Enzymes Used in Industries and their Uses!
Of the 1000 and more enzymes isolated, fewer than 20 are now commercially used on a scale that has significant impact on either the enzyme industry or the user industries. Only the more important will be treated here.
It is worth remembering that commercial enzyme preparations represent mixtures in which the principal enzyme activity is only one among many others. In enzyme applications this fact may imply that desired effects are interfered with by contaminating enzymes, or that effects observed trace back to such activities. The situation becomes even more complex when one considers the varying qualities and quantities of accompanying enzymes depending on the microbial source, the cultivation method, and the recovery process.
This is one of the reasons for variable suitability of different products of the market or even of different prices for the same product. In some applications, it is invariably necessary to eliminate the interfering activity unless a mutant unable to produce this activity has been found. The success of such a procedure can decisively influence the development of a special enzyme market.
Enzymes Used in Industries and their Uses and Application!
Amylolytic enzymes represent a group of catalytic proteins of great importance to the food industry. They were also one of the first enzymes to be produced commercially by microorganisms. Since industrial starch processing requires certain physical properties or a specific carbohydrate composition of the final starch hydrolysate, great efforts have been made to find microbial enzymes with the specific characteristics required. Consequently, a remarkable number of new starch-degrading enzymes have been discovered, most of which have characteristics and properties that clearly distinguish them from all amylolytic enzymes previously known.
Starch-degrading enzymes can be divided into two main groups, α-1,4- glucanases and α-1,6-glucanases (debranching enzymes).
This enzyme (α-1,4-glucan-glucanohydrolase, EC 188.8.131.52) acts on starch Components, which contain at least three α-1,4-linked glucose units, as an endoase, i.e., in an essentially random manner, with the production of reducing sugars. Mode of action, properties, and products of hydrolysis differ somewhat, depending on the source of the enzyme. Two types of microbial α-amylases have been recognized, termed “liquefying” and “saccharifying” α-amylases. The main difference between them is that the saccharifying enzyme produces a higher yield of reducing sugars than the liquefying enzyme.
Bacterial α-amylase might be produced by a number of Bacillus species, Pseudomonas saccharophila, and Clostridium species, but on an industrial scale specially selected strains of Bacillus subtilis seem to be preferred. Fungal α-amylases for commercial purposes are derived from Aspergillus oryzae. Certain strains of Aspergillus niger and particularly of Aspergillus oryzae produce large amounts of the saccharifying enzyme.
In industrial fermentation the cultivation of bacterial α-amylase producers is principally conducted by the submerged culture technique. The media employed are generally based on the use of natural raw materials, because of their cheapness and stimulatory effect. The latter effect is attributed to enzyme inducers or certain growth factors, such as trace elements, vitamins, or suitable combinations of amino acids.
A careful balance of carbohydrate and nitrogen ingredients of the medium is most important. From reports in the literature it appears that the concentration of the N source should normally be higher than necessary for meeting full growth of the organism. Mostly, organic N is supplied, but inorganic N can also serve as the major nitrogenous component- it is, however, usually combined with small amounts of proteinaceous compounds.
Fungal α-amylase was originally and is still produced in significant amounts in solid substrate culture. Wheat bran serves as the basic component of the medium. Most publications on this procedure mention weakly acidic solutions as a means of moistening the bran, but acid has been shown to be harmful to amylase production. The moisture content of the culture depends on the height of the culture and on incubation temperature. The incubation time required to reach maximum yield of enzyme varies between “overnight” and about 4 days.
While Dunn et al. (1959) succeeded in selecting strains capable of producing good yields of α-amylase under submerged culture conditions; fungal a-amylase is also produced in deep tank fermentation. Usually flour or starch serves as raw material, supplemented with inorganic salts.
Addition of stillage, corn steep liquor, or yeast extract contributes stimulating agents for some strains. In most cases of fungal α-amylase fermentation, the enzyme accumulates largely in the stationary phase, and quite often during the phase of autolysis. The amount of a-amylase that is formed during these growth phases is very strongly dependent on environmental and internal factors.
Published work indicates the existence of common features in both bacterial and fungal a-amylase fermentations. It has been shown quite generally that in both cases a proper course of pH change toward alkalinity during cultivation is of great importance. Salts of organic acids, e.g., citrate, gluconate, or acetate, can serve as pH regulators acting in the desired sense.
The necessary amounts depend on the carbohydrate concentration or on the kind of concentration of organic and inorganic N sources. The same effect call is achieved by alkalizing nitrogenous compounds, such as nitrates, urea, and proteinaceous matter. When ammonium salts are used as the N source, organic salts have to be given preference due to their capability of maintaining the pH of the culture at neutral or of resulting in a rising pH, as shown by Horvath and Inczefi (1972).
Proteinaceous matter, e.g., peptone, exerts the same effect. Fully synthetic media must not contain ammonium salts, when lacking salts of organic acids, but should contain nitrates or urea. In contrast to the acidifying effect of inorganic NH4+ salts, ammonium acetate tends to maintain the pH of the culture at neutral or to result in a rising pH. This can be attributed to the fact that the organisms produce non-utilizable organic acids.
The question of the type of mechanism that regulates the formation of α-amylase cannot be answered clearly or generally. While Coleman (1967) stated that in Bacillus subtilis α-amylase is constitutive and controlled by the size of the pool of nucleic acid precursors, Schaeffer (1969) and Meers (1972) found in bacilli that biosynthesis of this enzyme is governed by catabolite repression. Using a carbon-limited medium it was demonstrated that the presence of an inducer was not necessary for a-amylase biosynthesis.
Dunn et al. (1959) found that addition of Ca phytate to natural and synthetic media increased the yield of dextrinogenic α-amylase from a Bacillus strain. Similarly, Yamada (1961) observed an increase in production of both dextrinogenic and saccharogenic amylase as a response to the incorporation of 0.01 -0.05% phytic acid in the medium used for Aspergillus oryzae or Aspergillus awamori.
Commerical preparations of bacterial α-amylase are commonly produced with a minimum of purification. Highly active or purified preparations are obtained by precipitation and/or adsorption techniques. For some applications the absence of other enzymes, especially proteinase, is essential.
For this purpose several methods are available, e.g., adsorption procedures, fractional precipitation, and selective inactivation.
Bacterial α-amylase has a molecular size on the order of 50,000, each molecule containing 1 gram atom of Ca+ +. In the presence of zinc, a dimer is formed containing 1 atom of zinc. The calcium maintains the enzyme molecule in the optimum conformation for maximum activity and stability; it does not participate directly in the reaction. The addition of Ca salts is generally recommended to achieve maximum heat stability of the enzyme.
The maximum activities of a-amylase are in the slightly acidic region between pH 4,5 and 7.0, with differences depending on the enzyme source. The α-amylases from Bacillus subtilis, and especially Bacillus stearothermophilus, are particularly heat stable. In contrast, thermal stability of fungal α-amylase is relatively low.
