Other Reactions of Proteins in Foods

Other Reactions of Proteins in Foods Reactions with Lipids Oxidation of unsaturated lipids leads to formation of alkoxy and peroxy free radicals. These free radicals in turn react with proteins, forming lipid-protein free radicals. These lipid-protein conjugated free radicals can undergo polymerization crosslinking of proteins. In addition, the lipid free radicals can also induce formation of protein free radicals at cysteine and histidine side chains, which may then undergo cross-linking and polymerization reactions. Lipid peroxides (LOOH) in foods can decompose, resulting in liberation of aldehydes and ketones, notably malonaldehyde. These carbonyl compounds react with amino groups of proteins via carbonyl-amine reaction and Schiff’s base formation. As discussed earlier, reaction of malonaldehye with lysyl side chains leads to cross-linking and polymerization of proteins. The reaction of peroxidizing lipids with proteins generally has deleterious effects on nutritional value of proteins. Noncovalent binding of carbonyl compounds to proteins also imparts off flavors. Reactions with Polyphenols Phenolic compounds, such as p-hydroxybenzoic acid, catechol, caffeic acid, gossypol, and quercein, are found in all plant tissues. During maceration of plant tissues, these phenolic compounds can be oxidized by molecular oxygen at alkaline pH to quinones. This can also occur by the action of polyphenoloxidase, which is commonly present in plant tissues. These highly reactive quinones can irreversibly react with the sulfhydryl and amino groups of proteins. Pag e 415 ———————————————————————————————————————- Reaction of quinones with SH and a-amino groups (N-terminal) is much faster than it is with e-amino groups. In addition, quinones the oxidation products of phenolic compound, can also undergo condensation reactions, resulting in formation of high molecular weight brown color pigments sometimes referred to as tannins. Tannins remain highly reactive and readily combine with SH and amino groups of proteins. Quinone-amino group reactions decrease the digestibility and bioavailability of proteinbound lysine and cysteine. Reactions with Halogenated Solvents Halogenated organic solvents are often used to extract oil and some antinutritive factors from oilseed products, such as soybean and cottonseed meals. Extraction with trichloroethylene results in formation of a small amount of S-dichlorovinyl-L-cysteine, which is toxic. On the other hand, the solvents dichloromethane and tetrachloroethylene do not seem to react with proteins. 1,2- Dichloroethane reacts with Cys, His, and Met residues in proteins. Certain fumigants, such as methyl bromide, can alkylate Lys, His, Cys, and Met residues. All of these reactions decrease the nutritional value of proteins, and some are of concern from a safety standpoint. Reactions with Nitrites Reaction of nitrites with secondary amines, and to some extent with primary and tertiary amines, results in formation of Nnitrosoamines, which are among the most carcinogenic compounds formed in foods. Nitrites are usually added to meat products to improve color and to prevent bacterial growth. The amino acids (or residues) primarily involved in this reaction are Pro, His, and Trp. Arg, Tyr, and Cys also can react with nitrites. The reaction occurs mainly under acidic conditions and at elevated temperatures. (108) The secondary amines produced during the Maillard reaction, such as Amadori and Heyns products, also can react with nitrites. Formation of N-nitrosamines during cooking, grilling, and broiling of meat has been a major concern, but additives, such as ascorbic acid and erythorbate, are effective in curtailing this reaction. Reaction with Sulfites Sulfites reduce disulfide bonds in proteins to yield S-sulfonate derivatives. They do not react with cysteine residues. (109) In the presence of reducing agents, such as cysteine or mecaptoethanol, the S-sulfonate deriva- Pag e 416 tives are converted back to cysteine residues. S-Sulfonates decompose under acidic (as in stomach) and alkaline pH to disulfides. The S-sulfonation does not decrease the bioavailability of cysteine. The increase in electronegativity and the breakage of disulfide bonds in proteins upon S-sulfonation causes unfolding of protein molecules, which affects their functional properties. 6.7.2 Changes in the Functional Properties of Proteins The methods or processes used to isolate proteins can affect their functional properties. Minimum denaturation during various isolation steps is generally desired because this helps retain acceptable protein solubility, which is often a prerequisite to functionality of these proteins in food products. In some instances, controlled or partial denaturation of proteins can improve certain functional properties. Proteins are often isolated using isoelectric precipitation. The secondary, tertiary, and quaternary structures of most globular proteins are stable at their isoelectric pH, and the proteins readily become soluble again when dispersed at neutral pH. On the other hand, protein entities such as casein micelles are irreversibly destabilized by isoelectric precipitation. The collapse of micellar structure in isoelectrically precipitated casein is due to several factors, including solubilization of colloidal calcium phosphate and the change in the balance of hydrophobic and electrostatic interactions among the various casein types. The compositions of isoelectrically precipitated proteins are usually altered from those of the raw materials. This is because some minor protein fractions are reasonably soluble at the isoelectric pH of the major component and are therefore do not precipitate. This change in composition affects the functional properties of the protein isolate. Ultrafiltration (UF) is widely used to prepare whey protein concentrates (WPC). Both protein and nonprotein composition of WPC are affected by removal of small solutes during UF. Partial removal of lactose and ash strongly influences the functional properties of WPC. Furthermore, increased protein-protein interactions occur in the UF concentrate during exposure to moderate temperatures (50–55°C), and this decreases solubility and stability of the ultrafiltered protein, which in turn changes its water binding capacity and alters its properties with respect to gelation, foaming, and emulsification. Among the ash constituents, variations in calcium and phosphate content significantly affect the gelling properties of WPC. Whey protein isolates prepared by ion exchange contain little ash, and because of this they have functional properties that are superior to those of isolates obtained by ultrafiltration/diafiltration. Calcium ions often induce aggregation of proteins. This is attributable to formation of ionic bridges involving Ca2+ ions and the carboxyl groups. The extent of aggregation depends on calcium ion concentration. Most proteins show maximum aggregation at 40–50 mM Ca2+ ion concentration. With some proteins, such as caseins and soy proteins, calcium aggregation leads to precipitation, whereas in the case of whey protein isolate a stable colloidal aggregate forms (Fig. 27). Exposure of proteins to alkaline pH, particularly at elevated temperatures, causes irreversible conformational changes. This is partly because of deamidation of Asn and Gln residues, and b-elimination of cystine residues. The resulting increase in the electronegativity and breakage of disulfide bonds causes gross structural changes in proteins exposed to alkali. Generally, alkalitreated proteins are more soluble and possess improved emulsification and foaming properties. Hexane is often used to extract oil from oilseeds, such as soybean and cottonseed. This treatment invariably causes denaturation of proteins in the meal, and thus impairs their solubility and other functional properties. Pag e 417 FIGURE 27 Salt concentration versus turbidity of whey protein isolate (5%) in CaCl2 ( ) and Mg Cl2 ( ) solutions after incubating for 24 h at ambient temperature. (From Ref. 121.) The effects of heat treatments on chemical changes in, and functional properties of, proteins are described in Section 6.6. Scission of peptide bonds involving aspartyl residues during severe heating of protein solutions liberates low-molecular-weight peptides. Severe heating under alkaline and acid pH conditions also causes partial hydrolysis of proteins. The amount of lowmolecular-weight peptides in protein isolates can affect their functional properties. 6.8 Chemical and Enzymatic Modification of proteins 6.8.1 Chemical Modifications The primary structure of proteins contains several reactive side chains. The physicochemical properties of proteins can be altered, and their functional properties can be improved by chemically modifying the side chains. However, it should be cautioned that although chemical derivatization of amino acid side chains can improve functional properties of proteins, it can also impair nutritional value, create some amino acid derivatives that are toxic, and pose regulatory problems. Since proteins contain several reactive side chains, numerous chemical modifications can be achieved. Some of these reactions are listed in Table 6. However, only a few of these reactions may be suitable for modification of food proteins. The e-amino groups of lysyl residues and the Pag e 418 SH group of cyteine are the most reactive nucleophilic groups in proteins. The majority of chemical modification procedures involve these groups. Alkylation The SH and amino groups can be alkylated by reacting them with iodoacetate or iodoacetamide. Reaction with iodoacetate results in elimination of the positive charge of the lysyl residue, and introduction of negative charges at both lysyl and cyteine residues. (110) The increase in the electronegativity of the iodoacetate-treated protein may alter its pH-solubility profile, and may also cause unfolding. On the other hand, reaction with iodoacetamide results only in the elimination of positive charges. This will also cause a local increase in electronegativity, but the number of negatively charged groups in proteins will remain unchanged. Reaction with iodoacetamide effectively blocks sulfhydryl groups so disulfide-induced protein polymerization cannot occur. Sulfhydryl groups also can be blocked by reaction with N-ethylmaleimide (NEM). ——————————————————————————————————————————————————————————————————————————————– Amino groups can also be reductively alkylated with aldehydes and ketones in the presence of reductants, such as sodium borohydride (NaBH4) or sodium cyanoborohydride (NaCNBH3). In this case, the Schiff base formed by reaction of the carbonyl group with the amino group is subsequently reduced by the reductant. Aliphatic aldehydes and ketones or reducing sugars can be used in this reaction. Reduction of the Schiff base prevents progression of the Maillard reaction, resulting in a glycoprotein as the end product (reductive glycosylation). ———————————————————————————————————————- The physicochemical properties of the modified protein will be affected by the reactant used. Hydrophobicity of the protein can be increased if an aliphatic aldehyde or ketone is selected for the reaction, and the degree of hydrophobicity can be varied by changing the chain length of the aliphatic group. On the other hand, if a reducing sugar is selected as the reactant, then the protein will become more hydrophilic. Since glycoproteins exhibit superior foaming and emulsifying Pag e 419 properties (as in the case of ovalbumin), reductive glycosylation of proteins should improve solubility and interfacial properties of proteins. Acylation ———————————————————————————————————————- Amino groups can by acylated by reacting them with several acid anhydrides. The most common acylating agents are acetic anhydride and succinic anhydride. Reaction of protein with acetic anhydride results in elimination of the positive charges of lysyl residues, and a corresponding increase in electronegativity. Acylation with succinic or other dicarboxylic anhydrides results in replacement of positive charge with a negative charge at lysyl residues. This causes an enormous increase in the the electronegativity of proteins, and causes unfolding of the protein if extensive reaction is allowed to occur. Acylated proteins are generally more soluble than native proteins. In fact, the solubility of caseins and other less soluble proteins can be increased by acylation with succinic anhydride. However, succinylation, depending on the extent of modification, usually impairs other functional properties. For example, succinylated proteins exhibit poor heat-gelling properties, because of the strong electrostatic repulsive forces. The high affinity of succinylated proteins for water also lessens their adsorptivity at oil-water and air-water interfaces, thus impairing their foaming and emulsifying properties. Also, because several carboxyl groups are introduced, succinylated proteins are more sensitive to calcium induced precipitation than is the parent protein. Acetylation and