limits therapeutic utility. Furthermore,to reduce the risk of systemic side effects, the activity of anoptimal tTG inhibitor should be specifically limited to thecompartment where gliadin encounters the immune system,that is, in the gut. Therefore, blocking the transamidatingactivity of tTG represents an attractive tool to preventimmune activation. Similar approaches have already beeninvestigated in other diseases where tTG is involved, suchas in the neurodegenerative disorders such as Parkinson,Huntington, and Alzheimer diseases, as well as in cancer andin fibrotic/scarring conditions such as diabetic nephropathy.Consequently, tTG inhibitory drugs can be predicted to havea wide medical application.

6. Concluding Remarks

The high incidence of CD in the worldwide population is achallenging task, given the negative impact of a strict gluten-free diet on the perceived quality of life of celiac patientsfor several reasons. A life-long gluten-free diet is not easyto maintain since gluten is the most common ingredientin the human diet. Multidisciplinary research efforts arecurrently being carried out in several directions to findnew treatment strategies in order to reduce gluten toxicity.The use of oral proteases capable of detoxifying ingestedgluten and new food grade fermentation technologies usingbacterial-derived endopeptidases currently represent themost advanced and promising strategies.

Abbreviations

CD: Celiac diseaseSS: DisulphidetTG: Tissue transglutaminaseHLA: Human leukocyte antigenIFN: InterferonIL: Interleukin.

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[71] K. N. Dolyuchuk, “Topical putrescine (fibrostat) in treatmentof hypertrophic scars: phase II study,” Plastic and Reconstruc-tive Surgery, vol. 98, no. 4, p. 752, 1996.

[72] K. Hoffmann, M. Alminger, T. Andlid, T. Chen, O. Olsson, andA.-S. Sandberg, “Blocking peptides decrease tissue transglu-taminase processing of gliadin in vitro,” Journal of Agriculturaland Food Chemistry, vol. 57, no. 21, pp. 10150–10155, 2009.

SAGE-Hindawi Access to ResearchEnzyme ResearchVolume 2010, Article ID 918761, 8 pagesdoi:10.4061/2010/918761

Review Article

Uses of Laccases in the Food Industry

Johann F. Osma,1 Jose L. Toca-Herrera,2 and Susana Rodrıguez-Couto3, 4

1 Department of Electrical and Electronics Engineering, University of the Andes, Carrera 1 No. 18A-12, Bogota, Colombia2 Department of Nanobiotechnology, University of Natural Resources and Applied Life Sciences (BOKU), Muthgasse 11,1190 Vienna, Austria

3 Unit of Environmental Engineering, CEIT, Paseo Manuel de Lardizabal 15, 20018 San Sebastian, Spain4 IKERBASQUE, Basque Foundation for Science, Alameda Urquijo 36, 48011 Bilbao, Spain

Correspondence should be addressed to Susana Rodrıguez-Couto, ladysue212@yahoo.es

Received 15 June 2010; Accepted 22 August 2010

Academic Editor: Raffaele Porta

Copyright © 2010 Johann F. Osma et al. This is an open access article distributed under the Creative Commons AttributionLicense, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properlycited.

Laccases are an interesting group of multi copper enzymes, which have received much attention of researchers in the last decadesdue to their ability to oxidise both phenolic and nonphenolic lignin-related compounds as well as highly recalcitrant environmentalpollutants. This makes these biocatalysts very useful for their application in several biotechnological processes, including thefood industry. Thus, laccases hold great potential as food additives in food and beverage processing. Being energy-saving andbiodegradable, laccase-based biocatalysts fit well with the development of highly efficient, sustainable, and eco-friendly industries.

1. Introduction

Laccases (p-diphenol:dioxygen oxidoreductases; EC 1.10.3.2)are particularly abundant in white-rot fungi, which are theonly organisms able to degrade the whole wood components[1]. Fungal laccases are secreted, glycosylated proteins withtwo disulphide bonds and four copper atoms distributed inone mononuclear termed T1 (where the reducing substrateplace is) and one trinuclear cluster T2/T3 (where oxygenbinds and is reduced to water) [2]. Thus, electrons aretransferred from substrate molecules through the T1 copperto the trinuclear T2/T3 centre. After the transfer of fourelectrons, the dioxygen in the trinuclear centre is reduced totwo molecules of water [3, 4] (Figure 1).

From a mechanistic point of view, the reactions catalysedby laccases can be represented by one of the schemesshown in Figure 2. The simplest case (Figure 2(a)) isthe one in which the substrate molecules are oxidised tothe corresponding radicals by direct interaction with thecopper cluster. Frequently, however, the substrates of interestcannot be oxidised directly by laccases, either because theyare too large to penetrate into the enzyme active site orbecause they have a particularly high redox potential. Bymimicking nature, it is possible to overcome this limitation

with the addition of so-called “redox mediators”, which arelow-weight molecular compounds that act as intermediatesubstrates for laccases, whose oxidised radical forms are ableto interact with the bulky or high redox potential substratetargets (Figure 2(b)).

In nature, the role of laccases is to degrade lignin in orderto gain access to the other carbohydrates in wood (celluloseand hemicellulose). Their low substrate specificity allowslaccases to degrade compounds with a structure similar tolignin, such as polyaromatic hydrocarbons (PAHs), textiledyes, and other xenobiotic compounds [2]. This togetherwith the simple requirements of laccase catalysis (presence ofsubstrate and O2) makes laccases both suitable and attractivefor industrial applications.

Typical fungal laccases are extracellular proteins ofapproximately 60–70 kDa with acidic isoelectric pointaround pH 4.0 [5]. They are generally glycosylated, with anextent of glycosylation ranging between 10 and 25% andonly in a few cases higher than 30% [6, 7]. This feature maycontribute to the high stability of the enzyme [8].

A few laccases are at present in the market for textile,food and other industries (Table 1), and more candidatesare being actively developed for future commercialisation[9]. A vast amount of industrial applications for laccases

2 Enzyme Research

OH

OHOH

O2 2H2O

O

O

444

O•

Figure 1: Reactions on phenolic compounds catalysed by laccases(extracted from [10]).

H2O

O2

Laccase(ox)

Laccase(red) Substrate(ox)

Substrate(red)

(a)

H2O

O2

Laccase(ox)

Laccase(red)

Substrate(ox)

Substrate(red)Mediator(ox)

Mediator(red)

(b)

Figure 2: Schematic representation of laccase-catalysed redox cyclesfor substrates oxidation in the absence (a) or in the presence (b)of redox mediators (extracted from [11], with kind permission ofElsevier Ltd.)

have been proposed which include pulp and paper, textile,organic synthesis, environmental, food, pharmaceutical, andnano-biotechnology. Being energy-saving and biodegrad-able, laccase-based biocatalysts fit well with the developmentof highly efficient, sustainable, and eco-friendly industries.

This paper reviews the potential application of laccases inthe food industry. The utilisation of whole laccase-producingmicroorganisms is not considered in the present paper.

2. Application of Laccases in the Food Industry

Many laccase substrates, such as carbohydrates, unsaturatedfatty acids, phenols, and thiol-containing proteins, areimportant components of various foods and beverages.Their modification by laccase may lead to new functionality,quality improvement, or cost reduction [12, 13].

2.1. As Additives in Food and Beverage Processing. Laccasescan be applied to certain processes that enhance or modifythe colour appearance of food or beverage.

2.1.1. Wine Stabilisation. Wine stabilisation is one of themain applications of laccase in the food industry as alterna-tive to physical-chemical adsorbents [12]. Musts and winesare complex mixtures of different chemical compoundssuch as ethanol, organic acids (aroma), salts, and phenoliccompounds (colour and taste). Polyphenol removal mustbe selective to avoid an undesirable alteration in the wine’sorganoleptic characteristics. Laccase presents some impor-tant requirements when used for the treatment of polyphenolremoval in wines such as stability in acid medium and

reversible inhibition with sulphite [14]. Additionally, alaccase has been commercialised for preparing cork stoppersfor wine bottles [15]. The enzyme oxidatively reducesthe characteristic cork taint and/or astringency, which isfrequently imparted to aged bottled wine.

2.1.2. Beer Stabilisation. The storage life of beer depends ondifferent factors such us haze formation, oxygen content,and temperature. The former is produced by small quantitiesof naturally-occurring proanthocyanidins, polyphenols thatgenerate protein precipitation and, therefore, the formationof haze [16]. This type of complex is commonly found aschill-haze and appears during cooling processes but may re-dissolve at room temperature or above [12]. Even productsthat are haze-free at the time of packing can develop this typeof complex during long-term storage. Thus, the formation ofhaze has been a persistent problem in the brewing industry[17]. The use of laccases for the oxidation of polyphenols asan alternative to the traditional treatment has been tested bydifferent authors [16, 18, 19]. However, laccases have alsobeen used for the removal of oxygen at the end of the beerproduction process. According to Mathiasen [16], laccasecould be added at the end of the process in order to removethe unwanted oxygen in the finished beer, and thereby thestorage life of beer is enhanced. Also, a commercialisedlaccase preparation named “Flavourstar”, manufactured byNovozymes A/S, is marketed for using in brewing beerto prevent the formation of off-flavour compounds (e.g.,trans-2-nonenal) by scavenging the oxygen, which otherwisewould react with fatty acids, amino acids, proteins andalcohol to form off-flavour precursors [20] (Table 1).

2.1.3. Fruit Juice Processing. Enzymatic preparations havebeen studied since the decade of the 1930s for juice clarifica-tion [21]. The interaction between proteins and polyphenolsresults in the formation of haze or sediment in clear fruitjuices. Therefore, clear fruit juices are typically stabilised todelay the onset of protein-polyphenol haze formation [22].Several authors have proposed the use of laccase for the sta-bilisation of fruit juices [23–30]; however, results are contra-dictory. On one hand, Sammartino et al. [24] compared thetreatment of apple juice with a conventional method (SO2

added as metabisulfite, polyvinylpolypyrrolidone (PVPP),bentonite) with the use of free and immobilised laccase. Theyshowed that the enzymatically treated juice was less stablethan the one conventionally treated. Also, Giovanelli andRavasini [25] and Gokmen et al. [31] showed by stabilitytests of ultrafiltrated samples that laccase treatment increasedthe susceptibility of browning during storage. On the otherhand, Cantarelli [30] used a mutant laccase from Polyporusversicolor to treat black grape juice. He showed a removal of50% of total polyphenols and higher stabilisation than thephysical-chemical treatment.

The use of laccase in conjunction with a filtrationprocess has shown better results. Thus, Ritter et al. [27]and Maier et al. [29] obtained a stable and clear apple juiceby applying laccase in conjunction with cross-flow-filtration(ultrafiltration) in a continuous process without the addition

Enzyme Research 3

Table 1: Commercial preparations based on laccases for industrial processes.

Main application Brand name Manufacturer

Food industryBrewing Flavourstar Advanced Enzyme Technologies Ltd. (India)

Colour enhancement in tea, etc. LACCASE Y120 Amano Enzyme USA Co. Ltd.

Cork modification Suberase Novozymes (Denmark)

Paper industryPulp bleaching Lignozym-process Lignozym GmbH (Germany)

Paper pulp delignification Novozym 51003 Novozymes (Denmark)

Textile Industry

Denim bleaching Bleach Cut 3-S Season Chemicals (China)

Denim finishing Cololacc BB Colotex Biotechnology Co. Ltd. (Hong Kong)

Denim bleaching DeniLite Novozymes (Denmark)

Denim finishing Ecostone LC10 AB Enzymes GmbH (Germany)

Denim finishing IndiStar Genencor Inc. (Rochester, USA)

Denim finishing Novoprime Base 268 Novozymes (Denmark)

Denim bleaching and shading Primagreen Ecofade LT100 Genencor Inc. (Rochester, USA)

Denim bleaching ZyLite Zytex Pvt. Ltd. (India)

of finishing agents. Cantarelli and Giovanelli [28] reportedthat the use of laccase followed by “active” filtration orultrafiltration, by the addition of ascorbic acid and sulphites,improved colour and flavour stability in comparison toconventional treatments. Also, Stutz [26] used laccase andultrafiltration to produce clear and stable juice concentrateswith a light colour.

Artik et al. [32] studied the effect of laccase applicationon clarity stability of sour cherry juice. They found that highclarity was obtained by adding laccase in case of heating to50◦C for 6 h and filtering through 20 kDa membrane after 1 hof oxidation. Also, the phenolic content decreased by around70%.

More recently, Neifar et al. [23] used a combined laccase-ultrafiltration process for controlling the haze formation andbrowning of the pomegranate juice. The optimised treatmentwith laccase (laccase concentration 5 U/mL; incubationtime 300 min; incubation temperature 20◦C) followed byultrafiltration led to a clear and stable pomegranate juice.

2.1.4. Baking. Laccases are currently of interest in baking dueto their ability to cross-link biopolymers. The use of laccasein baking is reported to result in an increased strength,stability, and reduced stickiness and thereby improvedmachinability of the dough; in addition, an increased volumeand an improved crumb structure and softness of the bakedproduct were observed [33, 34].

Selinheimo et al. [35] showed that a laccase from thewhite-rot fungus Trametes hirsuta increased the maximumresistance of dough and decreased the dough extensibility inboth flour and gluten doughs. It was concluded that the effectof laccase was mainly due to the cross-linking of the esterifiedferulic acid (FA) on the arabinoxylan (AX) fraction of doughresulting in a strong AX network. Gluten dough treated withlaccase also showed some hardening suggesting that laccasecan also act to some extent on the gluten protein matrix. Thehardening effect of laccase was, however, clearly weaker ingluten dough. Thus, the AX fraction in flour dough is thepredominant substrate for laccase, and its activity caused the

hardening effect. Interestingly, laccase-treated flour doughsoftened as a result of prolonged incubation: the extentof softening increasing as a function of laccase dosage. Itis proposed that softening phenomenon is due to radicalcatalysed breakdown of the cross-linked AX network.

Renzetti et al. [36] showed that a commercial laccasepreparation significantly improved the bread-making perfor-mances of oat flour and the textural quality of oat breadby increasing specific volume and lowering crumb hardnessand chewiness. The improved bread-making performancescould be related to the increased softness, deformability andelasticity of oat batters with laccase supplementation.

2.1.5. Improving of Food Sensory Parameters. The physico-chemical deterioration of food products is a major problemrelated to the evolution of storing and distribution systemsand influences the consumer’s perception of the productquality. Thus, different uses of laccase have promotedodour control, taste enhancement, or reduction of undesiredproducts in several food products.

Takemori et al. [37] used crude laccase from Coriolusversicolor to improve the flavour and taste of cacao niband its products. Bitterness and other unpleasant tasteswere removed by the laccase treatment, and the chocolatemanufactured from the cacao mass tasted better than thecontrol.

Another type of food products that may use laccase toimprove sensory parameters is oil. Oil products may bedeoxygenated by adding an effective amount of laccase [38].Oils, especially vegetable oils (e.g., soybean oil), are present inmany food items such us dressings, salads, mayonnaise, andother sauces. Soybean oil contains a large amount of linoleicand linolenic acids that can react with dissolved oxygenin the product producing undesirable volatile compounds.Therefore, the flavour quality of some oils may be improvedby eliminating the oxygen present in the oils. Other foodproducts (e.g., juices, soups, concentrates, puree, pastes, andsauces) can also be deoxygenated by the mean of laccase [39].

4 Enzyme Research

Bouwens et al. [40, 41] reported that the colour of tea-based products could be enhanced when treated with laccasefrom a Pleurotus species. In the same way, chopped olivesin an olive-water mixture were treated with laccase fromTrametes villosa. In this case, the bitterness was considerablyreduced while the colour turned darker compared to thecontrols (Novo Nordisk A/S, 1995).

Tsuchiya et al. [42] used a recombinant laccase fromMyceliophthora thermophilum and chlorogenic acid to con-trol the malodour of cysteine. They showed that enzymat-ically treated cysteine presented a very weak odour whilethe nontreated cysteine presented a strong characteristic H2Sodour. HPLC analysis showed the reduction of more than50% of cysteine.

2.1.6. Sugar Beet Pectin Gelation. The sugar beet pectin is afunctional food ingredient that can form thermo-irreversiblegels. These types of gels are very interesting for the foodindustry as can be heated while maintaining the gel structure.

Norsker et al. [43] analysed the gelling effect of twolaccases and a peroxidase in food products. They found thatlaccases were more efficient as gelling agents in luncheonmeat and milk than peroxidase. In addition, in manycountries it is prohibited to add hydrogen peroxide to foodproducts making it impossible to use peroxidases as gellingagents. Hence, it is more realistic to add laccase to foodproducts.

Kuuva et al. [44] reported that by using laccases as cross-linking agents together with calcium, the ratio of covalentand electrostatic cross-links of sugar beet pectin gels can bevaried and it can be possible to tailor different types of gelstructures.

Littoz and McClements [45] showed that laccase could beused to covalently cross-link beet pectin molecules adsorbedto the surfaces of protein-coated lipid droplets at pH 4.5,thus suggesting that emulsions with improved functionalperformance could be prepared using a biomimetic approachthat utilised enzymes (laccases) to cross-link adsorbedbiopolymers.

2.2. Determination of Certain Compounds in Beverages. Theuse of laccases for improving the sensing parameters of foodproducts is not limited to treatment processes but also todiagnosis systems. In this regard, different amperometricbiosensors based on laccases have been developed to measurepolyphenols in different food products (e.g., wine, beer, andtea). Thus, Ghindilis et al. [46] showed the practical validityof a biosensor based on immobilised laccase in analysingtannin in tea of different brands.

Montereali et al. [47] reported the detection of polyphe-nols present in musts and wines from Imola (Italy) throughan amperometric biosensor based on the utilisation of tyrosi-nase and laccase from Trametes versicolor. Both enzymes wereimmobilised on graphite screen-printed electrodes modi-fied with ferrocene. Biosensors exhibited a good samplingbehaviour compared to that obtained from spectropho-tometric analysis; however, the presence of SO2 clearly

inhibited the enzymatic activity, and, thus, the measurementson musts and wines recently bottled were seriously affected.

Di Fusco et al. [48] reported the development of anamperometric biosensor based on laccases from T. versicolorand T. hirsuta for the determination of polyphenol indexin wines. Enzymes were immobilised on carbon nanotubesscreen-printed electrodes using polyazetidine prepolymer(PAP). They showed that biosensor performance dependedon the laccase source. Thus, values obtained by using T.hirsuta laccase were close to those determined by Folin-Ciocalteu method whereas polyphenol index measured withT. versicolor laccase was discordant to that found with thereference assay.

Prasetyo et al. [49] studied the use of tetramethoxyazobismethylene quinone (TMAMQ) for measuring theantioxidant activity of a wide range of structurally diversemolecules present in food and humans. TMAMQ wasgenerated by the oxidation of syringaldazine with laccasesand used to detect the antioxidant activity present in differentfood products.

Ibarra-Escutia et al. [50] developed and optimised anamperometric biosensor based on laccase from T. versicolorfor monitoring the phenolic compounds content in tea infu-sions. The biosensor developed showed an excellent stabilityand exhibited good performance in terms of response time,sensitivity, operational stability, and manufacturing processsimplicity and can be used for accurate determination of thephenolic content without any pretreatment of the sample.

2.3. Bioremediation of Food Industry Wastewater. The pres-ence of phenols in agroindustrial effluents has attractedinterest for the application of laccase-based processes inwastewater treatment and bioremediation. The presence ofphenolic compounds in drinking and irrigation water or incultivated land represents a significant health and/or envi-ronmental hazard. With government policies on pollutioncontrol becoming more and more stringent, industries havebeen forced to look for more effective treatment technologiesfor their wastewater.

Some fraction of beer factory wastewater represents animportant environmental concern due to its high content inpolyphenols and dark brown colour.

Distillery wastewater is generated during ethanol pro-duction from fermentation of sugarcane molasses (vinasses).It produces a serious ecological impact due to its highcontent in soluble organic matter and its intense dark browncolour. In fact, vinasses represent a major environmentalproblem for the ethanol production industry and they areconsidered as the most aggressive by-product generated bysugar-cane factories. Most of the organic matter present inthe vinasses can be diminished by conventional anaerobic-aerobic digestion, but the colour is hardly removed bythese treatments [51] making this effluent a potential waterpollutant blocking out light from rivers and streams therebypreventing oxygenation by photosynthesis and provokingtheir eutrophication.

Strong and Burgess [52] studied the fungal (Trametespubescens) and enzymatic (laccase from T. pubescens) reme-diation of different distillery wastewater and found that the

Enzyme Research 5

Table 2: Some prices of commercially available laccases (extracted from [12], with kind permission of Elsevier Ltd).

Quantity (Units)a Price

From Agaricus bisporus10.000 305.00 (US$)

100.000 1.560.00 (US$)

From Coriolus versicolor10.000 250.00 (US$)

100.000 1.290.00 (US$)

From Pleurotus ostreatus

10.000 (concentrate) 150.00 (US$)

10.000 (purified) 400.00 (US$)

100.000 (concentrate) 650.00 (US$)

100.000 (purified) 1,600.00 (US$)

USBiological

(www.usbio.net/)

From heterologus expression of Trametes versicolor laccase in Saccharomyces cerevisiae 100 (purified) 169 (US$)

Sigma-Aldrich

From Rhus vernicfiera 10,000 72.30 (US$)

From Agaricus bisporus (≥1.5 U/mg) 1 g 30.50 (US$)

From Coriolus versicolor (≥1 U/mg) 5 g 120.90 (US$)

1 g 44.00 (US$)

10 g 358.20 (US$)

Jena BioScience

From Trametes versicolor, Coprinus cinereus and Pycnoporus cinnabarinus100 U 15.00 (EUR)

1000 U 75.00 (EUR)aThe methodology and expression of laccase activity (Units) are different among the companies.

fungal culture displayed much better properties than laccasealone in removing both the total phenolic compounds andcolour.

Olive mill wastewater (OMW) is a characteristic by-product of olive oil production and a major environmentalproblem in the Mediterranean area. Thus, 30 million m3 ofOMW is produced in the Mediterranean area [53] whichgenerate 2.5 litres of waste per litre of oil produced [54].OMW contains large concentrations of phenol compounds(up to 10 g/L) [54, 55], which are highly toxic [52, 56]. Also,it has high chemical and biochemical oxygen demands (CODand BOD, resp.) [57].

OMW is characterised by a colour variable from darkred to black depending on the age and type of oliveprocessed [58], low pH value (∼5), high salt content andhigh organic load with elevated concentrations of aromaticcompounds [59], fatty acids, pectins, sugar, tannins andphenolic compounds, in particular polyphenols [58]. Thepresence of a large number of compounds, many withpolluting, phytotoxic, and antimicrobial properties [60],renders OMW a waste with high harmful effects towardshumans and environment and makes its disposal one of themain environmental concerns in all producing countries.

Martirani et al. [61] reported that the treatment of anOMW effluent collected at an olive oil factory in Abruzzo(Italy) with a purified laccase from Pleurotus ostreatussignificantly decreased its phenolic content (up to 90%) butno reduction of its toxicity was observed when tested onBacillus cereus.

Gianfreda et al. [62] showed that laccase from Cerrenaunicolor was able to oxidise different phenolic substances

usually present in OMW with oxidation percentages rangingfrom 60 to 100% after 24 h of laccase incubation.

D’Annibale et al. [63] used a laccase from the white-rotfungus Lentinula edodes immobilised on chitosan to treatOMV from an olive oil mill located in Viterbo (Italy). Theyfound that the treatment of the OMW with immobilisedlaccase led to a partial decolouration as well as to significantabatements in its content in polyphenols, and orthodiphe-nols combined with a decreased toxicity of the effluent. Theyalso showed that an oxirane-immobilised laccase from L.edodes efficiently removed the OMW phenolics [64].

Casa et al. [65] investigated the potential of a laccasefrom L. edodes in removing OMW phytotoxicity. For this,they performed germinability experiments on durum wheat(Triticum durum) in the presence of different dilutions ofraw or laccase-treated OMW. The treatment with laccaseresulted in a 65% and an 86% reduction in total phenolsand orthodiphenols, respectively, due to their polymerisationas revealed by size-exclusion chromatography. In addition,germinability of durum wheat seeds was increased by 57% ata 1 : 8 dilution and by 94% at a 1 : 2 dilution, as compared tothe same dilutions using untreated OMW.

Attanasio et al. [66] studied the application of a non-isothermal bioreactor with laccases from T. versicolor immo-bilised on a nylon membrane to detoxify OMW and showedthat the technology of non-isothermal bioreactors was veryuseful in the treatment of OMW.

Jaouani et al. [67] studied the role of a purified laccasefrom Pycnoporus coccineus in the degradation of aromaticcompounds in OMW. They found that the treatment ofOMW with laccase showed similar results to those reported

6 Enzyme Research

with the fungus indicating that laccase plays an importantrole in the degradative process. Berrio et al. [68] studiedthe treatment of OMW with a laccase from P. coccineusimmobilised on Eupergit C 250L. Gel filtration profiles of theOMW treated with the immobilised enzyme (for 8 h at roomtemperature) showed both degradation and polymerisationof the phenolic compounds.

Quaratino et al. [69] reported that phenols were the maindeterminants for OMW phytotoxicity and showed that theuse of a commercial laccase preparation (DeniLite, NovoNordisk, Denmark) might be very promising for a saferagronomic use of the wastewater.

Iamarino et al. [70] studied the capability of a laccasefrom Rhus vernicifera to degrade and detoxify two OMWsamples of different complexity and composition.