Bacterial a-amylases preparations are mainly used in the continuous process for desizing of textile fabrics. Other applications include modification of starches suitable for preparation of adhesives, sizes and coatings for the paper industry, as well as manufacture of glucose and glucose syrups and brewing processes. Fungal α-amylases are extensively used in flour treatment for supplementing the diastatic activity of flour.
The enzyme amyloglucosidase (glucoamylase, α-1, 4-glucan glucohydrolase, EC 184.108.40.206) acts as an exoase that liberates the α-1, 4-linked glucose units consecutively from the nonreducing ends of the starch chains. Terminal α-1, 6-bonds are also cleaved, but much more slowly than α-1, 4-linkages. Maltose is only slowly attacked; increasing the chain length up to 5 or 6 glucose units gives faster attack.
Amyloglucosidase occurs in many microorganisms, particularly in starch degrading molds (Aspergillus, Mucor, Rhizopus, Endomyces) and certain bacteria (Aerobacter, Clostridium). The mold species commonly used for large-scale production is Aspergillus niger. However, this species also produces transglucosidase, which transfers glucose and higher saccharides to oligosaccharides, resulting in synthesis of polysaccharide.
These products cause serious reduction in glucose yield and impede its crystallization. Strains are chosen which produce low levels of transglucosidase. Some organisms, e.g., members of the Mucor, Rhizopus, and Aspergillus phoenicis groups, usually produce amyloglucosidase without simultaneous formation of transglucosidase.
When Aspergillus niger is used as the production strain, the submerged fermentation is the cultivation method of choice. Mucor and Rhizopus strains are used in solid substrate fermentation because they are obviously unsuitable for submerged culture. The reason is that when the nonseptate hyphae of these species are damaged by the strong shearing action of high-speed impellers at one single point the protoplasm of the whole filament will be extruded.
Media employed for amyloglucodsidase production in submerged fermentation contain usually high solid contents of organic matter (of the order of 12-20%). Starchy material, such as maize, wheat, barley, rye, and sorghum, are the raw materials of choice. These cereals provide also the N source, the content of which is usually sufficient. Inorganic nitrogen can be utilized, ammonium salts being preferred.
Stimulation of amyloglucosidase formation may be obtained by incorporation of yeast extract or stillage into the medium. An acid reaction of the mash seems to be necessary for maximal production of amyloglucosidase. In Aspergillus niger fermentations the pH of the culture decreases to values as low as 2.8 at the end of the cultivation, caused by the excretion of organic acids.
Removal of transglucosidase activity is an important step in the processing of amyloglucosidase. One of the oldest methods to free the culture filtrate from transglucosidase takes advantage of the fact that amyloglucosidase is highly acid resistant. Adsorption techniques using synthetic water-insoluble hydrous magnesium silicate or clay minerals (e.g., attapulgite and bentonite) are said to be either unreliable or not sufficiently selective.
Barton (1967) claimed coprecipitation with both maleic anhydride copolymers and heteropolyacids to be commercially practicable for removal of transglucosidase. Another claim of Sternberg (1967) suggests the precipitation of transglucosidase by chloroform, but the conditions of the precipitation process are very critical.
Amyloglucosidase has been bound covalently to a variety of matrices and also has been physically adsorbed on or bound to various materials. Further, ultrafiltration reactors have been used for the continuous conversion of starch to glucose but obviously there is little inclination in industry to adopt processes based on immobilized amyloglucosidase.
Amyloglucosidase has largely replaced acid hydrolysis in glucose production. Following the solubilization of starch by acid or, preferably by a heat stable bacterial α-amylase, further degradation is achieved using amyloglucosidase.
The enzyme pullulanase (amylopectin 6-glucanohydro- lase, EC 220.127.116.11) splits α-1, 6-glucosidic linkages at a branch point as well as in a linear chain of a number of oligo- and polysaccharides. It degrades pullulan completely to maltotriose and is active on amylopectin, glycogen, and their β-dextrins. The minimum chain which pullulanase liberates is maltose; a 3 unit chain (maltotriose) is optimal.
Pullulanase has been found in Aerobacter aerogenes and other bacteria, Streptomyces included. Production of the enzyme is inducible by growth on α-glucans ranging from maltose to glycogen. Yokobayashi et al. (1968) found that starch or liquefied starch with a low dextrose equivalent (D.E. 5-10) was particularly inducive.
Highest yields occurred when this low D.E. syrup was used in all steps of inoculum buildup as well as in the production stages. Further, for optimal synthesis, a comparatively high pH is required. Ueda et al. (1971) found that in Streptomyces the pH of the culture rose up to 8.5 and the enzyme was abundantly produced in the autolytic phase.
This enzyme (amylopectin 6-glucanohydrolase, EC 3.2.1. 68) is able to completely debranch glycogen, but unable to act on pullulan. It differs from pullulanase in that it acts more readily on the native polymers. The α-1, 6-linkages are only split when present at branch points in oligo and polysaccharides. Isoamylase requires a minimum of 3 glucose units on the A-chain of amylopectin, while pullulanase requires only 2.
Isoamylase occurs in yeast and bacteria. It is produced commercially from Cytophaga and Pseudomonas, and is an inducible enzyme.
Other Starch-Degrading Enzymes:
Among these, the following are worthy of mention – An exo-amylase from Pseudomonas stutzeri which splits off maltotetraose from the ends of starch chains; an exo-amylase releasing maltohexaose, which is produced by Aerobacter aerogenes; an enzyme from Bacillus licheniformis that forms maltopentaose as the principal product; isopullulanase produced by certain species of Aspergillus niger.
This enzyme degrades pullulan to isopanose. It shows no action on amylopectin but is able to hydrolyze terminal isomaltosyl groups as produced by amyloglucosidase.
The name “cellulase” is given to all enzymes which cleave β-1, 4-gluco- sidic linkages in cellulose and chemically or physically modified cellulose, in cellodextrin, and in cellobiose.
It is well established that cellulase is a multi-enzyme complex, the different components of which bring about the complete degradation of cellulose to monosaccharide residues. A classification scheme of cellulolytic enzymes is given in Table 15.10. The terms C1-cellulase and Cx-cellulase were originally proposed by Reese et al. (1950) for two components of the cellulase complex that differed in their substrate specificities against cotton fiber.
According to this concept, C1 can attack native cellulose of higher crystallinity (e.g., cotton fiber), while Cx cannot attack such cellulose but can split in turn the cellulose fragments which have been produced by the action of C1. The mode of action of the C1– enzyme has long been questioned, but now many investigators tend to identify it with β-1,4-glucan cellobiohydrolase, which liberates cellobiose units from the non-reducing end of the cellulose chain.