succinylation reactions are irreversible. The succinyl-lysine isopeptide bond is resistant to cleavage catalyzed by pancreatic digestive enzymes. Furthermore, succinyl-lysine is poorly absorbed by the intestinal mucosa cells. Thus, succinylation and acetylation greatly reduce the nutritional value of proteins. The amphiphilicity of proteins can be increased by attaching long-chain fatty acids to the e-amino group of lysyl residues. This can be accomplished by reacting a fatty acylchloride or N-hydroxysuccinimide ester of a fatty acid with a protein. This type of modification can enhance lipophilicity and fat binding capacity of proteins, and can also facilitate formation of novel micellar structures and other types of protein aggregates. rylation Several natural food proteins, such as caseins, are phosphoproteins. Phosphorylated proteins are highly sensitive to calcium-ioninduced coagulation, which may be desirable in simulated cheese-type products. Proteins can be phosphorylated by reacting them with phosphorus oxychloride POCl3. Phosphorylation occurs mainly at the hydroxyl group of seryl and threonine residues and at the amino group of lysyl residues. Phosphorylation greatly increases protein electronegativity. Phosphorylation of amino groups results in addition of two negative charges for each positive charge eliminated by the modification. Under certain reaction conditions, especially at high protein concentration, phosphorylation with POCl3 can lead to polymerization of proteins, as shown here. Such polymerization reactions tend to minimize the increases in electronegativity and calcium sensitivity of the modified protein. The N-P bond is acid labile. Thus, under the conditions prevailing in the stomach, the N-phosphorylated proteins would be expected to undergo dephosphorylation and regeneration of lysyl residues. Thus, digestibility of lysine is probably not significantly impaired by chemical phosphorylation. (117) (118) Pag e 421 Sulfitolysis Sulfitolysis refers to conversion of disulfide bonds in proteins to the S-sulfonate derivative using a reduction-oxidation system involving sulfite and copper (CuII) or other oxidants. The mechanism is shown here. Addition of sulfite to protein initially cleaves the disulfide bond, resulting in the formation of one and one free thiol group. This is a reversible reaction, and the equilibrium constant is very small. In the presence of an oxidizing agent, such as copper (II), the newly liberated SH groups are oxidized back to either intra- or intermolecular disulfide bonds, and these, in turn, are again cleaved by sulfite ions present in the reaction mixture. The reduction-oxidation cycle repeats itself until all of the disulfide bonds and sulfhydryl groups are converted to the S-sulfonate derivative [41]. (119) Both cleavage of disulfide bonds and incorporation of groups cause conformational changes in proteins, which affect their functional properties. For example, sulfitolysis of proteins in cheese whey dramatically changes their pH-solubility profiles (Fig. 28) [41]. Amino acids Several plant proteins are deficient in lysine and methionine. The nutritional value of such proteins can be improved by covalent attachment of methionine and lysine at the e-amino group of lysyl residues. This can be accomplished by using the carbodiimide method or by reaction of N-carboxy anhydride of methionine or lysine with protein. Of these two approaches, the N-carboxy anhydride coupling method is preferable, because use of the carbodiimide is regarded as too toxic. N-Carboxy anhydrides of amino acids are extremely unstable in aqueous solutions, and they readily convert to the corresponding amino acid form even at low moisture levels. However, covalent coupling of amino acids with proteins can be achieved by mixing the protein directly with the anhydride. The isopeptide bond formed with the lysyl residues is susceptible to hydrolysis by pancreatic peptidases; thus, lysyl residues modified in this manner are biologically available. (120) Esterification Carboxyl groups of Asp and Glu residues in proteins are not highly reactive. However, under acidic conditions, these residues can be esterified with alcohols. These esters are stable at acid pH; but are readily hydrolyzed at alkaline pH. Pag e 422 FIGURE 28 The pH versus protein solubility profile of ( ) raw sweet whey and ( ) sulfonated sweet whey. (From Ref. 40.) 6.8.2 Enzymatic Modifications Several enzymatic modifications of proteins/enzymes are known to occur in biological systems. These modifications can be grouped into six general categories: glycosylation, hydroxylation, phosphorylation, methylation, acylation, and cross-linking. Such enzymatic modifications of proteins in vitro can be used to improve their functional properties. Although numerous enzymatic modifications of proteins are possible, only a few of them are practical for modifying proteins intended for food use. Enzymatic Hydrolysis Hydrolysis of food proteins using proteases, such as pepsin, trypsin, chymotrypsin, papain, and thermolysin, alters their functional properties. Extensive hydrolysis by nonspecific proteases, such as papain, causes solubilization of even poorly soluble proteins. Such hydrolysates usually contain low-molecular-weight peptides of the order of two to four amino acid residues. Extensive hydrolysis damages several functional properties, such as gelation, foaming, and emulsifying properties. These modified proteins are useful in liquid-type products, such as soups and sauces, where solubility is a primary criterion, and also in feeding persons who might not be able to digest solid foods. Partial hydrolysis of proteins either by using site-specific enzymes (such as trypsin or chymotrypsin) or by control of hydrolysis time often improves foaming and emulsification properties, but not gelling properties. With some proteins, partial hydrolysis may cause a transient decrease in solubility because of exposure of buried hydrophobic regions. Pag e 423 Certain oligopeptides released during protein hydrolysis have been shown to possess physiological activities, such as opioid activity, immunostimulating activity, and inhibition of angiotension-converting enzyme. The amino acid sequences of bioactive peptides found in peptic digests of human and bovine caseins are shown in Table 22. These peptides are not bioactive in the intact protein, but become active once they are released from the parent. Some of the physiological effects of these peptides include analgesia, catalepsy, sedation, respiratory depression, hypotension, regulation of body temperature and food intake, suppression of gastric secretion, and modification of sexual behavior [17]. When hydrolyzed, most food proteins liberate bitter-tasting peptides, which affect their acceptability in certain applications. The bitterness of peptides is associated with their mean hydrophobicity. Peptides that have a mean hydrophobicity value above 5.85 kJ/mol are bitter, whereas those with less than 5.43 kJ/mol are not. The intensity of bitterness depends on the amino acid composition, and sequence and the type of protease used. Hydrolysates of hydrophilic proteins, such as gelatin, are less bitter than the hydrolysates of hydrophobic proteins, such as caseins and soy protein. Proteases that show specificity for cleavage at hydrophobic residues produce hydrolysates that are less bitter than those enzymes that have broader specificity. Thus, thermolysin, which specifically attacks the amino side of hydrophobic residues, produces hydrolysates that are less bitter than those produced by low specificity trypsin, pepsin, and chymotrypsin [1]. Plastein Reaction The plastein reaction refers to a set of reactions involving initial proteolysis, followed by resynthesis of peptide bonds by a protease (usually papain or chymotrypsin). The protein substrate, at low concentration, is first partially hydrolysed by papain. When the hydrolysate containing the enzyme is concentrated to 30–35% solids and incubated, the enzyme randomly recombines the peptides, generating new polypeptides. The plastein reaction also can be performed in a one-step process, in which a 30–35% protein solution (or a paste) is incubated with papain in the presence of L-cysteine [116]. Since the structure and amino acid sequence of plastein products are different from those of the original protein, they often display altered functional properties. When L-methionine is included in the reaction mixture, it is covalently incorporated into the newly formed polypeptides. Thus, the plastein reaction can be exploited to improve the nutritional quality of methionine- or lysine-deficient food proteins. TABLE 22 Opioid Peptides from Caseins Peptides Name Orig in and position in the amino acid sequence Tyr-Pro-Phe-Pro-Gly-Pro-lle b-Casomorphin 7 Bovine b-casein (60–66) Tyr-Pro-Phe-Pro-Gly b-Casomorphin 5 Bovine b-casein (60–64) Arg -Tyr-Leu-Gly-Tyr-Leu-Glu a-Casein exorphin Bovine a s1-casein (90–96) Tyr-Pro-Phe-Val-Glu-Pro-lle-Pro Human b-casein (51–58) Tyr-Pro-Phe-Val-Glu-Pro Human b-casein (51–56) Tyr-Pro-Phe-Val-Glu Human b-casein (51–55) Tyr-Pro-Phe-Val Human b-casein (51–54) Tyr-Gly-Phe-Leu-Pro Human b-casein (59–63) Source: Ref. 17. Pag e 424 TABLE 23 Covalent Attachment of Lysine and Methionine into Food Proteins by the Transg lutaminase-Catalyzed Reaction Amino acid content (g /100 g protein) Protein Control Trang lutaminase-treated Incorporation of methionine as1-Casein 2.7 5.4 b-Casein 2.9 4.4 Soybean 7S protein 1.1 2.6 Soybean 11S protein 1.0 3.5 Incorporation of lysine W heat g luten 1.5 7.6 Source: Ref. 49. Protein Cross-Linking Transglutaminase catalyzes an acyl-transfer reaction that leads to covalent cross-linking of lysyl residues (acyl acceptor) with glutamine residues (acyl donor) via an isopeptide bond. This reaction can be used to cross-link different proteins, and to produce new forms of food proteins that might have improved functional properties. At high protein concentration, transglutaminase-catalyzed cross-linking leads to formation of protein gels and protein films at room temperature [72,77,78]. This reaction also can be used to improve nutritional quality of proteins by cross-linking lysine and/or methionine at the glutamine residues (Table 23) [49]. ———————————————————————————————————————- Bibliography Bodwell, C. E., J. S., Adkins, and D. T. Hopkins (eds.)(1981). Protein Quality in Humans: Assessment and In Vitro Estimation„ AVI Publishing Co., Westport, CT. Ghelis, C., and J. Yon (1982). Protein Folding, Academic Press, New York. Hettiarachchy, N. S., and G. R. Ziegler (eds.)(1994). Protein Functionality in Food systems, Marcel Dekker, New York. Kinsella, J. E., and W. G. Soucie (eds.)(1989). Food Proteins, American Oil Chemists’ Society, Champaign, IL. Mitchell, J. R., and D. A. Ledward (eds.)(1986). Functional Properties of Food Macromolecules, Elsevier Applied Science, New York. Parris, N., and R. Barford (1991). Interactions of Food Proteins„ ACS Symposium Series 454, American Chemical Society, Washington, DC. Phillips, R. D., and J. W. Finley (eds.) (1989). Protein Quality and the Effects of Processing, Marcel Dekker, New York. Schulz, G. E., and R. H. Schirmer (1980). Principles of Protein Structure, Springer-Verlag, New York. Whitaker J. R., and S. R. Tannenbaum (1977). Food Proteins, AVI Publishing Co., Westport, CT. Whitaker, J. R., and M. Fujimaki (eds.) (1980). Chemical Deterioration of Proteins, ACS Symposium Series, American Chemical Society, Washington, DC. Pag e 425 References 1. Adler-Nissen, J. (1986). Relationship of structure to taste of peptides and peptide mixtures. In Protein Tailoring for Food and Medical Uses R. E. Feeney, and J. R. Whitaker, (eds.), Marcel Dekker, New york, pp. 97–122. 2. Ahren, T. J. and A. M. Klibanov (1985). The mechanism of irreversible enzyme inactivation at 100°C. Science 228:1280–1284. 3. Arakawa, T., and S. N. Timasheff (1982). Stabilization of protein structure by sugars. Biochemistry 21:6536–6544. 4. Arakawa, T., and S. N. Timasheff (1984). Mechanism of protein salting in and salting out by divalent cation salts: Balance between hydration and salt binding. Biochemistry 23:5912–5923. 5. Asakura, T., K. Adachi, and E. Schwartz (1978). Stabilizing effect of various organic solvents on protein. J. Biol. Chem 253:6423–6425. ———————————————————————————————————————- 6. Bigelow, C. C. (1967). On the average hydrophobicity of proteins and the relation between it and protein structure. J. Theor. Biol. 16:187–211. 7. Bigelow, C. C., and M. Channon (1976). Hydrophobicities of amino acids and proteins, in Handbood of Biochemistry and Molecular Biology (G. D. Fasman, ed.), 3rd ed. CRC Press, Boca Raton, FL pp. 209–243. 8. Blundell, T. L., and L. N. Johnson (1976). Protein Crystallography, Academic Press, London, p. 32. 9. Brandts, J. F. (1967). In Thermobiology (A. H. Rose, ed.), Academic Press, New York, pp. 25–72. 10. Brems, D. N. (1990). Folding of bovine growth hormone, in Protein Folding (L. M. Gierasch and J. King, eds.), American Association for the Advancement of Science, Washington, DC, p. 133. 