Pant and Adholeya [71] used a concentrated enzymaticextract from solid-state fermentation (SSF) cultures ofdifferent fungi on wheat straw to decolourise a distilleryeffluent. They reported a maximum decolouration of 37% inthe undiluted distillery effluent using the extract of Pleurotusflorida EM1303 which was attributed to its high laccaseproduction.

3. Future Trends and Perspectives

This paper shows that laccase has a great potential applica-tion in several areas of food industry. However, one of thelimitations to the large-scale application of laccases is the lackof capacity to produce large volumes of highly active enzymeat an affordable cost (Table 2). The use of inexpensive sourcesfor laccase production is being explored in recent times. Inthis regard, an emerging field in management of industrialwastewater is exploiting its nutritive potential for productionof laccase enzymes. Besides solid wastes, wastewater from thefood processing industry is particularly promising for that.

Acknowledgment

This paper was financed by the Spanish Ministry of Scienceand Innovation (Project CTM2008-02453/TECNO).

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Enzyme Research 7

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[37] T. Takemori, Y. Ito, M. Ito, and M. Yoshama, “Flavor and tasteimprovement of cacao nib by enzymatic treatment,” JapanKokai Tokkyo Koho JP 04126037 A2, 1992.

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[45] F. Littoz and D. J. McClements, “Bio-mimetic approach toimproving emulsion stability: cross-linking adsorbed beet

pectin layers using laccase,” Food Hydrocolloids, vol. 22, no. 7,pp. 1203–1211, 2008.

[46] A. L. Ghindilis, V. P. Gavrilova, and A. I. Yaropolov, “Laccase-based biosensor for determination of polyphenols: determi-nation of catechols in tea,” Biosensors and Bioelectronics, vol. 7,no. 2, pp. 127–131, 1992.

[47] M. R. Montereali, L. Della Seta, W. Vastarella, and R.Pilloton, “A disposable Laccase-Tyrosinase based biosensor foramperometric detection of phenolic compounds in must andwine,” Journal of Molecular Catalysis B, vol. 64, no. 3-4, pp.189–194, 2010.

[48] M. Di Fusco, C. Tortolini, D. Deriu, and F. Mazzei, “Laccase-based biosensor for the determination of polyphenol index inwine,” Talanta, vol. 81, no. 1-2, pp. 235–240, 2010.

[49] E. N. Prasetyo, T. Kudanga, W. Steiner, M. Murkovic, G. S.Nyanhongo, and G. M. Guebitz, “Laccase-generated tetram-ethoxy azobismethylene quinone (TMAMQ) as a tool forantioxidant activity measurement,” Food Chemistry, vol. 118,no. 2, pp. 437–444, 2010.

[50] P. Ibarra-Escutia, J. Juarez Gomez, C. Calas-Blanchard, J. L.Marty, and M. T. Ramırez-Silva, “Amperometric biosensorbased on a high resolution photopolymer deposited onto ascreen-printed electrode for phenolic compounds monitoringin tea infusions,” Talanta, vol. 81, no. 4-5, pp. 1636–1642,2010.

[51] E. Valdez and M. C. Obaya, “Estudio del tratamiento anaer-obico de los residuales de la industria alcoholera,” RevistaICIDCA, vol. 19, no. 1, pp. 7–10, 1985.

[52] P. J. Strong and J. E. Burgess, “Treatment methods for wine-related and distillery wastewaters: a review,” BioremediationJournal, vol. 12, no. 2, pp. 70–87, 2008.

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8 Enzyme Research

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SAGE-Hindawi Access to ResearchEnzyme ResearchVolume 2010, Article ID 149748, 10 pagesdoi:10.4061/2010/149748

Review Article

Fungal Laccases: Production, Function, andApplications in Food Processing

Khushal Brijwani, Anne Rigdon, and Praveen V. Vadlani

Bioprocessing Laboratory, Department of Grain Science and Industry, Kansas State University, Manhattan, KS 66506, USA

Correspondence should be addressed to Praveen V. Vadlani, vadlani@ksu.edu

Received 1 July 2010; Accepted 22 August 2010

Academic Editor: Raffaele Porta

Copyright © 2010 Khushal Brijwani et al. This is an open access article distributed under the Creative Commons AttributionLicense, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properlycited.

Laccases are increasingly being used in food industry for production of cost-effective and healthy foods. To sustain this trendwidespread availability of laccase and efficient production systems have to be developed. The present paper delineate the recentdevelopments that have taken place in understanding the role of laccase action, efforts in overexpression of laccase in heterologoussystems, and various cultivation techniques that have been developed to efficiently produce laccase at the industrial scale. The roleof laccase in different food industries, particularly the recent developments in laccase application for food processing, is discussed.

1. Introduction

Laccase (benzenediol: oxygen oxidoreductase, EC 1.10.3.2) isa part of broad group of enzymes called polyphenol oxidasescontaining copper atoms in the catalytic center and areusually called multicopper oxidases. Laccases contain threetypes of copper atoms, one of which is responsible for theircharacteristic blue color. The enzymes lacking a blue copperatom are called yellow or white laccases. Typically laccase-mediated catalysis occurs with reduction of oxygen to wateraccompanied by the oxidation of substrate. Laccases are thusoxidases that oxidize polyphenols, methoxy-substituted phe-nols, aromatic diamines, and a range of other compounds[1].

Laccases are widely distributed in higher plants and fungi[2] and have also been found in insects and bacteria [3].Laccases are distributed in Ascomycetes, Deuteromycetes,and Basidiomycetes, being particularly abundant in manywhite rot fungi that are involved in lignin metabolism[4, 5]. Owing to the higher redox potential (+800 mV)of fungal laccases compared to plants or bacterial laccasesthey are implicated in several biotechnological applicationsespecially in the degradation of lignin [6]. For instance,redox potentials of laccases from common laccase pro-ducing fungi are reported as 790 mV (Trametes villosa),

450 mV (Myceliophthora thermophila), 750 mV (Pycnoporuscinnabarinus), and 780 mV (Botrytis cinerea) [7]. Laccases,therefore, possess excellent potential to be used as processingaids for the food industry.

The successful application of laccases in food processingwould require production of high amounts at reducedcosts. Several production strategies can be adopted alongwith media and process optimization to achieve betterprocess economics. Concomitantly, overexpression of laccasein suitable host organisms would provide means to achievehigh titers. Use of inducers could also enhance productioncapabilities [8].

The main objective of this paper is to summarize thewealth of information available in the literature with regardto laccase mechanism, production, and overexpression andeventually its role in food processing. Laccase has severalfood-based applications including bioremediation, beverage(fruit juice, wine and beer) stabilization, uses in bakingindustry, and role in improvement of overall food quality.The versatility of laccase in its action and its wide occurrencein several species of fungi contribute to the easy applicabilityin biotechnological processes. The present review, therefore,should help to shed light on the general characteristics oflaccase in an effort to create a database that could aid usageof laccase in the food processing industry.

2 Enzyme Research

2. Mechanism of Laccase Action

The catalysis of laccase occurs with reduction of onemolecule of oxygen to water accompanied with one electronoxidation of a wide range of aromatic compounds whichincludes polyphenols [9], methoxy-substituted monophe-nols, and aromatic amines [4]. This oxidation results ingeneration of oxygen-centered free radical that can beconverted to quinone in a second enzyme catalyzed reaction.Laccase catalysis occurs in three steps: (1) type I Cu reductionby substrate; (2) electron transfer from type I Cu to the typeII Cu and type III Cu trinuclear cluster; (3) reduction ofoxygen to water at the trinuclear cluster [10].

The laccase mediated catalysis can be extended to non-phenolic substrates by the inclusion of mediators. Mediatorsare a group of low molecular weight organic compounds thatcan be oxidized by laccase first forming highly active cationradicals capable of oxidizing nonphenolic compounds thatlaccase alone cannot oxidize (Figure 1). The most commonsynthetic mediators are 1-hydro-xybenzotriazole (HOBT),N-hydro-xyphthalimide (NHPI), and 2,2′-azinobis-3-ethyl-thiazoline-6-sulfonat (ABTS) [11].

3. Occurrence of Laccase in Fungal Systems

Laccase activity has been demonstrated in several fungalspecies leading to the notion that most of all fungi producelaccase. This, however, should not be generalized as thereare several physiological groups of fungi that apparentlydo not produce laccase. Laccase production has neverbeen demonstrated in lower fungi, that is, Zygomycetesand Chytridiomycetes [12]. Several reports can be referred,in the literature on production of laccase in ascomycetessuch as Gaeumannomyces graminis [13], Magnaporthe grisea[14], and Ophiostoma novo-ulmi [15], Mauginella [16],Melanocarpus albomyces [17], Monocillium indicum [18],Neurospora crassa [19], and Podospora anserina [20]. Inaddition to plant pathogenic species, laccase productionwas also reported for some soil ascomycete species from thegenera Aspergillus, Curvularia and Penicillium [21–23], andin some freshwater ascomycetes [24, 25].

Wood degrading ascomycetes like Trichoderma andBotryosphaeria have been shown to have some laccaseactivity. While Botryosphaeria produces constitutively adimethoxyphenol oxidizing enzyme that is probably truelaccase [26] there are only some strains of Trichodermathat exhibit low level production of a syringaldazine oxi-dizing enzyme [27]. In case of wood rotting xylariaceousascomycetes, two strains of Xylaria sp. and one of Xylariahypoxylon exhibited syringaldazine oxidation [28]. In com-plex liquid media, the fungi X. hypoxylon and Xylariapolymorpha produced appreciable titers of an ABTS oxidiz-ing enzyme [29]. Furthermore, ascomycete species closelyrelated to wood-degrading fungi which participate in thedecay of dead plant biomass in salt marshes have been shownto contain laccase genes and to oxidize syringaldazine [30].Basidiomycete yeast like Cryptococcus neoformans produces atrue laccase capable of oxidation of phenols and aminophe-nols and is unable to oxidize tyrosine [31]. The production of

laccase was not demonstrated in ascomycetous yeasts, but theplasma membrane bound multicopper oxidase Fet3p fromSaccharomyces cerevisiae shows both sequence and structuralhomology with fungal laccase [32, 33].

Wood rotting basidiomycetes causing white rot and arelated group of litter decomposing saprotrophic fungi arethe most widely known species that produce appreciablequantity of laccase. Almost all species of white rot fungiwere reported to produce laccase to varying degree [34].In case of Pycnoporus cinnabarinus laccase was described asthe only ligninolytic enzyme produced by this species thatwas capable of lignin degradation [35]. Brown-rot fungi onthe other hand are not known, in general, to carry laccaseproduction capabilities. A DNA sequence with relatively highsimilarity to that of laccase was detected in Gloeophyllumtrabeum that was capable of oxidizing ABTS [36]. Thoughno laccase protein has been purified from brown-rot species,the oxidation of syringaldazine has recently been detectedin the brown-rot fungus Coniophora puteana [37] and theoxidation of ABTS was reported in Laetiporus sulphureus[38].

4. Overexpression of Laccase

Due to the ability of fungal laccases to oxidize phenolicand nonphenolic aromatic compounds, increased interestin the application of these enzymes for various industrialapplications, including food, pulping, textile, wastewatertreatment, and bioremediation, is growing greatly [8]. Tosuccessfully utilize laccases in these applications, productionof large quantities at a low cost is essential.

To make laccases available for industrial applications,methods to reduce costs include fermentation media opti-mization, novel fermentation methods, and genetic mod-ification for large scale production via eukaryotic recom-binant strains. Much research has been done to identifyeffective methods for mass production of laccase usingthe above mentioned methods. Determination of optimumfermentation media can easily be achieved but cofactors andinducer compounds can cause an undesirable increase incost during growth at industrial scale. Novel fermentationmethods can also cause undesirable increases to cost due tomodifications to preexisting facilities. Genetic modificationpresents a promising method of overexpression of laccasefor large applications. However, fungal laccases requireposttranslational modifications (glycosylation), which onlyeukaryotic microorganisms are capable of carrying out cre-ating limitations for genetic manipulation for overexpressionof laccase. Laccase genes have been successfully cloned andheterologously expressed in the filamentous fungi Aspergillusniger, Aspergillus oryzae, and Trichoderma reesei [8]. Only afew bacterial laccases have been thoroughly studied to revealindustrial advantages over fungal laccases. Bacterial laccaseshave been found to be highly active and have higher stabilityat higher temperatures and pH values compared to fungallaccases [39]. Laccase-like enzymes isolated from bacterialcultures have been found to be very similar to fungal laccases;however, they vary in activity [39].

Enzyme Research 3

Phenolic substrate

Laccase

Laccase

Oxidized phenolic substrate

O2

O2

H2O

H2O

Non-phenolic substrate Oxidized Non-phenolic substrate

Oxidized mediator Mediator

Mediator

Figure 1: Mechanism of laccase action for both phenolic and nonphenolic substrates.

Research by [40] focused on optimizing media con-ditions using multiple micronutrients for maximum pro-duction of laccase by a previously identified fungal strainbelonging to the genus Gandoderma and referred to asWR-1. Strain WR-1, a white-rot fungus, was isolated fromtree bark using tissue culture techniques and was found toproduce high amounts of laccase during fermentation. WR-1 was naturally found to show laccase activity of 124 U/ml,compared to typical strains which show activity in a rangeof 4−100 U/ml. The experimental design for determiningthe optimum media conditions included the use of theorthogonal matrix method. This allowed for the statisticalevaluation of the relative importance of various nutrientsfor the highest production of laccase using submerged fer-mentation methods. It was concluded that WR-1 producedincreasing amounts of laccase when grown in a starch-basedmedium with the addition of copper sulphate and 2, 5-xylidine, as a laccase production inducer. WR-1 was able toincrease laccase production to 692 U/ml during fermentationin the optimized media, a significant increase compared toother strains under similar fermentation conditions [40].

To help increase laccase production, research has focusedon using recombinant fungal strains for maximum produc-tion. Research by [41] successfully transferred laccase genesfrom the basidiomycete Tramete hirsuta into the ascomycetePenicillium canescens for heterologous expression. The fungalstrain from the genus Penicillium was chosen due to its abilityto secrete large amounts of enzyme into culture media and ithas been demonstrated that synthesized enzymes are safe forhuman consumption. After successful transformation, it wasfound that 98% of the target enzyme activity was detectablein the liquid culture medium. It was also found that the

molecular weight of the recombinant enzyme matched thenative laccase produce by T. hirsuta [41]. Further research isstill needed to ensure high laccase production using large-scale fermentation methods.

Additionally, research by [53] focused on transforminglaccase genes from Trametes versicolor into the methyltrophicyeast Pichia pastoris for heterologous expression. The P.pastoris expression system is commonly used to achieve highexpression levels of heterologous proteins. This yeast hasbeen found to achieve high cell densities during growth ina minimal media in a short period of time. Furthermore,P. pastoris has been found to be an efficient secretionsystem and capable of posttranslational modifications (e.g.,glycosylation). After successful transformation of the P.pastoris expression system, it was found that utilizing a solid-state fermentation (SSF) method produced similar laccaseproduction results compared to submerged fermentation(SmF) methods [53].

Recently, a few bacterial laccases have been isolated fromEscherichia coli, Bacillus halodurans, Thermus thermophilus,and several species of Streptomycetes. Little is known aboutthe function of laccases in bacterial physiology but they arebelieved to play a role in melanin production, spore coatresistance, morphogenesis, and detoxification of copper [54].The bacterial laccase CotA isolated from Bacillus subtilis wasfound to be an endospore coat protein with high thermosta-bility [55]. Utilizing bacterial laccases for industrial pro-duction would allow for new biotechnological applicationsdue to the ease of genetic improvements to expression level,activity, and selectivity [56]. The combination of randomand site-directed mutagenesis was used to produce a doublemutant with improved functional expression in E. coli and

4 Enzyme Research

improved specific activity for different dyes [56]. Wu etal. [39] isolated a new strain of Aeromonas hydrophiliadesignated WL-11 from activated sludge in an effluenttreatment plant of a textile and dyeing industry. The geneencoding laccase was cloned from the newly isolated strainand successfully expressed in E. coli BL21(DE3). The recom-binant strain produced a high level of laccase compared tothe wild type. The recombinant laccase was characterizedand could be used as a biocatalyst in biotechnologicalapplications requiring large quantities of laccase [39].

5. Production Systems for Laccase

Laccases are extracellular enzymes secreted into the mediumby filamentous fungi [57]. Laccases are generally producedduring the secondary metabolism of different fungi. Severalfactors including type of cultivation (submerged or solidstate), carbon limitation, nitrogen source, and concentrationof microelements can influence laccase production [58].Subsequent sections delineate the role of different processparameters in laccase production.

6. Influence of Carbon and Nitrogen Source onLaccase Production

The excessive concentrations of glucose are inhibitory tolaccase production in various fungal strains [37]. An excessof sucrose also reduced the production of laccase by blockingits induction and only allowed constitutive production ofenzyme. Use of polymeric substrates like cellulose was ableto alleviate this problem [37]. Fungal laccases are oftentriggered by nitrogen depletion [59], but it was also foundthat in some strains nitrogen had no effect on enzyme activity[60]. High laccase activity was reported in some studies usinglow carbon to nitrogen ratio [61], but other studies showedthat higher laccase production was achieved at high carbon tonitrogen ratio [62]. Laccase was also produced earlier whenthe fungus was cultivated in nitrogen rich media rather thannitrogen-limited media [63].

7. Induction of Laccase

Production of laccase can be considerably enhanced byaddition of various supplements to the media [64]. The addi-tion of xenobiotic compounds such as xylidine, lignin, andveratryl alcohol increased and induced laccase activity [65].In one study by Lu et al. [66] it was observed that additionof cellobiose can induce appreciable laccase activity in somespecies of Trametes. Low concentration of copper was alsoshown to exhibit inducible effect on laccase activity [67].Various basidiomycetes, ascomycetes, and deuteromycetesgrown in sugar rich liquid medium were induced for laccaseproduction by the addition of 2,5-xylidine. It was explicitlydemonstrated that cultures of Fomes annosus, Pholiotamutabilis, Pleurotus ostreatus, and Trametes versicolor werestimulated for laccase production by addition of xylidine,and in the case of Podospora anserina rather decrease inactivity was observed by xylidine addition [68].

8. Influence of pH and Temperature onLaccase Production

The information on effect of pH and temperature effectson laccase production is scarce, but most reports indicateinitial pH between 4.5 and 6.0 that is suitable for enzymeproduction [6]. The optimum temperature for laccaseproduction is between 25◦C and 30◦C [69]. When fungi werecultivated at temperatures higher than 30◦C the activity ofenzyme was reduced [70].

9. Type of Cultivation

Laccases have been produced vividly in both submergedand solid state modes of fermentation. Table 1 lists thedifferent cultivation techniques that have been adopted forlarge-scale production of laccase using wild-type filamentousfungi. In forthcoming sections, important features of laccaseproduction into two modes, submerged and solid, state willbe discussed.

10. Submerged Fermentation

Submerged fermentation involves the cultivation of microor-ganisms in liquid medium containing appropriate nutrientswith high oxygen concentrations when operated in aerobicconditions. One of the major challenges in fungal submergedfermentations is viscosity of broth. Mycelium formationduring growth of fungal cells can also impede impeller actioncausing blockades resulting in oxygen and mass transferlimitations. Different strategies have been employed to dealwith oxygen and mass transfer limitations. A pulsed systemdeveloped by [71] to contain overcontrolled growth hasbeen employed in decoloration of synthetic dye by the whiterot fungus Trametes versicolor [72–75] allowing bioreactorto operate in continuous mode for prolonged times withhigh efficiency. Cell immobilization is another techniqueto alleviate problems associated with broth viscosity, andoxygen and mass transfer. Schliephake et al. [42] producedlaccase by Pycnoporus cinnabarinus immobilized on cubesof nylon sponge in a 10-L packed bed bioreactor operatedin a batch mode. Luke and Burton [44] reported thatthe immobilization of the fungus Neurospora crassa onmembrane supports allowed the continuous production oflaccase for the period of four months without enzymedeactivation. Sedarati et al. [76] compared the free cellcultures of T. versicolor with immobilized cultures usingnylon mesh for the bioremediation of pentachlorophenol(PCP) and 2,4-dichlorophenol (2,4 DCP). Authors observedthat immobilized cultures led to efficient removal. Coutoet al. [77, 78] investigated different synthetic materials ascarriers for the immobilization of the white rot fungusTrametes hirsuta in fixed bed bioreactors operated in batch.They found that among the different materials tested,stainless steel sponge led to the highest laccase activities.Park et al. [79] found that immobilization of the whiterot fungus Funalia trogii in Na-alginate beads allowed theefficient decolouration of dye Acid Black 52. Other factors

Enzyme Research 5

Table 1: Production of laccases in different cultivation modes.

Fungi Type of cultivation Inducer Laccase Activity (U/L) Reference

Pycnoporus cinnabarinus Submerged 10 mM Veratryl alcohol (VA) 280 [42]

Trametes pubescens Submerged 2 mM Cu2+ 333,000 [43]

Neurospora crassa Submerged 1 μM cyclohexamide 10,000 [44]

T. versicolor SSF (Immersion, nylon sponge) Tween 80 229 [45]

T. versicolor SSF (Immersion, barley bran) Tween 80 600 [45]

T. versicolor SSF (Expanded bed, nylon sponge) Tween 80 126 [45]

T. versicolor SSF (Expanded bed, barley bran Tween 80 600 [45]

T. versicolor SSF (Tray, nylon sponge) Tween 80 343 [45]

T. versicolor SSF (Tray, barley bran) Tween 80 3500 [45]

T. hirsuta SSF (Tray, grape seeds) — 18,715 [46]

affecting the laccase production is agitation. Hess et al.[80] found that laccase production by Trametes multicolordecreased considerably when the fungus was grown in stirredtank reactor, presumably because of damage to mycelia.Mohorcic et al. [81] found that it was possible to cultivatethe white rot fungus Bjerkandera adusta in a stirred tankreactor after its immobilization on a plastic net, althoughvery low activities were attained. Tavares et al. [82] oncontrary observed that agitation did not play an importantrole in laccase production by T. versicolor. Fed-batch mode ofoperation is also shown to be an effective way of producinglaccase. Galhaup et al. [43] found that operating in fed batchincreased the laccase production of T. pubescens by twofoldand obtained a higher laccase activity.

11. Solid State Fermentation

Solid state fermentation (SSF) is defined as fermentationprocess occurring in absence or near absence of free liquid,employing an inert substrate (synthetic materials) or anatural substrate (organic materials) as a solid support [83].SSF is shown to be particularly suitable for the productionof enzymes by filamentous fungi because they mimic theconditions under which the fungi grow naturally [83, 84].The use of natural solid substrates, especially lignocellulosicagricultural residues as growth substrates has been studiedfor various enzymes like cellulases [85, 86] including laccases[87]. The presence of lignin and cellulose/hemicellulose actas natural inducers and most of these residues are rich insugar promoting better fungal growth and thus making theprocess more economical [8]. The major disadvantage withSSF is lack of any established bioreactor designs. There areseveral bioreactor designs that exist in the literature that haveaddressed the major limitations of heat and mass transfer insolid media. Nevertheless lot of progress is still to be made.Different bioreactor configurations have been studied forlaccase production. Couto et al. [45] tested three bioreactorconfigurations immersion, expanded bed and tray for laccaseproduction by T. versicolor using, and inert (nylon) andnoninert support (barley bran). They found that the trayconfiguration led to the best laccase production. Couto etal. [46] also compared tray and immersion configurations

Table 2: Laccase producing organism and biotechnological appli-cation for use.

Laccase ProducingOrganism

Application Reference

Trametes versicolor Filtration aid [47]

Trametes versicolor Wine stabilization [48]

Myceliophthorathermophilia

Dough conditioner [49]

Rhizoctonia praticola Phenolic compound removal [50]

Trametes versicolor;Rhizoctonia praticola

Soil decontamination [50]

Coriolopsis gallica Beer factory waste water [51]

Trametes sp. Distillery waste water [51]

Trametes versicolor;Pleurotus ostreatus

Olive Mill wastewaters [51]

Trametes hirsuta Dough Conditioner [52]

for production of laccase by T. hirsuta using grape seedsas substrate. Tray configuration gave the best results hereas well, and in a similar study by Rosales et al. [88] trayconfiguration produced higher laccase activity in T. hirsutacultures raised on orange peels.

12. Applications of Laccase in Food Processing

Table 2 shows the multiple applications of laccases in thefood industry. Areas of the food industry that benefitfrom processing with laccase enzymes include baking, juiceprocessing, wine stabilization, and bioremediation of wastewater [52]. The use of laccase enzymes allows for theimprovement of functionality along with sensory properties.Laccase can also be utilized for analytical applications includ-ing biosensors, enzymatic, and immunochemical assays [51].

The baking industry utilizes a variety of enzymes toimprove bread texture, volume, flavor, and freshness alongwith improving machinability of dough during processing.Bt the addition of laccase to dough used for baked products,the enzyme exhibits an oxidizing effect resulting in improvedstrength of gluten structures in dough and baked products.It has also been found that the addition of laccase results in

6 Enzyme Research

increased volume, improved crumb structure, and softness ofbaked products. Machinability of dough was also found to beimproved due to increased strength and stability along withreduced stickiness with the addition of laccase. Improvedbread and dough qualities with the addition of laccase werealso seen when used with low quality flours [89].