Many bacteria and fungi are cellulolytic, but preparations marketed for industrial applications are derived only from Aspergillus niger, Trichoderma viride, Neurospora, and some other organisms. The Aspergillus enzyme exerts good activity on carboxymethylcellulose (CMC), but fails to attack solid cellulose because it lacks C1-cellulase. In contrast, Trichoderma viride produces an enzyme complex with high levels of C1-cellulase, which extensively degrades insoluble cellulose.
Cellulase in fungi is an inducible enzyme. It is only produced when the cells are grown on cellulose, on glucans of mixed linkages including the β-1,4 bond, and on a few oligosaccharides, The inducing effect of cellulose is due to soluble hydrolysis products of the cellulose, in particular cellobiose.
Lactose, a β-1, 4-galactoside, and sophorose, a β-1, 2-glucoside, are the only known cellulase inducers that do not have a β-1, 4-glucoside bond. The inductive action of sophorose is limited to Trichoderma viride. However, in spite of the impressive inductive power of this rare sugar, the levels of enzyme produced are not equal to those on cellulose.
Cellobiose plays a complex role- in low concentrations (0.1%) it serves as an inducer of cellulase; in high concentrations (0.5-1.0%) it represses cellulase formation, and, in addition, it can also act as an inhibitor of cellulase action.
Cellulase yields can be increased by various additives to the medium. Reese and Maguire (1971) observed that Tween 80 and Tween 40 doubled the cellulase yield in Trichoderma cultures. The mechanism of the action of these surfactants is not understood but may be related to increased permeability of the cell membrane.
Never the less, Tween 80 has proven useful in the fermentation industry and is routinely incorporated into the culture medium. Further, enhancement of enzyme production can be achieved by supplying peptone at one-tenth the cellulose concentration. This leads to a decrease in the lag of growth and cellulase synthesis.
In actual large-scale fermentation Aspergillus niger is mostly cultured by the wheat bran-tray method. This process has no problems and leads to high yields of cellulase. The extraction of cellulase from solid substrate cultures is performed by percolation of the dried mold bran with 0.02 to 0.1 M lactic acid.
Neurospora and Trichoderma are grown by submerged culture. For continuous culture it is advantageous that Trichoderma viride produces a suspension of short mycelial threads, rarely forming pellets. Mitra and Wilke (1975) proposed a 2-stage operation in continuous stirred tank reactors. The first stage utilizes glucose for biomass production and the second stage utilizes pure spruce wood cellulose for enzyme formation. A significant increase in enzyme productivity was obtained.
Bran, straw and other plant materials pretreated with alkali serve as cellulose-containing raw materials for submerged cultivations. Ammonium ions can be used as suitable N source. A correct pH profile is necessary to give optimum enzyme yields in batch culture. It has been shown that a drop to the range of pH 3.5-3.0 is optimal for Trichoderma viride. This is usually achieved by an empirical procedure which involves medium composition and initial pH.
Length of cultivation affects the relative amount of the various cellulase components present in the medium. This has been demorstrated for Myrothecium verrucaria with CMC and with swollen substrates. In both cases C1 appeared prior to Cx.
Concentration and purification of the enzyme is carried out by precipitation, adsorption, or gel filtration techniques. Granulated preparations can be obtained by mixing with salt hydrates (e.g., Na2SO4.H2O) and subsequent vacuum drying.
Current use of cellulase is limited to improving texture and palatability of poor quality vegetables. It is also useful for accelerating drying of vegetables. A potential application of cellulase is the conversion of cellulosic materials to glucose and other sugars which in turn can be used as microbial substrates to produce single cell protein or a variety of fermentation chemicals (alcohol, etc.).
Many plant pathogenic bacteria and fungi have long been known to produce pectolytic enzymes and it is widely accepted that the production of these enzymes is a major means by which microorganisms invade the host tissue. Moreover, pectolytic enzymes are essential in the decay of dead plant material by nonpathogenic microorganisms and thus assist in recycling carbon compounds in the biosphere. Lastly, these enzymes play a decisive role, in the microbial spoilage of fruits and vegetables.
Several types of enzymes are involved in the degradation of pectic materials. They are divided into 2 main groups, depolymerizing enzymes and saponifying enzymes or pectinesterases. According to the scheme of Neukom (1963), the depolymerizing pectolytic enzymes are further classified by applying the following 3 criteria- preference for pectic acid or pectin as substrate, hydrolytic or transeliminative cleavage of the glycosidic linkages, and endo- or exo-types of the action mechanism.
By various combinations of these characteristics, 8 groups of depolymerizing enzymes can be listed, but the existence of the exo-polymethylgalacturonase and exo- polygalacturonate lyase types is doubtful. In 1971 Hatanaka and Ozawa proposed a scheme which included only those enzymes whose existence had been demonstrated (Table 15.11).
Occurrence of pectolytic enzymes has been reported in a large number of bacteria and fungi. Commercial enzymes are generally obtained from fungal sources since the pH optima of these enzymes are in the range found naturally in materials to be processed. Most potent strains are selected from Aspergillus niger. Japanese enzyme manufacturers also use Sclerotinia libertiana and Coniothyrium diplodiella as producers of pectolytic enzymes.
In the majority of cases microorganisms produce a variety of pectolytic enzymes and, for this reason, commercial preparations are mixtures of these enzymes. The relative amount of the single components varies considerably with the particular strain employed, medium composition, and culture conditions. Careful observation of the factors responsible for the promoted synthesis of certain enzyme fractions or limited formation of others enables the manufacturer to “control” the composition of the preparation and to meet the need for specific formulations.
Biosynthesis of pectolytic enzymes is constitutive or controlled by the mechanisms of induction or catabolite repression. No uniformity exists among the various organisms and the various components of the enzyme complex. Phaff (1947) found pectolytic enzymes to be adaptive in Penicillium chrysogenum, but constitutive in Aspergillus foetidus. Saito (1955) showed that in Aspergillus niger endopolygalacturonase (endo-PGase) is adaptive. For Clostridium felsineum, a plant retting organism, Osman et al. (1969) demonstrated pectinmethylesterase (PME) to be adaptive and PGase constitutive.
On an industrial scale pectolytic enzymes are produced by the solid substrate method as well as by submerged culture. Wheat bran or defatted rice bran have been recognized as satisfactory basic substrates in solid substrate cultures. It is well known that some by-products of the food industry, such as beet pulp, apple pulp, or grape pulp, exert a promoting effect on enzyme formation. Other ingredients, e.g., nutrient salts, acid, or buffers, are also incorporated to regulate the pH during the growth of fungi. The time of cultivation can extend up to 7 days, but when Aspergillus niger is used, the desired enzyme level is normally reached within 36 to 72 hr.
After fermentation the mold bran is dried and can be used as such. For obtaining concentrates the dried mold bran is extracted with suitable aqueous solutions and concentrated under vacuum or by ultrafiltration. Crude or refined solid concentrates are obtained by spray drying or precipitation with neutral salts or solvents.