11. Bull, H. B., and K. Breese (1973). Thermal stability of proteins. Arch. Biochem. Biophys. 158:681–686. 12. Bushuk, W., and F. MacRitchie (1989. Wheat Proteins: Aspects of structure that determine bread-making quality, in Protein Quality and the Effects of Processing (R. Dixon Phillips and J. W. Finley, eds.) Marcel Dekker, New York, pp. 345–369.——————————————————————————————————————————————————————————————————————————————– 13. Cameron, D. R., M. E., Weber, E. S. Idziak, R. J. Neufeld, and D. G. Cooper (1991). Determination of interfacial areas in emulsions using turbidimetric and droplet size data: Correction of the formula for emulsifying activity index. J. Agric. Food Chem. 39:655–659. 14. Chen, C., A. M. Pearson, and J. I. Gray (1990). Meat mutagens. Adv. Food Nutr Res. 34: 387–449. 15. Chen, B., and J. A. Schellman (1989). Low Temperature unfolding of a mutant of phage T4 lysozyme. 1. Equilibrium studies. Biochemistry 28:685–691. 16. Cherkin, A. D., J. L. Davis, and M. W. Garman (1978). D-Proline: Sterospecific-sodium chloride dependent lethal convulsant activity in the chick. Pharmacol. Biochem. Behav. 8:623–625. 17. Chiba, H., and M. Yoshikawa (1986). Biologically functional peptides from food proteins: New opioid peptides from milk proteins, in Protein Tailoring for Food and Medical Uses (R. E. Feeney and J. R. Whitaker, eds.), Marcel Dekker, New York, pp. 123–153. 18. Cuq, J. L., M. Provansal, F. Uilleuz, and C. Cheftel (1973). Oxidation of methionine residues of casein by hydrogen peroxide. Effects on in vitro digestibility. J. Food Sci. 38:11–13. 19. Damodaran, S. (1988). Refolding of thermally unfolded soy proteins during the cooling regime of the gelation process: Effect on gelation. J. Agric. Food Chem. 36:262–269. 20. Damodaran, S. (1989). Influence of protein conformation on its adaptability under chaotropic conditions. Int. J. Biol. Macromol. 11:2–8. 21. Damodaran, S. (1989). Interrelationship of molecular and functional properties of food proteins, in Food Proteins (J. E. Kinsella and W. G. Soucie, eds.), American Oil Chemists’ Society, Champaign, IL, pp. 21–51. 22. Damodaran, S. (1990). Interfaces, protein films, and foams. Adv. Food Nutr. Res. 34:1–79. 23. Damodaran, S. (1994). Structure-function relationship of food proteins, in Protein Functionality in Food Systems (N. S. Hettiarachchy and G. R. Ziegler, eds.), Marcel Dekker, New York, pp. 1–38. 24. Damodaran, S., and J. E. Kinsella (1980). Flavor-protein interactions: Binding of carbonyls to bovine serum albumin: Thermodynamic and conformational effects. J. Agric. Food Chem. 28:567–571. 25. Damodaran, S., and J. E. Kinsella (1981). Interaction of carbonyls with soy protein: Thermodynamic effects. J. Agric. Food Chem. 29:1249–1253. Pag e 426 26. Damodaran, S., and K. B. Song (1988). Kinetics of adsorption of proteins at interfaces: Role of protein conformation in diffusional adsorption. Biochim. Biophys. Acta 954:253–264. 27. Dickinson, E., and Y. Matsummura (1991). Time-dependent polymerization of b-lactoglobulin through disulphide bonds at the oil-water interface in emulsions. Int. J. Biol. Macromol. 13:26–30. 28. Eggum, B. O., and R. M. Beames (1983). The nutritive value of seed proteins, in Seed Proteins (W. Gottschalk and H. P. Muller, eds.), Nijhoff/Junk, The Hague, pp. 499–531. 29. Erbersdobler, H. F., M. Lohmann, and K. Buhl (1991). Utilizaiton of early Maillard reaction products by humans, in Nutritional and Toxicological Consequences of Food Processing (M. Friedman, ed.), Advances in Experimental Medicine and Biology, vol. 289, Plenum Press, New York, pp. 363–370. 30. FAO/WHO/UNU (1985). Energy and Protein Requirements, Report of a Joint FAO/WHO/UNU Expert Consultation. World Health Organization Technical Rep. Ser. 724, WHO, Geneva. 31. FAO/WHO (1991). Protein Quality Evaluation, Report of a Joint FAO/WHO Expert Consultation. FAO Food Nutr. Paper 51, FAO, Geneva, pp. 23–24. 32. Fay, L., U. Richli, and R. Liardon (1991). Evidence for the absence of amino acid isomerization in microwave-heated milk and infant formulas. J. Agric. Food Chem. 39:1857–1859. 33. Ferry, J. D. (1961). Viscoelastic Properties of Polymers, Wiley, New York, p. 391. 34. Ford, J. E. (1981). Microbiological methods for protein quality assessment, in Protein Quality in Humans: Assessment and in vitro estimation (C. E. Bodwell, J. S. Adkins, and D. T. Hopkins, eds.), AVI Publishing Co., Westport, CT. pp. 278–305. 35. Fransen, K., and J. E. Kinsella (1974). Parameters affecting the binding of volatile flavor compounds in model food system. I. Proteins. J. Agric. Food Chem. 22:675–678. 36. Friedman, M., and M. R. Gumbmann (1986). Nutritional improvement of legume proteins through disulfide interchange. Adv. Exp. Med. Biol. 199:357–390. 37. Fujita, Y., and Y. Noda (1981). The effect of hydration on the thermal stability of ovalbumin as measured by means of differential scanning calorimetry. Bull. Chem. Soc. Jpn. 54:3233–3234. 38. Gekko, K., and Y. Hasegawa (1986). Compressibility-structure relationship of globular proteins. Biochemistry 25:6563–6571. 39. Ghelis, C., and J. Yon (1982). Protein Folding, Academic Press, New York, p. 51. 40. Gonzalez, J. M., and S. Damodaran (1990). Recovery of proteins from raw sweet whey using a solid state sulfitolysis. J. Food Sci. 55:1559–1563. 41. Gonzalez, J. M., and S. Damodaran (1990). Sulfitolysis of disulfide bonds in proteins using a solid state copper carbonate catalyst. J. Agric. Food Chem. 38:149–153. 42. Gonzalez, O. N., and R. H. Tanchuco (1977). Chemical composition and functional properties of coconut protein isolate. Phil. J. Coco. Studies. 11:21–29. 43. Gumbmann, M. R., W. L. Spangler, G. M. Dugan, and J. J. Rackis (1986). Safety of trypsin inhibitors in the diet: Effects on the rat pancreas of long-term feeding of soy flour and soy protein isolate. Adv. Exp. Med. Biol. 199:33–79. 44. Harwalkar, V. R. and C.-Y. Ma (1989). Effects of medium composition, preheating, and chemical modification upon thermal behavior of oat globulin and b-lactoglobulin, in Food Proteins (J. E. Kinsdella, and W. G. Soucie, eds.), American Oil Chemists’ Society, Champaign, IL, pp. 210–231. 45. Hayase, F., H. Kato, and M. Fujimaki (1973). Racemization of amino acid residues in proteins during roasting. Agric. Biol. Chem. 37:191–192. 46. Hayashi, R. (1989). In Engineering and Food (W. E. L. Spiess and Schubert, H, eds.), Elsevier Applied Science, London, pp. 815–826. 47. He, X. M., and D. C. Carter (1992). Atomic structure and chemistry of human serum albumin. Nature 358:209–214. 48. Heremans, K. (1982). High pressure effects on proteins and other biomolecules. Annu. Rev. Biophys. Bioeng. 11:1–21 49. Ikura, K., M. Yoshikawa, R. Sasaki, and H. Chiba (1981). Incorporation of amino acids into food proteins by transglutaminase. Agric. Biol. Chem. 45:2587–2592. 50. Israelachvili, J., and R. Pashley (1982). The hydrophobic interaction is long range, decaying exponentially with distance. Nature 300:341–342. Enzymes are proteins with catalytic activity due to their power of specific activation and conversion of substrates to products: (1) Some of the enzymes are composed only of amino acids covalently linked via peptide bonds to give proteins that range in size from about 12,000 MW to those that are near 1,000,000 MW (Table 1). Other enzymes contain additional components, such as carbohydrate, phosphate, and cofactor groups. Enzymes have all the chemical and physical characteristics of other proteins (see Chap. 6). Composition-wise, enzymes are not different from all other proteins found in nature and they comprise a small part of our daily protein intake in our foods. However, unlike other groups of proteins, they are highly specific catalysis for the thousands of chemical reactions required by living organisms. Enzymes are found in all living systems and make life possible, whether the organisms are adapted to growing near 0°C, at 37°C (humans), or near 100°C (in microorganisms found in some hot springs). Enzymes accelerate reactions by factors of 103 to 1011 times that of non-enzyme-catalyzed reactions (108 to 1020 over uncatalyzed reactions; Table 2). In addition, they are highly selective for a limited number of substrates, since the substrate(s) must bind stereospecifically and correctly into the active site before any catalysis occurs. Enzymes also control the direction of reactions, leading to stereospecific product(s) that can be very valuable by-products for foods, nutrition, and health or the essential compounds of life. Enzymes are synthesized in vivo by living organisms, based on expression (translation) of specific genes. They are wild type (“nature” dictated sequences) enzymes that fit the definition already given. A few enzymes, such as ribonuclease, have been synthesized in the laboratory from the amino acids. However, the time, cost, and inefficiency of the process are prohibitive. Isozymes are enzymes from a single organism that have qualitatively the same enzymatic activity but differ quantitatively in activity and structure because of differences in amino acid sequences (different gene products) or differ quantitatively due to posttranslational modification (glycosylation, proteolytic activation, etc). They are found very frequently in organisms. There are also a few naturally occurring ribozymes, composed of ribonucleic acids, that slowly catalyze (one or two bonds per minute) the specific hydrolysis of phosphodiester bonds within the same TABLE 1 Molecular W eig hts of a Few Selected Enzymes Enzyme Molecular weig ht Enzyme Molecular weig ht Ribonuclease 13,683 Fumarase 194,000 Lysozyme 14,100 Catalase 232,000 Chymotrypsinog en 23,200 Aspartate transcarbamylase 310,000 b-Lactog lobulin 35,000 Urease 483,000 Alkaline phosphatase 80,000 b-Galactosidase 520,000 Polyphenol oxidase (mushrooom) 128,000 Glutamate dehydrog enase 2,000,000 Source:Adapted from Ref. 111, p. 41. Pag e 434 TABLE 2 Nature of Effect of Catalyst on E a and on Relative Rates of Some Reactions Substrate Catalyst Ea (kcal/mol) n´/na (25°C) Relative rates (25°C) H2O2 None 18.0 5.62 × 10-14 1.00 13.5 1.16 × 10-10 2.07 × 103 Catalase 6.4 1.95 × 10-5 3.47 × 108 Sucrose H + 25.6 1.44 × 10-19 1.00 Invertase 11.0 8.04 × 10-9 5.58 × 1010 Carbonic acid None 20.5 8.32 × 10-16 1.00 Carbonic anhydrase 11.7 2.46 × 10-9 2.98 × 106 ———————————————————————————————————————- 24.5 9.33 × 10-19 1.00 Urease 8.7 3.96 × 10-7 4.25 × 1011 aFraction of molecules with Ea or g reater. Source: Ref. 111, p. 324. ribonucleic acid or in different ribonucleic and deoxyribonucleic acids. These are important in vivo catalysts, but they will not be discussed further in this chapter. Some enzymes result from in vivo or in vitro chemical modification of one or more nucleotide bases in genes of the wild-type enzymes or by chemical synthesis. These mutated enzymes may result from environmentally caused random in vivo modification (by free radicals, for example) of one or more nucleotide bases of a gene or in vitro by selective and deliberate chemical changes in the DNA nucleotide sequence of a gene. The latter is accomplished using specific restriction enzymes and ligases or by the polymerase chain reaction (PCR). When deliberately modified or synthesized in the laboratory, enzyme mimics are called synzymes. Other types of synzymes include the abzymes and the pepzymes. The abzymes are laboratory- made catalytic antibodies. The binding sites of catalytic antibodies are tailored in vivo by use of a specific hapten, while the catalytic groups are added in vitro by specific chemical modification of one or more of the amino acid side chains. The pepzymes are synthesized in the laboratory to mimic the sequence and stereochemistry of the active site of an enzyme. Enzymes may also be chemically modified by deliberate or unintentional changes (Maillard reaction, for example) in one or more side chain groups of a specific amino acid. This can lead to qualitatively different multiple forms of enzymes or total loss of activity. 7.1.2 Catalysis Enzymes are positive catalysts; that is, they increase the rates of reactions by 103 to 1011 that of non-enzyme-catalyzed reactions (Table 2). A minimum of two events must occur for enzyme activity to be observed. First, the enzyme must bind (noncovalently) a compound stereo- specifically into the active site. Second, there must be chemical conversion of the initial compound into a new compound. This is shown in Equation 2, where E is free enzyme, S is free substrate, E.S is a noncovalent complex of enzyme and substrate, and P is the new compound (product) formed. (2) The E is generated and participates repeatedly in the reaction. Some compounds only bind to the Pag e 435 active site of the enzyme; they are not converted to new products. They are called competitive inhibitors, and no chemical changes occur in I. (3) 7.1.3 Regulation of Enzyme Reactions Enzyme activity can be controlled in a number of ways that are very important to food scientists. The velocity of an enzymecatalyzed reaction is usually directly proportional to the active enzyme concentration, and dependent (in a complex way) on substrate, inhibitor, and cofactor concentrations, and on temperature and pH [111]. As an example, it is well known that enzyme-catalyzed reactions occur more slowly when a food is placed in a refrigerator (~4°C). But the reactions do not stop at 4°C (or at 0°C). Most enzyme-catalyzed reactions decrease 1.4–2 times/10°C decrease in temperature. Therefore, at 5°C, the velocities would be 0.5 to 0.25 times the velocity at 25°C. Changing the pH of the system by 1 or 2 pH units from the pH-activity optimum can decrease enzymatic velocity to 0.5 or 0.1, respectively, of that at the pH optimum. Decreasing (by heating, breeding, or genetic engineering) the concentration of enzyme to 0.1 that normally present would decrease the enzyme activity to 0.1 that originally present. 