Food Effect of Equilibria

Several of the constituents of an enzyme reaction system ionize or dissociate, depending on pH. Constituents that may ionize include the buffer, the substrate, the cofactor (if required), and the essential ionizable groups in the active site of the enzyme. Association/dissociation occurs in the binding of substrate and cofactor into the active site. Nothing can be done with respect to ionization of the buffer. Certainly, the ionic strength of the solution should be kept constant at all levels by use of NaCl or KCl. When different buffers must be used, it is essential to overlap the two buffers, for one or more pH levels. Alternatively, the buffers can be combined at all pH levels. For fundamental studies, the substrate should be neutral, so that ionization does not affect combination of substrate with enzyme. Practically, this may be impossible. The control of ionic strength is again very important. If a cofactor is required, the enzyme should be saturated with the cofactor at all pH levels ([CoF] >> KCoF). For further details on this subject see Whitaker [111]. 7.5.3.4 The Relationship of [S]o to Km This is a major problem in most experiments reported in the literature. Consider the minimum steps involved in enzyme-catalyzed reactions (Eq. 7) and the possible effect of pH on these steps. First, there is binding of substrate and enzyme to form the enzyme-substrate complex (E.S) as Pag e 464 controlled by k1. Second, there is the dissociation of the E.S complex to E + S (controlled by k-1) and there is the catalytic conversion of E.S complex to E + P, controlled by k2 (or more than one rate constant if there are additional intermediate steps). The ionization of each of the reactants (E, S, E.S, and P) will affect the pH versus no curve. The substrate should be selected (if possible) so as not to undergo ionization in the pH range used. By proper design of the experiments, the effect of pH on stability of enzyme, the binding of substrate into the active site of the enzyme, and the pKa values of the ionizable groups in the active site of the free enzyme, the enzyme-substrate complex, and of other intermediate species (such as acylenzyme) can be determined [111]. Improperly designed experiments give data, such as in Figure 14 for alkaline phosphatase, but the results are more confusing than enlightening as to what is going on between the enzyme and the substrate. 7.5.4 Effect of Temperature Experimental data on the effect of temperature on velocities of enzyme-catalyzed reactions can be just as confusing and uninterpretable as the effect of pH, unless the experiments are designed properly. Temperature affects not only the velocity of catalysis of , but also stability of the enzyme; the equilibria of all association/dissociation reactions (ionization of buffer, substrate, product and cofactors (if any); association/disassociation of enzyme- substrate complex; reversible enzyme reactions ; solubility of substrates, especially gases; and ionization of prototropic groups in the active site of the enzyme and enzyme-substrate complex. To

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 some extent, proper design of the experiments for temperature effects is easier than for pH effects. There are usually three reasons temperature effects on enzymes are studied: (a) to determine stability of the enzyme; (b) to determine the activation energy, Ea, of the enzyme-catalyzed reaction; and (c) to determine the chemical nature of the essential prototropic groups in the active site of the enzyme. The design of the experiments is not different, except the buffer must always be made up to have the same pH at all temperatures used. This requires the buffer to be made up at the temperature to be used. Otherwise, the pH of the buffer is an uncontrolled variable. 