Submerged cultures, in contrast to solid substrate cultures, seem to have the disadvantages of poor yields and undesirable composition. Brooks and Reid (1955), for example, found that Aspergillus foetidus produced endo- PGase and exo-PGase in surface culture, but only endo-PGase in submerged culture.
The production of pectolytic enzymes by submerged fermentation has been described by Nyiri (1968, 1969). As an example, the method reported for Aspergillus alliaceus can be cited. The fungus is grown in a liquid medium composed of 2% wheat bran, 2% (NH4)2SO4, 0.25% KH2PO4, 0.25% yeast extract, 0.1% pectin (degree of esterification = 59%). The initial pH is 3.8, adjusted with HCl.
Traces of silicon serve as antifoam. After inoculation with conidia, the medium is agitated and aerated, with the pressure inside the fermentor maintained at 141.8 kPa (1.4 atm). The fermentation is completed within 72 hr and the mycelium separated by filtration. The filtrate is cooled to 0°-1°C and the enzyme precipitated by addition of (NH4)2SO4 during a period of 4 hr. After standing for 12 hr, the precipitate is washed and dried to give a solid concentrate.
Initial pH of the medium and pH development in the growing culture play an important role with regard to both enzyme composition and yield of the enzyme fractions. Low pH values, in particular a decreasing pH during cultivation, are favorable for the production of pectinases used in the fruit juice industry, according to Hauptmann (1951), pH 2-3 at culture maturity of Aspergillus niger being desirable. In contrast, Tuttobello and Mill (1961) allowed the pH to rise up to 4 at the end of the culture following a drop to about 3 on the fifth day of cultivation.
Tuttobello and Mill (1961) found an aqueous extract of non-defatted peanut flour strongly stimulated the production of pectolytic enzymes. They also observed a strong influence of inoculum on enzyme production. The kind of inoculum and, particularly, its size have to be standardized carefully. With Aspergillus niger Tuttobello and Mill (1961) found that the production of pectolytic enzymes was strongly influenced by inoculum size in the range of 104 to 2 x 105 conidia per ml, while mycelium formation remained unchanged.
From many reports in the literature it can be concluded that there is a strong variation in relative activity of the various components of the pectolytic system in the course of fermentation, indicating their sequential production.
Extensive use of pectolytic enzymes is made in processing fruit juices for increasing juice yields on pressing, as aid in clarification of juices, and for depectinizing in order to obtain high density fruit juice concentrates. Fungal enzymes are widely used in producing apple juice, grape juice, and wine. In the production of coffee beans the residual mucilaginous coating surrounding the bean can be liquefied by commercial pectolytic enzyme preparations, thus offering an alternative to the usually used fermentation process.
The curing or fermentation of cocoa, tea, and tobacco also can involve pectolytic enzymes. One of the oldest applications of these enzymes is the process of retting, in which textile fibers, such as flax, hemp, and jute, are loosened from their plant stems. The enzyme system of Clostridium felsineum, an organism that is involved in aerobic retting, contains endopoly- galacturonate trans-eliminase, but not pectinesterase.
Recently, pectolytic enzymes have been proposed as a means to make commercial softwoods, such as Sitka and Norway spruce, more permeable to preservatives. It has been demonstrated that treatment with enzyme preparations as well as with the specific bacteria that produce them is possible.
Plant cell wall polysaccharides other than cellulose and pectic substances are referred to as hemicelluloses. They are complex compounds and very few of their chemical structures have been clarified. During recent years many enzymes have been recognized which specifically act on different types of hemicelluloses. By far the best known group is that of the xylanases. Other groups of hemicellulases are, for example- mannanases, galactanases, etc.
Many strains of bacteria and fungi are known to produce hemicellulases inducibly or constitutively, but on industrial scale only fungal strains seem to be used as enzyme sources. Even in these cases hemicellulases are mostly obtained as side activities in the production of other enzymes such as cellulase. Therefore, as a rule, the hemicellulase activity of the commercially available enzyme systems is low. This fact reflects either an inherent instability of these enzymes or lack of knowledge of how to produce them.
Sucrase and invertase are two of the older names of the enzyme β- fructofuranosidase (EC 18.104.22.168). It catalyzes the hydrolysis of the terminal non-reducing β-fructofuranoside residues in β-fructofuranosides. The name “invertase” was derived from the action in splitting sucrose, which is optically dextrorotatory, to form glucose and fructose, a mixture that is levorotatory-
This reaction can also be carried out by a-D-glucosidase (so-called glucosi- doinvertase), but this enzyme is unable to split off fructose from the trisaccharide raffinose as is β-fructosidase (so-called fructosidoinvertase).
β-Fructosidase can be prepared from a variety of microbial sources, but only the enzymes from Saccharomyces cerevisiae and Saccharomyces carlsbergensis have industrial importance.
Biosynthesis of invertase is controlled by a catabolite repression mechanism in Saccharomyces fragilis and by repression through unknown effectors in Saccharomyces cerevisiae. For a long time invertase was considered to be totally an intracellular enzyme.
It has, however, been established that in derepressed cells only a small proportion of the invertase is located inside the cytoplasmic membrane, most of it being retained externally within the cell wall or between the wall and the cell membrane. In fully repressed cells all the enzyme is intracellular.
The release of the invertase from yeast is achieved by destruction of the structures responsible for the retention of the enzyme. There are various ways in which the separation from the cells can be accomplished. One method is autolysis with chloroform, toluene, or ethylacetate at 30°C for not over 3 hr.
Following extraction from yeast, comparatively high purification of invertase is necessary for its application in foods because the enzyme preparation usually has an undesirable, irritating taste originating from yeast. These procedures include common methods for purification of enzymes such as ultrafiltration, precipitation, and adsorption techniques.
The commercial preparation of invertase usually starts with an accumulation step. For this purpose pressed bottom yeast is suspended in a 20-fold amount of nutrient broth containing 4 parts (NH4)2.HPO4, 4 parts KH2PO4,1 part Mg(NO3)2, and 1 part KNO3.
The mixture is aerated for 3-8 hr, while the temperature is maintained at 28° to 30°C and at a pH of 4.5. During the same period, 3 to 20% sucrose in solution is added continuously, a procedure that ensures reduced catabolite repression. At the end of the process the invertase activity of the yeast is increased up to 15-fold.
The intracellular yeast invertase has a molecular weight of 135,000 and is free of carbohydrate, whereas the external enzyme has a molecular weight of 270,000. Approximately half of the external β-fructofuranoside consists of mannan. The pH activity curve of invertase is rather broad between pH 3.5 and 5.5, with an optimum between 4 and 4.5.
This is the same range within which the enzyme exhibits its highest stability. Yeast invertase is strongly inhibited by heavy metal ions (especially Ag+); they combine with the histidine side chains of the enzyme molecule, not with its thiol groups.
Invertase has a number of interesting uses in the confectionery industry for soft center candies, fondant, and chocolate coatings. Its use in the preparation of invert sugar by hydrolyzing sucrose has been restricted by glucose isomerase which permits production of invert sugar from cheaper sources.