7.1.4 Historical Aspects Enzymes are essential for all living organisms; therefore, they have existed since the beginning of life. They are important in ripening of fruits, fermentation of fruit juices, souring of milk, tenderization of meat, etc., and humans were aware of these changes early on. 7.1.4.1 Nature of Enzymes During the 1870–1890 period, there was a major debate between Pasteur [85], who believed that enzymes functioned only when associated with living organisms, and Liebig [72], who believed that enzymes continued to function in the absence of cells. This argument was settled in 1897 when Büchner [19] separated enzymes, including invertase, from yeast cells and showed that they were still active. The debate between Pasteur and Liebig had resulted in the terms organized ferments and unorganized ferments to differentiate between enzymes in yeasts and enzymes in the stomach. In 1878 Kühne proposed use of en zyme (Greek, “in yeast”) to describe both types of enzymes [68]. 7.1.4.2 Enzyme Specificity During the period of 1890–1940 much work was done on the specificity of enzymes. Emil Fischer and his students, Bergmann, Fruton, and others, synthesized many carbohydrates, peptides, and other compounds of known structures and determined whether they were hydrolyzed, and at what rate, by carbohydrases and proteases. They found that small changes, such as removal or change in configuration of one group, could lead to marked effects on rates of hydrolyses. This led Fischer [38] to hypothesize the lock-and-key analogy of enzyme- substrate interactions in 1894 in which the active site of the enzyme was considered to be rigid. This concept remained essentially unchallenged until 1959, when Koshland [65] suggested the induced fit hypothesis to be better supported by currently available data, since there appeared to be some flexibility in accommodation between enzyme and substrate when they complexed. The induced fit hypothesis retained the concept of a stereospecific complex Pag e 436 formation between enzyme and substrate, but it rejected the idea that the binding locus of the active site is rigid structure that retains its exact shape even in the absence of substrate. Much data subsequently proved the induced fit concept to be more appropriate than the lock-and-key concept [78]. 7.1.4.3 Enzyme Kinetics Study of enzyme activities requires kinetic approaches, not equilibrium approaches. Therefore, the development of the kinetics to deal with these reactions was very important. In 1902 Henri [46;46a] and Brown [18] independently suggested that the hyperbolic relationship between velocity (-dS/dt or dp/dt) and substrate concentration (Fig. 1) was due to an obligatory enzyme-substrate complex intermediate prior to conversion of the substrate to product (Eq. 2). When all the enzyme is saturated with substrate, the enzyme is operating at its maximum capacity and further increase in substrate concentration cannot increase the velocity further (Fig. 1). In 1913, Michaelis and Menten [79] showed that the hyperbolic relationship between velocity and substrate concentration (Fig. 1) can be expressed by (4) where n is the measured velocity, Vmax is the maximum velocity when enzyme is saturated with substrate, Km [=(k-1+k2)/k1 for Eq. 2] is the substrate concentration at which n=0.5Vmax, and [S]t is the substrate concentration at any time t in the reaction. For reasons that will be detailed later, the initial velocity, no, determined early in the reaction (5% substrate conversion to product) is used, so that [S][email protected][S]o, the starting substrate concentration. Lineweaver and Burk [76] in 1934 developed a reciprocal transform (1/n vs. 1/[S]) of Equation 4 that gives a linear rather than a hyperbolic relationship, making it easier to determine Km and Vmax. Beginning in 1958, kinetic equations dealing with multisubstrate-multiproduct Fig ure 1 Variation of observed velocity with substrate concentration [A o] for an enzyme-catalyzed reaction that follows Michaelis-Menten kinetics. (From Ref. 111, p. 168.) Pag e 437 enzyme-catalyzed reactions were developed for more complex cases than that of Equation 2 [22, 27, 115]. However, Equation 4 still fits all Michaelis-Menten type reactions, where steady-state concepts are valid. The numerical definitions of Vmax and Km change as the complexity of the reaction changes. In 1965, Monod et al. [80] and Koshland et al. [66] developed equations for allosteric-behaving enzymes. Other workers [44] developed equations for pre-steady-state treatment of enzyme-catalyzed reactions. 7.1.4.4 Enzyme Purification Purification of enzymes began only after 1920. Before 1920, it was generally thought that enzymes were proteins. However, during the period of 1922–1928, Willstätter and his colleagues purified horseradish peroxidase to the point where it had appreciable activity even when no protein could be detected by existing methods. Therefore, they concluded, incorrectly, that enzymes could not be proteins. In 1926, Sumner [98], at Cornell University, crystallized urease from jack beans and showed it to be a protein. This led to much debate between Willstätter, a noted German chemist, and Summer, with many European biochemists and chemists siding with Willstätter. Sumner was later awarded the Noble prize for crystallization of the first enzyme and for showing that enzymes are proteins. Beginning in the 1930s, and continuing to the present, much work has been devoted to enzyme purification using techniques such as chromatography (ion exchange, gel filtration, chromatofocusing, affinity, hydrophobic), several types of electrophoresis [regular polyacrylamide gel electrophoresis (PAGE), SDS-PAGE, isoelectric focusing], and techniques based on solubility and stability differences. Now nearly 3000 enzymes have been purified. All are proteins. 7.1.4.5 Enzyme Structure The first primary sequence of a protein, insulin (of 6000 MW), was determined by Sanger and co-workers in 1955 [90] after almost 10 years of methods development and application. In 1960 [49, 95], the primary sequence of the first enzyme, ribonuclease (13,683 MW), was determined. Many primary sequences of enzymes are now known, many via gene sequencing. The secondary and tertiary structures of ribonuclease were determined in 1967 [58]. Now several hundred secondary and tertiary structures of enzymes are known, including structures in solutions, by nuclear magnetic resonance (NMR). The first enzyme, ribonuclease, was completely synthesized chemically from amino acids in 1969 by two groups. Now, wild-type and mutant enzymes can be synthesized almost overnight by the PCR (polymerase chain reaction) method, once the gene has been isolated. Those of us who have observed advances in protein and enzyme chemistry since the 1940s are astounded and awed by the remarkable ease and speed at which enzyme sequences and structures can now be determined. 7.1.5 Literature on Enzymology Perhaps no other single topic has received so much attention from researchers as enzymes. Papers on enzymes can be found in almost any journal in the physical, chemical, or biological sciences, since they are of interest to chemists, physicists, mathematicians, chemical engineers, and the life scientists, including food scientists and nutritionists. The bibliography at the end of this chapter includes selected journals, books, and monographs on enzymes. Obviously, no scientist can keep up completely in this field. Food scientists should, by all means, be familiar with current progress in fundamental aspects of enzymes, as well as with applications. Pag e 438 7.2 Enzyme Nomenclature The first enzyme to be named was catalase, which converts hydrogen peroxide to water and O2 (Eq. 5). Its name was derived by using the stem of catalyst and adding ase. This generic type name is unfortunate considering the many enzymes that have since been discovered. ———————————————————————————————————————- The name of the second enzyme, diastase, was derived from the Greek word diastasis, meaning to separate. The name of the third enzyme, peroxidase, was based on one of the substrates being peroxide. Polyphenol oxidase was so named because it oxidizes numerous phenols, while invertase inverts the optical rotation of a solution of sucrose ([a]=+66.5°) to [a]=-19.7° due to formation of equimolar concentrations of glucose ([a]=+52.7°) and fructose ([a]=-92°). Note that two general principles began to be used in naming some enzymes: -ase to designate an enzyme, while in two cases the stem was derived from the name of one of the substrates (polyphenol oxidase from phenols and peroxidase from peroxide). In the case of polyphenol oxidase, it is clear the enzyme catalyzes oxidation of phenols. 7.2.1 The Enzyme Commission As more enzymes were discovered nomenclature became an increasingly severe problem. Therefore, an International Commission on Enzymes of the International Union of Biochemistry was established and its charge was “to consider the classification and nomenclature of enzymes and coenzymes, their units of activity and standard methods of assays, together with the symbols used in the description of enzyme kinetics.” The Report of the Commission (1961) contained 712 enzymes. Subsequent updates were issued, the most recent being Enzyme Nomenclature (1992), which contains 3,196 enzymes [35]. 7.2.1. Rules for Naming The basis for the classification adopted by the Enzyme Commission was to divide the enzymes into groups on the basis of the type of reaction catalyzed, and this, together with the name(s) of the substrate(s), provided a basis for naming individual enzymes. There are six types of reactions catalyzed by enzymes: (a) oxidoreduction, (b) transfer, (c) hydrolysis, (d) formation of double bonds without hydrolysis, (e) isomerization, and (f) ligation. The names of the type of enzymes are formed by adding -ase to the stem of the type of reaction catalyzed. Thus the corresponding six groups of enzymes are (a)oxidoreductases, (b) transferases, (c) hydrolases, (d) lyases, (e) isomerases, and (f) ligases. 7.2.1.2 Overall Reaction as Basis of Nomenclature The overall reaction, as expressed by the formal chemical equation (eq. 6), is the basis for the nomenclature. Intermediate steps in the reaction reflecting the mechanism are not taken into account. Therefore, an enzyme cannot be named properly by this system until the substrate(s) and chemical nature of the reaction catalyzed are known. (6) Pag e 439 7.2.1.3 Three-Tier Classification The Enzyme Commission assigned each enzyme three designations: a systematic name, a trivial name, and an Enzyme Commission (EC) number. Generally, the systematic name is composed of two principal parts. The first consists of the name(s) of the substrate(s); in the case of two or more substrates (or reactants), the name of the substrates are separated by a colon. The second part of the systematic name, ending in -ase, is based on one of the six types of chemical reactions catalyzed (see Sec. 7.2.1.1). When the overall reaction involves two different chemical reactions, such as oxidative demethylation, the second type of reaction is listed in parenthesis following the first chemical reaction—for example, “sarcosine:oxygen oxidoreductase (demethylating).” The trivial name is one that is generally recognized and in common use such as a-amylase, cellulase, trypsin, chymotrypsin, peroxidase, or catalase. It is usually a shorter name than the systematic name. The number system derives directly from the classification scheme, and each number contains four digits, separated by periods and preceded by EC. The numbers are permanent. Newly discovered enzymes are placed at the end of the list under appropriate headings. If the classification of an enzyme is changed, the number remains in the listing, but the user is directed to the new listing, including the new number of the enzyme. 