Due to the growing awareness of celiac disease (CD),increased interest has focused on the development of gluten-free baked products. CD is an immune-mediated enteropa-thy triggered by the ingestion of gluten, contained in manycereal flours including wheat, rye, and barley, by geneticallysusceptible individuals. Cereal flours, like oats and starchessuch as rice, potato, and corn, have been the focus forthe development of gluten-free baked products [90]. Theseflours and starches lack the protein matrix responsiblefor dough formation and physical characteristics found inwheat-based baked products. Mimicking the protein matrixformed by the gluten proteins during dough formationof wheat flour has become exceedingly complex. Recentresearch has focused on using gluten-free oat flour alongwith enzymes to produce baked products acceptable for CDpatients. The addition of laccase and proteolytic to oat flourlead to a significant improvement to texture quality of oatbread, due to increased loaf specific volume and loweringcrumb hardness and chewiness. Chemical analysis of oatflour batter treated with laccase and proteolytic enzyme werefound to cause a β-glucan depolymerisation and proteinpolymerization, resulting in improved rheological propertiesand positively contribute to improved bread making perfor-mance by oat flour [49].

Laccase is also commonly used to stabilize fruit juices.Many fruit juices contain naturally occurring phenolics andtheir oxidation products, which contribute to color and taste.The natural polymerization and cooxidation reactions ofphenolics and polyphenols over time results in undesirablechanges in color and aroma. The color change, referred toas enzymatic darkening, increases due to a higher concen-tration of polyphenols naturally present in fruit juices [91].Research by Giovanelli and Ravasini [47] utilized laccase incombination with filtration in the stabilization of apple juice.Treatment with laccase caused the removal of phenols withhigh efficiency compared to other methods, like activatedcoals. The substrate-enzyme complex is then removed viamembrane filtration, a critical treatment process. Colorstability was found to be greatly increased after treatmentwith laccase and active filtration, although turbidity waspresent. The phenolic content of juices has been found tobe greatly reduced after treatment with laccase along withan increase in color stability [91]. Laccase treatment has alsobeen found to be more effective for color and flavor stabilitycompared to conventional treatments, such as the additionof ascorbic acid and sulphites [89].

The high concentration of phenolics and polyphenolsalso come into play during wine production, particularlythe crushing and pressing stages. The high concentration ofpolyphenols from stems, seeds, and skins contribute to colorand astringency and are dependent on grape variety andvinification conditions [48]. The complex sequence of eventsresulting in the oxidation of polyphenols occurs in musts and

wines causing flavor alterations and intensification of colorin red wines. This phenomenon is also known as maderiza-tion [92]. Multiple methods can be utilized to prevent made-irization and they include catalytic factors, block oxidizers orthe removal of polyphenols via proteinaceous, clarification,polyvinylpolypyrrolidone (PVPP) and high doses of sulfurdioxide [48]. However, research by Minussi et al. [48] foundthat treatment with laccase for the removal of polyphenolsshould be selective, as indiscriminate removal can result inundesirable organoleptic characteristics. Minussi et al. [48]further concluded that treatment of white wines with laccaseis feasible and could diminish processing costs and increasestorability of white wines over extended periods of time.

The use of laccase for stabilization is not limited to wine;the beer industry has potential to benefit from laccase treat-ment. Classic haze formation in beer is attributed proteinprecipitation stimulated by proanthocyanidins polyphenols,which are naturally present in small quantities [92]. Thiscomplex formed is commonly referred to as chill haze,which occurs upon cooling of the beer. The complex canbe redissolved by warming of the beer to room temperatureor above. However, after extended periods of time, proteinsulphydryl groups replace phenolic rings and lead to per-manent haze that does not redissolve at room temperature[89]. Traditionally, excess polyphenols are removed viaPVPP treatment, however, PVPP is difficult to handle andcreates problems in waste water treatment due to its lowbiodegradability. Laccase has been identified as easier tohandle and safer for the oxidation of polyphenols in wort[89]. The addition of laccase at the end of processing hasthe added benefit of the removal of polyphenols and excessoxygen present; reduced oxygen content results in a longershelf life of beer [92].

Since laccase are capable of degrading phenolic com-pounds, utilization for bioremediation of food industrywastewaters is vital. Bioremediation includes processes andactions used to biotransform an environment altered bycontaminants back to its original status [93]. Many countriesheavily regulate pollutants, including the class of aromaticcompounds, which includes phenols and amines [50].Research by Minussi et al. [48] reported the removal ofnaturally occurring and xenobiotic aromatic compoundsfrom aqueous suspensions using immobilized laccase onorganogel supports. The application laccase for bioremedi-ation of wastewater streams is particularly of interest to beerfactories. Fractions of wastewater released from beer factoriescontain a large amount of polyphenols and are dark brownin color. Research by Yague et al. [94] found laccase producedby the white rot fungus Coriolopsis gallica was capable ofdegrading polyphenols present in wastewater. Other researchby Gonzalez et al. [95] utilized laccase from Trametes sp.for the bioremediation of distillery wastewater generatedfrom the ethanol production from the fermentation ofsugarcane molasses with a high content of organic matter andan intense dark-brown color. Bioremediation of olive millwastewaters via immobilized laccase has also been reported.It has also been found that utilizing olive oil mill wastewatershas been beneficial in the cultivation of fungi for laccaseproduction [89].

Enzyme Research 7

13. Conclusions

Laccases are versatile oxidases, and their versatility lies inthe high reduction potential that makes them potentialcandidate for biotechnological applications, especially forthe food industry. Laccases have the potential to make foodprocessing more economical and environmental friendly. Toproficiently realize this potential it would require more effi-cient laccase production systems and better understanding oftheir mode of action. With the use of mediators it is possibleto extend the role of laccase to nonphenolic substrates.Extensive occurrence of laccase in various fungal generaensures their widespread availability, and especially the woodrotting basidiomycetes also referred as white rot fungi arethe excellent laccase producers. Overexpression of laccases inheterologous systems has been actively pursued to enhancetheir titers and to improve their catalytic activity. Mediaoptimization and use of appropriate inducers could bringadditional benefits of higher production with expenditureof minimum resources. Both submerged and solid statecultivation techniques have been embraced by the researchersfor laccase production. Submerged fermentation, though,leads the SSF for industrial production of laccase. Futureefforts in improving the SSF bioreactor designs can make SSFmore potent and competitive. With plethora of applicationsin food processing including baking and role in gluten-freebreads, beverage (wine, juice and beer) stabilization, andbioremediation, laccases certainly have important role toplay in green food processing.

Acknowledgments

The authors gratefully acknowledge financial support fromdepartment of Grain Science and Industry, Kansas StateUniversity. This paper is contribution no. 10-389-J from theKansas Agricultural Experiment Station, Manhattan, KS.

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[65] A. M. R. B. Xavier, D. V. Evtuguin, R. M. P. Ferreira, and F.L. Amado, “Laccase production for lignin oxidase activity,” inProceedings of 8th International Conference on Biotechnology,Helsinki, Finland, 2001.

[66] S. X. F. Lu, C. L. Jones, and G. T. Lonergan, “Correlationbetween fungal morphology and laccase expression under theinfluence of cellobiose,” in Proceedings of the 10th InternationalBiotechnology Symposium, Sydney, Australia, 1996.

[67] G. Palmieri, P. Giardina, C. Bianco, A. Scaloni, A. Capasso, andG. Sannia, “A novel white laccase from Pleurotus ostreatus,”Journal of Biological Chemistry, vol. 272, no. 50, pp. 31301–31307, 1997.

[68] J.-M. Bollag and A. Leonowicz, “Comparative studies ofextracellular fungal laccases,” Applied and EnvironmentalMicrobiology, vol. 48, no. 4, pp. 849–854, 1984.

[69] S. B. Pointing, E. B. G. Jones, and L. L. P. Vrijmoed, “Opti-mization of laccase production by Pycnoporus sanguineus insubmerged liquid culture,” Mycologia, vol. 92, no. 1, pp. 139–144, 2000.

[70] F. Zadrazil, A. Gonser, and E. Lang, “Influence of incuba-tion temperature on the secretion of extracellular lignolyticenzymes of Pleurotus and Dichomitus squalus into soil,” inProceedings of the Conference on Enzymes in the Environment,Granada, Spain, 1999.

[71] J. M. Lema, E. Roca, A. Sanroman, M. J. Nunez, M. T.Moreira, and G. Feijoo, “Pulsating bioreactors,” in MultiphaseBioreactor Design, J. M. S. Cabral, M. Mota, and J. Tramper,Eds., pp. 309–329, Taylor and Francis, London, UK, 2001.

[72] P. Blanquez, N. Casas, X. Font et al., “Mechanism of textilemetal dye biotransformation by Trametes versicolor,” WaterResearch, vol. 38, no. 8, pp. 2166–2172, 2004.

[73] P. Blanquez, M. Sarra, and M. T. Vicent, “Study of the cellularretention time and the partial biomass renovation in a fungaldecolourisation continuous process,” Water Research, vol. 40,no. 8, pp. 1650–1656, 2006.

[74] S. Romero, P. Blanquez, G. Caminal et al., “Differentapproaches to improving the textile dye degradation capacityof Trametes versicolor,” Biochemical Engineering Journal, vol.31, no. 1, pp. 42–47, 2006.

[75] P. Blanquez, G. Caminal, M. Sarra, and T. Vicent, “The effectof HRT on the decolourisation of the Grey Lanaset G textiledye by Trametes versicolor,” Chemical Engineering Journal, vol.126, no. 2-3, pp. 163–169, 2007.

[76] M. R. Sedarati, T. Keshavarz, A. A. Leontievsky, and C. S.Evans, “Transformation of high concentrations of chlorophe-nols by the white-rot basidiomycete Trametes versicolorimmobilized on nylon mesh,” Electronic Journal of Biotechnol-ogy, vol. 6, no. 2, pp. 27–37, 2003.

[77] S. R. Couto, M. A. Sanroman, D. Hofer, and G. M. Gubitz,“Production of laccase by Trametes hirsuta grown in animmersion bioreactor and its application in the decolorizationof dyes from a leather factory,” Engineering in Life Sciences, vol.4, no. 3, pp. 233–238, 2004.

[78] S. R. Couto, M. A. Sanroman, D. Hofer, and G. M. Gubitz,“Stainless steel sponge: a novel carrier for the immobilisationof the white-rot fungus Trametes hirsuta for decolourizationof textile dyes,” Bioresource Technology, vol. 95, no. 1, pp. 67–72, 2004.

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[83] A. Pandey, P. Selvakumar, C. R. Soccol, and P. Nigam, “Solidstate fermentation for the production of industrial enzymes,”Current Science, vol. 77, no. 1, pp. 149–162, 1999.

[84] M. Moo-Young, A. R. Moreira, and R. P. Tengerdy, “Principlesof solid sate fermentation,” in The Filamentous Fungi, J. E.Smith, D. R. Berry, and B. Kristiansen, Eds., pp. 117–144,Edward Arnold, London, UK, 1983.

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SAGE-Hindawi Access to ResearchEnzyme ResearchVolume 2010, Article ID 473137, 16 pagesdoi:10.4061/2010/473137

Review Article

Potential Applications of Immobilized β-Galactosidase inFood Processing Industries

Parmjit S. Panesar, Shweta Kumari, and Reeba Panesar

Biotechnology Research Laboratory, Department of Food Engineering & Technology, Sant Longowal Institute ofEngineering and Technology, Longowal, Punjab, 148 106, India

Correspondence should be addressed to Parmjit S. Panesar, pspanesarrr@yahoo.com

Received 16 June 2010; Revised 22 September 2010; Accepted 21 November 2010

Academic Editor: Cristina M. Rosell

Copyright © 2010 Parmjit S. Panesar et al. This is an open access article distributed under the Creative Commons AttributionLicense, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properlycited.

The enzyme β-galactosidase can be obtained from a wide variety of sources such as microorganisms, plants, and animals. The useof β-galactosidase for the hydrolysis of lactose in milk and whey is one of the promising enzymatic applications in food and dairyprocessing industries. The enzyme can be used in either soluble or immobilized forms but the soluble enzyme can be used onlyfor batch processes and the immobilized form has the advantage of being used in batch wise as well as in continuous operation.Immobilization has been found to be convenient method to make enzyme thermostable and to prevent the loss of enzyme activity.This review has been focused on the different types of techniques used for the immobilization of β-galactosidase and its potentialapplications in food industry.

1. Introduction

The enzyme β-galactosidase (EC.3.2.1.23), most commonlyknown as lactase, which hydrolyses lactose into its monomersthat is glucose and galactose has potential applications infood processing industry. Because of low levels of the enzymein intestine, large fraction of the population shows lactoseintolerance and they have difficulty in consuming milk anddairy products. Lactose has a low relative sweetness andsolubility, and excessive lactose in large intestine can leadto tissue dehydration due to osmotic effects, poor calciumabsorption due to low acidity, and fermentation of the lactoseby microflora resulting in fermentative diarrhea, bloating,flatulence, blanching and cramps, and watery diarrhea [1].Furthermore, lactose is a hygroscopic sugar and has a strongtendency to absorb flavours and odours and causes manydefects in refrigerated foods such as crystallization in dairyfoods, development of sandy or gritty texture, and depositformation [2].

Technologically, lactose gets easily crystallized, which setsthe limits of its applications to certain processes in thedairy industry. Cheese manufactured from hydrolyzed milkripens more quickly than that made from normal milk.

Treatment of milk and milk products with lactase to reducetheir lactose content seems to be an appropriate method toincrease their potential uses and to deal with the problems oflactose insolubility and lack of sweetness. Furthermore, thistreatment could make milk, a most suitable food, availableto a large number of adults and children that are lactoseintolerant. Moreover, the hydrolysis of whey converts lactoseinto a very useful product like sweet syrup, which can beused in various processes of dairy, confectionary, baking,and soft drink industries [3, 4]. Therefore, lactose hydrolysisnot only allows the milk consumption by lactose intolerantpopulation but can also solve the environmental problemslinked with whey disposal [5–7].

The enzyme β-galactosidase can also be used in transg-lycosylation of lactose to synthesize galacto-oligosaccharides(GOSs). These were widely recognized as the nondigestibleoligosaccharides, not hydrolyzed or absorbed in the upperintestinal tract, and they pass onto the colon where they arefermented selectively by beneficial intestinal bacteria. Besidestheir prebiotic effects, these GOSs have low cariogenicity, lowcaloric values, and low sweetness [8, 9]. GOSs occur naturallyin trace amounts in breast milk, cow milk, honey, and avariety of fruits and vegetables [10]. As a result, increased

2 Enzyme Research

production of GOS is useful. GOS can be readily manufac-tured by enzymatic transgalactosylation of β-galactosidasefrom whey lactose, which is available in abundance as a by-product of cheese industry.

Thus, the application of β-galactosidase in the hydrolysisof lactose in dairy industry has attracted the attention ofresearchers. Although most industries still hydrolyze lactosewith free enzyme, the immobilization of β-galactosidase isan area of great interest because of its potential benefits[11]. The use of immobilization technology is of signif-icant importance from economic point of view since itmakes reutilization of the enzyme and continuous operationpossible and also precludes the need to separate the cellsfrom the whey following processing. It can also help toimprove the enzyme stability. Nowadays, immobilized β-galactosidase is intensively being used in lactose hydrolysis ofmilk/whey and has been tested for the production of galacto-oligosaccharides.

2. Microbial Sources of Enzyme

The enzyme β-galactosidase can be obtained from a widevariety of sources such as microorganisms, plants, andanimals; however, according to their sources, their propertiesdiffer markedly [11, 12]. Enzymes of plants and animalorigin have little commercial value but several microbialsources of β-galactosidase are of great technological interest.Microorganisms offer various advantages over other avail-able sources such as easy handling, higher multiplicationrate, and high production yield. As a result of commercialinterest in β-galactosidase, a large number of microorgan-isms [13–26] have been assessed as potential sources of thisenzyme (Table 1).

2.1. Production and Purification. Microorganisms are con-sidered potential source of β-galactosidase for industrialapplications. However, they differ in their optimum con-ditions for the enzyme application especially pH range.A recovery cost of the enzyme depends on the level ofproduction and purification. Therefore, there has beenincreasing interest in finding microorganisms with adequateproperties for industrial use, higher production capacity, andless expensive purification methods of this enzyme. A widevariety of bacterial, yeast, and fungal cultures have beenreported for β-galactosidase production.

2.1.1. Bacterial Enzymes. The enzyme β-galactosidase can beproduced by a large number of bacteria but Streptococcusthermophilus and Bacillus stearothermophilus are consideredas potential bacterial sources. The enzyme from Escherichiacoli serves as a model for understanding the catalyticmechanism of β-galactosidase action, but it is not consideredsuitable for use in foods due to toxicity problems associatedwith the host coliform [11]. Hence, the β-galactosidase fromE. coli is generally not preferred for use in food industry [13–15].

β-galactosidase has been isolated from an extremelythermophilic Gram-negative anaerobe. Thermoanaerobac-

ter has been purified by chromatography through DEAE-cellulose [27]. The optimization of the ultrasonicationmethods for the maximum cell disruption of Escherichiacoli for the release of β-galactosidase has also been reported[28]. Lactobacillus delbrueckii subsp. bulgaricus cultures weresubjected to treatments using sonication, a high-speed beadmill, and a high-pressure homogenizer for the release of β-galactosidase [29].

β-galactosidase has also been purified from psychotropicPseudoalteromonas sp. isolated from Antarctica and a highyield of purification has been reported by a rapid purificationscheme using extraction in an aqueous two-phase systemfollowed by hydrophobic interaction chromatography andultrafilteration techniques [30].

An intracellular β-galactosidase from a thermoaci-dophilic Alicyclobacillus acidocaldarius subsp. rittmannii hasbeen purified using precipitation (with ammonium sul-phate), gel permeation, ion-exchange, and affinity chro-matography and preparative electrophoresis [26]. Further,a thermostable β-galactosidase gene bgaB from Bacillusstearothermophilus was cloned and expressed in B. subtilisWB600 and recombinant enzyme has been purified by acombination of heat treatment, ammonium sulfate frac-tionation, ion exchange, and gel filtration chromatographytechniques [31]. The intracellular β-galactosidase from ther-mophile B1.2 was purified by ion-exchange and affinitychromatography with a fold purification of 2.2 and 3.9,respectively. The molecular mass of the purified enzyme asdetermined by native PAGE was approximately 215 kDa, bySDS-PAGE was 75 kDa, and by gel filtration was 215 kDa[32]. The efficiency of different cell disruption methodson the extraction of intracellular β-galactosidase enzymefrom Streptococcus thermophilus and Lactobacillus delbrueckiisubsp. thermophilus has been tested [33]. Lysozyme enzymetreatment was determined as the most effective method,which resulted in approximately 1.5 and 10 times higherenzyme activity than glass bead and homogenization treat-ment, respectively.

The activity and the stability of the partially purified β-galactosidases from Thermus sp strain T2 and K. fragilis havebeen compared [34]. Both enzymes showed a remarkablehydrolytic activity and a weak transgalactosilation activity,even in the presence of high concentrations of lactose.The thermophilic enzyme showed a higher resistance tohydrophobic agents and a higher stability at different temper-atures, pHs, and chemical conditions. However, the enzymeof Thermus was less stable in the presence of oxygen peroxide,showing that some residues important for its stability wereaffected by oxidation. The enzyme from K. fragilis wasstrongly inhibited by o-nitrophenol in a acompetitive waybut it was weakly and competitively inhibited by galactose.The thermophilic enzyme was competitively inhibited bygalactose much strongly than its mesophilic counterpart butthe inhibition did not change with the temperature. A novelthermostable chimeric β-galactosidase was constructed byfusing a poly-His tag to the N-terminal region of the β-galactosidase from Thermus sp. strain T2 to facilitate itsoverexpression in E. coli and its purification by immobi-lized metal-ion affinity chromatography [35]. To improve

Enzyme Research 3

Table 1: Microbial sources of β-galactosidase.

Source Microorganism (s)

Bacteria

Alicyclobacillus acidocaldarius subsp. rittmannii

Arthrobacter sp.

Bacillus acidocaldarius, B. circulans, B. coagulans, B.subtilis, B. megaterum, B. stearothermophilus

Bacteriodes polypragmatus

Bifidobacterium bifidum, B. infantis

Clostridium acetobutylicum, C. thermosulfurogens

Corynebacterium murisepticum

Enterobacter agglomerans, E. cloaceae

Escherichia coli

Klebsiella pneumoniae

Lactobacillus acidophilus, L. bulgaricus,L. helviticus, L.kefiranofaciens, L. lactis, L. sporogenes, L. themophilus, L.delbrueckii

Leuconostoc citrovorum

Pediococcus acidilacti, P. pento

Propioionibacterium shermanii

Pseudomonas fluorescens

Pseudoalteromonas haloplanktis

Streptococcus cremoris, S. lactis, S. thermophius

Sulfolobus solfatarius

Thermoanaerobacter sp.

Thermus rubus, T. aquaticus

Trichoderma reesei

Vibrio cholera

Xanthomonas campestris

Fungi

Alternaria alternate, A. palmi

Aspergillus foelidis, A. fonsecaeus, A. fonsecaeus, A.Carbonarius, A. Oryzae

Auerobasidium pullulans

Curvularia inaequalis

Fusarium monilliforme, F. oxysporum

Mucor meihei, M. pusillus

Neurospora crassa

Penicillum canescens, P. chrysogenum, P. expansum

Saccharopolyspora rectivergula

Scopulariapsis sp

Streptomyces violaceus

Yeast

Bullera singularis

Candida pseudotropicalis

Saccharomyces anamensis, S. lactis, S. fragilis

Kluyveromyces bulgaricus, K. fragilis, K. lactis, K.marxianus

Source: [12–26].

the enzyme purification a selective one-point adsorptionwas achieved by designing tailor-made low-activated Co-iminodiacetic acid (Co-IDA) or Ni-IDA supports. The newenzyme was not only useful for industrial purposes but alsohas become an excellent model to study the purification

of large multimeric proteins via selective adsorption ontailor-made immobilized metal-ion affinity chromatographysupports. Furthermore, β-galactosidase from Thermus sp.Strain T2 has been purified and immobilized in a singlestep, combining the excellent properties of epoxy groupsfor enzyme immobilization with the good performanceof immobilized metal-chelate affinity chromatography forprotein purification [36].

2.1.2. Fungal Enzymes. The optimum pH range for thefungal enzyme is 2.5–5.4, which makes them suitable forprocessing of acid whey and its ultrafiltration permeate.The optimum temperatures for these enzymes are high andcan be typically used at temperatures up to 50◦C. Thepurification of β-galactosidase from different fungal sourceshas been carried using a variety of purification techniques.

β-galactosidase from Aspergillus niger has been purifiedand resolved into three multiple forms, using molecular siev-ing, ion exchange, and hydrophobic chromatography [37].The purification of β-galactosidase has been carried from acellular extract of Fusarium oxysporum var. lini by heat shockand successive chromatography on DEAE-cellulose DE-52and sephadex G-100 [18]. The purification of β-galactosidaseby ammonium sulphate precipitation and CM-sephadexchromatography from cell-free extracts of fungus Beauveriabassiana has also been reported [38]. An extracellular β-galactosidase from a themophilic fungus Rhizomucor sp. hasbeen purified by successive DEAE-cellulose chromatography,followed by gel filtration on sephacryl S-300 [39]. Theprecipitation with ammonium sulfate, ion-exchange chro-matography on DEAE-sephadex, affinity chromatographyand chromatofocusing has also been used for purification ofβ-galactosidase from Penicillium chrysogenum NCAIM 00237[40].

β-galactosidase produced by submerged culture ofAspergillus japonicus showed 2.95 U mg−1 protein specificactivities with an approximate molecular weight of 27 kDa[41]. The enzyme was purified 6.43-fold with 24.02%yield and a specific activity of 18.96 U mg−1 protein. Anintracellular β-glycoside hydrolase with β-glucosidase andβ-galactosidase activity designated β-glucosidase BGL1 hasbeen purified from the thermophilic fungus Talaromycesthermophilus. The monomeric enzyme has a molecular massof 50 kDa (SDS-PAGE), an isoelectric point of 4.5-4.6. β-galactosidase activity of β-glucosidase BGL1 is activated byvarious mono and divalent cations including Na+, K+, andMg2+, and it is moderately inhibited by its reaction productsthat is glucose and galactose [42].

2.1.3. Yeast Enzymes. Yeast has been considered as an impor-tant source of β-galactosidase from industrial point of view.With neutral pH optima, these are well suited for hydrolysisof lactose in milk and are widely accepted as safe for use infoods. Much work has been carried on the production ofβ-galactosidase from different yeast strains for its potentialuse. The most commercially available yeast β-galactosidaseunder the trade name of Maxilact (DSM Food Specialties,Delft, The Netherlands) and Lactase (SNAM Progetti, Italy)

4 Enzyme Research

is preparations extracted from K. lactis and Lactozym (Novo,Nordisk A/S, Bagsvaerd, Denmark) from K. fragilis [11, 14,17].

Biermann and Glantz [43] first attempted the purifi-cation of β-galactosidase from Sacharomyces lactis by gelfiltration on sephadex G-100, followed by ion exchange chro-matography on DEAE-sephadex. The homogenizer (800 barpressure, pH 7.5) was used for the disruption and extractionof β-galactosidase of K. marxianus [44]. Further, the partialpurification of the enzyme from K. marxianus was carriedusing DEAE-sepharose column. The studies on the use of fedbatch culture techniques to achieve high culture productivityin K. fragilis have also been carried out [45].