Lactose or milk sugar is enzymically split to glucose and galactose by the action of enzymes called β-galactosidases or, more commonly, lactases (β-D-galactoside galactohydrolase, EC 22.214.171.124). Lactase is very specific for the galactose residue but much less specific for the aglycone moiety of 0-galactosides. The enzyme is also responsible for transfer activities which occur with the formation of oligosaccharides.
Lactase is widely distributed in microorganisms. Some strains of Escherichia coli are very poteht producers, but are not suitable for food purposes. Available commercial preparations are derived from lactose fermenting yeasts such as Saccharomyces fragilis, Zygosaccharomyces lactis, and Candida pseudotropicalis or from fungi like Aspergillus niger, and particularly a mutant strain of Aspergillus foetidus.
The biosynthesis of β-galactosidase has been extensively investigated, largely in Escherichia coli in connection with studies on the biosynthesis of proteins and its genetic control. The enzyme is of the inducible type, with lactose serving as an inducer. In Fusarium oxysporum and Verticillium alboatrum the lactase can be induced by D-galacturonic acid and, to a lesser extent, by D-galactose.
On an industrial scale the enzyme is obtained, for example- by growing yeast on a lactose medium or on whey. The separated yeast is autolyzed or extracted, and a cell-free extract is obtained by centrifugation or filtration. The enzyme may then be further processed by salt or solvent precipitation.
Another procedure, described by Stimpson (1954), involved spray drying of the washed yeast at temperatures which destroy any residual alcoholic fermentation activity, thus leading to crude products which can immediately be used as lactase preparations.
The lactases from various microbial sources differ in properties such as pH optima, etc. For example, the pH optimum of the bacterial enzymes is around 7.0; that of the fungal preparations near 5.0; and that of the yeast enzymes near 6.0; the lactase from Corticium rolfsii is distinguished by its unusual maximum activity and stability at pH 1.8-2.0.
Yeast lactase is activated by potassium and ammonium ions and is inhibited by certain metals such as copper and iron. Metal-chelating agents do not stimulate the enzyme, indicating little sensitivity to trace heavy metals. The addition of reducing compounds, e.g., cysteine’, sodium sulfide, or potassium metasulfite, is able to overcome the effect of metal inactivators and to activate the enzyme.
Hereditary intolerance to lactose precludes use of milk as a valuable protein source in large areas of Asia and Africa. In addition, lactose causes a number of problems in the dairy and allied industry because of its poor solubility, resulting in crystallization in concentrated dairy products.
Enzymic hydrolysis of the milk sugar is helpful in overcoming these problems. Moreover, lactase is used in the production of sweet syrups from sources of lactose such as cheese whey and in making waste whey a better substrate for growing microorganisms for single cell protein.
When treating milk it is preferred to employ lactase from yeast because of legal reasons, although this enzyme is less stable than the bacterial one. During past years the use of skim milk powders in bread has been considerably reduced. This has reduced the interest in lactase for the formation of fermentable sugars in baking.
The microbial proteases which are of interest for application in the food industry are all of the endopeptidase type and are all extracellular enzymes. There are many different types of proteases produced by an extraordinarily large number of microorganisms, but in actual practice the enzymes prepared commercially are of a very limited number of types and they are derived from very few organisms (see Table 15.12).
The proteolytic enzymes from microorganisms are classified into 4 main groups according to the Scheme of Hartley (1960) and based on the mechanism of their action- serine proteinases, thiol proteinases, metalloproteinases, and acid proteinases. Further subgroupings refer to the side-chain specificity of the proteinases and to the properties of their active centers. A classification scheme of microbial endopeptidases is given in Table 15.13.
Industrial production of microbial proteases is carried out on a large scale by a number of companies in Europe, Japan, and the United States. For cultivation of the microorganisms the submerged fermentation is the preferred method; with bacteria it is the exclusively used process. However, fungi usually give higher yields when cultured on solid media so this method continues to play a role.
As in most fermentation there is a trend to use highly concentrated media. The reason for this is that one can expect higher enzyme yields per unit volume with a larger cell concentration, although there is no direct correlation between growth and protease production. With regard to serine and metalloproteinases it seems that low concentrations of purely carbonaceous substrates and high concentrations of proteinaceous N sources stimulate production.
Many of the organisms excrete more than one kind of protease. The type of proteolytic enzyme formed may depend on the composition of the medium. For example- Bacillus NRRL B-3411 produces the preferable neutral protease when grown on a grain medium, but mainly alkaline protease when cultured on a fishmeal-enzose-cerelose medium. The biosynthesis of proteases is often correlated with particular growth phases of the microbial culture.
Under most growth conditions, Bacillus species produce extracellular protease during the post-exponential growth phase. Mandelstam (1958) attributed this behavior to an increased need for turnover of cell proteins at the slower growth rate. Other bacilli synthesize proteases during the exponential growth phase. However, this kinetics depends on the composition of the medium.
For all protease preparations the degree of purification depends on the intended use. A number of purification procedures are in existence. Of course, various combinations are possible. Particular care is necessary during the drying process in order to avoid the formation of dust. For this reason protease preparations are pelleted or coated with some suitable material.
Bacterial proteases are used on a large scale in enzyme-containing washing powders, but they are not widely used in food processing. Minor uses are in the chill proofing of beer, in the production of protein hydrolysates, in the production of condensed fish solubles, and as feed supplement. In contrast to bacterial preparations fungal proteases are the more interesting group for the food industry. They are used, for example- for the modification of wheat proteins in bread doughs, in meat tenderizing, and in several less important applications.
These proteases are widespread in bacteria and fungi. They show maximum activity at neutral to alkaline pH and are inhibited by diisopropyl fluorophosphate (DFP) or phenylmethane sulfonylfluoride (PMSF). They can be classified into at least 5 groups- trypsin like proteinases, alkaline proteinases, Myxobacter α-lytic proteinase, staphylococcal proteinases, and serine neutral proteinases. In this article only the serine alkaline proteinases will be treated because of their superior economic importance.
Proteases of this type are most active at pH 9.5-10.5; they are sensitive to DFP and potato inhibitor, but not to tosyl-L-lysine chloromethyl ketone. Their specificity is similar to that of a α-chymotrypsin but somewhat broader. All alkaline serine proteases show specificity toward aromatic or hydrophobic amino acid residues, such as tyrosine, phenylalanine, or leucine, at the carboxyl side of the cleavage point.
The molecular weights are 26,000-34,000, slightly below the range of neutral metalloproteases. The isoelectric points are about pH 9. Most of the alkaline proteases are stable from pH 5 to 10 at low temperatures, but show rapid loss of activity at 65°C. Certain strains of Bacillus, showing alkalophilic properties, synthesize a serine alkaline proteinase that is most active at pH 11-12.
Serine alkaline proteinase is produced by numerous species of bacteria and fungi. The best known representatives of this type are the subtilisins, which are produced by Bacillus subtilis and related species.