7.2.1.4 Six Main Types of Enzymes The six main types of enzymes, based on the chemical reaction catalyzed, are further explained in this section. Oxidoreductases Oxidoreductases are enzymes that oxidize or reduce substrates by transfer of hydrogens or electrons or by use of oxygen. The systematic name is formed as “donor:acceptor oxidoreductase.” An example, including systematic name followed by trivial name and EC number in parenthesis, is [35] H2O2 + H2O2 = O2 + 2H2O which gives hydrogen peroxide:hydrogen peroxide oxidoreductase (catalase, EC 1.11.1.6). Transferases Transferases are enzymes that remove groups (not including H) from substrates and transfer them to acceptor molecules (not including water). The systematic name is formed as “donor:acceptor group-transferred-transferase.” An example is: ATP + D-glucose = ADP + D-glucose 6-phosphate which gives ATP:D-glucose 6-phosphotransferase (glucokinase, EC 2.7.1.2). Note that the position to which the group is transferred is given in the systematic name when more than one possibility exists. Hydrolases Hydrolases are enzymes in which water participates in the breakage of covalent bonds of the substrate, with concurrent addition of the elements of water to the principles of those bonds. The systematic name is formed as “substrate hydrolase.” Water is not listed as a substrate, even though it is, because it is 55.6 M and the concentration does not change significantly during the Pag e 440 reaction. When the enzyme specificity is limited to removal of a single group, the group is named as a prefix, for example, “adenosine aminohydrolase.” Another example is Triacylglycerol + H2O = diacylglycerol + a fatty acid anion catalyzed by triacylglycerol acylhydrolase (triacylglycerol lipase, EC 3.1.1.3). Lyases Lyases are enzymes that remove groups from their substrates (not by hydrolysis) to leave a double bond, or which conversely add groups to double bonds. The systematic name is formed as “substrate prefix-lyase.” Prefixes such as “hydro-” and “ammonia-” are used to indicate the type of reaction—for example, “L-malate hydro-lyase” (EC 4.2.1.2). Decarboxylases are named as carboxy-lyases. A hyphen is always written before “lyase” to avoid confusion with hydrolases, carboxlases, etc. An example is: (S)-Malate = fumarate + H2O using the enzyme (S)-malate hydro-lyase (fumarate hydratase, EC 4.2.1.2; formerly known as fumarase). Isomerases Isomerases are enzymes that bring about an isomerization of substrate. The systematic name is formed as “substrate prefixisomerase.” The prefix indicates the types of isomerization involved, for example, “maleate cis-trans- isomerase” (EC 5.2.1.) or “phenylpyruvate keto-enolisomerase” (EC 5.3.2.1). Enzymes that catalyze and aldose-ketose interconversion are known as “ketol-isomerases,” for example, “L-arabinose ketol-isomerase” (EC 5.3.1.4). When the isomerization consists of an intramolecular transfer of a group, such as 2-phospho-D-glycerate = 3-phospho-D-glycerate, the enzyme is named a “mutase,” for example, “D-phosphoglycerate 2,3-phosphomutase” (EC 5.4.2.1). Isomerases that catalyze inversions of asymmetric groups are termed “racemases” or “epimerases,” depending on whether the substrate contains one or more than one center of asymmetry, respectively. A numerical prefix is attached to the word “epimerase” to show the position of inverstion. An example is L-Alanine = D-alanine with alanine racemase (alanine recemase, EC 5.1.1.1). Ligases Ligases are enzymes that catalyze the covalent linking together of two molecules, coupled with the breaking of a pyrophosphate bond as in ATP. This group of enzymes has previously been referred to as the “synthetases.” The systematic name is formed as “X:Y ligase (Z),” where X and Y are the two molecules to be joined together. The compound Z is the product formed from the triphosphate during the reaction. An example is: ATP + L-aspartate + NH3 = AMP + pyrophosphate + L-asparagine with L-aspartate:ammonia ligase (AMP-forming) (aspartate-ammonia ligase, EC 6.3.1.1). 7.2.2 Numbering and Classification of Enzymes A key to the numbering and classification of enzymes, in the extensive listings in Enzyme Nomenclature (1992) [35], is given in Table 3. The subclassification of the enzymes is best Pag e 441 determined from this type of key. For example, it is easy to see how the number system is derived. As noted previously, the first EC number indicates one of the six types of chemical reactions catalyzed by enzymes of that type. Using the EC 1. catagory, the second number indicates the nature of the donor substrate, the third number the nature of the acceptor molecule (not shown in Table 3) and the fourth number (not shown in Table 3) is the number of the specific enzyme. For example, EC 1.1.1.1 is alcohol dehydrogenase (alcohol:NAD+ oxidoreductase), where the second number indicates the donor substrate is a primary alcohol, the third number that the acceptor molecule is NAD+ , or NADP+ , and the fourth number is the specific enzyme alcohol dehydrogenase. EC 1.6.2.x would be NADH (or NADPH): cytochrome oxidoreductases. 7.3 Enzymes in Organisms All organisms contain many enzymes. The raw materials that are converted to foods contain hundreds dreds of different enzymes. These enzymes are important in the growth and maturation of the raw materials, and the enzymes continue to be active after harvest until all the substrate is exhausted, or the pH changes to one where the enzyme is no longer active, or the enzyme is denatured by an imposed treatment (pH, heating, chemicals, etc.). Often, enzyme activity increases during storage because of distintegration of cellular structure that separates enzymes and their substrates. As an example, softening of tomatoes acelerates following optimum maturity due to increased enzyme activity. Plant tissues contaminated with microorganisms, such as brown rot fungi containing polyphenol oxidase, can literally be lost overnight due to the rapid growth and multiplication of enzymecontaining microorganisms. Bruising of a fruit (i.e. apple) or vegetable (i.e. potato) leads to rapid browning due to polyphenol oxidase. This is because O2 from the atmosphere, one of the required substrates, can more easily penetrate the damaged skin of the fruit or vegetable. An intact skin is one of the best protections against browning. Unfortunately, up to 50% of tropical fruits are lost due to improper handling, shipping, and storage, and this is caused by uncontrolled activity of endogenous enzymes or exogenous enzymes present in attached microorganisms. 7.3.1 Location of Enzymes ———————————————————————————————————————- Enzymes are not uniformly distributed in organisms. A particular enzyme sometimes is found only in one type of organelle within a cell, as shown in Table 4. The nucleus (Fig. 2 and Table 4) contains primarily enzymes involved with nucleic acid biosynthesis and hydrolytic degradation. The mitochondria (Fig. 2) contain oxidoreductases associated with oxidative phosphorylation and formation of ATP, and the lysosomes and the pancreatic zymogen granules contain primarily hydrolases (Table 4). Each type of organelle in a cell is specialized in carrying out limited types of enzyme-catalyzed reactions. Organs may also be specialized in the types of enzymes found. In animals, the gastrointestinal tract contains primarily hydrolases designed to hydrolyze complex a-1,4-glucose-type carbohydrates, lipids, proteins, and nucleic acids to glucose, glycerol and fatty acids, amino acids, and purine and pyrimidines, respectively. Hydrolysis (digestion) of starch and glycogen by a-amylase begins in the mouth and is completed in the small intestine. Hydrolysis of proteins begins in the stomach (with pepsin) and is completed in the small intestine (with trypsin, chymotrypsin, carboxypeptidases A and B, aminopeptidases, and di- and tripeptidases). The first four proteolytic enzymes are synthesized in and secreated by the pancreas into the small intestine. Lipids are hydrolyzed by gastric (stomach) lipases and small-intestinal lipases. Nucleic acids are hydrolyzed by nucleases in the small intestine. Plant organs also contain specialized enzymes. For example, the seed contains substantial amounts of hydrolytic enzymes to hydrolyze starch and proteins, especially to feed the new seedling following seed germination. Pag e 442 TABLE 3 Key to Numbering and Classification of Enzymes a 1. Oxidoreductases 1.1 Acting on the CH—OH g roup of donors 1.2 Acting on the aldehyde or oxo g roup of donors 1.3 Acting on the CH—CH g roup of donor 1.4 Acting on the CH—NH2 g roup of donors 1.5 Acting on the CH—NH g roup of donors 1.6 Acting on NADH or NADPH 1.7 Acting on other nitrog enous compounds as donors 1.8 Acting on a sulfur g roup of donors 1.9 Acting on a heme g roup of donors 1.10 Acting on diphenols and related substances as donors 1.11 Acting on hydrog en peroxide as acceptor 1.12 Acting on hydrog en as donor 1.13 Acting on sing le donors with incorporation of molecular oxyg en (oxyg enases) 1.14 Acting on paired donors with incorporation of molecular oxyg en 1.15 Acting on superoxide radicals as acceptor 1.16 Oxidizing metal ions 1.17 Acting on CH2 g roups 1.18 Acting on reduced ferredoxin as donor 1.19 Acting on reduced flavodoxin as donor 1.97 Other oxidoreductases 2. Transferases 2.1 Transferring one-carbon g roups 2.2 Transferring aldehyde or ketone residues 2.3 Acyltransferases 2.4 Glycosyltransferases 2.5 Transferring alkyl or aryl g roups, other than methyl g roups 2.6 Transferring nitrog enous g roups 2.7 Transferring phosphorus-containing g roups 2.8 Transferring sulfur-containing g roups 3. Hydrolases 3.1 Acting on ester bonds 3.2 Glycosidases 3.3 Acting on ether bonds 3.4 Acting on peptide bonds (peptidases) 3.5 Acting on carbon-nitrog en bonds, other than peptide bonds 3.6 Acting on acid anhydrides 3.7 Acting on carbon-carbon bonds 3.8 Acting on halide bonds 3.9 Acting on phosphorus-nitrog en bonds 3.10 Acting on sulfur-nitrog en bonds 3.11 Acting on carbon-phosphorus bonds 4. Lyases ———————————————————————————————————————- 4.1 Carbon-carbon lyases 4.2 Carbon-oxyg en lyases 4.3 Carbon-nitrog en lyases 4.4 Carbon-sulfur lyases 4.5 Carbon-halide lyases 4.6 Phosphorus-oxyg en lyases 4.99 Other lyases (table continued on next page) Pag e 443 (table continued from previous page) 5. Isomerases 5.1 Racemases and epimerases 5.2 cis-trans-Isomerases 5.3 Intramolecular oxidoreductases 5.4 Intramolecular transferases (mutases) 5.5 Intramolecular lyases 5.99 Other isomerases 6. Lig ases 6.1 Forming carbon-oxyg en bonds 6.2 Forming carbon-sulfur bonds 6.3 Forming carbon-nitrog en bonds 6.4 Forming carbon-carbon bonds 6.5 Forming phosphoric ester bonds aThe third and fourth levels of classification are g iven in Ref. 35. Source: Ref.35, pp. v-xi, by courtesy of Academic Press. If we want to find a particular enzyme in plants, animals, and microorganisms, it is important to know in which organelle or organ an enzyme is located. Two techniques are important for this. The organs containing a particular enzyme can be determined by histochemical techniques (Fig. 3). As shown in Figure 3, alkaline phosphatase and polyphenol oxidase are not distributed uniformly in cells of porcine logissimus muscle and the beet, respectively. These sensitive enzyme histochemical techniques are used routinely by medical and veterinary histologists and pathologists [100]. Unfortunately, they are rarely used by food scientists. TABLE 4 Subcellular Location of Several Enzymes in Animal Cells Location Enzymes Nucleus DNA-dependent RNA polymerase, a polyadenylate synthetase Mitochondria Succinate dehydrogenase, cytochrome oxidase, g lutamate dehydrog enase, malate dehydrog enase, a-ketog lutarate dehydrog enase, a-g lyerol phosphate dehydrog enase, pyruvate decarboxylase Lysosomes Cathepsins A, B, C, D, and E, collag enase, acid ribonuclease, acid phosphatase,b-g alactosidase, sialidase, lysozyme, trig lyceride lipase Peroxisomes (microbodies) Catalase, urate oxidase, D-amino acid oxidase Reticulum, g olg i, etc. Glucose 6-phosphatase, nucleoside diphosphatase, TPNH-linked lipid peroxidase, nucleoside phosphatases Soluble Lactate dyhydrogenase, phosphofructokinase, glucose 6-phosphate dehydrogenase, transketolase, transaldolase Pancreatic zymog en g ranules Trypsinogen,b chymotrypsinogen, b lipase, amylase, ribonuclease aEnzymes in italics are most frequently used as indicator enzymes for type of cellular org anelles present in a homog enate. bZymog ens activated to trypsin and chymotrypsin, respectively. Source: Ref. 111, p. 68. Pag e 444 Fig ure 2 “Typical” cell. Diag ram of a typical cell based on data from electron microg raphs. (From Ref. 14, p. 55, by courtesy of Scientific American, Inc.) The second method used to determine which organelle contains a particular enzyme is to use differential centrifugation, following careful disintegration of the tissue in a buffer to control pH and osmotic pressure so that the organelles remain intact. The separated organelles are then disintegrated and the enzymes present are determined by use of specific substrates to detect their presence and concentration. The results of this type of study are shown in Table 4. 