The optimization of β-galactosidase production by K.lactis using deprotienized whey as fermentation mediumhas been reported. The optimized condition for theenzyme production was reported as follows: temperature30.3◦C, pH 4.68, agitation speed 191 rpm, and fermentationtime 18.5 hours [46]. The studies on two commercial β-galactosidase (Lactozym and Maxilact) preparations indi-cated that the enzyme activities of both enzyme preparationspresent similar behaviour with different pH and temperatureand similar kinetic parameter values, which suggest thatboth enzymes are probably the same [47]. Response surfacemethodology has also been applied in the production of β-galactosidase using K. lactis NRRL Y-8279 [48] and maxi-mum specific enzyme activity of 4218.4 U g−1was obtainedat the optimum levels of process variables (pH 7.35, agitationspeed 179.2 rpm, initial sugar concentration 24.9 g L−1, andincubation time 50.9 hours).

3. Immobilization of β-Galactosidase

Although the enzyme β-galactosidase has numerous appli-cations in the food and dairy industries, but the moderatestability of enzyme is one of the limitations that hindergeneral implementation of biocatalysts at industrial scale.Thus, there is a need to explore their full potential as catalystby adopting suitable strategies for enzyme stabilization.The multimeric enzyme can be stabilized by using properexperimental conditions and genetic tools to cross linkor to strengthen the subunit-subunit interaction [49]. Thestability of monomeric or multimeric enzymes can also beenhanced by multipoint and multi-subunit covalent immo-bilization and enzyme engineering via immobilization [50].The enzyme has been immobilized by various methods suchas physical absorption, entrapment, and covalent bindingmethod [51–85] on different supports (Table 2).

3.1. Physical Adsorption. Physical adsorption is consideredas the simplest method of immobilization in which anenzyme is immobilized onto a water-insoluble carrier andthe biocatalysts are held on the surface of the carri-ers by physical forces (van der waals forces). Frequently,however, additional forces are involved in the interactionbetween carrier and biocatalyst principally hydrophobicinteractions, hydrogen bridges, and heteropolar (ionic)bonds [86]. This method has the advantage of being

simple to carry out and has little influence on the con-formation of the biocatalyst. However, the disadvantage ofthis technique is the relative weakness of the adsorptivebinding forces. Different inorganic (alumina, silica, porousglass, ceramics, diatomaceous earth, clay, bentonite, etc.)and organic (cellulose, starch, activated carbon and ion-exchange resins, such as Amberlite, Sephadex, Dowex) sup-port materials can be used for enzyme adsorption. Furtheradsorption of enzyme may be stabilized by glutaraldehydetreatment.

Immobilization of β-galactosidase on hydrophobic cot-ton cloth indicated that the enzyme adsorbed on the clothwas about 50% active as free enzymes [87]. The immobiliza-tion of β-galactosidase active yeast K. fragilis and K. lactisonto chitosan showed an enzyme activity of 0.9–2.2 U/mgdry cell wt [88]. Enzyme activity of immobilized enzymefrom K. fragilis was higher but the operational stabilityof A. oryzae enzyme was 5–14 times higher dependingupon the mode of immobilization [89]. When adsorptionmethod was used, the highest activity was obtained withyeast enzyme and support Ostsorb-DEAE. The enzyme fromA. oryzae immobilized on polyvinyl chloride (PVC) andsilica gel membrane has been used for the hydrolysis oflactose in skim milk in an axial-annular flow reactor [51].Further, maximum immobilization occurred at pH 5.5 andoptimal results were obtained with citrate/phosphate bufferduring immobilization of β-galactosidase from E. coli byphysical adsorption on chromosorb-W [51]. A novel reactorconsisting of β-galactosidase from B. circulans immobilizedon a ribbed membrane composed of PVC and silica hasalso been used for skim milk lactose hydrolysis [53]. Theimmobilization of partially purified Bullera singularis β-galactosidase in Chitopearl BCW 3510 bead (970 GU/gresin) by simple adsorption has also been carried out[90].

The studies on the kinetic behaviour of β-galactosidasefrom Kluyveromces marxianus (Saccharomyces) lactis, immo-bilized on to different oxide supports, such as alumina,silica, and silicated alumina indicated that the immobilizedenzyme activity strongly depends on the chemical natureand physical structure of the support [53]. In particular,when the particle sizes of the support are increased, theenzymatic activity strongly decreases. Immobilization of β-galactosidase from Thermus sp. preceded very rapidly ontoPEI-Sepabeads and conventional DEAE-Agarose. However,the adsorption strength was much higher in the case of PEI-Sepabeads [53].

A recombinant thermostable B. stearothermophilus β-galactosidase was immobilized onto chitosan using Tris(hydroxymethyl) phosphine (THP) and glutaraldehyde, anda packed bed reactor was utilized to hydrolyze lactose inmilk. The thermostability and enzyme activity of THP-immobilized β-galactosidase during storage was superior tothat of free and glutaraldehyde-immobilized enzymes. TheTHP-immobilized β-galactosidase showed greater relativeactivity in the presence of Ca2+ than the free enzyme andwas stable during the storage at 4◦C for 6 weeks, whereasthe free enzyme lost 31% of the initial activity under thesame storage conditions [91]. Response surface methodology

Enzyme Research 5

Table 2: Different sources of β-galactosidase and methods of immobilization.

Immobilization method Source of β-galactosidase Immobilizing agents References

(1) Physical adsorption

K. fragilis and K. lactis Chitosan [6]

A. oryzae Phenol-formaldehyde resin [69]

A. oryzae Polyvinyl chloride and Silica gel membrane [51]

E. coli Chromosorb-W [52]

B. circulans Polyvinyl chloride and Silica [53]

B. stearothermophilus Chitosan [54]

A. niger Porous ceramic monolith [70]

K. fragilis Chitosan bead [2]

K. fragilis Chitosan [71]

K. lactis CPC-silica and agarose [72]

Thermus sp. T2 PEI- sepabeads, DEAE-agarose [55]

K. fragilis Cellulose beads [14]

A. oryzae Celite and chitosan [73]

Pisum sativum Sephadex G-75 and chitosan beads [56]

(2) Entrapment

K. bulgaricus Alginate using BaCl3 [57]

E. coli Polyacrylamide gel [58]

A. oryzae Nylon-6 and zeolite [62]

Thermus aquaticus YT-1 Agarose bead [59]

A. oryza Spongy polyvinyl alcohol Cryogel [60]

Penicillium expansum F3 Calcium alginate [23]

K. lactis, A. oryzae Saccharomyces cerevisiae Poly(vinylalcohol) hydrogel [7]

(3) Covalent Binding

L. bulgaricus Egg shells [61]

S. anamensis Calcium alginate [74]

E. coli Hen egg white [75]

E. coli Polyvinyl alcohol [76]

A. oryzae Silica gel activated with TiCl3 and FeCl3 [77]

E. coli (Recombinant β-galactosidase) Cyanuric chloride-activated cellulose [66]

K. lactis Corn grits [78]

E. coli Gelatin [63]

K. lactis Thiosulfinate/thiosulfonate [79]

B. circulans Eupergit C (Spherical acrylic polymer) [65]

K. fragilis Silica-alumina [64]

K. lactis Graphite surface [68]

A. oryzae Chitosan bead and nylon membrane [80]

A. oryzae Cotton cloth and activated [81]

With tosyl chloride

A. oryzae Amino-epoxy sepabead [67]

K. latis Cotton fabric [82]

A. niger Magnetic polysiloxane-polyvinyl alcohol [83]

A. oryzae Silica [84]

A. oryzaePolyvinylalcoheol hydrogel and magneticFe3O4-chitosan as supporting agent

[85]

(RSM) and centre composite design (CCD) have beenused to optimize immobilization of β-galactosidase (BGAL)from Pisum sativum onto two matrices: Sephadex G-75 andchitosan beads. The immobilization efficiency of 75.66%and 75.19% was achieved with Sephadex G-75 and chitosan,respectively [56].

A. oryzae β-galactosidase was immobilized on aninexpensive bioaffinity support, concanavalin A-cellulose.Concanavalin A-cellulose adsorbed and cross-linked β-galactosidase preparation retained 78% of the initial activity.The optimum temperature was increased from 50 to 60◦C forthe immobilized β-galactosidase. The cross-linked adsorbed

6 Enzyme Research

enzyme retained 93% activity after 1-month storage whilethe native enzyme showed only 63% activity under similarincubation conditions [92].

3.2. Entrapment Method. Entrapment method is the phys-ical enclosure of enzymes in a small space. Matrix andmembrane entrapment (including microcapsulation) are themajor methods of entrapment. The major advantage of theentrapment technique is the simplicity by which sphericalparticles can be obtained by dripping a polymer-cell suspen-sion into a medium containing positively charged ions orthrough thermal polymerization [86]. Further, beads formedparticularly from alginate are transparent and generallymechanically stable. The major limitation of this techniquefor the immobilization of enzymes is the possible slowleakage during continuous use in view of the small molecularsize compared to the cells. However, improvements can bemade by using suitable linking procedures. The matrices usedfor the immobilization are usually made up of polymericmaterials such as Ca-alginate, agar, k-carragenin, polyacry-lamide, and collagen. However, some solid matrices such asactivated carbon, porous ceramic, and diatomaceous earthcan also be used for the immobilization. The membranescommonly used for the entrapment of enzymes are nylon,cellulose, polysulfone, and polyacrylmide.

Fungal β-galactosidase immobilized in polyvinyl alcoholgel was more thermostable than free enzyme and retained70% of activity after 24 h at 50◦C and 5% activity at 60◦C[93]. The glutaraldehyde-treated K. bulgaricus cells having β-galactosidase were entrapped in alginate using BaCl2 solution[57]. The alginate beads obtained after treatment withpolyethyleneimine followed by glutaraldehyde solution werestable.

E. coli β-galactosidase has been immobilized in poly-acrylamide gels and through the preparation of cross-linked derivatives of E. coli β-galactosidase by treating theenzyme with bisimidoesters. The combination of threeprotective agents, namely, bovine serum albumin, cysteine,and lactose, during immobilization gave an increased yieldof 190% in the case of dimethyladipimidate (DMA) cross-linked preparation [58]. K. marxianus cells having lactaseactivity were entrapped in calcium pectate gel (CPG) and incalcium alginate gel (CAG) hardened by polyethyleneimineand glutaraldehyde. Permeabilized cells entrapped in CPGhydrolyzed lactose more than 80% in semicontinuous andcontinuous processes [94].

The comparison of the various methods of immobiliza-tion of β-galactosidase from Thermus aquaticus indicatedthat immobilization by cross-linking followed by entrapmentin agarose beads can be beneficial for high enzyme loadingwith good activity yield [59]. The entrapment of A. oryzae β-galactosidase in a spongy polyvinyl alcohol cryogel increasedthe stability towards temperature, pH, and ionic strengthmore than the free enzyme [60]. The fibers composed ofalginate and gelatin hardened with glutaraldehyde retained56% relative activity of β-galactosidase for 35 days withoutany decrease. Moreover, the optimum conditions were alsonot affected by immobilization [95]. Another approach for

Table 3: Cross-linking reagents used in β-galactosidase immobi-lization.

Cross-linking reagent References

Bis-oxirane [102]

Carbodiimide [103]

Chromium (III) acetate [63]

Glutaraldehyde [14, 20, 69, 70, 104–107]

Polyethyleneimine [57, 101, 108]

Sulfate-dextran [109]

Transglutaminase [110]

Tris(hydroxymethyl)phosphine [54]

immobilization of β-galactosidase is the use of liposomes andin this direction response surface methodology was appliedto optimize the entrapment of the enzyme in liposomes bydehydration-rehydration vesicle method, which resulted inan entrapment efficiency of 28% [96].

It has been observed that entrapped cross-linked con-canavalin A-β-galactosidase complex preparation was moresuperior in the continuous hydrolysis of lactose in a batchprocess as compared to the other entrapped preparationsbecause it retained 95% activity after seventh repeated useand 93% of its original activity after 2-month storage at4◦C [97]. A. oryzae β-galactosidase was immobilized onthe surface of a novel bioaffinity support: concanavalin Alayered calcium alginate-starch beads. The maximum activityof the immobilized β-galactosidase has been obtained at60◦C, approximately 10 degrees higher than that of the freeenzyme. It has been also observed that the immobilized β-galactosidase exhibited significantly higher stability to heat,urea, MgCl2, and CaCl2 than the free enzyme [98]. Calciumalginate-entrapped β-galactosidase preparations have beenused for the hydrolysis of lactose from synthetic solution,milk, and whey in batch processes as well as in continuouspacked bed columns. From the kinetic studies, it wasobserved that the Michaelis constant (Km) for the free andimmobilized β-galactosidase was 2.51 mM and 5.18 mM,respectively. Moreover, the Vmax for the soluble and immobi-lized enzyme was 4.8×10−4 mol/min and 4.2×10−4 mol/min,respectively [99].

The main problems associated with this type of immo-bilization process are desorption of β-galactosidase fromimmobilization matrix and the leakage of the entrappedenzyme due to a small molecular weight compared to poresof gel in matrices, which can be overcome by cross-linkingusing bifunctional or multifunctional reagents [100]. Theconditions for polyethyleimine- (PEI-) coating of agarosesupports to achieve a β-galactosidase derivative have beenoptimized that allows a high lactose conversion from wheyin a steady bed-reactor with no enzyme leakage, togetherwith good elution properties [101]. Various cross-linkingreagents used for improvement of β-galactosidase stability inimmobilized state are described in Table 3 [102–110].

3.3. Covalent Binding Method. Covalent binding is theretention of enzymes on support surface by covalent bond

Enzyme Research 7

formation. Enzyme molecules bind to support material viacertain functional groups such as amino, carboxyl, hydroxyl,and sulfydryl groups. These functional groups must not bein the active site. It is often advisable to carry out the immo-bilization in the presence of its substrate or a competitiveinhibitor so as to protect the active site Functional groupson support material usually activated by using chemicalreagents such as cyanogen bromide, carbodimide, andglutaraldehyde.

Eggshells ground into pieces can be good carrier forimmobilization of β-galactosidase because of its low cost,good mechanical strength, and resistance to microbial attack[61]. Fungal enzyme from A. oryzae has been immobilizedonto powdered nylon-6 and zeolite [62]. Zeolites were non-ideal since its coupling yield was low whereas nylon resultedin a stable matrix. The derivatives obtained either by diazoor by carbodiimide coupling showed the highest activitiesduring immobilization of the enzyme on glycophase-coatedporous glass [103].

E. coli β-galactosidase immobilized onto gelatin usingchromium (III) acetate and glutaraldehyde retained therelative activities of 25% and 22% for glutaraldehyde andchromium (III) acetate immobilized enzyme, respectively[64]. The enzyme immobilized on silica-alumina was morestable than the free form at acidic pH [65]. The ratioof protein to polymer also plays an important role dur-ing enzyme immobilization and 100% binding of proteinto polymer can be obtained using optimal conditions[66].

The performance of immobilization of the thermostableβ-galactosidase from Thermus sp. strain T2 on a stan-dard Sepabeads-epoxy support with other Sepabeads-epoxysupports partially modified with boronate, iminodiacetic,metal chelates, and ethylenediamine was compared [109].Immobilization yields depended on the support, rangingfrom 95% using Sepabeads-epoxy-chelate, Sepabeads-epoxy-amino, or Sepabeads-epoxy-boronic to 5% using Sepabeads-epoxy-IDA. The immobilized β-galactosidase derivativesshowed very improved but different stabilities after favor-ing multipoint covalent attachment by long-term alkalineincubation, the enzyme immobilized on Sepabeads-epoxy-boronic being the most stable. The optimal derivative wasvery active in lactose hydrolysis even at 70◦C, maintaining itsactivity after long incubation times under these conditions.Recently, the cross-linking of β-galactosidase on magneticbeads prepared from different sources (Artemisia seed gum,chitosan, and magnetic fluid) was done in the presenceof glutaraldehyde and the effects of various preparationconditions on the activity of the immobilized β-galactosidasewere studied. The immobilized β-galactosidase resulted in anincrease in enzyme stability [110].

The heat stability of lactase can be increased throughimmobilization [66, 111, 112]. The effect of temperature andpH on the catalytic activity of immobilized β-galactosidasefrom K. lactis indicated that the temperature-activity curvesare similar for both the free and immobilized enzymes [113].However, the maximum activity of the immobilized enzymewas shifted from 40◦C to 50◦C compared with the freeenzyme.

The comparison of a new and commercially availableamino-epoxy support (amino-epoxy-Sepabeads) to conven-tional epoxy supports to immobilize β-galactosidase from A.oryzae showed that the enzyme stability can be significantlyimproved by the immobilization on this support, suggestingthe promotion of some multipoint covalent attachmentbetween the enzyme and the support [67]. The immobiliza-tion of thermophilic β-galactosidase on Sepabeads for lactosehydrolysis showed decrease in product inhibition, which canbe helpful in improving the industrial performance of theenzyme [55].

Alginate-chitosan core-shell microcapsules have alsobeen used for the immobilization of β-galactosidase [68].The rate of 2-nitrophenyl β-galactopyranoside conversionto 2-nitrophenol was faster in the case of calcium alginate-chitosan microcapsules as compared to barium alginate-chitosan microcapsules. Barium alginate-chitosan microcap-sules, however, did improve the stability of the enzyme at37◦C relative to calcium alginate-chitosan microcapsules orfree enzyme.

Among the three different models (without protectionand molecular imprinting technique pretreatment) accom-plished for the encapsulation of β-galactosidase, the highestenzymatic activity of enzyme was obtained with molecularimprinting technique [114]. The free lactase has beencross-linked into Fe3O4-chitosan magnetic microspheres forlactulose synthesis by dual-enzymatic method in organic-aqueous two-phase media using lactose and fructose as theraw materials [115]. The organic-aqueous media can signifi-cantly promote the transglycosidation activity of lactase andtherefore improves the lactulose yield.

Immobilization technology has shown promising role inreducing the product inhibition of β-galactosidase. A. oryzaeenzyme immobilized on chitosan beads was more effectiveas compared to nylon membranes to reduce the galactoseinhibition [116]. Immobilization of the enzyme on hetero-functional epoxy Sepabeads (boronate-epoxy-Sepabeads andchelate-epoxy-Sepabeads) has shown considerable results inreducing the product inhibition [55]. The effect of internalmass transfer and product (galactose) inhibition on asimulated immobilized enzyme-catalyzed reactor for lactosehydrolysis has been studied [104]. A general mathematicalmodel has been developed for predicting the performanceand simulation of a packed-bed immobilized enzyme reactorperforming lactose hydrolysis, which follows Michaelis-Menten kinetics with competitive product (galactose) inhi-bition. The performance characteristics of a packed-bed-immobilized enzyme reactor have been analyzed taking intoaccount the effects of various diffusional phenomena likeaxial dispersion and internal and external mass transferlimitations. The effects of intraparticle diffusion resistances,external mass transfer, and axial dispersion have been studiedand their effects were shown to reduce internal effective-ness factor. A. oryzae β-galactosidase was immobilized onthe surface of a novel bioaffinity support: concanavalinA layered calcium alginate-starch beads. The immobilizedβ-galactosidase had a much higher Kiapp value than thefree enzyme, which indicated less susceptibility to productinhibition by galactose [98].

8 Enzyme Research

4. Applications of Immobilized β-Galactosidase

Immobilized β-galactosidases can be used in a number ofways to hydrolyze lactose in milk, whey/whey permeate, andoligosaccharides synthesis. The choice of process technologydepends on the nature of the substrate, the characteristics ofthe enzyme, economics of production, and marketing of theproduct. The primary characteristic, which determines thechoice and application of a given enzyme, is the operationalpH range. Acid-pH enzymes from fungi are suitable forprocessing of acid whey and whey permeate whereas theneutral-pH enzymes from yeasts and bacteria are suitable forprocessing of milk and sweet whey.

4.1. Hydrolysis of Milk/Whey. Lactose-hydrolyzed milk hasbeen used for the preparation of flavoured milk, cheese,and yoghurt. The hydrolysis of lactose in milk for foodprocessing also prevents lactose crystallization in frozen andcondensed milk products. Moreover, the use of hydrolyzedmilk in yoghurt and cheese manufacture accelerates theacidification process, because lactose hydrolysis is normallythe rate-limiting step of the process, which reduces theset time of yoghurt and accelerates the development ofstructure and flavour in cheese [1]. The quality of icemilk and ice-cream was significantly improved by additionof lactozyme. It prevents the crystallization of lactose bybreaking into glucose and galactose and reduces sandiness[117].

High concentration of lactose in whey is a majorenvironmental problem since its disposal in local waterstreams increases the biological oxygen demand manifolds.The hydrolysis of whey lactose is another important appli-cation of β-galactosidase in dairy industry. Concentratedhydrolyzed whey or whey permeates can be used as asweetener in products such as canned fruit syrups and softdrinks [1]. Various immobilizing agents employed for theimmobilization of β-galactosidase along with hydrolysis oflactose have been summarized in Table 4 [118, 119].

Fungal β-galactosidase (Miles Chemie) immobilized inpolyvinyl alcohol gel was found more thermostable thansoluble enzyme, retaining 70% of the activity after 24 h at50◦C and 5% activity at 60◦C. A lactose hydrolysis of 75%was obtained in 5-6 h and the degree of conversion decreasedto 50% after 30 runs [93]. The studies on the hydrolysis oflactose using immobilized β-galactosidase (Aspergillus niger)on phenol-formaldehyde resin indicated that the optimaltemperature was found to be dependent on the operatingtime but not on the lactose concentration or the conversion[69].

The immobilized β-galactosidase from A. niger displayed70% hydrolysis in skim milk at 40◦C, with a space time of10 min [51]. The β-galactosidase enzyme of fungal sourcesimmobilized on hydrogels was used for whey hydrolysisand 70%–75% hydrolysis was achieved within 7 h [118].The immobilization of β-galactosidase from A. oryzae onactivated silica gel resulted in the most active immobilizedpreparation from TiCl3 and FeCl3-activated silica gel andresulted in 81 and 84% hydrolysis, respectively, in 4% lactosesolution [77].

The β-galactosidase from Bacillus circulans immobilizedonto Duolite ES-762 displayed lactose conversion of >70%in a continuous stirred tank reactor [120]. The immobilizedβ-galactosidase from A. oryzae in a packed bed reactordisplayed 80% of lactose hydrolysis in whey [121] whereasthe immobilized β-galactosidase from Saccharomyces fragilisresulted in a hydrolytic rate of 50% within 3 h in a recyclingpacked bed reactor [78]. Further, the operational stabilitywas tested, with the system being used up to 5 times beforeany significant drop in the activity. The addition of Mg2+

and Mn2+ enhances the hydrolysis of ONPG and lactose byβ-galactosidase from K. lactis, but the rates of activation byeach metal on both substrate were not the same [122]. Theimmobilized K. lactis β-galactosidase from onto CPC-silica(silanizated and activated with glutaraldehyde) and agarose(activated with cyanylating agent) displayed 90% lactoseconversion in packed bed minireactors [72]. β-galactosidaseentrapped in a copolymer gel of N-isopropylacrylamide andacrylamide was effective in hydrolysis of lactose at 5◦C forproduction of low lactose milk. It has been observed thatlactose conversion decreased the stability of milk caseinparticles and increased its dispersity [123].

The kinetic model for the lactose using immobilized β-galactosidase from K. fragilis has also been developed. Theimmobilized enzyme was active at a low temperature of 5◦Cand it could also be applied for the production of freeze dairyproducts to avoid lactose crystallization and to enhance thedigestibility and flavour of such products [64].

K. fragilis β-galactosidase immobilized on silanizedporous glass modified by glutraldehyde binding retainedmore than 90% of its activity [124]. A lactose saccharificationof 86%–90% in whey permeate was achieved both in abatch process and in a recycling packed-bed bioreactor. K.lactis β-galactosidase immobilized onto graphite surface andglutaraldehyde has been used as the cross-linking reagentwith the specific activity yield of 17% and 25% while theenzyme loading was 1.8 and 1.1 U/cm2 of the graphite exter-nal surface area, respectively. It was observed that specificactivity yield decreased with the increase of the enzymeloading [113]. Lactose hydrolysis by a β-galactosidase fromThermus sp. both in solution and immobilized on a com-mercial silica-alumina was studied [34]. Both the free andthe immobilized enzymes are competitively inhibited bygalactose, while glucose inhibited only the action of freeenzyme, in an uncompetitive way. The immobilization stephelped to eliminate the inhibition by glucose. Moreover, theimmobilization reduced to a half the inhibitory action ofgalactose. In general, the immobilization reduced the activityof the enzyme but increased its thermal stability.

The Lactozym (a commercially available enzyme prepa-ration of β-galactosidase obtained from K. fragilis) immo-bilized on cellulose beads has been used for the hydrolysisof whey lactose (>90% conversion) and milk lactose (60%conversion) in 5 h and the immobilized enzyme could bereused three times without any change in the performance ofthe fluidized bed reactor [14]. The immobilized preparationsof β-galactosidase from Thermus sp. resulted in hydrolysisyield higher than 99%. These immobilized forms of β-galactosidase could be used in the total hydrolysis of lactose

Enzyme Research 9

Table 4: Hydrolysis of lactose with various immobilizing techniques of β-galactosidase.