Due to their great economic importance, the biosynthesis of Bacillus alkaline proteases has been well investigated. Keay and Moser (1969) have proposed that alkaline serine proteinases produced by different bacilli or different strains of Bacillus subtilis can be divided into two groups- subtilisin Carlsberg and subtilisin Novo. These enzymes are quite distinct from each other, but possess many similar properties. It may be mentioned that a similar situation has also been observed with various alkaline serine proteinases from the genus Aspergillus.
The synthesis of these enzymes is linked to particular phases of development of the microbial culture. Some strains, e.g., those of Bacillus megaterium, produce the protease during the log phase of growth, while others, like those of Bacillus subtilis and Bacillus cereus, produce it in the stationary phase.
However, the relationship between growth cycle and enzyme formation depends on the ingredients of the substrate. It is generally valid that the time of biosynthesis is genetically determined and can be changed or extended by selecting proper mutants.
In most species production can be inhibited by certain components in the growth media, such as free ferric ions, amino acids, carbon sources, or several of these. Catabolite repression and availability of nucleic acid precursors are also thought to play a role in alkaline protease synthesis. The concentration of purely carbonaceous medium components should normally be kept on a low level.
This can be achieved by incremental feeding of the C source, e.g., glucose, keeping its concentration at a range of 0.4 to 1%. High concentrations of C sources yield excess organic acids leading to a decrease in pH, which is accompanied by a decrease in alkaline protease production. On the other hand, Keay et al. (1972) have shown that the enzyme yield can be largely enhanced when the synthesis of protease is accompanied by an increase in pH of the culture.
Such an increase in pH can be reached, for example, by using an organic acid (or its salts) as major C source. Niwa et al. (1971) observed good yields of protease under this condition, and the results of Kline et al. (1944), Dion (1950), and Maxwell (1952) confirm this assumption.
Bacterial alkaline proteases are produced exclusively by the submerged culture methods. Amounts of more than 1 g protease per liter culture liquor are quite usual. With specially selected strains markedly higher yields are possible; e.g., Bacillus subtilis strain AJ 3266 can produce more than 10 g enzyme per liter.
Continuous fermentation techniques do not seem to have been employed in the industrial production of alkaline protease, but there are several publications describing continuous culture on the laboratory scale. Heine- ken and O’Connor (1972) observed that a continuous fermentation of Bacillus subtilis yielded mutants with lower protease productivity.
Fungal alkaline proteases are mainly produced from Aspergillus species, in both solid substrate and deep tank fermentations. Solid substrate cultures, extensively used in Japan, are carried out with wheat or rice bran or whole grains as the basic substrate. It has been shown that NH4+ ions strongly inhibit production of the enzyme, while nitrates and Na salts of aspartic and glutamic acids promote its formation. Na salts of organic acids had the same effect. Probably the effect of all these compounds on protease synthesis is produced through their influence on pH development during cultivation.
The processing of culture filtrates or clarified extracts follows the general description of recovery.
Serine alkaline proteases of bacterial origin are used in large amounts in laundering and to a lesser extent in leather tanning and the food industry.
Enzymes of this type play a less important role in commercial applications than the serine and acid proteinases. This is mainly due to their relatively poor stability. Metalloproteinases exhibit maximum activity at pH 7 to 8. In the majority of cases they contain a Zn atom in their active center. They are inhibited by metal chelating agents such as ethylenediaminetetraacetate (EDTA) or o-phenanthroline (OP), but not by DFP or thiol reagents.
Regarding their pH activity metalloproteinases are divided into neutral and alkaline types. The neutral enzymes all have pH optima around pH 7. Their molecular weights are in the range of 35,000-45,000. The isoelectric points of the proteinases from Bacillus subtilis, Pseudomonas aeruginosa, and Streptomyces have been determined to be at pH 9.0, 5.9, and 4.2, respectively. Neutral metalloproteinases of bacterial and fungal origin are specific toward hydrophobic or bulky amino acid residues on the amino side of the cleavage point.
In general, these enzymes are the least stable of the microbial proteases. The enzyme from Bacillus subtilis retains only about 10% of its activity after treatment at 60°C and pH 7 for 15 min. Bacillus thermoproteolyticus produces a very stable neutral protease (thermolysin) that retains 50% of its activity after 60 min at 80°C. Neutral proteases tend to undergo very rapid autolysis, which makes their recovery and application difficult.
Neutral proteases are widespread in microorganisms, both fungi and bacteria. Strains used for industrial production belong to the genera Aspergillus, Bacillus, and Streptomyces. Many of the organisms used for commercial production of neutral proteases also produce alkaline or acid proteases.
Only few strains have been found which synthesize neutral protease free of accompanying serine and acid proteases. Such strains are, for example, Bacillus cereus ATCC 14579 and NCTC 945, Bacillus megaterium ATCC14 581 and MA, and Bacillus polymyxa ATCC 842.
The formation of neutral proteases by bacteria does not seem to be correlated with sporulation. In Bacillus subtilis enzyme synthesis is subject to catabolite repression. Fogarty and Griffin (1973) found that the enzyme was produced irrespective of the C source used, but that the nature of the peptone had a marked effect on protease accumulation.
Without pH adjustment during the fermentation, the culture produced the neutral protease parallel to growth, and enzyme formation reached its maximum toward the end of the log phase. The yield was 15 times that obtained when the culture was run with a “fixed” pH of 6.8. According to Kalunyants et al. (1974), the best medium pH (6.9) can be achieved by separate sterilization of carbohydrate and N-containing compounds of the medium.
Variable temperature during fermentation (45°C at the beginning and lowering it to 43°, 40°, and 37°C, respectively during the 1st, 2nd, and 3rd 2-hr period) was preferable to a constant temperature. Zn+ +, Ca+ +, and Mn++ exert a beneficial effect on the level of the metalloproteinase.
In cultures of Aspergillus species, biosynthesis and externalization of neutral protease are repressed by low molecular weight sources of C, N, and S. Protease production and release occur when the medium is deficient for any of these elements. For Aspergillus terricola it has been shown that the enzyme accumulation in the medium was maximal when the N/C ratio was 0.5.
Due to their high instability, processing of metalloproteinases may lead to high activity losses. Therefore, the main problem in the concentration and purification of the enzyme is its stabilization. This can be achieved by strictly observing the tolerated range of pH, by the presence of metal ions (Zn++ for activity, Ca++ for stability), and by elimination of alkaline protease activity. One has also to take into consideration that the pigment complex produced by an organism can act as an inhibitor of the neutral protease of this organism, as shown for Bacillus mesentericus by Velcheva and Kolev (1975).
The application of neutral metalloproteinase is very limited because of the mentioned instability of these enzymes. Actual and potential uses are treatment of beer, application in bakeries, and reduction of dental plaque in humans.