7.3.2 Compartmentation and Access to Substrate The activities of enzymes in intact cells are controlled by compartmentation in subcellular membranes, by organelles, by membrane-or cell-wall-bound enzymes and/or substrates, and by Pag e 445 Fig ure 3 Histochemical localization of enzyme activity. (a) Alkaline phosphatase activity in frozen section of porcine long issimus muscle detected with a-naphthyl phosphate and diazotized 4′-amino-2′, 5′-diethoxybenzanilide. (From Ref. 24, p. 300, by courtesy of Institute of Food Technolog ists.) (b) Transverse section of fresh beef tissue after treatment with dihydroxyphenylalanine for detection of polyphenol oxidase. (From Ref. 13, p. 579, by courtesy of the Institute of Food Technolog ists.) separation of the substrate from the enzyme, including exclusion of free O2 from the tissues. Other controls of activity include proenzyme (inactive until activated) biosynthesis (such as found in the blood and pancreas of animals) and by physiologically important endogenous enzyme inhibitors. Partial distintegration of the tissue, by aging or bruising, by insects or microorganisms, by intentional peeling, cutting, slicing, or blending, or by freezing and thawing, brings the enzymes and substrates together, allowing the enzymes to act very rapidly on their substrates. This can cause rapid changes in the color, texture, flavor, aroma, and nutritional qualities of food. Heat treatment, storage at low temperature, and/or use of enzyme inhibitors is necessary to stabilize the product. 7.3.3 Typical Concentrations of Enzymes in Some Foods Relative amounts of three enzymes, polygalacturonase, lipoxygenase, and peroxidase, in plants are shown in Table 5. Polygalacturonase, responsible for softening, varies widely in content in the plant sources listed, being very high in concentration in tomato and zero in cranbery, carrot, and grape. Lipoxygenase is very high in concentration in soybeans, accounting for the beany taste of cooked soybeans, but occurs at barely detectable levels in wheat and peanuts. Peroxidase is found in essentially all fruits, but varies some sevenfold from English green peas to lima beans. Polyphenol oxidase is one of the most noticeable enzymes in plants. It is present at high concentrations in some grapes (dark raisins), prune plums, Black Mission figs, dates, tea leaves, and coffee beans, where its action is desired. It is present at moderate concentrations in peaches, apples, bananas, potatoes, and lettuce, where its activity is undesirable, and is not present in peppers. Pag e 446 Table 5 Relative Amounts of Some Enzymes in Various Sources Enzyme Source Relative amounts Polyg alacturonase a Tomato 1.00 Avocado 0.065 Medlar 0.027 Pear 0.016 Pineapple 0.024 Cranberry 0 Carrot 0 Grape 0 Lipoxyg enase b Soybeans 1.00 Urd beans 0.60 Mung beans 0.47 Peas 0.35 W heat 0.02 Peanuts 0.01 Peroxidase c Green peas 1.00 Pea pods 0.72 String beans 0.62 Spinach 0.32 Lima beans 0.15 aAdapted from Ref. 51. bAdapted from Ref. 93. cAdapted from Ref. 54. ———————————————————————————————————————- The levels of a given enzyme activity can be highly variable in raw foods, since a cultivar of the same fruit, for example, can be bred to have more or less enzyme. The age (maturity) of an organism (Fig. 4), and the environmental conditions of growing (especially plants), including temperature, water supply, soil, and fertilization, all affect the level of enzyme activity. This makes it very difficult for the food processor to produce uniform-quality food products. Fortunately, the rate of denaturation of enzymes is generally first order; therefore, the time required to inactivate a fixed percentage of the enzyme is independent of its concentration. However, at different initial concentrations of active enzyme, the absolute concentrations of active enzyme left will be different. For example, at 5 half-lives, there will be 3.13% of the original active enzyme left regardless of initial active enzyme concentration. But at initial concentrations of 1 × 10-3M and 1 × 10-5M active enzyme, after 5 half-lives there will be 3.13 × 10- 5M and 3.13 × 10-7 Mactive enzyme left, respectively. So both the two initial and two final concentrations of active enzyme differ by 100. It takes longer to inactivate all the enzyme with higher initial concentration. The conditions just listed also affect the relative levels of isozymes (for example, those of peroxidase or lipoxygenase). These isozymes often have different temperature stabilities, as shown for the isoenzymes of peroxidase and lipoxygenase (Fig. 5). The presence of two or more isozymes is indicated by the inability to inactivate completely each enzyme at 60°C; at 70°C, lipoxygenase is completely inactivated but peroxidase still retains about 40% activity. Pag e 447 Fig ure 4 Chang es in the polyg alacturonic acid lyase activity of Bacilus pumilis as a function of ag e of culture: , absorbance at 600 nm of a 1:20 dilution as a measure of cell population; , activity of enzyme (units/ml). (From Ref. 28, p. 41, by courtesy of B.A. Dave.) 7.4 Rates of Enzyme-Catalyzed Reactions 7.4.1 Rate Determination ———————————————————————————————————————- The distinguishing feature of enzymes from other proteins is that enzymes bind their substrates stereospecifically into the active site and convert the substrates to products. Therefore, the presence of an enzyme is determined by whether it converts a compound to a product faster than that which occurs in its absence. Assays for enzyme activity are based on physical or chemical FIGURE 5 Rates of inactivation of peroxidase and lipoxyg enase in Eng lish g reen pea homog enates incubated at 60 and 70°C. (From Ref. 114, p. 132, with permission of the Institute of Food Technolog ists.) Pag e 448 changes in substrate(s) and products(s) in the reaction mixture. Continuous methods (change in absorbance, fluorescence, or pH) are very much preferred over methods that require taking aliquots at selected times, stopping the reaction by inactivating the enzyme, and adding one or more reagents to form a measurable derivative of either the substrate or product (preferable). Since an enzyme is highly specific for only one or a few substrates, it follows that it is relatively easy to measure that enzyme selectively even among hundreds of other enzymes of different specificities. The enzyme activity can be used to determine not only the presence of an enzyme (qualitative), but also how much enzyme is present (quantitative), based on its rate under controlled conditions. Some substrates and products do not differ greatly in physical and/or chemical properties; therefore it is difficult to determine the first-order rate constant, k1 (Eq. 7). In these cases, coupled enzyme assays should be considered, where the product, P1, of the first enzyme is a substrate of an added second enzyme. The product, P2, of the second enzyme is then chosen for measurement (Eq. 7). In order to measure k1 correctly, the relation k2[E2] @ 100 k1[E1] should be used. (7) 7.4.2 Steady-State Rates The velocity of a typical enzyme-catalyzed reaction is shown in Fig. 6, where concentration of product formed is plotted versus time. In the first few milliseconds, the velocity (dP/dt) of product formation accelerates depending on how fast the enzyme and substrate combine to give the enzyme. substrate complex (Eq. 2). This is the pre-steady-state period (0.1–2 msec) and can only be measured by very fast mixing and measuring system (i.e., stopped-flow spectrophotometers). After ~2 msec, the concentration of enzyme-substrate complex [E.S] reaches a steady FIGURE 6 Typical enzyme-catalyzed reaction showing the pre-steady-state (msec), the constant rate, and the declining rate parts of the prog ress curve. (From Ref. 111, p. 161.) Pag e 449 state where d[E.S]/dt =-d[E.S]/dt over the time required to measure a change in [P] or [S]. The steady-state concept applies from the end of the pre-steady-state period to the end of conversion of all substrate. Enzyme activity is measured under steadystate conditions in most laboratories (see later discussion). Why then does the velocity of the reaction becomes slower and approach zero as shown in Figure 6? The decrease in velocity with time can be a result of these: (a) the [S] becomes the ratelimiting factor (i.e., [S] < 100Km); (b) as [P] increases it may partially inhibit the enzyme; (c) the reaction is reversible and the reverse reaction becomes more noticeable as [P] increases; and/or (d) instability of the enzyme. These possible causes can be distinguished by appropriate methods [111]. Since steady-state velocities of enzyme-catalyzed reactions decreases with reaction time, often in a complex manner for the reasons already given, enzymologists generally determine the initial velocity, no, of enzyme-catalyzed reactions. This is done as shown in Figure 7. If done with proper control(s), all reactions should begin at zero time with zero [P]. The no is determine from a tangent drawn (dashed lines) to the initial part of the reaction. No more than 5% of the substrate should be converted to product during the time required to obtain the tangent. Note that accurate tangents are easier to obtain when the velocity is relatively slow. Experimentally, the velocity can be controlled by controlling the enzyme concentration, and this must be done if accurate, reproducible no values are expected. Please note that the slope of two point assays (zero at zero time and another point later on) does not yield accurate values of no! 7.4.2.1 Kinetics and Reaction Order Reaction order is determined from the dependence of velocity (dP/dt, or -dS/dt) on the concentration(s) of the reactant(s). Therefore, we begin with an equation. For an enzyme. FIGURE 7 Method of determining initial velocities of reactions. The solid lines are the experimentally determined data, and the dashed lines are tang ents drawn to the initial slope of the experimental data. Note the marked difference between the actual concentration of product formed at the time indicated by vertical dashed line for reactions (Ao) 4 and (Ao)5 compared with that predicted from initial rates. (From Ref. 111, p. 163.) Pag e 450 appropriate kinetic description is given by Equation 8 (same as Eq. 1), where E is enzyme, S is substrate, E.S is the enzymesubstrate complex, and P is product. The velocity of (8) formation of E.S (d[E.S]/dt) isk1 [E][S], while the velocity of disappearance of E.S (-d[E.S]/dt) is k-1(E.S) + k2 [E.S]. Steadystate conditions exist when ever d(E.S)/dt =-d(E.S)/dt for ~5 msec. The Michaelis-Menten equation (Eq. 9), derived from Equation 8, is based on the ———————————————————————————————————————- following assumptions [111]: 1. Initial velocity, no, is used so that [S]o [S]. As discussed earlier, this is common practice. 2. [S]o>>[E]o, so that there is little change is [S] o. Practically, this is the case with most enzymes where [S]o is of the order of 10- 4 to 10-2M since Km is generally within this range and [E]o is on the order of 10-8 -10-6M. 3. The step controlled by k2 (Eq. 8) is irreversible in reality or because no is used in ([P] is essentially zero). Therefore, k-2 is ~0. 4. d(E.S/dt=-d(E.S)/dt, so steady-state conditions prevail. 5. k2 controls the velocity of formation of product (dP/dt). If k2 > k1 then k1 controls velocity of product formation. Note that Vmax is not really a constant in Equation 9, since it is dependent on [E]o (Vmax = k2 [E]o); k2 is a constant, but Vmax will change when [E]o is changed. 6. If any one of these assumptions is not true then the form of the Michaelis-Menten equation will be more complex, even through a plot of no versus [S]o will be hyperbolic (Fig. 1). For example, if there is an additional intermediate in the reaction, such as an acylenzyme (Eq. 10); [111], then Equation 10 applies: and the Michaelis-Menten equation is ———————————————————————————————————————- In the preceding equations, [E]o is total enzyme concentration, [E] is free enzyme concentration, [E.S] is enzyme-substrate concentration, [S]o is initial substrate concentration, [S] is substrate concentration at any time t, and Km = (k2 + k-1)/k-1 for Equation 9 and (k-1 + k2)k3/k1 (k2 + k3) for Equation 11. What is the order of an enzyme-catalyzed reaction and how does one determine the order? Since [E]o is constant throughout the reaction, assuming it is stable, the order of an enzyme-catalyzed reaction is determined by the relationship between [S]o and Km (as long as [S]o >> [E]o; see assumption 2). When [S] o<0.01 Km, the order of the reaction is first order with respect to [S]o. This can be readily seen from Equation 9. When [S]o <0.01 Km, [S]o in the denominator can be ignored and no=Vmax[S]o/Km ,(no=k´[S]o), where k´ = Vmax/Km, showing direct dependence on [S]o. A plot of In([S]o/[S]) versus time gives a straight-line plot over several half-lives Pag e 451 of substrate concentration disappearance, with the slope equal to Vmax/Km, and the half-life time independent of [S]o. When [S]o >100Km, the order of the reaction with respect to [S]o is zero. This also can be seen from Equation 9. When [S]o >>Km, Equation 9 reduces to no = Vmax, indicating that no is independent of [S]o and all enzyme is saturated with substrate. A plot of [P] versus time gives a straight-line plot, with a slope Vmax (=k2[E]o).