Microbial source Immobilizing agent% Lactosehydrolysis

Time ofhydrolysis

References

Fungal galactosidase (MilesChemie)

Polyvinyl-alcohol 75% 5-6 h [93]

E. coli Polyacrylamide gel 47% 6 h [58]

K. lactis Thiosulfinate/thiosulfonae 85%–90% 2.5 h [79]

K. fragilis Cellulose beads >90% 5 h [14]

K. lactis Cotton fabric 95% 2 h [82]

Fungal β-galactosidase Hydrogels 70%–75% 7 h [116]

K. marxianus Calcium alginate 84.8% 2.5 h [117]

A. oryzaeConcanavalin A layered calcium alginate-starch hybridbeads

89% 3 h [98]

Bacillus stearothermophilus Chitosan >80% 2 h [54]

in milk or dairy whey even at 70◦C [55]. The hydrolysisof lactose by immobilized β-galactosidase has also beenstudied in a continuous flow capillary bed reactor by varioustemperatures. Based on the observed thermal deactivationrate constants, at an operating temperature of 40◦C, only10% of the enzyme activity loss could be there in one year[125].

The β-galactosidase entrapment in liposomes showedsuperior thermal stability at various ranges of temperature.Moreover, the proteolytic stability of the β-galactosidasewas enhanced by encapsulation in liposomes [126]. Theentrapment of β-galactosidase in liposomes by dehydration-rehydration vesicle method has also been used to prevent animmediate hydrolysis of lactose in milk [96]. A. oryzae β-galactosidase was immobilized by three different techniques:adsorption on celite, covalent coupling to chitosan, andaggregation by cross-linking and comparing the yield ofimmobilized preparation, enzymatic characteristics, stability,and efficiency in oligosaccharide synthesis. Cross-linkedenzyme aggregates of β-galactosidase were found effectivein lactose hydrolysis yielding 78% monosaccharide in 12 h[75]. K. lactis β-galactosidase immobilized on cotton fabricusing glutaraldehyde as the cross-linking reagent was usedfor hydrolysis of lactose in whole milk and 95% of lactoseconversion has been observed after 2 h of batch operation[23].

K. lactis β-galactosidase was covalently immobilized ontoa polysiloxane-polyvinyl alcohol magnet, using glutaralde-hyde as activating agent that presented a higher operationaland thermal stability than the soluble enzyme; so thisimmobilized β-galactosidase was also effectively used forthe hydrolysis of lactose from milk [83]. A. oryzae β-galactosidase was immobilized on silica, the enzyme activityas well as stability has been evaluated, and the best immo-bilization results were obtained by using glutaraldehydeas support’s activator and enzyme stabilizer. Among thedifferent treatments (microfiltration, thermal treatment, andultrafiltration) of whey, ultrafiltration was the best treatmenttowards a proper substrate solution for feeding the reactor[84].

A recombinant thermostable β-galactosidase from Bacil-lus stearothermophilus immobilized onto chitosan usingTris (hydroxymethyl) phosphine (THP) and glutaraldehyderesulted in >80% lactose hydrolysis in milk after 2 h ofoperation in a packed bed reactor. Thus, THP-immobilizedrecombinant thermostable β-galactosidase from Bacillusstearothermophilus has the potential application for the pro-duction of lactose-hydrolyzed milk [54]. Calcium alginateentrapped β-galactosidase used for the hydrolysis of lactosefrom solution, milk, and whey in batch processes as wellas in continuous packed bed column. It was also observedthat entrapped cross-linked concanavalin A-β-galactosidasewas more efficient in the hydrolysis of lactose present inmilk (77%) and whey (86%) in batch processes as comparedto the entrapped soluble β-galactosidase [93]. Among thetwo matrices (Sephadex G-75 and chitosan beads) testedfor immobilization β-galactosidase (BGAL) from Pisumsativum for lactose hydrolysis, chitosan-PsBGAL displayedhigher rate of lactose hydrolysis in milk and whey at roomtemperature and 4◦C than Sephadex-PsBGA and is bettersuited for industrial application based on its broad pH andtemperature optima, high temperature stability, reusability,and so forth [56]. β-galactosidases (from K. lactis and A.oryzae) were also immobilized in poly(vinylalcohol) hydrogellens-shaped capsules LentiKats used for the production ofD-galactose from lactose (200 g L−1) in the batch mode ofa simultaneous saccharification and fermentation process[7].

A. oryzae β-galactosidase was immobilized on an inex-pensive bioaffinity support, and concanavalin A-cellulosewas used for the continuous hydrolysis of lactose frommilk and whey. It was observed that the optimum pHfor soluble and immobilized β-galactosidase is 4.8 but theoptimum temperature increased from 50 to 60◦C for theimmobilized β-galactosidase. The immobilized enzyme hadhigher thermal stability at 60◦C [92]. Recently, a packed bedreactor together with alginate entrapped permeabilized cells(K. marxianus) has been used for hydrolysis of milk lactosein a continuous system, which resulted in 87.2% hydrolysisof milk lactose [127].

10 Enzyme Research

4.2. Synthesis of Galacto-Oligosaccharides (GOSs). Besideshydrolytic action, β-galactosidase has also transferase activityby which the enzyme produces and hydrolyses a seriesof oligosaccharides, which have a beneficial effect on thegrowth of desirable intestinal microflora [12]. Moreover, thetransferase reaction can be used to attach galactose to otherchemicals, resulting in formation of galacto-oligosaccharides(GOSs), and consequently have potential application inthe production of food ingredients, pharmaceuticals, andother biological active compounds. Nowadays, oligosaccha-ride production becomes the interesting subject for theresearchers, because the oligosaccharides have beneficialeffect on human intestinal as “bifidus factor”—promotinggrowth of desirable intestinal microflora. Oligosaccharidesare recognized as useful dietary tools for the modula-tion of the colonic microflora toward a healthy balance.This usually involves selectively increasing the levels ofgut Bifidobacteria and Lactobacilli at the expense of less-desirable organisms such as Escherichia coli, Clostridia, andproteolytic bacteroides [128]. The amount and nature ofoligosaccharides formed depend upon the several factorsincluding the enzyme source, the concentration and natureof the substrate, and reaction conditions [12, 129, 130]. Theyield of oligosaccharides can be increased by using highersubstrate concentrations or decreasing the water content[31].

Although β-galactosidase catalyzes both hydrolysis andtransgalactosylation reactions, however, the process condi-tions for lactose hydrolysis and GOS synthesis are different.The reaction conditions for transgalactosylation should behigh lactose concentration, elevated temperature, and lowwater activity in the reaction medium [130]. The tempera-ture, concentration of substrate, and enzyme origin play animportant role in the enzymatic synthesis of oligosaccharides[131]. However, the influence of the initial lactose concen-tration can be much larger [132, 133]. In general, more andlarger galacto-oligosaccharides (GOSs) can be produced withhigher initial lactose concentrations. The higher tempera-tures can be beneficial in higher oligosaccharide yields. Thehigher yield at higher temperatures is an additional advan-tage when operating at high initial lactose concentrations andconsequently elevated temperatures. Hence, immobilized β-galactosidase should be stable at high temperature, low watercontent, and giving high transgalactosylation activity [134].

Partially purified β-galactosidase from Bullera singularisATCC 24193 immobilized in Chitopearl BCW 3510 beadhas been used for the production of galacto-oligosaccharides(GOSs) from lactose in a packed bed reactor, which resultedin 55% (w/w) oligosaccharides with a productivity of4.4 g/(L-h) during a 15-day operation [132]. The enzymeimmobilized on tosylate cotton cloth was used in plug-flowreactor for continuous production of galacto-oligosaccharidefrom lactose. In general, more and larger GOS can be pro-duced with higher initial lactose concentrations. A maximumGOS production of 27% (w/w) of initial lactose was achievedat 50% lactose conversion with 500 g/L of initial lactoseconcentration. Tri-saccharides were the major types of GOSformed, accounting for more than 70% of the total GOSsproduced in the reactions. The chitosan-immobilized A.

oryzae β-galactosidase gave maximum trisaccharides yield(17.3% of the total sugar) using 20% (w/v) of lactose, within2 h as compared to 10% with free enzyme and 4.6% withcross-linked aggregates [73].

An immobilized-enzyme system using polyethylene-imine, glutaraldehyde, and cotton cloth was studied andcompared the galacto-oligosaccharide production in free-enzyme ultrafiltration and in immobilized-enzyme systems[135]. In the immobilization process, approximately 50%to approximately 90% enzyme inactivation was found withthe combination of PEI and GA. Equivalent free- andimmobilized-enzyme systems showed very similar maximumGOS production of approximately 22% and approximately20% (w/v) at approximately 15 to 17 min, 50% conversionfor free- and immobilized-systems, respectively.

The synthesis of galacto-oligosaccharides was optimizedwith respect to lactose concentration and enzyme to sub-strate ratio using immobilized A. oryzae β-galactosidase[136]. In the sequential batch production of galacto-oligosaccharides, biocatalyst efficiency was increased by190% with respect to the free enzyme in solution, and 8500 gof galacto-oligosaccharides per gram of enzyme prepara-tion were produced after 10 batches. The immobilized A.oryzae β-galactosidase enzyme on magnetic Fe3O4–chitosan(Fe3O4–CS) nanoparticles as support resulted in 15.5%(w/v) maximum yield of galacto-oligosaccharides [137].The synthesis of galacto-oligosaccharides (GOSs) using A.oryzae β-galactosidase (free and immobilized) on magneticpolysiloxane-polyvinyl alcohol (mPOS-PVA) has also beencarried out [138]. A maximum of 26% (w/v) of total sugarswas achieved at near 55% lactose conversion from 50%(w/v) lactose solution at pH 4.5 and 40◦C. Trisaccharidesaccounted for more than 81% of the total GOS produced.GOS formation was not considerably affected by pH andtemperature. The concentrations of glucose and galactoseencountered near maximum GOS concentration greatlyinhibited the reactions and reduced GOS yield.

The packed bed reactor and a plug-flow reactor havebeen successfully used for continuous production of GOSfrom lactose using immobilized β-galactosidase [63, 90].The selectivity for GOS synthesis can be increased several-fold under microwave irradiation, using immobilized β-glucosidase and with added cosolvents such as hexanol [139].Recently, a new type of ceramic membrane reactor systemusing immobilized β-galactosidase (Kluyveromyces lactis)has been proposed for continuous enzymatic productionof galactosyl-oligosaccharides (GOSs) from lactose, whichresulted in maximum oligosaccharide (38%, w/w) when theaverage residence time was 24 min, with an initial 30% (w/w)lactose concentration [140].

5. Scale-Up Issues

For the production of lactose-free milk, the enzyme β-galactosidase can be added directly to whole milk, butafter complete lactose hydrolysis at a desired level, theenzyme can be deactivated by heat treatment. Since the freeenzyme cannot be reused, thus the resulting operation is

Enzyme Research 11

not cost effective. To overcome this problem, immobilized β-galactosidase is used for the hydrolysis of skim milk. After thedesired lactose hydrolysis is achieved, cream is added to thehydrolysed milk to adjust its fat content. Although numeroushydrolysis systems have been investigated, only few of themhave been scaled up and even fewer have been applied at anindustrial or semi-industrial level.

The first company for the commercial hydrolysis oflactose in milk by using immobilized lactase was Centraledel Latte of Milan, Italy, utilizing the SNAM Progettitechnology. The process used an immobilized Saccharomyces(Kluyveromyces) lactis lactase entrapped in cellulose triac-etate fibres. Sumitomo Chemical, Japan, has also developedan immobilization process to immobilize β-galactosidase offungal origin on the rugged surface of an amphoteric ion-exchange resin of phenol formaldehyde polymer and thistechnology was used by Drouin Cooperative Butter Factoryfor producing market milk and hydrolyzed whey [141].

Snow Brand’s factory has developed a rotary columnreactor that could be used both as a stirred tank reactorand as a packed bed reactor [142]. The reaction rate wasgreatly affected by the packing density of immobilized β-galactosidase in the rotary column. This reactor can alsoovercome the problem of channeling or severe pressure drop.If the hydrolysis of lactose was carried out in horizontalrotary column, 70%–80% lactose hydrolysis was observedand washing of the immobilized enzyme was carried outfor 36 cycles, which indicated that horizontal rotary columnreactor was well suited for hydrolyzing lactose in milk withfibrous immobilized enzyme. From the pilot plant experi-mentations, a commercial plant was set up at Snow Brand’sfactory [142]. Although the immobilized β-galactosidase waswashed with phosphate buffer solution and pasteurized withTego-51, the standard plate count of lactose hydrolyzed milkincreased sharply.

Thus, immobilized β-galactosidase technology is aneffective process for successful hydrolysis of lactose and itcan overcome the problems associated with costs of solubleenzyme. However, major problems associated with theimmobilized enzyme system are microbial contamination,protein adherence, and channeling. Therefore, for long-termoperations using immobilized system, periodic washing,and pasteurization are indispensable processes [143–145].In immobilized enzyme system, protein adhered to theenzyme can be easily dissolved by using high and lowpH solutions, because the immobilized enzyme has highdurability over a wide range of pH. The immobilized enzymecan be pasteurized with benzalkonium chloride (quaternaryammonium salt) after removing the proteins. The use ofacetic acid solution as a cleaning and pasteurizing agentinstead of lactic acid can also be effective. The problem ofchanneling observed in the packed column system can beovercome by changing the flow direction of feed during theoperation [142, 144, 145].

Galacto-oligosaccharides are produced simultaneouslyduring lactose hydrolysis due to transgalactosylation activityof β-galactosidase. Oligosaccharides/Bifidobacteria provide awide variety of health benefits, including anticarcinogeniceffects, reduction in serum cholesterol, improved liver

function, reduction of the colon cancer risk, and improvedintestinal health [146, 147]. Therefore, the public demandfor their production is significantly increased together withthe development of an effective and inexpensive GOSproduction. Major companies dealing with oligosaccharidesproduction (including GOS) are in Japan [148]. Recently,there is also an increasing trend of GOS production inEurope. Besides lactulose and soybean oligosaccharides, alloligosaccharides are prepared by transglycosylation frommono and disaccharides or by controlled hydrolysis ofpolysaccharides [147].

6. Conclusions

β-galactosidase is one of the most important enzymes usedin food processing, which offers nutritional, technological,and environmental applications. Enzyme immobilizationprovides enzyme reutilization and may result in increasedactivity by providing a more suitable microenvironment forthe enzyme. Moreover, immobilized systems can providebetter enzyme thermostability and pH tolerance. However,major problems associated with the immobilized enzymesystem are microbial contamination, protein adherence, andchanneling. The periodic washing and pasteurization andflow direction of feed can solve these problems to greatextent. The problem of microbial contamination can alsobe solved by exploiting the temperature property of theenzyme. The immobilized enzyme preparations showed upto 99% hydrolysis, and thus it can be applied successfullyfor the hydrolysis of lactose in milk or whey. The isolationof pyschrophilic bacteria with cold active β-galactosidase hasopened up the possibility of processing of milk and wheyeven at low temperatures. On the other side, thermostableenzymes have the unique ability to retain their activity athigher temperatures for prolonged periods, and the processis less prone to microbial contamination due to higheroperating temperature. Thus, cold active and thermostableenzymes will have the great potential in the lactose hydrolysisand of particular interest to the researchers. Thus, immo-bilization enzyme systems will certainly find greater role infuture times for the hydrolysis of milk, whey, and synthesisof galacto-oligosaccharides.

Acknowledgment

The authors acknowledge the financial support given by theCouncil for Scientific and Industrial Research (CSIR), NewDelhi, India.

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[107] A. Tanriseven and S. Dogan, “A novel method for theimmobilization of β-galactosidase,” Process Biochemistry, vol.38, no. 1, pp. 27–30, 2002.

[108] N. Albayrak and S. T. Yang, “Immobilisation of beta-galactosidase on fibrous matrix by polyethyleneiminefor production of galacto-oligosaccharides from lactose,”Biotechnology Progress, vol. 18, pp. 240–251, 2002.

[109] B. C. C. Pessela, C. Mateo, M. Fuentes et al., “Stabilizationof a Multimeric β-Galactosidase from Thermus sp. Strain T2by Immobilization on Novel Heterofunctional Epoxy Sup-ports Plus Aldehyde-Dextran Cross-Linking,” BiotechnologyProgress, vol. 20, no. 1, pp. 388–392, 2004.

[110] S. Zhang, S. Gao, and G. Gao, “Immobilization of β-galactosidase onto magnetic beads,” Applied Biochemistry andBiotechnology, vol. 160, no. 5, pp. 1386–1393, 2010.

[111] R. F. H. Dekker, “Immobilization of a lactase onto a mag-netic support by covalent attachment to polyethyleneimine-glutaraldehyde- activated magnetite,” Applied Biochemistryand Biotechnology, vol. 22, no. 3, pp. 289–310, 1989.

[112] C. S. Chen, C. K. Hsu, and B. H. Chiang, “Optimizationof the enzymic process for manufacturing low-lactose milk

containing oligosaccharides,” Process Biochemistry, vol. 38,no. 5, pp. 801–808, 2002.

[113] Q. Z. K. Zhou and X. D. Chen, “Effects of temperatureand pH on the catalytic activity of the immobilized β-galactosidase from Kluyveromyces lactis,” Biochemical Engi-neering Journal, vol. 9, no. 1, pp. 33–40, 2001.

[114] Z. Wu, M. Dong, M. Lu, and Z. Li, “Encapsulation of β-galactosidase from Aspergillus oryzae based on “fish-in-net”approach with molecular imprinting technique,” Journal ofMolecular Catalysis B, vol. 63, no. 1-2, pp. 75–80, 2010.

[115] X. Hua, R. Yang, W. Zhang, Y. Fei, Z. Jin, and BO. Jiang,“Dual-enzymatic synthesis of lactulose in organic-aqueoustwo-phase media,” Food Research International, vol. 43, no.3, pp. 716–722, 2010.

[116] M. Portaccio, S. Stellato, S. Rossi et al., “Galactose com-petitive inhibition of β-galactosidase (Aspergillus oryzae)immobilized on chitosan and nylon supports,” Enzyme andMicrobial Technology, vol. 23, no. 1-2, pp. 101–106, 1998.

[117] D. K. Stevenson, J. S. Crawford, and J. O. Carroll, “Enzymatichydrolysis of lactose in ice-cream,” Journal of Dairy Science,vol. 66, pp. 75–78, 1983.

[118] A. Kozhucharova, K. Baztosolova, Y. Papova, N. Kirova, D.Klysursky, and D. Simeonov, “Properties and employmentof β-galactosidase immobilized on hydrogel from silica,”Bioteknologiya, vol. 1, pp. 61–64, 1990.

[119] R. Panesar, P. S. Panesar, R. S. Singh, and M. B. Bera, “Appli-cability of alginate entrapped yeast cells for the production oflactose-hydrolyzed milk,” Journal of Food Process Engineering,vol. 30, no. 4, pp. 472–484, 2007.

[120] K. Nakanishi, R. Matsuno, K. Torii, K. Yamamoto, andT. Kamikubo, “Properties of immobilized β-d-galactosidasefrom Bacillus circulans,” Enzyme and Microbial Technology,vol. 5, no. 2, pp. 115–120, 1983.

[121] H. Plainer, B. G. Sprossler, and H. Uhlig, “Technical applica-tion of immobilized lactase and amino acid acylase,” Hoppe-Seyler’s Zeitschrift fur Physiologische Chemie, vol. 365, pp.1043–1047, 1984.

[122] Y.-S. Kim, C.-S. Park, and D.-K. Oh, “Lactulose productionfrom lactose and fructose by a thermostable β-galactosidasefrom Sulfolobus solfataricus,” Enzyme and Microbial Technol-ogy, vol. 39, no. 4, pp. 903–908, 2006.

[123] A Miezeliene, A Zubriene, S Budriene, G Dienys, and J.Sereikaite, “Use of native and immobilized β-galactosidase inthe food industry,” Progress in Biotechnology, vol. 17, pp. 171–175, 2000.

[124] J. Szczodrak, “Hydrolysis of lactose in whey permeate byimmobilized β-galactosidase from Kluyveromyces fragilis,”Journal of Molecular Catalysis B, vol. 10, no. 6, pp. 631–637,2000.

[125] R. S. Peterson, C. G. Hill, and C. H. Amundson, “Effects oftemperature on the hydrolysis of lactose by immobilized β-galactosidase in a capillary bed reactor,” Biotechnology andBioengineering, vol. 34, no. 4, pp. 429–437, 1989.

[126] J. M. Rodriguez-Nogales and A. Delgadillo, “Stabilityand catalytic kinetics of microencapsulated β-galactosidasein liposomes prepared by the dehydration-rehydrationmethod,” Journal of Molecular Catalysis B, vol. 33, no. 1-2,pp. 15–21, 2005.

[127] R. Panesar, P. S. Panesar, R. S. Singh, and J. F. Kennedy,“Hydrolysis of milk lactose in a packed bed reactor systemusing immobilized yeast cells,” Journal of Chemical Technol-ogy & Biotechnology, vol. 86, no. 1, pp. 42–46, 2011.

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[128] R. Fuller and G. R. Gibson, “Probiotics and prebiotics:microflora management for improved gut health,” ClinicalMicrobiology and Infection, vol. 4, no. 9, pp. 477–480, 1998.

[129] J. E. Prenosil, E. Stuker, and J. R. Bourne, “Formation ofoligosaccharides during enzymatic hydrolysis of lactose—part 1: sate of art,” Biotechnology and Bioengineering, vol. 30,no. 9, pp. 1019–1025, 1987.

[130] S. Zarate and M. H. Lopez-Leiva, “Oligosaccharide formationduring enzymatic lactose hydrolysis: a literature review,”Journal of Food Protection, vol. 53, pp. 262–268, 1990.

[131] M. A. Boon, A. E. M. Janssen, and K. Van ’t Riet, “Effect oftemperature and enzyme origin on the enzymatic synthesisof oligosaccharides,” Enzyme and Microbial Technology, vol.26, no. 2–4, pp. 271–281, 2000.

[132] N. Albayrak and S. T. Yang, “Production of galacto-oligosaccharides from lactose by Aspergillus oryzae β-galactosidase immobilized on cotton cloth,” Biotechnologyand Bioengineering, vol. 77, no. 1, pp. 8–19, 2002.

[133] S. Chockchaisawasdee, V. I. Athanasopoulos, K. Niranjan,and R. A. Rastall, “Synthesis of galacto-oligosaccharidefrom lactose using β-galactosidase from kluyveromyces lactis:studies on batch and continuous UF membrane-fitted biore-actors,” Biotechnology and Bioengineering, vol. 89, no. 4, pp.434–443, 2005.

[134] S. X. Chen, D. Z. Wei, and Z. H. Hu, “Synthesis of galacto-oligosaccharides in AOT/isooctane reverse micelles by β-galactosidase,” Journal of Molecular Catalysis B, vol. 16, no.2, pp. 109–114, 2001.

[135] N. J. Matella, K. D. Dolan, and Y. S. Lee, “Comparison ofgalactooligosaccharide production in free-enzyme ultrafil-tration and in immobilized-enzyme systems,” Journal of FoodScience, vol. 71, no. 7, pp. C363–C368, 2006.

[136] L. M. Huerta, C. Vera, C. Guerrero, L. Wilson, and A.Illanes, “Synthesis of galacto-oligosaccharides at very highlactose concentrations with immobilized β-galactosidasesfrom Aspergillus oryzae,” Process Biochemistry, vol. 46, no. 1,pp. 245–252, 2011.

[137] C. Pan, B. Hu, W. Li, YI. Sun, H. Ye, and X. Zeng, “Novel andefficient method for immobilization and stabilization of β-d-galactosidase by covalent attachment onto magnetic Fe3O4-chitosan nanoparticles,” Journal of Molecular Catalysis B, vol.61, no. 3-4, pp. 208–215, 2009.

[138] D. F. M. Neri, V. M. Balcao, R. S. Costa et al., “Galacto-oligosaccharides production during lactose hydrolysis byfree Aspergillus oryzae β-galactosidase and immobilized onmagnetic polysiloxane-polyvinyl alcohol,” Food Chemistry,vol. 115, no. 1, pp. 92–99, 2009.

[139] T. Maugard, D. Gaunt, M. D. Legoy, and T. Besson,“Microwave-assisted synthesis of galacto-oligosaccharidesfrom lactose with immobilized β-galactosidase fromKluyveromyces lactis,” Biotechnology Letters, vol. 25, no. 8, pp.623–629, 2003.

[140] M. Ebrahimi, L. Placido, L. Engel, K. S. Ashaghi, and P.Czermak, “A novel ceramic membrane reactor system forthe continuous enzymatic synthesis of oligosaccharides,”Desalination, vol. 250, no. 3, pp. 1105–1108, 2010.

[141] Y. Honda, M. Kako, K Abiko, and Y. Sogo, “Hydrolysis oflactose in milk,” in Industrial Application of ImmobilizedBiocatalysts, A. Tanaka, T. Tosa, and T. Kobayashi, Eds., pp.209–234, Marcel Dekker, New York, NY, USA, 1993.

[142] Y Honda, H Hashiba, K Ahiko, and H. Takahashi, “Hydroly-sis of immobilized β-galactosidase using a horizontal rotary

column reactor,” Nippon Shokuhin Kogyo Gakkaish, vol. 38,p. 384, 1991.

[143] H. Hirohara, H. Yamamoto, E. Kawano, and T. Nagase,“Continuous hydrolysis of lactose in skim milk and acid wheyby immobilized lactose of Aspergillus oryzae,” in Proceedingsof the 6th International Enzyme Engineering Conference, vol.6, pp. 295–297, 1982.

[144] Y. Honda and S. Takafuji, “Hydrolysis of lactose in recon-stituted skimmed milk using immobilized β-galactosidase(immobilized Sumylact). 1. Development of bioreactor,”Report of Research Laboratory (Snow Brand Milk ProductsCo Ltd). 83: 85, 1987.