These enzymes are without doubt the most interesting group of proteases with respect to use in the food industry. They are characterized by maximum activity and stability at pH 2.0-5.0. The molecular weight is around 35,000. Acid proteinases are low in basic amino acid content and have low isoelectric points.
They are insensitive to SH- reagents, metal chelators, heavy metals, and DFP and are generally stable in the acid pH range (pH 2-6), but are rapidly inactivated at higher pH values. The acid proteases exhibit limited esterase activity, but split a wide range of peptide bonds.
Acid proteinases of commercial importance are prepared exclusively from’ fungal sources and are tentatively divided into two subgroups by their physiological characteristics- pepsin-like acid proteinases and rennin-like proteinases.
Pepsin-like acid proteinases have usually been reported in the group of black aspergilli, such as Aspergillus niger, Aspergillus awamori, Aspergillus usamii, and Aspergillus saitoi, but also occur in species of Penicillium, Rhizopus, and others. To a large extent they are produced in solid substrate cultures. Biosynthesis of these enzymes is favored by high C/N ratios.
Inorganic N sources show an inhibiting effect on the production of acid proteinases, whereas peptone was found highly effective in inducing this enzyme in a strain of Aspergillus niger. The inducing action of peptone was much more remarkable when this material was added to a culture during the growth phase than in the stationary phase.
From this finding it is evident that the increase in acid proteinase activity observed when growth has ceased represents de novo synthesis. It has been shown to be due to exhaustion of adenine-group growth substances. Adipic and glutaric acids were also highly effective in supporting enzyme formation, but amino acids tested and some dipeptides were less effective than peptone.
Acid proteinases of the pepsin type play an important role in the production of fermented foods by molds from soybeans, rice, and other cereals. They are further used in the baking industry for the modification of wheat proteins in bread doughs.
Rennin-like acid proteinases are produced by strains of Mucor miehei, Mucor pusillus, Endothia parasitica, and Trametes sanguinea. The enzyme from the Mucor species has been, and is now, produced by the solid substrate culture method. However, Aunstrup (1974) isolated a strain of Mucor miehei, which he succeeded in growing in submerged culture for rennin production.
Microbial rennet substitutes must be freed of lipase to avoid rancidity of the cheese. This can be achieved by controlled heating or by adjusting to a low pH. Unspecific proteolytic enzymes, which may cause bitter taste of the cheese, must also be removed.
Their separation is obtained by adsorption on aluminosilicates. These adsorbents are particularly advantageous because they can be mixed into the culture liquid at the end of the fermentation, even in the presence of the medium, with a good separation effect and without any loss of milk clotting activity. Bentonite, permutite, and attapulgite are also suitable.
Because of their particular properties, rennet-like microbial proteases are used for clotting of milk in cheese manufacture. The process is based on the coagulation of casein under the influence of the rennet-like protease. It is known that the casein in milk is mainly composed of αs-, β-, and κ-casein. In particular, κ-casein plays an important role in the coagulation process, because it keeps the casein micelles present in milk in solution and protects them against flocculation by calcium ions. The clotting effect of rennins consists of the destabilization of the casein complex.
Two phases can be distinguished:
(1) The primary or enzymatic phase, in which the protective colloid (κ-casein) of the casein micelle is broken down and a glycomacro- peptide is split off as follows-
(2) The secondary or non-enzymatic phase, in which the coagulum is formed under the influence of calcium ions
The primary phase has a temperature coefficient, Q10, of about 2 (like most enzyme reactions), whereas the secondary phase has a Q10 of about 15. Therefore, it is reasonable to develop a system for continuous clotting of milk employing immobilized enzymes.
Consequently, in a 2-stage enzyme reactor the enzymatic phase is conducted in the first stage at low temperatures in order to inhibit the non-enzymatic phase. In the second stage subsequent warming clots the milk by the action of calcium ions. The whole process of cheese manufacturing is schematically illustrated in Fig. 15.5.
The enzyme lipase catalyzes the reaction:
This reaction goes to completion, i.e., until glycerol and free fatty acids are formed.
Many, perhaps most, bacteria and fungi produce lipase. Potent producers are among the fat-producing microorganisms. However, no distinct relationship between capacity of fat production and lipase production has been found. The enzyme from Candida cylindracea is commercially available.
Other producers of lipase are, e.g., Geotrichum candidum, Rhizopus arrhizus, and Aspergillus niger. Geotrichum candidum lipase is unique with respect to its specificity properties. The enzyme from the thermophile, Humicola lanuginosa, exhibits better thermostability.
The production of lipase is markedly affected by many factors. Generally, synthetic media produce lower yields of lipase than complex media. Lipase formation is highly dependent on nutrient and physical conditions. In a number of cases, addition of lipid material or fatty acids to the culture medium was found to enhance lipase production.
In contrast, Smith and Alford (1966) observed inhibition of lipase formation in media supplied with lard, sodium oleate, or salts of other unsaturated fatty acids. These authors also reported that the inhibition was prevented, but not reversed, for example, by some divalent cations and Tweens.
Glucose is unsuitable as the C source and ammonium ions seem to be unsuitable as the N source. Incorporation of CaCO3 acts differently, depending on the strain used. In some cases promotion of lipase production was observed, whereas in other cases investigators found an inhibitory effect using CaCO3.
Sometimes lipase preparations prove to be unstable. This can be due to the presence of proteases, the removal of which will lead to stable products.
Despite the fact that there is a considerable industry based on fats there is little industrial application of lipases. The main use is as digestive aid.
Glucose oxidase (β-D-glucose-oxygen oxidoreductase, EC 126.96.36.199), also known as notatin, acts in the presence of molecular oxygen to convert glucose to gluconic acid and hydrogen peroxide-
It is highly specific for β-D-glucose, although slight activity is found with 2-deoxyglucose.
At present, glucose oxidase is commercially prepared from Aspergillus niger and Penicillium amagasakiense in submerged culture. It has also been reported that Penicillium notatum and Penicillium chrysogenum synthesize glucose oxidase on liquid media in surface culture, but not in submerged culture.
During the growth of the fungal culture, the enzyme occurs in the phase following the lag phase. By feeding glucose this phase can be extended and thus the enzyme yield enhanced. The special culture conditions, however, depend markedly on microbial strains used.
For example- beet molasses has proved to be a suitable carbon source in Penicillium purpurogenum, but not suitable in Penicillium chrysogenum; high aeration rates supported enzyme synthesis in Penicillium purpurogenum, but did not in Penicillium chrysogenum.
For the purpose of concentration and purification, glucose oxidase must be separated from cells by extraction. The crude solutions also contain catalase which may interfere with glucose oxidase in some applications. In these cases separation is conducted by adsorption of the catalase on alumina or kaolin. For preparation of solid products, glucose oxidase can be precipitated by neutral salts or solvents, but liquid preparations are preferred.