——————————————————————————————————————————————————————————————————————————————– When [S]o > 0.01 Km, but < 100 Km the order of the reaction with respect to [S]o is a mixture of first and zero order and the integrated form of the Michaelis-Menten equation is: k2[E]ot = Km In([S]o/[S]) + ([S]o - [S]) (12) The relative contributions of the terms In([S]o/[S]) and ([S]o - [S]) will depend on the relation of [S]o to Km. Section 7.5 on factors influencing enzyme reactions will provide experimental procedures for evaluating the effect of [S]o on no. Some typical rate constants of enzyme-catalyzed reactions are given in Table 6. The value of k1, the rate constant for formation of the enzyme-substrate complex, ranges from 109 to 104 M-1sec-1 (where 109 is the limiting rate of diffusion-controlled reactions). The value of k-1, the rate constant for dissociation of the enzyme-substrate complex, ranges from about 4.5 × 104 to about 1.4 sec-1. The value of ko, the observed rate constant for conversion of enzyme-substrate complex to product, ranges from 107 to 101 sec-1. In reactions that follow Equation 8 ko = k2, but not for those that follow Equation 10. 7.4.3 Pre-Steady-State Reactions [17, 44] Pre-steady-state kinetic experiments are rarely done in food science, yet they provide details regarding additional intermediate steps in enzyme reactions that are difficult to obtain by other TABLE 6 Rate Constants for Some Selected Enzymes Enzyme Substrate (M-1 (sec-1) (sec-1 (sec-1) Fumarase Fumarate >109 >4.5 ×104 ———————————————————————————————————————- Acetylcholinesterase Acetylcholine 109 — 103 Liver alcohol NAD+ 5.3 × 105 74 103 dehydrog enase NADH 1.1 × 107 3.1 Ethanol >1.2 × 104 >74 Catalase H2O2 5 × 106 — 107 Peroxidase H2O2 9 × 106 <1.4 106 Hexokinase Glucose 3.7 × 106 1.5 × 103 103 Urease Urea >5 × 106 — 104 Chymotrypsin 102 to 103 Trypsin 102 to 103 Ribonuclease 102 Papain 101 aRate constant for formation of enzyme-substrate complex. bRate constant for dissociation of enzyme-substrate complex. cOrder of mag nitude of the turnover number in moles of substrate converted to product per second per mole of enzyme: k o ( k cat) is the observed rate constant and may or may not involve a sing le ratelimiting step. Source: Ref. 33, p. 14, courtesy of Academic Press. ——————————————————————————————————————————————————————————————————————————————– methods. Why are they not used in food science? Primarily because of requirements of relatively large amounts of pure enzyme, specialized instrumentation, and lack of appreciation of the nature and simplicity of interpreting the results. The instrumentation for following pre-steady-state reactions requires mixing times of <1 msec for substrate and enzyme and recording times of milliseconds. This is because concentrations of enzyme and substrate must be in the range of 10-4 M and both must be present at about equal concentration in order to detect intermediate steps. Bray et al. [17] used low-temperature pre-steady-state kinetic methods to identify and place in order the intermediate steps in the reaction of milk xanthine oxidase with xanthine (Eq. 13), a very important reaction in milk and in gout. (13) Another way in which intermediate steps in enzyme-catalyzed reactions can be studied is by doing steady-state kinetics using several substrates that have different rate-determining steps. 7.4.4 Immobilized Enzyme Reactions [52, 88, 117] The preceding discussions apply to systems where enzyme and substrate are soluble and both are free to diffuse independently until the two collide with enough energy and proper orientation to form the enzyme-substrate complex. However, enzymes are frequently attached to cell walls and membranes in vivo or are bound to insoluble supports for commercial conversion of substrate to product. When the enzyme is immobilized, additional factors affect formation of the enzyme-substrate complex and catalysis. These include: (a) only the substrate is free of diffuse; (b) the enzyme support is surrounded by the Nerst (diffusion) layer, which acts as a boundary, decreasing the substrate concentration in proximity to the enzyme, compared to the bulk phase; (c) there are electrostatic factors due to charges on the substrate, enzyme, and support that may increase binding (opposite charges) or decrease binding (same charges); and (d) initial velocity, no, conditions do not apply as maximum conversion of substrate to product is desired. Hornby et al. [52] developed equations to take factors (a)–(c) into account. The velocity for a flow column reactor is given in Equation 14: -------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------- and is the apparent Michaelis constant, x is the thickness of the Nerst boundary layer, D is the diffusion coefficient for substrate, T is the temperature (Kelvin), z is the valence of substrate, F is Faraday's constant, R is the universal gas constant, and V is the potential gradient around the enzyme support. Pag e 453 Since n is not no and as much substrate conversion as possible is desired, zero-, mixed-, and first-order rate kinetics are observed in the reactor at different times of reaction. Therefore, (16) describes the kinetic process in the column reactor. The fraction p of S reacted at any time is (17) Also required is the void volume, Vo, of the column and the flow rate, Q, of the substrate through the column. The ratio of Vo/Q is equal to the residence time t in the reactor. Therefore, the reactor output can be expressed as (18) Equation 18 is the equation of a straight line, where y=[S]op and x=In(1-p). The slope of the plot is Lilly and Sharp [73] developed an analogous kinetic equation applicable to a continuous- feed stirred tank reactor: (19) In some cases, such as the hydrolysis of insoluble cellulose, biomass, lipid micelles, and cell walls, the substrate is insoluble and the soluble enzyme must diffuse to and bind to the insoluble substrate as shown in Equation 20 where E is the soluble enzyme and Es is the bound to the insoluble substrate: (20) The size of the insoluble substrate (micelle droplets or insoluble particles) also affects and Verger et al. [102] and others have shown that the kinetics of soluble enzyme action on insoluble substrate fits Michaelis-Menten kinetics where (21) The value of [S] is dependent on the surface of the insoluble substrate and (22) where kd and kp are from Equation 20. The values of and can be determined from Lineweaver-Burk plots. The and will be greatly dependent on experimental conditions. 7.5 Factors Influencing Enzyme Reactions In the previous sections, we developed the basic concepts and kinetic equations required to quantify the velocity (no and n) of enzyme-catalyzed equations. In developing Section 7.4, only the substrate concentration was considered as a variable. All other conditions were assumed to Pag e 454 be held constant. In Section 7.5, we will expand further on the effect of substrate concentration on the velocity of enzymecatalyzed reactions. In addition, the effect of enzyme concentrations, pH, temperature, water activity, and organic solvents on velocity of the reactions will be discussed. The effect of activators and inhibitors on velocity of enzyme-catalyzed reactions will be discussed in Sections 7.6 and 7.7, respectively. 7.5.1 Substrate Concentration The effect of substrate concentration on velocity of product formation is shown in Figure 1 for one-substrate reactions when the reactions follow Michaelis-Menten kinetics. Determination of Vmax and Km from data plotted as in Figure 1 is difficult at best because Vmax is achieved only when [S]o > 100Km (zero order with respect to substrate concentration). The substrate may be insoluble and/or expensive at the concentrations needed, the substrate may inhibit the reaction at high substrate concentrations (Fig.8), or it may activate the reaction at high substrate concentrations (Fig. 9). For these and other reasons, Lineweaver and Burk [76] in 1934 showed that the Michaelis-Menten equation (Eq. 9) can be transformed from a right hyperbola to a straight line by taking the reciprocal to gi——————————————————————————————————————————————————————————————————————————————–Effect of substrate inhibition on the initial velocity of an enzyme-catalyzed reaction. The dashed line shows the normal curve in absence of inhibition, and the solid line shows inhibition by a second substrate molecule for which the dissociation constant, is 10K m. The solid line is calculated according to the equation v o = Vmax/{[1 + Km/(Ao)] + [Ao/ ]}. E.A2 does not form product. Larg e g raph plotted by Line weaver-Burk method; the insert is a Michaelis-Menten plot. (From Ref. 111, p. 193). Pag e 455 ——————————————————————————————————————————————————————————————————————————————– Effect of substrate activation on initial velocity of an enzyme-catalyzed reaction. The dashed line shows a normal reaction in absence of activation, and the solid line shows a reaction in the presence of activation. The solid line is calculated on the assumptions that for the second substrate molecule is 10Km, that Vmax is doubled when all the enzyme is saturated with a second substrate molecule (i.e., E.S 2 g oes to product twice as fast as E.S), and that the second substrate molecule does not form product. The larg e g raph is plotted by the Line weaver-Burk method; the insert is a Michaelis-Menten plot. (From Ref. 111, p. 194.) where 1/no = y,1/[S]o = x, Km/Vmax is the slope and 1/Vmax is the y-intercept of the straight-line plot, and -1/Km is the x-intercept (Fig. 10). Therefore, Vmax and Km can be readily determined, using all the experimental data. For best results the [S]o used to obtain data for Figure 10 should range from 0.2Km to 5Km, if possible. The Lineweaver-Burk method for calculating Vmax and Km is, by far, the most used. Other linear transforms include the methods of Augustinsson (Eq. 22) [111] and of Eadie-Hofstee (Eq. 23) [111]. (22) (23) It is often useful to plot the experimental data by all three methods; easily done with available computer software programs for enzyme kinetics. Linear plots by all three methods are Pag e 456 FIGURE 10 Plot of substrate-velocity data according to the Lineweaver-Burk method as shown in Equation 21. (From Ref. 111, p. 176.) not always found, indicating complexities (multiple intermediate steps, etc.) not readily apparent from only one of the plots. 7.5.1.1 Multiple Substrate Reactions Most enzyme-catalyzed reactions involve more than one substrate. Hydrolase-catalyzed reactions require water as the second substrate, but under most conditions where the water concentration is 55.6 M, its effect is ignored in developing the kinetic equations. However, in all other cases, the effect of concentration of the second substrate cannot be ignored. Several investigators have developed kinetic equations for handling these multi-substrate reactions [22, 27, 115]. Whitaker [111] has summarized the experimental procedures and kinetic approaches to be used. The experimental procedures are no more complex than those described for the one substrate reactions, and most of the reactions fit Michaelis-Menten type kinetics. Experiments must be done so that the effects of all substrates can be determined, varying the concentration of one of them at a time. The interpretation of Vmax and Km is more complex but the individual rate constants can be determined readily by the King-Altman method [62], among other methods. The multiple substrate reactions that fit Michaelis-Menten kinetics belong to three main types [22]. Using examples for twosubstrate/two-product reactions, two of these types are (a) ordered sequential bi bi mechanism, in which the two substrates must bind with the active site of the enzyme in an ordered manner and the two products are released from the enzyme in an ordered manner (Eq. 24), and (b) random sequential bi bi mechanism, in which the two substrates can bind into the active site of an enzyme in either order and the two products come off the enzyme in either order (Eq. 25). In the sequential mechanisms, all substrates must bind into the active site of the enzyme before any products are formed. In the third type, (c) the ping-pong bi bi mechanism, the first substrate must bind in the active site and one product is released before the second substrate can bind into the active site and be catalyzed to product (Eq. 26). These different types of mechanisms are best determined by plotting 1/no versus 1/[S]o according to the Pag e 457 ——————————————————————————————————————————————————————————————————————————————– Lineweaver-Burk method. The two sequential mechanisms [(a) and (b)] are distinguished by use of equilibrium dialysis to determine the order of substrate binding into the active site. Ordered sequential bi bi mechanism (24) Random sequential bi bi mechanism (25) Ping-pong bi bi mechanism (26) The reader is referred to Whitaker [111] or Cleland [22] for further details on experimental procedures and analyses of data. 7.5.1.2 Substrate Inhibition One substrate molecule binds into the active site of the enzyme and is catalytically converted to product according to normal kinetics. At higher substrate concentrations, a second molecule binds at a site different than the active site. It is not catalyzed to product, but its binding results in a decrease in catalytic efficiency of the enzyme (Fig. 8). Note that when plotted by the Michaelis-Menten method (insert), no reaches a maximum but is lower than the true Vmax, and no decreases as [S]o is increased beyond the maximum no The Lineweaver-Burk plot shows an upward deviation from linearity at high substrate concentrations. Isozymes sometimes show differences in substrate inhibition that appear to be a regulatory control of activity. 