[145] Y. Honda, K. Murata, S. Takafuji, S. Fujiwara, T. Iwasaki,and M. Kuwazuru, “Hydrolysis of lactose in reconstitutedskimmed milk using immobilized β-galactosidase (immobi-lized Sumylact). 2. Results of long term experiments in a pilotplant,” Report of Research Laboratory (Snow Brand MilkProducts Co Ltd). 85: 33, 1987.

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SAGE-Hindawi Access to ResearchEnzyme ResearchVolume 2010, Article ID 324184, 7 pagesdoi:10.4061/2010/324184

Research Article

Screen-Printed Carbon Electrodes Modified by Rhodium Dioxideand Glucose Dehydrogenase

Vojtech Polan, Jan Soukup, and Karel Vytras

Department of Analytical Chemistry, Faculty of Chemical Technology, University of Pardubice, Studentska 573,532 10 Pardubice, Czech Republic

Correspondence should be addressed to Karel Vytras, karel.vytras@upce.cz

Received 3 June 2010; Accepted 15 December 2010

Academic Editor: Raffaele Porta

Copyright © 2010 Vojtech Polan et al. This is an open access article distributed under the Creative Commons Attribution License,which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

The described glucose biosensor is based on a screen-printed carbon electrode (SPCE) modified by rhodium dioxide, whichfunctions as a mediator. The electrode is further modified by the enzyme glucose dehydrogenase, which is immobilized on theelectrode’s surface through electropolymerization with m-phenylenediamine. The enzyme biosensor was optimized and tested inmodel glucose samples. The biosensor showed a linear range of 500–5000 mg L−1 of glucose with a detection limit of 210 mg L−1

(established as 3σ) and response time of 39 s. When compared with similar glucose biosensors based on glucose oxidase, the mainadvantage is that neither ascorbic and uric acids nor paracetamol interfere measurements with this biosensor at selected potentials.

1. Introduction

There exists today an ever-increasing demand for fast, selec-tive, reliable, and, above all, inexpensive analytical methods.For food products, it is necessary to monitor whether ornot microbial, or some other form of, contamination hasoccurred. Furthermore, it is necessary to monitor compli-ance with given technological procedures and whether thestated raw materials were used [1]. These requirements placevery great demands on the analysis of given samples. Theanalysis itself should be very fast, sufficiently sensitive andaccurate, but also inexpensive. To meet these criteria, anapplication of electrochemical biosensors seems to be a goodalternative.

Electrochemical biosensors combine two advantages:specificity of the enzyme to the given molecule and transferof the biochemical signal to an electrochemical signal [2].As a result, these biosensors are selective in establishinga specific substrate [3, 4]. By using these biosensors, it ispossible to determine a large number of substances even incomplex matrices.

Electrochemical biosensors often use redox enzymesduring catalysis of substrate splitting reactions. Most usedredox enzyme’s are oxidases and dehydrogenases. There areseveral methods for establishing a substrates concentration.

The most methods often used involve detecting hydrogenperoxide (a product of most oxidases) and nicotinamide ade-nine dinucleotide (NADH) (a product of dehydrogenases)resulting during the catalytic process. NADH oxidation oncarbon electrodes requires high overvoltage (around 1.0 V).This is a highly unfavorable phenomenon, as the impactof interferents (e.g., uric acid, ascorbic acid, paracetamol)that are easily oxidized at a given overvoltage becomemost evident at such potentials. High overvoltage can besuppressed by using a so-called mediator [5–9] that enablesthe transfer of electrons between the enzymes active center,or the product of the enzyme reaction, and the electrodessurface. As the mechanism of NADH oxidation has not beenfully explained, we have written it according to the generallyrecognized mechanism [10], as shown in the formula below(1):

NADH−e−−−→ NADH+• −H+−−→ NAD• −e−� NAD+

slow medium fast(1)

The most important step in preparing a biosensor is thatof enzyme immobilization. Should an inadequate procedurefor enzyme entrapment be chosen, its denaturation, indirectinactivation, or washing from the electrode may occur. Many

2 Enzyme Research

established immobilization techniques are currently usedthat include physical and chemical immobilization. Choice ofenzyme immobilization method depends upon the proper-ties of the enzyme, type of the mediator, conditions in whichthe biosensor is to work, and, last but not least, the physicalproperties of the analyte (or possibly the size of the moleculesto be determined). Due to its simplicity, immobilizationusing electropolymerization [11–14] is one of the mostcommonly used techniques. Electropolymerization proceedsin a buffer solution that contains both a certain monomerand the enzyme itself which will be immobilized. A majoradvantage of this technique is the possibility to regulate thethickness of the membrane formed.

This article describes the preparation, optimization, andanalytical properties of an enzyme biosensor prepared usingthe screen-printing technique, modified by rhodium dioxide,and containing glucose dehydrogenase immobilized in alayer of m-phenylenediamine (the main reason for thisselection was price of the substance when compared withanalogous o- or p-derivatives). The main advantage of thisbiosensor is that it works even at low input potentials,where contributions from other easily oxidizable or reduciblemolecules are negligible.

2. Experimental

2.1. Instrumentation. A modular electrochemical system,AUTOLAB, equipped with modules PGSTAT 30 and ECD(Ecochemie, Utrecht, Holand) was used in combination withcorresponding software (GPES, Ecochemie).

The flow injection system consisted of a peristaltic pump(Minipuls 3, Gilson SA., France), a sample injection valve(ECOM, Ventil C, Czech Republic), and a self-constructedthin-layer electrochemical flow-through cell. The workingelectrode was fixed via rubber gaskets (thickness 0.6 mm)directly to the back plate of the thin layer cell. The referenceelectrode was Ag/AgCl/3M KCl (RE-6, BAS, USA), and thestainless steel back plate represented the counter electrode ofthe cell. The responses were evaluated using the peak heights(differences between background and response current of theanalyte). Corresponding pH values were measured using aportable pH-meter (CPH 52 model, Elteca, Turnov, CzechRepublic) equipped with a combined glass pH-sensor (OP-0808P, Radelkis, Budapest, Hungary). The measuring cellwas calibrated using buffer solutions of the conventionalactivity scale.

2.2. Chemicals, Reagents, and Solutions. Glucose oxidase (EC1.1.3.4. from Aspergillus niger, specific activity 198 U mg−1;GOx), glucose dehydrogenase (EC 1.1.1.47 from Pseu-domonas sp., specific activity 277 U mg−1; GDH), Nafion(5% m/m solution in lower aliphatic alcohols), nicoti-namide adenine dinucleotide (NAD+) and its reducedform (NADH), rhodium dioxide, acetate cellulose (M∼37000 g moL−1), m-phenylenediamine, glutaraldehyde so-lution (GA, 50 wt. % in H2O), bovine serum albumin (BSA;5% solution) and pyrrole (98% solution) were purchasedfrom Aldrich. All other chemicals used for the preparation

of buffer, stock, and standard solutions were of analyticalreagent grade and purchased from Lachema (Brno, CzechRepublic). Phosphate buffer was prepared by mixing aqueoussolutions of sodium dihydrogenphosphate and disodiumhydrogenphosphate (both 0.1 M) to achieve solutions of therequired pH values. The glucose stock solution (2.5 g L−1)was prepared and diluted appropriately. Solutions of ascorbicacid and uric acid (both Aldrich, 50 mg L−1) were preparedimmediately before use.

2.3. Electrode Preparation. Carbon ink (0.95 g, GwentC50905D1, Pontypool, UK) and corresponding catalyst(0.05 g) were thoroughly mixed manually for 5 min andsubsequently sonicated for 5 min. The resulting mixturewas immediately used for the fabrication of electrodes.The working electrodes were prepared by screen-printing ofmodified ink onto an inert laser pre-etched ceramic support(113 × 166 × 0.635 mm, no. ADS96R, Coors Ceramics,Chattanooga, TN, USA). Thick layers of the modified carbonink were formed by brushing the ink through an etchedstencil (thickness 100 μm, electrode printing area 105 mm2)with the aid of the spatula provided with the screen-printingdevice (SP-200, MPM, Franklin, MA, USA and/or UL1505 A, Tesla, Czech Republic) onto the ceramic substrates.The resulting plates were dried at 60◦C for 2 h.

2.4. Enzyme Immobilization. Several types of immobilizationmethods were tested with glucose oxidase, comprisingentrapment in Nafion, cross-linking with glutaraldehyde,immobilization using cellulose acetate, and electropolymer-ization of pyrrole or m-phenylenediamine. Subsequently, aGDH enzyme together with cofactor NAD+ were immobi-lized using the best method, in terms of retaining enzymeactivity, response time, sensitivity, and dynamic range ofconcentrations.

2.4.1. Entrapment in Nafion. An enzyme (GOx, 1 mg) wasdissolved in 20 μL of 0.1 M phosphate buffer (pH 7.5) andmixed with an equal amount of 0.05%, 0.5%, or 5% Nafionsolution neutralized with ammonia to pH ∼7. The resultingmixture (5 μL) was applied directly onto the active area of theSPCE/RhO2 surface and air-dried for 30 min.

2.4.2. Immobilization in Cellulose Acetate. An enzyme (GOx,1 mg) was dissolved in 40 μL of 0.1 M phosphate buffer (pH7.5), and a volume of 3 μL of this solution was applied ontothe active area of the SPCE/RhO2 surface and air-dried.Subsequently, volumes of 3 μL of cellulose acetate solutionin acetone (0.05%, 0.5%, 1.5%, or 3.0%) were applied ontothe aforementioned enzyme layer and dried for 5 min.

2.4.3. Cross-Linking with Glutaraldehyde. Volumes of 5, 10or 20 μL of 5% glutaraldehyde (diluted with 0.1 M phosphatebuffer, pH 7.5) were mixed with 1 μL of 5% BSA and with35 μL, 30 μL, or 20 μL of the enzyme solution (1 mg of GOxin 0.1 M phosphate buffer, pH 7.5). After thorough mixing,a volume of 3 μL was applied onto an SPCE/RhO2 and air-dried for 30 min. As a variant, cross-linking of the enzyme

Enzyme Research 3

was also performed with GA vapor, whereby a volume of3 μL of the enzyme solution (1 mg of GOx in 40 μL of 0.1 Mphosphate buffer, pH 7.5) was applied onto the SPCE, air-dried (30 min), and then the SCPE/RhO2 so treated wasenclosed overnight (17 hours) in a vial over 5% GA.

2.4.4. Electropolymerization with Pyrrole. An enzyme solu-tion (3 μL, 1 mg of GOx in 40 μL of 0.1 M phosphate buffer,pH 7.5) was applied onto an SPCE/RhO2. After drying for30 min, the electrode was dipped into the 5 mM solution ofpyrrole in 0.1 mM phosphate buffer, pH 6.0. Electropolymer-ization was performed at +0.75 V versus Ag/AgCl for 0.25,0.5, 1.0, or 2.5 min, respectively. Finally, the electrode waswashed with the phosphate buffer.

2.4.5. Electropolymerization with m-Phenylenediamine—GOx. The procedure applied was similar to that describedin the previous paragraph, but, concerning deposition time,the electrode was polarized in 5 mM m-phenylenediaminefor 0.5, 1.0, 5, 10, or 20 min, respectively. Additionally,electropolymerization was performed at +0.75 V versusAg/AgCl from 5 mM m-phenylenediamine GOx solution(10 mL containing 1 mg of GOx) for 5 min.

2.4.6. Electropolymerization with m-Phenylenediamine—GDH. A volume of 3 μL of NAD+ solution (3 mg in 40 μLof 0.1 M pH 7.5 phosphate buffer) was applied onto theSPCE/RhO2 electrode surface. After drying, the surface wasoverlayered with GDH solution (3 μL, 1 mg in 40 μL of0.1 M phosphate buffer) and dried for 45 min. An electrodewas then dipped into the 5 mM solution (10 mL) of m-phenylenediamine in 0.1 mM phosphate buffer (pH 6.0)containing the remaining 37 μL of NAD+ and 37 μL of GDHsolution, and it was left there for electropolymerization(5 min at +0.75 V versus Ag/AgCl). After washing with buffer,the electrode was prepared for measurements.

2.5. Procedure. Measurements were performed by DCamperometry using both flow injection and batch arrange-ments. All operational variables were optimized, that is,applied potential (from +0.6 to −0.3 V versus Ag/AgCl), pHof phosphate buffer (5–9), and flow rate (0.1–1.5 mL min−1).Responses were evaluated using the peak heights (differencesbetween background and response current of the analyte).Injections of analyte were repeated at least three times.

2.6. Sample Processing. A sample of honey was preparedby dissolving the given amount of honey (3.4 g or 4.4 g offorest honey) in 50 mL of 0.1 M phosphate buffer of pH7.5. Similarly, a sample of syrup was prepared, that is, 2.9 gof orange-flavored syrup was dissolved in 50 mL of 0.1 Mphosphate buffer of pH 7.5. For analysis, 200 μL of thesamples thus prepared were always taken.

3. Results and Discussion

3.1. Effect of Enzyme Immobilization on Biosensor Response.Glucose oxidase was chosen as a test enzyme because of its

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Figure 1: Immobilization using Nafion. Measurement condition:input potential −0.2 V (versus Ag/AgCl); 0.1 M phosphate buffer(pH 7.5); measured with SPCE/RhO2/GOx; analysis in a batcharrangement; concentration of Nafion: 1–0.5%, 2–5%.

stability and sensitivity to glucose [15]. For immobilizingglucose oxidase, the following methods and substanceswere used: immobilization in polymer—Nafion or celluloseacetate; immobilization using cross-linking—glutaraldehydeand BSA; electropolymerization—pyrrole or phenylenedi-amine. The entire study devoted to entrapment of theenzyme was performed in a batch arrangement in a cell witha volume of 10 mL. Selected key factors were monitored foreach system: sensitivity, response time, and dynamic range.

3.1.1. Entrapment in Nafion. Figure 1 shows calibrationdependences obtained in immobilization of 0.5% GOxand 5% Nafion. The concentration of 0.05% was notsufficient to properly entrap the enzyme and the enzymewas shortly washed into the solution, which preventedfurther measurements. The dynamic ranges for Nafionconcentrations of 0.5% and 5% were almost identical. The0.5% Nafion, however, shows greater sensitivity to glucoseand the response time here was the shortest, hovering around28 s.

3.1.2. Immobilization of GOx by Cross-Linking with Glu-taraldehyde and BSA. This immobilization procedure isvery popular and well-proven for the design of enzymeelectrochemical biosensors, and, therefore, it was includedin this study. Concentrations of 0.625%, 1.25%, and 2.5%glutaraldehyde were compared here in a mixture withthe enzyme. The possibility for enzyme immobilizationusing glutaraldehyde saturated vapors was examined aswell (Figure 2). The response time was shortest in thecase of enzyme immobilization using saturated vapors—30 s. While the response sensitivity to glucose decreased(poorer permeability of the analyte to the enzyme and poorerpermeability of the metabolic product to the electrode’ssurface) with increasing thickness of the GA layer, thedynamic range of the setting increased at the same time.

4 Enzyme Research

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Figure 2: Immobilization using glutaraldehyde and BSA. Mea-surement condition: input potential −0.2 V (versus Ag/AgCl);0.1 M phosphate buffer (pH 7.5); measured with SPCE/RhO2/GOx;analysis in a batch arrangement; concentration of glutaraldehyde:1–0.625%, 2–1.25%, 3–2.5%, 4—immobilization with vapour ofglutaraldehyde.

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Figure 3: Immobilization using cellulose acetate. Measurementcondition: input potential −0.2 V (versus Ag/AgCl); 0.1 M phos-phate buffer (pH 7.5); measured with SPCE/RhO2/GOx; analysis ina batch arrangement; concentration of cellulose acetate in acetone:1–1.5% and 2–0.5%.

3.1.3. Immobilization Using Cellulose Acetate. For this study,solutions at concentration of 0.05%, 0.5%, 1.5%, and 3% ofcellulose acetate in acetone were used. The first disadvantageof this method of biosensor preparation is the need to usethe relatively volatile acetone, which vaporized very quicklywhile being pipetted and spread onto the electrodes surface.This resulted in an uneven distribution of the celluloseacetate layer. Acetone further dissolved the binder in carbonink (of a resin type), which caused partial washing of theelectrode.

Likewise in Nafion, the concentration of 0.05% was notsufficient to entrap properly the enzyme and no response toglucose was thus observed. By contrast, at the concentrationof 3% the response to glucose was observed only for lowconcentrations of glucose up to 50 mg L−1; there was noincrease in response above this concentration. The crucial

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Figure 4: Immobilization using electropolymerization with pyr-role. Measurement condition: input potential −0.2 V (versusAg/AgCl); 0.1 M phosphate buffer (pH 7.5); measured withSPCE/RhO2/GOx; analysis in a batch arrangement; time of elec-tropolymerization 1–0.5 min, 2–1 min, 3–0.25 min, and 4–2.5 min.

disadvantage of this method, however, is its relatively longresponse time of 240 s. Another problem is the existence ofa very narrow interval for usable concentrations of celluloseacetate for the enzyme immobilization within a range of1% (Figure 3)—compared, for example, to Nafion with thechoice of 0.5–5.0%.

3.1.4. Immobilization by Pyrrole Electropolymerization. Pyr-role was polymerized on the electrodes surface for periodsof 0.25, 0.5, 1.0, and 2.5 min (Figure 4). The results showthat for the period of 2.5 min, a very strong polypyrrolemembrane is created which causes a slow transport ofglucose molecules to the GOx enzyme and subsequently,the transport of H2O2 to the electrodes surface. This isevidenced by lower responses to glucose and longer responsetime. Another situation occurs for the period of 0.25 min.The enzyme is not sufficiently entrapped in this case, and,therefore, it is partially washed into the solution, whichis again shown by very low responses. The best resultwas achieved using electropolymerization of pyrrole lasting0.5 min. The responses are the highest here and the responsetime of 35 s is also acceptable.

3.1.5. Immobilization Using Electropolymerization with m-Phenylenediamine. The m-phenylenediamine was polymer-ized onto the electrodes surface for periods of 0.5, 1, 5, 10,and 20 min. Furthermore, GOx was incorporated directlyinto the phosphate buffer solution with m-phenylenediamineand the electropolymerization was performed for 5 min.Figure 5 shows that the best response to glucose was achievedwhen the m-phenylenediamine was electropolymerized for1 min. Shorter times were insufficient to entrap the enzymeinto the polymeric membrane. Longer times, however,created a thicker membrane which slowed the processes,transporting the analyte to the enzyme and the metaboliteto the electrodes surface, which was similar to the situationfor pyrrole.

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5

6

Concentration (mg L−1)

Figure 5: Immobilization using electropolymerization with m-phenylenediamine. Measurement condition: input potential−0.2 V(versus Ag/AgCl); 0.1 M phosphate buffer (pH 7.5); measuredwith SPCE/RhO2/GOx; analysis in a batch arrangement; time ofelectropolymerization 1–1 min, 2–10 min, 3–5 min, 4–0.5 min, 5–5 min with addition of 1 mg GOx to electropolymerization mixture,and 6–20 min.

In the case of electropolymerization with m-phenylene-diamine together with the enzyme directly in a phosphatebuffer solution, the sensitivity was almost the lowest andthe response time was relatively long (100 s). However thedynamic range was greatest in this case

3.1.6. Comparison of the Immobilization Techniques. Table 1compares the various methods of immobilization. Ideally, abiosensor should have the shortest-possible response time,the largest dynamic range of concentrations, and highlysensitive responses to the given analyte. In practice, however,it is necessary to compromise and to favour one parameterover another according to the determination requirements.Since all the immobilizations listed show rather sensitiveresponses to glucose, the decisive criteria are response timeand dynamic range. The most appropriate method cantherefore, be considered the electropolymerization with m-phenylenediamine, which was used for immobilization of theglucose dehydrogenase enzyme.

3.2. Determination of Glucose by Glucose Dehydrogenase.From the methods of immobilization examined, that oneusing electropolymerization with m-phenylenediamine wasselected for preparation of the given biosensor. Whenworking with dehydrogenases, great emphasis must be givento correctly executing the immobilization, because not onlythe enzymes but also their cofactors (NAD+ or NADP+)are immobilized. These cofactors are soluble in aqueoussolutions and thus they wash rapidly into the solution, andespecially when using flow analysis. The entire procedurefor electrode preparation is described in Section2.4.6 (whileSection 3.1.5 stated that the best response to glucose wasreached where m-phenylenediamine was electropolymerizedfor 1 min, an electropolymerization time of 5 min was chosenhere due to better entrapment of the NAD+ cofactor.)

−0.1

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

−0.4 −0.2 0 0.2 0.4 0.6 0.8

Potential (V)

I(µ

A)

Figure 6: Effect of potential on the biosensor response. Measure-ment condition: glucose concentration 1000 mg L−1; batch volume200 μL; pH of the supporting electrolyte 7.5; flow rate 0.2 mL min−1;measured on SPCE/RhO2/GDH; analysis in a flow arrangement.

−0.1

−0.08

−0.06

−0.04

−0.02

0

0.02

0.04

−0.4 −0.2 0 0.2 0.4 0.6 0.8

Potential (V)

I(µ

A)

1

2

Figure 7: Effect of interferents on the biosensor response. Mea-surement condition: ascorbic acid, and uric acid concentrations10 mg L−1; batch volume 200 μL; pH of the supporting electrolyte7.5; flow rate 0.2 mL min−1; measured on SPCE/RhO2/GDH;analysis in a flow arrangement; 1—ascorbic acid, 2—uric acid.

3.2.1. Effect of the Potential on the Biosensor Response. Inputpotential is one of the most important parameters in theamperometric determination of analytes since its choiceaffects the selectivity of the given biosensor. Figure 6 showsthe dependence of response on the operating potential(dependence of the peak size on the potential was observedin the range of −0.3 to +0.6 V versus Ag/AgCl in 0.15 Vintervals). As is visible there, oxidation starts at around+0.15 V and the response increases with the increasingpotential. Oxidation is also observed in the vicinity of−0.3 V,but this response is very low and, therefore, unsuitable fordetermination of glucose. In the range of −0.2 V to +0.1 V,the biosensor records no catalytic activity. As this shows, themost appropriate area for determination of glucose is in therange of +0.15 to +0.6 V (taking into consideration the effectof interferents).

3.2.2. Effect of Interferents on the Biosensor Response. Therecan be many interfering substances in the samples (such as

6 Enzyme Research

Table 1: Comparison of individual immobilization methods and their parameters.

Type of immobilizationResponse∗

(μA)Linearity(mg L−1)

Response time(s)

Nafion (0.5%) 1.991 10–200 120

Glutaraldehyde vapors 1.054 10–200 30

Cellulose acetate (1.5%) 1.831 10–200 240

Pyrrole (0.5 min) 1.255 50–250 35

m-phenylenediamine (1 min) 1.260 10–500 25∗

Measured at glucose concentration of 200 mg L−1.

Table 2: Determination of glucose in real sample using SPCE/RhO2/GDH.

Proposed method Reference method

Sample n x ± R [%] n x ± R [%] u ucrit

Honey 4 33.84± 5.63 4 33.97± 2.16 0.017 0.406

Syrup 4 26.06± 4.70 4 24.31± 4.74 0.185 0.406

n: number of measurements; x: arithmetic mean; R: range; ucrit and u: critical and calculated values of Lord’s test (selected probability—95%).

0

0.04

0.08

0.12

0.16

0.2

0 0.2 0.4 0.6 0.8 1 1.2

Flow rate (mL min−1)

I(µ

A)

Figure 8: Effect of flow rate on the biosensor response. Mea-surement condition: glucose concentration 1000 mg L−1; batchvolume 200 μL; input potential 0.45 V (versus Ag/AgCl); pH of thesupporting electrolyte 7.5; measured on SPCE/RhO2/GDH; analysisin a flow arrangement.

blood and food). The most important interferents includeascorbic acid, uric acid and paracetamol. It has beenobserved that all of these are electroactive at the appliedpotential of +0.5 V, but, in the potential window of −0.2 to+0.45 V, their responses are negligible (Figure 7). For thisreason, potentials in the given range were chosen for furtherwork.

3.2.3. Effect of Flow Rate and pH on the Biosensor Response.Flow rate also belongs among the very important parametersthat must be optimized. It was done in the range of0.1 mL min−1 to 1 mL min−1. Figure 8 shows that the sizeof the response decreases with an increasing flow rate. Thisis due to the fact that if the flow rate is too high, theNADH+ on the electrode is not fast enough to react. Onthe other hand, at low flow rates, the biosensors response

0

0.01

0.02

0.03

0.04

0 1000 2000 3000 4000 5000 6000

Concentration (mg L−1)

I(µ

A)

Figure 9: Biosensor response to glucose at input potential +0.35 V.Measurement condition: batch volume 200 μL; input potential0.35 V (versus Ag/AgCl); pH of the supporting electrolyte 7.5;flow rate 0.5 mL min−1; measured on SPCE/RhO2/GDH; analysisin a flow arrangement; regression equation: y = 5.00 × 10−6x +0.0129,R2 = 0.991.

is unstable (decrease of the response by 20% over threedeterminations). This response instability was probablycaused by passivation of the electrode’s surface. For thisreason, a flow rate ranging from 0.4 to 0.6 mL min−1 seemedideal. For other measurements, the flow rate of 0.5 mL min−1

was chosen. That seems to be a good compromise betweenbuffer consumption, response stability, and speed of theexperiment.