Glucose oxidase is a good glycoprotein. The enzyme from Aspergillus niger contains 10.5% carbohydrate, which is believed to contribute to the stability and not to affect the overall structure. Two FAD molecules per molecule of enzyme act as the prosthetic group. The molecular weight of the Aspergillus niger enzyme is 186,000, that of Penicillium notatum is 152,000. The optimum pH of glucose oxidase is about 5.5.
The enzyme is fully stable between pH 4 and 6 at 40°C for 2 hr. Specially stabilized preparations for use at pH 2.5 are available. Use above pH 8.0 may be possible, but requires a high glucose concentration. Glucose oxidase is very unstable above 50°C, although glucose has some protective effect. Normal increase in activity caused by increased temperatures is counteracted by decrease in dissolved oxygen concentration at higher temperatures.
According to the reaction equation, glucose oxidase can be used in order to remove glucose or oxygen or to form hydrogen peroxide or gluconic acid. Indeed, in food processing glucose oxidase finds application for removal of residual glucose prior to the preparation of dried eggs or to remove it from other products in order to reduce non-enzymatic browning.
It is highly effective in removing residual oxygen from beer, wine, fruit juices, high fat products (mayonnaise), or packaged dehydrated foods. In the treatment of flour, when the formation of peroxide is desired, only catalase-free preparations can be used.
Catalase (EC 188.8.131.52) splits hydrogen peroxide to water and oxygen-
The enzyme is widely distributed in microorganisms. Its biological role has been studied by a number of investigators. In methanol-utilizing yeasts, it is generally accepted that catalase must be involved in the metabolism of methanol, since hydrogen peroxide is liberated during methanol oxidation by alcohol dehydrogenase. This suggestion is supported by the fact that catalase is markedly induced when the yeast cells are grown on methanol.
Commercially, catalase is prepared from Aspergillus niger, Penicillium vitale, or Micrococcus lysodeikticus. It is a hemo-protein containing 4 ferriprotoporphyrin prosthetic groups per molecule of enzyme, with a molecular weight of 250,000. The optimum pH of Aspergillus niger catalase is at pH 6.0; 75% of its optimum activity occurs between pH 3.0 and 9.0. The enzyme is inactivated by cyanides, phenols, alkali, urea, freezing, and by sunlight under aerobic conditions.
Production of catalase is conducted in deep tank cultivation. Biosynthesis occurs simultaneously with glucose oxidase formation. The ratio of these enzymes is controlled by quality and quantity of the inoculum, the composition of the medium, and by aeration conditions.
With Penicillium vitale, Nikolskaya et al. (1972) found that greater amounts of catalase were accumulated when the C:N ratio in the medium was as high as 12:1. Optimum pH was 4.5-5.5 during the first 48-72 hr of growth at 26°-27°C. L-Cysteine and DL-methionine promoted catalase production by the same fungus.
The stimulating effect observed with CaCO3 was demonstrated to be not the result of neutralization. Ca++ was believed to facilitate the transport of the enzyme from the mycelium into the medium.
Catalase produced by the directed biosynthesis can be separated selectively from extracts containing catalase and glucose oxidase. The recovery of catalase from Micrococcus lysodeikticus starts with lysing of the cells in a solution of sodium chloride (0.5 to 2%).
The next step is fractionation of the lysate by centrifugation of a mixture of the lysate, an organic solvent (ethanol in an end concentration of 40 to 50% v/v), and a salt (sodium or potassium chloride adjusted to a concentration of 1 to 2% either before or after addition of the solvent). The dissolved catalase can then be precipitated from solution by adding ethanol to a final concentration of 75%.
Separation of catalase from solution can also be conducted by adsorption methods using alumina or kaolin as adsorbents. For commercial application, liquid preparations are preferred.
Catalase finds application wherever the removal of hydrogen peroxide is required or the controlled release of oxygen from hydrogen peroxide is desired. Therefore, in the food industry catalase is employed to remove the excess of hydrogen peroxide used for cold sterilization in milk and cheese processing. Catalase may also be employed in cake baking as well as in irradiated foods, in the process of which hydrogen peroxide is formed.
This enzyme converts D-glucose to D-fructose. The main substrate of this enzyme, however, is xylose and, indeed, the glucose isomerizing enzyme is a D-xyloseketoisomerase (EC 184.108.40.206) with side activities to n-glucose and D-ribose, as shown in Fig. 15.6.
A large number of genera of bacteria and some yeast have been found to produce a glucose isomerizing enzyme. But the strains most widely used as sources for commercial production are members of the genus Streptomyces. Outtrup (1974) found thermophilic atypical variants of Bacillus coagulans to be particularly suited as the enzyme source due to the properties of its glucose isomerase.
The isomerase in Streptomyces is an inducible enzyme which requires the presence of D-xylose in the culture medium for its production. Sanchez and Quinto (1975) selected a double mutant strain of Streptomyces phaeochromogenes which was able to produce glucose isomerase constitutively and were insensitive to catabolite repression.
Diers (1976), who worked with Bacillus coagulans in chemostat cultures, found that glucose isomerase production of this strain was regulated mainly by catabolic repression. The latter occurred when inorganic compounds limited growth, whereas carbon limitation and particularly carbon-oxygen limitation were advantageous.
Media for the commercial production of glucose isomerase are based on xylan or xylan-containing raw materials such as wheat bran, maize husks, sulfite liquor, etc. As the enzyme is of the intracellular type, it can be used in the form of whole cells. In 1969 Takasaki et al. described the immobilization of Streptomyces cells by heating them to over 60°C for about 10 min.
This procedure prevents autolysis of the cells and “fixes” the glucose isomerase. Purified preparations with higher activities can be obtained by application of the usual methods of cell rupture and solubilizing of the enzyme. After discarding the cellular material, the glucose isomerase is then adsorbed on DEAE-cellulose or a similar material, recovered, and washed.
Co and Mg ions are well-known activators of glucose isomerase and essential for obtaining maximum activity, whereas copper, nickel, and zinc are strong inhibitors. The other characteristics such as pH and temperature activity and stability vary with the enzyme source and depend on whether the enzyme is in the native or an immobilized form. In order to prevent alkaline conversion of fructose produced during the isomerization process it is necessary that the employed glucose isomerase be highly active at pH 6.5.
Principal producers and users of glucose isomerase are found in the corn wet milling industry, where the enzyme is used to convert glucose in corn syrups to fructose. Crystalline D-glucose can be used as a substrate for D-fructose production. However, in industrial practice, high D.E. starch hydrolysates are a more economical source of D-glucose.
Such a hydrolysate can be prepared by the combined action of bacterial α-amylase, amyloglucosidase, and isoamylase on starch to yield a glucose syrup of about 95-98 D.E. Subsequent isomerization is carried out in agitated vessels at 65°C and pH 7 for 18 to 24 hr using, for example- immobilized cells.
Following conversion to D-fructose, the immobilized enzyme is removed and the liquor further treated to give invert sugar syrup of about 45% fructose and 55% glucose.