7.5.1.3 Substrate Activation Binding of one substrate molecule into the enzyme active site results in product formation kinetics in the normal manner. The binding of a second substrate molecule at another site does not result in conversion of the substrate molecule to product, but it does enhance the rate of conversion of the substrate molecule in the active site to product (Fig. 9). Note that substrate activation results in downward deviation of the line from linearity at higher substrate concentrations when plotted by the Lineweaver-Burk method. ——————————————————————————————————————————————————————————————————————————————– Substrate activation can be confused when two enzymes act on the same substrate, if the Km values for the two enzymes are somewhat different. This possibility can be verified by showing that two isozymes are present by electrophoretic separation (for example) of the two. 7.5.1.4 Allosteric Behavior Allosteric behavior is defined by a plot of no versus [S]o giving a sigmoidal shaped curve (Fig. 11; solid line), in contrast to a right hyperbola (Fig. 11; dashed line) found for Michaelis-Menten Pag e 458 FIGURE 11 Comparison of effect of substrate concentration on the intial velocities of two enzyme-catalyzed reactions, one that obeys Michaelis-Menten kinetics (dashed line) and one that shows allosteric behavior (solid line). The desig nations and indicate the substrate concentrations at which the initial velocity is 0.1 and 0.9 Vmax, respectively, for Michaelis-Menten kinetics, while and have the same meaning for the allosteric behaving system. The Hill value was 4 in the calculation of the allosteric curve (solid line). (From Ref. 111, p. 195.) behavior. There is a smaller than expected effect of increasing [S]o on no at [S]o < Km, followed by larger than expected effect of [S]o on no at [S] around Km. The example shown in Figure 11 is for positive allosteric kinetic behavior. In some cases, the effect of [S]o on no is greater than expected at lower [S]o, and then no does not increase as much as [S]o increases. This is negative allosteric kinetic behavior. Positive allosteric kinetic behavior was discovered by Monod et al. [80], who subsequently received the Nobel Prize. Koshland et al. [66] discovered negative allosteric kinetic behavior, and developed methods for distinguishing allosteric behavior from Michaelis-Menten behavior, including a mathematical verification for occurrence of allosteric behavior. To distinguish allosteric behavior from Michaelis-Menten behavior, Koshland et al. [66] proposed an equation for measuring cooperativity: Rs = [S]0.9Vmax/[S]0.1Vmax = cooperativity factor (27) where [S]0.9 is the substrate concentration required to give no = 0.9Vmax and [S]0.1 is the substrate concentration required to give vo = 0.1Vmax. These points are marked on Figure 11 for and for Michaelis-Menten behaving enzymes and and for positive allosteric behavior. For all Michaelis-Menten behaving enzymes, Rs is 81, while for positive allosteric behaving enzymes Rs < 81 (often as low as 30–40), and for negative allosteric behaving enzymes Rs > 81. Allosteric behavior is most often observed with multi-subunit enzymes. Koshland et al. [66] proposed that for allosteric behavior two or more subunits of an enzyme must show cooperativity: ——————————————————————————————————————————————————————————————————————————————– Consider K1 and K2 as binding constants. If K2 > K1 then the second substrate molecule binds more easily (tightly) than the first substrate molecule and positive allosteric behavior is seen. If K1 > K2 the first substrate molecule binds more tightly than the second one, and negative allosteric behavior is seen. If K1 = K2, the two substrate molecules bind equally well with no effect on each other’s binding, and Michael-Menten behavior is observed. Allosteric behavior is one method of regulating enzymatic activity by small changes in substrate concentration, especially around Km. The metabolic enzymes in the glycolytic and TCA cycles often show allosteric behavior. Allosteric behavior can also be elicited in some enzymes by non-substrate compounds that act as allosteric inhibitors or activators. 7.5.2 Enzyme Concentration——————————————————————————————————————————————————————————————————————————————– The relationship between no and [E]o is usually linear when all other factors, such as [S]o, pH, and temperature, are kept constant (Fig. 12). Doubling the [E]o doubles no; tripling the [E]o triples no. This is expected since no = k2[E.S] = dP/dt = -dS/dt (see Eq. 2). This linear relationship between [E]o and no at all [S]o is very valuable, since we can determine how much enzyme is present by measuring its activity under standard conditions without purifying the enzyme. Of course, the enzyme should be extracted from the tissues and should be free of any insoluble materials and/or other compounds that might interfere with determination of activity (such as those having intense absorbance at the same wavelength, in vivo activators and inhibitors that occur in variable amounts, or other enzymes that act on the same substrate). Enzyme activity determinations are important in clinical medicine, nutritional abnormalities, quality control in food processing, and in analytical uses of enzymes to determine concentration of compounds that are substrates, activators, or inhibitors (see Sec. 7.12). There are at least five exceptions to the expected linear relationship between [E]o and no. [111]. Two of these are (a) limitations on solubility of a substrate, such as O2, and (b) conversion of substrates to products that either are less good substrates or are competitive inhibitors. Both lead to a decrease in no as [P] increases. The third (c) is coupled enzyme FIGURE 12 Expected relationship between enzyme concentration and observed velocity of reaction. Substrate concentration, pH, temperature, and buffer are kept constant. (From Ref. 111, p. 202.) Pag e 460 reactions where the product of enzyme-1 is the substrate of enzyme-2. In this instance, product-2 is measured to determine no, yet the concentration of enzyme-1 is desired: (29) To obtain linearity between no (dP2/dt) and [E1]o, k2[E2]o > 100k1[E1]o should be used. There may also be (d) irreversible inhibitor, such as Hg2+, Ag+ , or Pb2+ in the substrate, buffer, or water, which inactivates a fixed amount of the enzyme; and (e) a dissociable essential cofactor in the enzyme solution. 7.5.3 Effect of pH The pH has a marked effect on activity of most enzymes. As shown in Figure 13, pepsin, peroxides, trypsin, and alkaline phosphatase have optimal activity at about pH 2, 6, 8, and 10, respectively. All the activity-pH curves are bell-shaped, with activity decreasing to near zero at 2 pH units below or above the pH optimum. Note that the left-hand side of the pepsin curve and the right-hand side of the alkaline phosphatase curve (Fig. 13) decrease abruptly as a function of pH (the bell-shaped curves are skewed). The pH optima of several enzymes found in raw food products are listed in Table 7. The pH optima range from 2 for pepsin to 10 for alkaline phosphatase. Catalase (Table 7) has maximum activity from pH 3 to 10. The apparent complexity of the effect of pH on enzyme-catalyzed reactions is shown in Figure 14, with milk alkaline phosphatase as an example. Figures 14a and 14d illustrate the effect of different concentrations of the substrate phenyl phosphate on the shape and height, respectively, of the pH versus no curves. The pH optimum shifts from pH 8.4 at 2.5 × 10-5 M phenyl phosphate to pH 10 at 2.5 × 10-2 and 7.5 × 10-2 M phenyl phosphate. The pH optimum also is different for different substrates (Fig. 14c). The pH optima are 9.3, 9.5, and 9.8 for 0.02 M FIGURE 13 pH effect on activity of several enzymes. (a) Pepsin acting on N-acetyl-L phenylalanyl-L-diodotyrosine at 36.7°C and 5 × 10-4 M pepsin; reaction time 15 min. (b), Ficus g labrata peroxidase acting on 0.03 M g uaiacol and 0.005 M H2O2 at 30.0°C. (c) Trypsin acting on 0.1% casein. (d) Hydrolysis of 5 × 10-4 M p-nitrophenyl phosphate by crude milk alkaline phosphatase at 35°C in 0.1 M sodium g lycinate buffer; reaction time 15 min. Times of reaction are g iven for one-point assays, not when no was determined. (From Ref. 111, p. 273.) Pag e 461 TABLE 7 pH-Activity Optimum of Several Enzymes a Enzyme pH optimum Acid phosphatase (prostate g land) 5 Alkaline phosphatase (milk) 10 a-Amylase (human salivary) 7 b-Amylase (sweet potato) 5 Carboxypeptidase A (bovine) 7.5 Catalase (bovine liver) 3–10 Cathepsins (liver) 3.5–5 Cellulase (snail) 5 a-Chymotrypsin (bovine) 8 Dextransucrase (Leuconostoc mesenteroides) 6.5 Ficin (fig ) 6.5 Glucose oxidase (Penicillium notatum) 5.6 Lactate dehydrog enase (bovine heart) 7 (forward reaction) 9 (backward reaction) Lipase (pancreatic) 7 Lipoxyg enase-1 (soybean) 9 Lipoxyg enase-2 (soybean) 7 Pectin esterase (hig her plants) 7 Pepsin (bovine) 2 Peroxidase (fig ) 6 Polyg alacturonase (tomato) 4 Polyphenol oxidase (peach) 6 Rennin (calf) 3.5 Ribonuclease (pancreatic) 7.7 Trypsin (bovine) 8 aThe pH optimum will vary with source and experimental conditions. These pH values should be taken as approximate values. Source: Ref. 111, p. 274. b-glycerophosphate, 0.015 M phosphocreatine, and 0.025 M phenyl phosphate, respectively. The pH optimum of alkaline phosphatase is dependent on cofactor type, with pH optima of 8.0 for Mn2+ and 9.4 for Mg2+. The in vivo cofactor is Zn2+ ——————————————————————————————————————————————————————————————————————————————– One can conclude from the data of Figures 13 and 14 that the pH optimum of an enzyme is dependent on the nature of the enzyme and the conditions used to measure the activity as a function of pH. But is it possible to get valuable information from pH versus no curves? The answer is yes, provided the experiments are properly done. The four factors that primarily affect the nature of the pH versus no curves are (a) whether no data are used, (b) pH stability of the enzyme, (c) equilibria (important ionizations and dissociations) of the system, and (d) the relationship of [S]o to Km. These factors deserve discussion. 7.5.3.1 How Velocity Data Are Obtained The importance of obtaining initial velocities, no, in the kinetic investigations of enzymes cannot be overstressed. The data of Figure 15 illustrate that the difference in determined pH optimum is about 0.5 pH unit when different times are used to measure n, for an enzyme that is unstable either below or above the pH optimum. Since the enzyme is more stable on the acid side of the ———————————————————————————————————————- ———————————————————————————————————————- Effect of experimental conditions on the pH optimum of calf intestinal mucosa alkaline phosphatase. (a) Effect of initial substrate concentration on pH optimum. The curves are for the following substrate (phenyl phosphate) concentrations (M): A, 2.5 × 10 -5; B, 5 ×10-5; C, 1 × 10-4; D, 5 × 10-4;; E, 7.5 × 10-4; F, 2.5 × 10-3; G, 2.5 × 10-2; H, 7.5 × 10-2. (From Ref. 82, p. 675, by courtesy of the Biochemical Society.) (b) Effect of nature of activating cation on pH optimum. The cations were: , 5 × 10-3 M Mg Cl2 and , 1 × 10-3 M MnCl2 with 2.5 × 10-4 M phenyl phosphate as substrate. (From Ref. 82, p. 677, by courtesy of the Biochemical Society.) (c) Effect of nature of substrate on pH optimum. The substrates were: , 0.015 M phosphocreatine: , 0.02 M b-g lycerophosphate; +, 0.025 M phenyl phosphate. (From Ref. 81, p. 235, by courtesy of the Biochemical Society.) (d) pH optima from data of part (a) plotted ag ainst -log [A] o. The lowest substrate concentration is at the rig ht of the fig ure. (From Ref. 82, p. 675, by courtesy of the Biochemical Society.) The temperature was 38.0°C in all cases. ——————————————————————————————————————————————————————————————————————————————– optimum than on the alkaline side, the observed pH optimum shifts to the left as a function of time between no and nt. 7.5.3.2 Stability of the Enzyme Enzymes generally are not stable at all pH levels. It is important to determine the pH range over which an enzyme is stable before determining the pH optimum. This can be done by at least three methods. The best way is to incubate the enzyme under the same conditions (pH, temperature, buffer, enzyme concentration, time) to be used to determine no at different pH, but in the absence of substrate. At various times, aliquots are removed, added to tubes containing substrate buffered at a pH at or near the pH optimum, and no is determined. The control (100% activity) is a sample

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