Optimization of pH was carried out in the range of 5 to9. Stable responses were observed at all measured pH valuesand the highest was achieved at pH 8, where at the same timethe maximum enzyme activity is seen. For further work, thepH of 7.5 was chosen because the given pH is close to thephysiological pH and that is optimal for the determinationof biological substances in food and especially in clinicalsamples.

Enzyme Research 7

3.2.4. Biosensor Response to Glucose. Calibration depen-dences were measured at two different potentials (+0.35 Vand +0.45 V). At the potential of +0.35 V, the biosensorshowed lower responses, but the dynamic range was greaterthan at the potential of +0.45 V. A big advantage is that atthe input potential of +0.35 V, the effects of interferents aremuch more suppressed. The proposed biosensor retained itsactivity after more than 50 injections. No loss of the originalsignal was achieved after 1 month, when stored at 6◦C in therefrigerator.

3.3. Real Samples. Honey and syrup samples were usedas real analytes. Measurement was performed under theseoptimized conditions: input potential +0.35 V; batch vol-ume 200 μL; 0.1 M phosphate buffer pH 7.5; flow rate0.5 mL min−1in a three-electrode arrangement in the pres-ence of SPCE/RhO2/GDH, where the enzyme was entrappedby m-phenylenediamine. The determined concentrations areshown in Table 2.

The amperometric determination with SPCE/RhO2/GOxwas used as a reference method (carbon printed electrodemodified by glucose oxidase and rhodium oxide—theenzyme immobilized by Nafion). Measurement conditions:−0.2 V (versus Ag/AgCl); phosphate buffer pH 7.5; flow rate0.2 mL min−1; batch volume 50 μL.

4. Conclusion

A biosensor containing rhodium dioxide and glucose dehy-drogenase enzyme was prepared using the screen-printingtechnique. Various methods of enzyme immobilization weretested, among which m-phenylenediamine electropolymer-ization proved the best. It excelled with its response time,sensitivity, and signal stability. The enzyme biosensor wasoptimized and tested in model glucose samples and alsoapplied to analyze real samples (honey, syrup).

Good results in the determination of glucose in realsamples indicate, among other things, that the biosensor wasnot affected by any complicated sample matrix (ascorbic acidand other oxidizable substances) and has prospects for usealso for similar applications in the food industry and clinicalpractice.

Acknowledgments

This paper was supported by the Ministry of Education,Youth, and Sports of the Czech Republic (project MSM0021627502) and the Czech Science Foundation (project203/08/1536).

References

[1] M. Nistor and E. Csoregi, “Biosensors for food analysis,”in Encyclopedia of Sensors, C. A. Grimes, C. E. Dickey, andM. V. Pishko, Eds., vol. 1, pp. 353–369, American ScientificPublishers, Stevenson Ranch, CA, USA, 2006.

[2] A. P. F. Turner, I. Karube, and G. S. Wilson, Biosensors:Fundamentals and Applications, Oxford University Press, NewYork, NY, USA, 1987.

[3] G. J. Moody, G. S. Sangbera, and J. D. R. Thomas, “Chemicallyimmobilised bi-enzyme electrodes in the redox mediatedmode for the low flow injection analysis of glucose andhypoxanthine,” Analyst, vol. 112, no. 1, pp. 65–70, 1987.

[4] H. Okuma, H. Takahashi, S. Sekimukai, K. Kawahara, andR. Akahoshi, “Mediated amperometric biosensor for hypox-anthine based on a hydroxymethylferrocene-modified carbonpaste electrode,” Analytica Chimica Acta, vol. 244, no. 2, pp.161–164, 1991.

[5] F. Ricci, A. Amine, D. Moscone, and G. Palleschi, “A probefor NADH and HO amperometric detection at low appliedpotential for oxidase and dehydrogenase based biosensorapplications,” Biosensors and Bioelectronics, vol. 22, no. 6, pp.854–862, 2007.

[6] M. C. Rodrıguez and G. A. Rivas, “An enzymatic glucosebiosensor based on the codeposition of rhodium, iridium, andglucose oxidase onto a glassy carbon transducer,” AnalyticalLetters, vol. 34, no. 11, pp. 1829–1840, 2001.

[7] P. Kotzian, P. Brazdilova, K. Kalcher, and K. Vytras, “Determi-nation of hydrogen peroxide, glucose and hypoxanthine using(bio)sensors based on ruthenium dioxide-modified screen-printed electrodes,” Analytical Letters, vol. 38, no. 7, pp. 1099–1113, 2005.

[8] J. Razumiene, A. Vilkanauskyte, V. Gureviciene et al., “Newbioorganometallic ferrocene derivatives as efficient mediatorsfor glucose and ethanol biosensors on PQQ-dependent dehy-drogenases,” Journal of Organometallic Chemistry, vol. 668, no.1-2, pp. 83–90, 2003.

[9] B. Prieto-Simon, J. Macanas, M. Munoz, and E. Fabregas,“Evaluation of different mediator-modified screen-printedelectrodes used in a flow system as amperometric sensors forNADH,” Talanta, vol. 71, no. 5, pp. 2102–2107, 2007.

[10] A. CH. Pappas, M. I. Prodromidis, and M. I. Karayannis,“Flow monitoring of NADH consumption in bioassays basedon packed-bed reactors bearing NAD-dependent dehydroge-nases: determination of acetaldehyde using alcohol dehydro-genase,” Analytica Chimica Acta, vol. 467, no. 1-2, pp. 225–232,2002.

[11] H.-Y Chen and J.-J Xu, “Amperometric enzyme biosensors,”in Encyclopedia of Sensors, C. A. Grimes, C. E. Dickey, andM. V. Pishko, Eds., vol. 1, pp. 145–167, American ScientificPublishers, Stevenson Ranch, CA, USA, 2006.

[12] S. A. Rothwell, C. P. McMahon, and R. D. O’Neill, “Effectsof polymerization potential on the permselectivity of poly(o-phenylenediamine) coatings deposited on Pt-Ir electrodes forbiosensor applications,” Electrochimica Acta, vol. 55, no. 3, pp.1051–1060, 2010.

[13] Y. Sha, Q. Gao, B. Qi, and X. Yang, “Electropolymerizationof azure B on a screen-printed carbon electrode and itsapplication to the determination of NADH in a flow injectionanalysis system,” Microchimica Acta, vol. 148, no. 3-4, pp. 335–341, 2004.

[14] X. G. Li, M. R. Huang, W. Duan, and Y. L. Yang, “Novel mul-tifunctional polymers from aromatic diamines by oxidativepolymerizations,” Chemical Reviews, vol. 102, no. 9, pp. 2925–3030, 2002.

[15] P. Kotzian, P. Brazdilova, S. Rezkova, K. Kalcher, and K. Vytras,“Amperometric glucose biosensor based on rhodium dioxide-modified carbon ink,” Electroanalysis, vol. 18, no. 15, pp.1499–1504, 2006.

SAGE-Hindawi Access to ResearchEnzyme ResearchVolume 2010, Article ID 415949, 5 pagesdoi:10.4061/2010/415949

Research Article

Preparation of Antioxidant Enzymatic Hydrolysates fromHoneybee-Collected Pollen Using Plant Enzymes

Margarita D. Marinova and Bozhidar P. Tchorbanov

Institute of Organic Chemistry with Centre of Phytochemistry, Bulgarian Academy of Sciences, Acad. G. Bonchev Street,Building 9, BG-1113 Sofia, Bulgaria

Correspondence should be addressed to Margarita D. Marinova, m.d.marinova@abv.bg

Received 14 June 2010; Accepted 16 December 2010

Academic Editor: A. Pandey

Copyright © 2010 M. D. Marinova and B. P. Tchorbanov. This is an open access article distributed under the Creative CommonsAttribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work isproperly cited.

Enzymatic hydrolysates of honeybee-collected pollen were prepared using food-grade proteinase and aminopeptidases entirelyof plant origin. Bromelain from pineapple stem was applied (8 mAU/g substrate) in the first hydrolysis stage. Aminopeptidase(0.05 U/g substrate) and proline iminopeptidase (0.03 U/g substrate) from cabbage leaves (Brassica oleracea var. capitata), andaminopeptidase (0.2 U/g substrate) from chick-pea cotyledons (Cicer arietinum L.) were involved in the additional hydrolysisof the peptide mixtures. The degree of hydrolysis (DH), total phenolic contents, and protein contents of these hydrolysateswere as follows: DH (about 20–28%), total phenolics (15.3–27.2 μg/mg sample powder), and proteins (162.7–242.8 μg/mgsample powder), respectively. The hydrolysates possessed high antiradical scavenging activity determined with DPPH (42–46%inhibition). The prepared hydrolysates of bee-collected flower pollen may be regarded as effective natural and functional dietaryfood supplements due to their remarkable content of polyphenol substances and significant radical-scavenging capacity withspecial regard to their nutritional-physiological implications.

1. Introduction

Natural products and preparations for food and nutritionalsupplementation or dietary purposes have gained increasedattention in recent years. Among them, honeybee-derivedapicultural products, such as pollen, have been appliedfor centuries in alternative medicine as well as in fooddiets and supplementary nutrition due to their nutritionaland physiological properties. Each pollen has its ownspecificity, mainly linked to the floral species or culti-vars. Bee-collected pollens contain nutritionally essentialsubstances including carbohydrates, proteins, amino acids,lipids, vitamins, mineral substances, and trace elements, butalso significant amounts of polyphenol substances mainlyflavonoids which, furthermore, are regarded as principalindicating ingredient substances of pollen and can beused for setting up quality standards in relation to theirnutritional-physiological properties and for quality controlof commercially distributed pollen preparations [1–4]. Itis well known that polyphenols are responsible for the

antioxidative and radical scavenging activity of plant food[5, 6]. An antioxidant defense system protects cells fromthe injurious effects of free radicals. Furthermore, thebiological, biochemical, physiological, pharmaceutical, andmedicinal properties of polyphenol compounds have beenextensively studied and have been reviewed by Rice-Evanset al. [7] in regard to their free-radical scavenging activityand multiple biological activities including vasodilatory,anticarcinogenic, anti-inflammatory, antibacterial, immune-stimulating, antiallergic, antiviral, and estrogenic effects, aswell as being inhibitors of specific enzymes.

On the basis of these reports, we have prepared water-soluble fractions from honeybee-collected pollen and inves-tigated their functional properties. As a result, high free-radical scavenging activities against the DPPH free radicalwere exhibited. These results are comparable to the resultsreported for the antioxidant activities in red grapes (Vitisvinifera, L.) extracts [8, 9]. Moreover, we showed thatenzymatic hydrolysates from honeybee-collected pollenspossessed even higher antioxidative properties. In the present

2 Enzyme Research

study, our aim was to prepare enzymatic hydrolysatesfrom honeybee-collected pollens using plant proteinase andaminopeptidases and, then, to investigate the antioxidantactivities in these peptide samples.

2. Materials and Methods

2.1. Materials. The pollen loads were collected in 2009 fromthe Ajtos area by honeybee colonies (Apis mellifera) settled inhives with bottom-fitted pollen traps. The aminopeptidaseand proline iminopeptidase from cabbage leaves (Bras-sica oleracea var. capitata) and the aminopeptidase fromchick-pea cotyledons (Cicer arietinum L.) were purified asdescribed by Marinova et al. [10, 11]. Bromelain frompineapple stems (EC 3.4.22.4, 2 mAnsonU/mg), L-aminoacid p-nitroanilides, Folin-Ciocalteu, and 1,1-diphenyl-2-picrylhydrazyl (DPPH), were purchased from Sigma-AldrichCo. (St. Louis, USA).

2.2. Methods

2.2.1. Preparation of Enzymatic Hydrolysates from Honeybee-Collected Pollen. Honeybee-collected pollens (28% protein)were added, suspended in 5 volumes of distilled water, andhomogenized (Ultra-Turax, IKA-Werke, Germany), and pHof the suspension was adjusted at 7.0 using NH4OH. Thedigestion was started by addition of 8 mAU/g bromelain at37◦C. After 4 hours, hydrolysis was stopped by boiling ina microwave for 2 minutes. The additional hydrolysis wascarried out by adding aminopeptidase (0.05 units/g sub-strate) and proline iminopeptidase (0.03 units/g substrate)from cabbage leaves as well as aminopeptidase from chick-pea cotyledons (0.2 units/g substrate) and incubating for twohours at 37◦C and pH 7.5 with constant stirring. Hydrolysiswas stopped by boiling in a microwave for 2 minutes.The obtained hydrolysates were centrifuged at 6000× g for30 min at 5◦C (MLW K24 D, Germany) to remove theresidue. The supernatant fractions were collected and freeze-dried.

2.2.2. Assays of Enzymes’ Activities, Total Nitrogen, TotalProtein, and Total Phenolic Compounds. Aminopeptidases’activities were determined using L-leucine-p-nitroanilidesas substrate [12]. After incubation for 10 min at 30◦C in0.05 M sodium phosphate buffer (pH 7.2–7.5), the liberatedp-nitroaniline was measured at 410 nm on a spectropho-tometer (UV-VIS Spectrophotometer, Shimadzu 1240). Theiminopeptidase activity was assayed spectrophotometricallyat 410 nm against L-proline-p-nitroanilide (Pro-p-NA) in0.1 M Tris/HCl buffer (pH 8.0) for 20 min at 30◦C [13]. Oneunit of enzyme activity was defined as the amount of enzymereleasing 1 μmol of p-nitroaniline per minute.

The total protein content of the honeybee-collectedpollen was determined by the method of Kjeldahl using theequation: N × 6.25, where N is the total Kjeldahl nitrogenmultiplied by a factor to arrive at protein content [14]. Theprotein concentration of the samples after hydrolysis wasmeasured according to the method of Lowry et al. [15],

using bovine serum albumin as standard. The total phenoliccontent was determined by the Folin-Ciocalteu colorimetricmethod using catechin as standard, and the absorbance wasmeasured at 760 nm [16].

2.2.3. Radical Scavenging Activity. The antiradical power ofbee-collected pollen and pollen hydrolysates was evaluatedin terms of the hydrogen-donating or radical-scavengingability by the DPPH method [17], which is related to theinhibition in the initiation step of free radical processes.DPPH (2,2-diphenyl-1- picrylhydrazyl) is a stable free radicalthat accepts an electron or hydrogen radical to become astable diamagnetic molecule and, accordingly, is reduced inpresence of an antioxidant (AH):

DPPH• + AH −→ DPPH−H + A•· (1)

For the evaluation of the antioxidant activity of specificcompounds or extracts, they are allowed to react with thestable DPPH radical in a methanol solution. In its radicalform, DPPH has a characteristic absorbance at 515 nm,which disappears upon reduction by H gained from anantioxidant compound.

For the test, appropriate methanol stock solutions of thepollen preparations (500 mg/L) and DPPH (6× 10−5 mol/L)were prepared. Immediately after adding 0.3 mL of thepollen extract solution to 2.7 mL of the DPPH solution, thereduction of the DPPH-radical was measured by monitoringcontinuously the decrease of absorption at 515 nm in thedark until stable absorption values were obtained (30 min).The antiradical activity was determined in terms of PIvalues (% inhibition) which was calculated by the ratio ofthe decrease of absorption of the DPPH-pollen extract testsolution after a 30-minute reaction time (stable phase) to theabsorption value of the reference sample where an equivalentvolume of methanol was added, as defined according to theformula:

PI (% inhibition) =[A0 − At

A0

]× 100, (2)

where A0 is the absorbency of the DPPH-methanol solution(reference) and At is the absorbency of the DPPH-pollenextract solution after 30 min of reaction time.

2.2.4. SDS-Polyacrylamide Gel Electrophoresis. Sodium dode-cyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE)was performed in 15% polyacrylamide gel using Tris-glycine buffer, pH 8.3, according to Laemmli [18]. Rabbitmuscle myosin (205 kDa), β-galactosidase (116 kDa), rabbitmuscle phosphorylase b (97 kDa), bovine serum albu-min (66 kDa), lactate dehydrogenase (36.5 kDa), carbonicanhydrase (29 kDa), trypsin inhibitor (20 kDa), lysozyme(14 kDa), aprotinin (6.1 kDa), insulin a (3.4 kDa), andinsulin b (2.4 kD) were used as molecular weight markerproteins. The gel was visualized by silver staining [19].

2.2.5. Determination of the Degree of Hydrolysis (DH) andAmino Acid Composition. The degree of hydrolysis was

Enzyme Research 3

Table 1: The contents of protein and total phenolic components of enzymatic hydrolysates from honeybee-collected pollen.

Sample Protein (μg/mg sample) Total phenols (μg/mg sample) PI-value (%)

Bee-pollen 162.7± 0.2 15.3± 0.3 28± 21BH 227.1± 0.3 21.5± 0.6 40± 12APH1 238.8± 0.5 25.6± 0.7 44± 23APH2 230.5± 0.6 24.1± 0.9 42± 24APH3 242.8± 0.5 27.2± 0.6 46± 1

1BH bromelain hydrolysate

2APH1 cabbage aminopeptidase and proline iminopeptidase hydrolysate3APH2 chick-pea aminopeptidase hydrolysate4APH3 cabbage and chick-pea aminopeptidases hydrolysate

determined using 2,4,6-trinitrobenzenesulfonic acid [20]. Asample solution (0.25 mL) is mixed with 2.0 mL of 0.2 Msodium phosphate buffer (pH 8.2) and 2.0 mL of 0.1%trinitrobenzenesulfonic acid, followed by incubation in thedark for 60 min at 50◦C. The reaction is quenched byadding 4.0 mL of 0.1 N HCl, and the absorbance is read at340 nm. A 1.5 mM L-leucine solution is used as the standard.Transformation of the measured leucine amino equivalentsto degree of hydrolysis is carried out by means of a standardcurve for each particular protein substrate.

The amino acid composition was determined after50 min of hydrolysis at 165◦C with 6 N HCl, and the analysiswas performed on HPLC Nova-Pak C18 (3.9× 150 mm,4 μm, Waters). The mobile phase consisted of eluent A(prepared from Waters AccQ·Tag Eluent A concentrate, byadding 200 mL of concentrate to 2 L of Milli-Q water andmixing), eluent B (acetonitrile, HPLC grade), and eluent C(Milli-Q water). The following conditions were used: lineargradient of 100–0% eluent A, 0–60% eluent B, and 0–40%eluent C in 30 min and then isocratic 100% of eluent A for20 min with a flow rate of 1 mL/min.

3. Results and Discussion

3.1. The Total Phenolic Contents and Protein Contents ofEnzymatic Hydrolysates from Bee Pollen. The enzymatichydrolysates from bee pollen were digested and preparedusing plant proteinase bromelain, and aminopeptidases fromcabbage leaves and chick-pea cotyledons. SDS-PAGE analysisindicated that the pollen was perfectly digested by theseenzymes (Figure 1). The degrees of hydrolysis of the bee-pollen hydrolysates were as follows: about 20% for thebromelain hydrolysate (BH), 26% for the cabbage aminopep-tidase and proline iminopeptidase hydrolysate (APH1), 24%for the chick-pea aminopeptidase hydrolysate (APH2), and28% for the hydrolysate obtained by the combinationof aminopeptidases from cabbage and chick-pea (APH3).Total phenolic contents of these hydrolysates were as fol-lows: 21.5 μg/mg sample (BH), 25.6 μg/mg sample (APH1),24.1 μg/mg sample (APH2), and 27.2 μg/mg sample (APH3),respectively (Table 1). On the other hand, the protein con-tents of these hydrolysates were as follows: 227.1 μg/mg sam-ple (BH), 238.8 μg/mg sample (APH1), 230.5 μg/mg sam-ple (APH2), and 242.8 μg/mg sample (APH3), respectively

1 2 3 4 5

2.5

3.4

6.1

14

20

29

36.55597

116220

Figure 1: SDS-polyacrylamide gel electrophoresis of molecu-lar weight markers and enzymatic hydrolysates from honeybee-collected pollen. (1) Molecular weight markers; (2) aminopepti-dases hydrolysate (cabbage and chick-pea); (3) aminopeptidase andproline iminopeptidase hydrolysate (cabbage); (4) aminopeptidasehydrolysate (chick-pea); (5) bromelain hydrolysate.

(Table 1). It suggests that the protein contents correlatedclosely with the contents of total phenolic components.

3.2. DPPH Radical Scavenging Ability of EnzymaticHydrolysates from Pollen. Although native bee-collectedpollen water extract shows considerable antiradical activity(PI= 28% inhibition), the reduction of the DPPH radicalwas significantly increased by applying the obtained pollenhydrolysates (PI= 42–46% inhibition) as is shown inTable 1, which indicate an elevated free-radical scavengingefficiency of the pollen hydrolysates. The highest degreeof radical scavenging capacity was assessed in the APH3

(PI= 46% inhibition) which, correspondingly, also has thehighest concentration of polyphenol substances (27.2 μg/mgsample powder). For this reason, it can be assumed thatthere is a general correlation between the content of totalpolyphenolics and the free-radical scavenging capacity of thepollen preparations.

Although, in comparison, tests with equivalent amountof the synthetic antioxidant gallic acid shows higher PI valuesof approximately 90%, it must be taken into consideration

4 Enzyme Research

Table 2: The amino acids composition of honeybee-collectedpollen and bee-pollen hydrolysate obtained by the combination ofaminopeptidases from cabbage and chick-pea (∗).

Amino acids C (%) C∗ (%)

Asp 7.5 6.6

Hyp 1.5 1.5

Glu 8.6 7.8

Ser 5.8 5.9

Gly 9.8 10.5

His + Thr 5.1 5.0

Ala 8.0 7.9

Arg 4.6 1.0

Pro 17.1 22.2

Tyr 1.7 1.4

Val 6.7 6.7

Met 2.2 2.1

Ile 5.3 5.1

Leu 7.9 7.5

Lys 3.4 2.6

Phe 8.1 5.5

that pollen extract represents a concentrated nature-derivedmixture of different active polyphenol compounds.

The results of the amino acids’ composition for the bee-pollen extract and the pollen hydrolysate obtained by combi-nation of aminopeptidases from cabbage and chick-pea areshown in Table 2. Relatively high content of hydrophobicamino acids Pro, Phe, and Gly is characteristic for the bee-pollen extract and the APH3. The amounts of these aminoacids in bee-pollen hydrolysate after a 6-hour hydrolysis areapproximately 22, 5.5 and 10.5%, respectively of the totalcontent.

The honeybee products are considered to be abundantsources of antioxidants. In honey, royal jelly, propolis, andbee pollens high antioxidant activity was found [21]. Inbee-collected pollen water extracts high radical-scavengingactivity, activity against superoxide anion, and hydroxylradical-scavenging activity were reported [22–25].

Antioxidative ability of pollen seems to be due tophenolic compounds. In the present investigations, a veryhigh antioxidant activity, expressed as radical-scavengingactivity corresponded to high levels of total phenols, wasfound in water-soluble extract and in plant proteinase andaminopeptidases hydrolysates.

4. Conclusion

The use of honeybee-collected pollens as an alternativemedicine is increasing due to their biologically active proper-ties that make them attractive as a source of essential aminoacids, vitamins, minerals, and antioxidants in human diets.The useful components from honeybee-collected pollen canbe fully digested using the food-grade enzymes such asbromelain, cabbage aminopeptidase and proline iminopep-tidase, and chick-pea aminopeptidase, although it is not easy

to digest honeybee-collected pollen with a hard cell wall. Inthis process, consumer demand of honeybee-collected pollenfor natural foods with medicinal effects such as antioxidativeactivity is increasing.

In conclusion, pollen extracts represent a concentratednature-derived mixture of different active polyphenol com-pounds which, according to practical applications as abioactive diet component, are usually applied and consumedin higher amounts than the pure synthetic antioxidant foodadditives.

Acknowledgment

The authors thank the National Foundation for ScientificResearch for financial support of Project TK-X 1608.

References

[1] B. M. Talpay, Der Pollen, Eigenverlag Institut fur Honig-forschung, Bremen, Germany, 1981.

[2] R. G. Stanley and H. F. Linskens, Pollen: Biologie, Biochemie,Gewinnung und Verwendung, Urs Freund, Greifenberg, Ger-many, 1985.

[3] G. Kroyer, “Flavonoids and phytosterols as bioactive sub-stances in dietary applied pollen products,” in Proceedings ofEuro Food Chem X: Functional Foods—A New Challenge forthe Food Chemists, pp. 102–108, Publishing Company of TUB,Budapest, Hungary, 1999.

[4] J. Kanner, E. Frankel, R. Granit, B. German, and J. E.Kinsella, “Natural antioxidants in grapes and wines,” Journalof Agricultural and Food Chemistry, vol. 42, no. 1, pp. 64–69,1994.

[5] N. Salah, N. J. Miller, G. Paganga, L. Tijburg, G. P. Bolwell,and C. Rice-Evans, “Polyphenolic flavanols as scavengers ofaqueous phase radicals and as chain-breaking antioxidants,”Archives of Biochemistry and Biophysics, vol. 322, no. 2, pp.339–346, 1995.

[6] J. A. Vinson and B. A. Hontz, “Phenol antioxidant index:comparative antioxidant effectiveness of red and white wines,”Journal of Agricultural and Food Chemistry, vol. 43, no. 2, pp.401–403, 1995.

[7] C. A. Rice-Evans, N. J. Miller, and G. Paganga, “Structure-antioxidant activity relationships of flavonoids and phenolicacids,” Free Radical Biology and Medicine, vol. 20, no. 7, pp.933–956, 1996.

[8] V. Briedis, V. Povilaityte, S. Kazlauskas, and P. R. Venskutonis,“Polyphenols and anthocyanins in fruits, grapes juices andwines, and evaluation of their antioxidant activityPolifenoliuir antocianinu kiekis vynuogese, vynuogiu sultyse ir raudon-uose vynuose bei ju antioksidacinio aktyvumo ivertinimas,”Medicina, vol. 39, pp. 104–112, 2003.

[9] C. H. Chen, M. C. Wu, C. Y. Hou, C. M. Jiang, C. M. Huang,and Y. T. Wang, “Effect of phenolic acid on antioxidant activityof wine and inhibition of pectin methyl esterase,” Journal of theInstitute of Brewing, vol. 115, no. 4, pp. 328–333, 2009.

[10] M. Marinova, A. Dolashki, F. Altenberend, S. Stevanovic,W. Voelter, and B. Tchorbanov, “Characterization of anaminopeptidase and a proline iminopeptidase from cabbageleaves,” Zeitschrift fur Naturforschung, vol. 63, no. 1-2, pp. 105–112, 2008.

[11] M. Marinova, A. Dolashki, F. Altenberend, S. Stevanovic,W. Voelter, and B. Tchorbanov, “Purification and character-ization of L-phenylalanine aminopeptidase from chick-pea

Enzyme Research 5

cotyledons (Cicer arietinum L.),” Protein and Peptide Letters,vol. 16, no. 2, pp. 207–212, 2009.

[12] M. J. Chrispeels and D. Boulter, “Control of storage proteinmetabolism in the cotyledons of germinating mung beans: roleof endopeptidase,” Plant Physiology, vol. 55, pp. 1031–1037,1975.

[13] T. Yoshimoto and D. Tsuru, “Proline iminopeptidase fromBacillus coagulans: purification and enzymatic properties,”Journal of Biochemistry, vol. 97, no. 5, pp. 1477–1485, 1985.

[14] P. L. Kirk, “Kjeldahl method for total nitrogen,” AnalyticalChemistry, vol. 22, no. 2, pp. 354–358, 1950.

[15] O. H. Lowry, N. J. Rosebrough, A. L. Farr, and R. J. Randall,“Protein measurement with the Folin phenol reagent,” TheJournal of Biological Chemistry, vol. 193, no. 1, pp. 265–275,1951.

[16] K. Slinkard and V. L. Singleton, “Total phenol analysis,”American Journal of Enology and Viticulture, vol. 28, pp. 49–55, 1977.

[17] W. Brand-Williams, M. E. Cuvelier, and C. Berset, “Use ofa free radical method to evaluate antioxidant activity,” FoodScience and Technology, vol. 28, no. 1, pp. 25–30, 1995.

[18] U. K. Laemmli, “Cleavage of structural proteins during theassembly of the head of bacteriophage T4,” Nature, vol. 227,no. 5259, pp. 680–685, 1970.

[19] D. W. Sammmons, L. D. Adams, and E. E. Nishizawa,“Ultrasensitive silver-based color staining of polypeptides inpolyacrylamide gels,” Electrophoresis, vol. 2, pp. 135–141, 1980.

[20] J. Adler-Nissen, “Determination of the degree of hydrolysisof food protein hydrolysates by trinitrobenzenesulfonic acid,”Journal of Agricultural and Food Chemistry, vol. 27, no. 6, pp.1256–1262, 1979.

[21] T. Nagai, M. Sakai, R. Inoue, H. Inoue, and N. Suzuki,“Antioxidative activities of some commercially honeys, royaljelly, and propolis,” Food Chemistry, vol. 75, no. 2, pp. 237–240, 2001.

[22] M. G. Campos, R. F. Webby, and K. R. Markham, “The uniqueoccurrence of the flavone aglycone tricetin in Myrtaceaepollen,” Zeitschrift fur Naturforschung C, vol. 57, no. 9-10, pp.944–946, 2002.

[23] M. G. Campos, R. F. Webby, K. R. Markham, K. A. Mitchell,and A. P. Da Cunha, “Age-induced diminution of free radicalscavenging capacity in bee pollens and the contributionof constituent flavonoids,” Journal of Agricultural and FoodChemistry, vol. 51, no. 3, pp. 742–745, 2003.

[24] M. Leja, A. Mareczek, G. Wyzgolik, J. Klepacz-Baniak, and K.Czekonska, “Antioxidative properties of bee pollen in selectedplant species,” Food Chemistry, vol. 100, no. 1, pp. 237–240,2007.

[25] T. Nagai, R. Inoue, N. Suzuki, T. Myoda, and T. Nagashima,“Antioxidative ability in a linoleic acid oxidation system andscavenging abilities against active oxygen species of enzymatichydrolysates from pollen Cistus ladaniferus,” InternationalJournal of Molecular Medicine, vol. 15, no. 2, pp. 259–263,2005.

SAGE-Hindawi Access to ResearchEnzyme ResearchVolume 2010, Article ID 517283, 5 pagesdoi:10.4061/2010/517283

Research Article

Characterization of Activity of a Potential Food-Grade LeucineAminopeptidase from Kiwifruit

A. A. A. Premarathne and David W. M. Leung

School of Biological Sciences, University of Canterbury, Private Bag 4800, Christchurch 8140, New Zealand

Correspondence should be addressed to David W. M. Leung, david.leung@canterbury.ac.nz

Received 14 June 2010; Revised 25 August 2010; Accepted 4 October 2010

Academic Editor: Raffaele Porta

Copyright © 2010 A. A. A. Premarathne and D. W. M. Leung. This is an open access article distributed under the CreativeCommons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided theoriginal work is properly cited.

Aminopeptidase (AP) activity in ripe but firm fruit of Actinidia deliciosa was characterized using L-leucine-p-nitroanilide as asubstrate. The enzyme activity was the highest under alkaline conditions and was thermolabile. EDTA, 1,10-phenanthroline,iodoacetamide, and Zn2+ had inhibitory effect while a low concentration of dithiothreitol (DTT) had stimulatory effect on kiwifruitAP activity. However, DTT was not essential for the enzyme activity. The results obtained indicated that the kiwifruit AP was athiol-dependent metalloprotease. Its activity was the highest in the seeds, followed by the core and pericarp tissues of the fruit. Theelution profile of the AP activity from a DEAE-cellulose column suggested that there were at least two AP isozymes in kiwifruit: oneunadsorbed and one adsorbed fractions. It is concluded that useful food-grade aminopeptidases from kiwifruit could be revealedusing more specific substrates.

1. Introduction

Kiwifruit (Actinidia spp.) is an important commercial cropin New Zealand. The fruit contains a high level of a cysteineendopeptidase called actinidin (E.C. 3.4.22.14) found in thecortex of the fruit [1]. Due to this proteolytic activity ofkiwifruit, it has been used to tenderize meat and preventgelatin-based jelly from setting.

Aminopeptidases (APs), particularly those from micro-bial sources, are important food processing enzymes andare widely used to modify proteins in food [2–4]. Animalwaste products were also investigated as a potential sourceof useful APs [5]. It is also possible that APs from plantscould be of use in the food processing industry [6]. Recently,it has been demonstrated that APs of cabbage leaves orchickpea cotyledons can be used to catalyze the hydrolysis ofpeptide bonds including those of hydrophobic bitter peptidesin soy protein hydrolysates, resulting in the less bitter orbland taste products which have food processing applications[7, 8]. However, for debittering protein hydrolysates or otherfood processing needs an attractive alternative would be touse APs from fruits grown commercially that are normally

consumed fresh such as kiwifruit, which have the merit ofalready being generally regarded safe for the food processingindustry.

Generally, there are many studies on seed aminopep-tidases [9–11] but there is a paucity of information onthe occurrence and characteristics of AP activities in fruits.Importantly, since there is no prior study on AP fromkiwifruit, a prerequisite towards the goal of evaluating useof APs from this fruit for food processing applicationsis an investigation into the occurrence and biochemicalcharacteristics of aminopeptidase (AP) activity of kiwifruit.Here, using L-leucine-p-nitroanilide (L-leu-p-NA) as a sub-strate, localization and some basic biochemical character-istics of AP activity within the fruit of Actinidia deliciosa,and an attempt to partially purify the enzyme that isnormally sufficient for food-grade enzymes are reportedhere.

2. Materials and Methods

2.1. Enzyme Extraction. Ripe but firm kiwifruit (Actinidiadeliciosa cv. Hayward) was obtained from a local supermarket

2 Enzyme Research

in Christchurch, New Zealand. Unless indicated otherwise,the whole kiwifruit was peeled and cut into small piecesbefore enzyme extraction. Kiwifruit tissues were ground ina mortar and pestle while adding 0.1 M of potassium phos-phate buffer pH 8.0 supplemented with 1% (w/v) insolublepolyvinyl polypyrrolidone (PVPP), 5% (v/v) glycerol and3 mM DTT. The ratio of weight of tissue (g) to volumeof extraction buffer (ml) was 2 : 1. The homogenate wasfiltered through 2 layers of synthetic cloth and centrifuged at10,000 × g for 20 min at 4◦C. The supernatant was carefullyremoved and used as crude extract of the whole fruit. Theextraction process was carried out in a cold room or on anice bath.

2.2. Determination of Total Protein Concentration. The pro-tein concentration in extracts was determined based on theCoomassie brilliant blue dye-protein binding principle [12].A protein standard curve was prepared using serial dilutionsof BSA (bovine serum albumin; BDH, England).

2.3. Determination of Aminopeptidase (AP) Activity. Amino-peptidase activity was determined as described below unlessindicated otherwise using L-leucine-p-nitroanilide (L-Leu-p-NA) as a substrate. The substrate solution was preparedby dissolving 20 mg of L-Leu-p-NA (Sigma, St. Louis, USA)in one ml of dimethyl sulfoxide (Sigma, St. Louis, USA)and adjusting the volume to 20 ml with 0.01 M potassiumphosphate buffer (pH 8.0). It was found to be stored better at−20◦C for use later if prepared at pH 8.0 than at higher pH.The reaction mixture contained 0.45 ml of 0.1 M potassiumphosphate buffer at pH 8.0, 0.45 ml substrate solution, and150 μl enzyme extract in Eppendorf tubes kept on ice. Thecontrol tube contained the same reaction mixture except thatthe enzyme extract had previously been boiled for 5 minin a water bath at 100◦C and centrifuged afterwards. Allthe tubes were vortexed, and incubated for 1 h in a waterbath at 37◦C. After the incubation period, they were placedin a water bath at 100◦C for 5 min to stop the enzymereaction. After this, 0.45 ml distilled water was added toall the tubes, vortexed. and then centrifuged for 10 min at10,000 × g at room temperature. The supernatants werecarefully transferred to the cuvettes and the absorbancewas measured at 410 nm. One unit of enzyme activity isdefined as a change in one unit of absorbance per h at37◦C.

2.4. Effect of Temperature on AP Activity. The effect oftemperature on AP activity was determined in three differentexperiments. To find the optimum temperature for theenzyme activity, AP activity in crude extracts of the wholefruit was determined at different incubation temperaturesranging from 25◦C to 70◦C for 1 h. In another experimentto investigate thermal stability, 150 μl of the enzyme extractswere pre-incubated with 0.45 ml of potassium phosphatebuffer (pH 8.0) for 30 min at the above testing temperatures.After preincubation, the substrate was added to initiate theenzyme reaction for AP activity determination at 37◦C for1 h.

2.5. Effect of pH on AP Activity. The effect of pH on APactivity in the crude extracts of fruit was determined byreplacing the potassium phosphate buffer at pH 8.0 in theassay mixture, with the three buffer mixtures (25.0 mM aceticacid, 25.0 mM MES, and 50.0 mM Tris) at different pH valuesranging from 6 to 10 as described in [13]. Then AP activitywas determined.

2.6. Effect of Different Classes of Proteolytic Enzyme Inhibitorsand Promoters on AP Activity. Crude enzyme extracts werepreincubated with 0.45 ml of 0.1 M potassium phosphatebuffer (pH 8) in the presence of different inhibitors oractivators for 30 min at 37◦C. After pre-incubation, theenzyme reaction was initiated by the addition of the substratesolution (L-leu-p-NA) and AP activity was determined. Con-centration of activators in the reaction mixture during pre-incubation was 1.0 or 10.0 mM. The chemicals tested wereEDTA, 1, 10-phenanthroline, PMSF, DTT, iodoacetamide,and NEM.

2.7. Effect of Divalent Cations on AP Activity. The crudeenzyme extracts were pre-incubated at 37◦C for 30 minwith 0.45 ml of 0.1 M potassium phosphate buffer in thepresence of the chlorides of Mn2+, Co2+, Ni2+, Mg2+, Ca2+, orZn2+. The concentration of divalent cations in the reactionmixture during pre-incubation was 1.0 or 10.0 mM. Afterpre-incubation, the substrate solution (L-leu-p-NA) wasadded to start the enzyme reaction and AP activity wasdetermined.

2.8. Partial Purification of Aminopeptidase. The wholekiwifruit (550 g) was cut into small pieces and homogenizedin 225 ml of 0.1 M of potassium phosphate buffer (pH8.0) supplemented with 1% (w/v) insoluble PVPP, 5%(v/v) glycerol, and 3 mM DTT (extraction buffer). Thehomogenate was filtered through 2 layers of synthetic cloth.The filtrate was centrifuged at 10,000 × g at 4◦C for 20 min,and the supernatant was removed and used as crude extract.Solid ammonium sulphate ((NH4)2SO4) was added to thecrude extract, and the resulting 25–75% precipitate wasdissolved in 7.5 ml of 0.01 M potassium phosphate buffercontaining 10% (v/v) glycerol and 0.2 mM DTT (buffer A).After dialysis of the 25–70% ammonium sulphate fractionagainst buffer A, a DEAE cellulose column (10 × 2 cm) wasused to separate the fractions. Unbound proteins were elutedwith buffer A, and then bound proteins were eluted with100 ml of the buffer A containing a linear gradient of 0.00–1.0 M KCl.

2.9. Statistical Analysis. Statistical analysis of the data wasperformed using STATISTIX 8.0 software. The comparisonbetween treatments was analysed using one-way analysisof variance (ANOVA). Where a statistical significance wasobserved, a Tukey’s Honest Significance Difference (HSD)test was performed to determine how significant from theappropriate zero the values were. Standard errors werecalculated and graphically represented as symmetrical errorbars.

Enzyme Research 3

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Figure 1: Effect of pH on aminopeptidase activity in extracts of thewhole fruit of A. deliciosa. The enzyme activity at pH 9 was taken as100%. Mean values of three different extracts ± standard errors arepresented.

3. Results and Discussion

3.1. Aminopeptidase (AP) Activity in Different Parts ofActinidia deliciosa Fruit. In preliminary experiments, whencrude extracts from the whole fruit had been prepared withsodium phosphate or potassium phosphate buffer (pH 7.0),AP activity was not detectable. Kiwifruit contains morethan 80 volatile aroma, and flavour compounds includingterpenses, esters, aldehydes, alcohols with varying levels ofmonoterpenes, and phenolic compounds [14, 15]. Thesecompounds could have interfered with aminopeptidaseisolation and activity. Here, a reliable protocol (as describedin Section 2) for extraction of AP from kiwifruit anddetermination of its activity using L-leucine-p-nitroanilide(L-leu-p-NA) as a substrate has been established. The presentstudy has established for the first time that kiwifruit hasAP activity and some useful parameters with respect to itsextraction, assay, stability, localization and purification.

AP activity was found in all parts of the fruit of A.deliciosa at different levels. The highest specific (units/mgsoluble protein) and total (units/g fresh weight) AP activitywas localized in the seed followed by the core, inner and outerpericarp, respectively, (Table 1). In contrast, higher enzymeactivities were found in the hypodermis of fully ripe grapeberries than in the seed or flesh [13].

3.2. Effects of pH and Temperature on Kiwifruit AP Activity.AP activity in crude extracts of the whole kiwifruit was mostactive at alkaline pH (Figure 1; ANOVA, P < .05). Similarly,hydrolysis of L-leu-p-NA by crude extracts was most activeat a range of alkaline pH values from many different plantsincluding potato [16], Arabidopsis thaliana [17], tomato [18],and daylily flowers [19].

Kiwifruit AP was most active at 37◦C and 50◦C, sug-gesting the presence of two aminopeptidase isozymes. At

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Figure 2: Effect of temperature on the aminopeptidase activity inthe crude extracts prepared from the whole fruit of A. deliciosa. Theenzyme activity at 37◦C was taken as 100%. Mean values of threedifferent extracts ± standard errors are presented.

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Figure 3: Effect of temperature on the stability of aminopeptidaseactivity in crude extracts prepared from the whole fruit of A.deliciosa. The enzyme activity at 40◦C was taken as 100%. Meanvalues of three different extracts ± standard errors are presented.

55◦C its activity was reduced to 63% (with the activity at37◦C designated as 100%) and then to about 20% at 60–70oC (Figure 2). It was most stable at 37–40◦C (Figure 3)but became unstable as only less than 15% of its activityremained at temperatures higher than 55◦C (ANOVA, P <.05).

3.3. Effects of Protease Inhibitors, Activators, and MetalIons. The presence of 1 mM of 1,10-phenanthroline, EDTA-Na2 and iodoacetamide inhibited kiwifruit aminopeptidaseactivity (Figure 4). In contrast, 1 mM of DTT had a slight

4 Enzyme Research

Table 1: Aminopeptidase activity in different parts of kiwifruita.

Type of tissue Total activity (units/g fresh weight) Specific activity (units/mg soluble protein)

Outer pericarp 0.38± 0.15 0.37± 0.10

Inner pericarp 0.64± 0.19 0.52± 0.07

Core 2.91± 0.68 3.68± 0.96

Seed 60.15± 7.99 5.78± 0.48aAminopeptidase (AP) activity was determined in extracts of each tissue from three different fruits of A. deliciosa. Mean values± standard errors are presented.

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Figure 4: Effect of proteolytic enzyme inhibitors and activatorson aminopeptidase activity in crude extract of the whole fruitof A. deliciosa. Enzyme activity in the absence of any chemical(control) was taken as 100%. Mean values of three different extracts± standard errors are presented.

stimulatory effect (ANOVA, P < .05). The same concentra-tion (1 mM) of NEM and PMSF had no effect. At 10 mM,1,10-phenanthroline, NEM, iodoacetamide, and EDTA-Na2

caused more inhibition. But 10 mM of DTT and PMSF hadneither stimulatory nor inhibitory effect (ANOVA, P < .05).

The observed inhibition of kiwifruit AP activity by metalchelators such as 1,10-phenanthroline and EDTA suggestedthe involvement of a metal ion in the active site of theenzyme. Similar effects were also reported in the studies onleucine aminopeptidases of potato [16], tomato, E. coli pep A,and porcine LAPs [18]. Furthermore, DTT (a thiol reducingagent) at a lower concentration (1 mM) had a stimulatoryeffect but an inhibitory effect at a higher concentration onkiwifruit AP activity suggesting that it was a thiol-dependentmetalloprotease rather than a cysteine protease [20]. On theother hand, iodoacetamide (1 mM) and NEM (10 mM), thespecific inhibitors of cysteine protease, had 60% and 40%inhibition of kiwifruit AP activity, respectively, suggestingthat cysteine residues were likely involved in the enzymeconformation rather than catalysis. A serine-type proteasemight not be a significant contributor to the kiwifruit APactivity as PMSF, a serine protease inhibitor, did not have anysignificant effect on its activity.

The effects on kiwifruit AP activity of Ca2+, Mg2+, Co2+,Ni2+, Mn2+, and Zn2+ with chloride as the counter ionwere studied (Figure 5). At metal ion concentrations of1 mM, only Zn2+ significantly inhibited kiwifruit AP activity

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1 mM10 mM

Figure 5: Effect of divalent cations on aminopeptidase activity incrude extracts of the whole fruit of A. deliciosa. Enzyme activity inthe absence of any cations (control) was taken as 100%. Mean valuesof three different extracts ± standard errors are presented.

(ANOVA, P < .05) whereas the other metal cations tested hadno significant effect. When the concentration of metal ionswas increased to 10 mM, the enzyme activity was stronglyinhibited by Zn2+ (ANOVA, P < .05), and inhibited to alesser extent by Ni2+, Co2+, and Mn2+. At this concentrationCa2+ and Mg2+ did not have any significant effects. Thissuggests that the AP activity might be different from that ofa previously studied protease in kiwifruit that was inhibitedby calcium ions [21]. Furthermore, kiwifruit AP activity wasdifferent from that in potato, Arabidopsis, tomato, porcineand E. coli pep A as they were highly activated by Mn2+

and Mg2+ ions but were also inhibited by Zn2+ ions [16–18]. The kiwifruit AP activity was also different from thatof grape berries which was not inhibited by EDTA, 1,10-phenanthroline, or metal ions [13].

3.4. Partial Purification of Kiwifruit Aminopeptidase. Twomajor peaks of AP activity were separated using DEAEcellulose column chromatography: the unadsorbed andadsorbed fractions (Figure 6), suggesting that there were atleast two isoforms of AP activity in A. deliciosa fruit. In thesefractions only a few low-molecular weight polypeptides werefound to be present following SDS PAGE (data not shown).

Enzyme Research 5

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Figure 6: Elution profile from a DEAE cellulose column ofaminopeptidase (AP) activity and protein content in a concentratedfraction of ammonium sulfate precipitation of crude extracts ofA. deliciosa fruit. The first 30 fractions were eluted with 10 mMpotassium phosphate buffer (pH 8) supplemented with 10%glycerol. The next fractions were eluted with a linear gradient of0 to 1.0 M KCl in the same buffer. One unit of enzyme activity wasdefined as a change in one unit of absorbance at 410 nm per h at37◦C. Protein content was measured at 280 nm.

This might be a facile route to obtain a relatively pure food-grade aminopeptidases from kiwifruit. Further studies, usingmore specific substrates, could lead to some useful food-grade aminopeptidases from kiwifruit. Recombinant DNAtechniques could also be applied to mass produce kiwifruit-originated APs.

References

[1] M. Rassam and W. A. Laing, “Purification and character-ization of phytocystatins from kiwifruit cortex and seeds,”Phytochemistry, vol. 65, no. 1, pp. 19–30, 2004.

[2] N. Izawa, K. Tokuyasu, and K. Hayashi, “Debittering of proteinhydrolysates using Aeromonas caviae aminopeptidase,” Journalof Agricultural and Food Chemistry, vol. 45, no. 3, pp. 543–545,1997.

[3] R. Raksakulthai and N. F. Haard, “Exopeptidases and theirapplication to reduce bitterness in food: a review,” CriticalReviews in Food Science and Nutrition, vol. 43, no. 4, pp. 401–445, 2003.

[4] S.-J. Lin, Y.-H. Chen, L.-L. Chen, H.-H. Feng, C.-C. Chen,and W.-S. Chu, “Large-scale production and application ofleucine aminopeptidase produced by Aspergillus oryzae LL1for hydrolysis of chicken breast meat,” European Food Researchand Technology, vol. 227, no. 1, pp. 159–165, 2008.

[5] S. Mane, M. Damle, P. Harikumar, S. Jamdar, and W. Gade,“Purification and characterization of aminopeptidase N fromchicken intestine with potential application in debittering,”Process Biochemistry, vol. 45, no. 6, pp. 1011–1016, 2010.

[6] M. D. Marinova and B. P. Tchorbanov, “Brassicaceae plants asa new source of food grade peptidases,” Bulgarian ChemicalCommunications, vol. 40, no. 4, pp. 397–400, 2008.

[7] M. Marinova, A. Dolashki, F. Altenberend, S. Stevanovic,W. Voelter, and B. Tchorbanov, “Characterization of anaminopeptidase and a proline iminopeptidase from cabbageleaves,” Zeitschrift fur Naturforschung Section C, vol. 63, no. 1-2, pp. 105–112, 2008.

[8] M. Marinova, A. Dolashki, F. Altenberend, S. Stevanovic,W. Voelter, and B. Tchorbanov, “Purification and character-ization of L-phenylalanine aminopeptidase from chick-peacotyledons (Cicer arietinum L.),” Protein and Peptide Letters,vol. 16, no. 2, pp. 207–212, 2009.

[9] D. W. M. Leung and J. D. Bewley, “Increased activity ofaminopeptidase in the cotyledons of red light-promotedlettuce seeds is controlled by the axis,” Physiologia Plantarum,vol. 59, no. 1, pp. 127–133, 1983.

[10] Y. Yamaoka, M. Takeuchi, and Y. Morohashi, “Purificationand partial characterization of an aminopeptidase from mungbean cotyledons,” Physiologia Plantarum, vol. 90, no. 4, pp.729–733, 1994.

[11] K. Tishinov, N. Stambolieva, S. Petrova, B. Galunsky, and P.Nedkov, “Purification and characterization of the sunflowerseed (Helianthus annuus L.) major aminopeptidase,” ActaPhysiologiae Plantarum, vol. 31, no. 1, pp. 199–205, 2009.

[12] M. M. Bradford, “A rapid and sensitive method for thequantitation of microgram quantities of protein utilizing theprinciple of protein dye binding,” Analytical Biochemistry, vol.72, no. 1-2, pp. 248–254, 1976.

[13] H.-C. Kang, T.-R. Hahn, I.-S. Chung, and J.-C. Park, “Char-acterization of an aminopeptidase from grapes,” InternationalJournal of Plant Sciences, vol. 160, no. 2, pp. 299–306, 1999.

[14] C. O. Perea, H. Young, and D. J. Beever, “Kiwifruit,” in Tropicaland Subtropical Fruits, P. E. Shaw, J. Chan, and S. Nagy, Eds.,pp. 336–385, Agscience, Auburndale, Fla, USA, 1998.

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