7.5.4.1 Stability of Enzyme This can be determined in the following manner. Tubes with a fixed concentration of enzyme (similar to those to be used in the kinetic experiments), in the buffer adjusted to pH desired when made up at each temperature to be used, are incubated at the selected temperatures (usually starting at 25°C). Aliquots are removed at various times, substrate in a buffer near the pH optimum is added, and the activity left is determined at a constant temperature and pH. The control enzyme activity (100%) is determined on enzyme maintained at 0°C. The data are plotted as shown in Figure 16, since the rate of loss of activity is usually first order for a pure enzyme (no isozymes present). The slope of the line is k, the rate constant for denaturation of the enzyme. As shown (Fig. 16), the enzyme is stable at 20–35°C, but the rates of loss of activity increase as temperature increases. Within the temperature range where there is loss of activity, a plot of In k versus 1/T(K) should give a linear relationship with a slope Ea/R, where Ea is the activation energy for denaturation and R is the universal gas constant in cal mol-1 deg-1 (or J mol-1 deg-1). Typical transition state denaturation constants for some enzymes are shown in Table 8 (where dH‡ = Ea – RT). Pag e 465 FIGURE 16 Rate of denaturation of an enzyme at various temperatures. The data are calculated for E a of 60,000 cal/mol, where the first-order rate constant, k, of denaturation is 0.005, 0.020, 0.090, 0.395, and 1.80 min-1 at 40, 45, 50, 55, and 60°C, respectively. Calories × 4.186 = Joules. (From Ref. 111, p. 305.) 7.5.4.2 Activation Energy of the Enzyme-Catalyzed Reaction To determine activation energies, two plots are required, the first is a plot of experimentally determined product concentration versus time at various temperatures (Fig. 17), and the second a plot of log k, the zero-order reaction rate constant versus 1/T (in K; Fig. 18). For the first plot, TABLE 8 Transition-State Denaturation Constants for Various Enzymes Substance DH‡ (cal/mol) DS‡ (eu)a Number of bonds broken b dG‡ (25°C) (cal/mol) Lipase, pancreatic 45,400 68.2 9 25,100 Amylase, malt 41,600 52.3 8 26,000 Pepsin 55,600 113.3 11 21,800 Peroxidase, milk 185,300 466.0 37 46,400 Rennin 89,300 208.0 18 27,300 Trypsin 40,200 44.7 8 26,900 Invertase, yeast pH 5.7 52,400 84.7 10 27,200 pH 5.2 86,400 185.0 17 31,300 pH 4.0 110,400 262.5 22 32,200 pH 3.0 74,400 152.4 15 29,000 aDS‡ in cal/mol deg ree. bNumber of noncovalent bonds broken on denaturation = DH‡/5000, where the averag e DH‡ per bond is assumed to be 5000 cal/mol. Calories × 4. 186 = Joules. Source: Modified from Ref. 96. Pag e 466 FIGURE 17 Effect of temperature on rate of product formation. The solid lines are for experimental data; the dashed lines are based on initial rates (i.e., lines drawn tang ent to experimental data at time close to zero). The data shown were calculated for the following conditions: [S]o >> Km; Ea for transformation of reactant to product, 12,000 cal/mol; Ea for denaturation of enzyme, 60,000 cal/mol; first-order rate constant, k, for denaturation of enzyme, 9.0 × 10-4 min-1 at 40°C. Calories × 4.186 = Joules. (From Ref. 111, p. 303.) temperatures must be low enough to obtain good no values before rate of denaturation of the enzyme becomes a problem, [S]o must be >> Km so the enzyme is saturated with substrate at all temperatures, and the pH (same value at all temperatures) must be at the pH optimum. If all enzyme is not in the form E.S, then the temperature dependence will also include Ks (association/dissociation of . If the pH is not at the pH optimum, then the effect of temperature on ionization of groups in the active site of the E.S complex will also be measured. Each slope of a dashed line in Figure 17 is a k value. Using the k values and temperatures from Figure 17, the second plot can be prepared (Fig. 18). Ea, the activation energy, is obtained from the slope of this plot (slope = -Ea/2.3R). FIGURE 18 Effect of temperatures on rate constant of a reaction. Plotted as log k versus 1/T(K) to permit determination of Ea as shown. (From Ref. 111, p. 315.) Pag e 467 Typical Ea values for enzyme-catalyzed reactions range from about 25,000 to 50,000 J/mol. A comparison of Ea values for noncatalyzed, non-enzyme-catalyzed, and enzyme-catalyzed reactions are shown in Table 2. The value of DH‡ at a fixed temperature can be determined from the relationship Ea = DH‡ + RT; DG‡ is calculated from DG‡ = -RT In(kh/kBT) (30) where kB is the Boltzman constant (1.380 × 10-16 erg deg-1) and h is the Planck constant (6.624 × 10-27 erg sec). The DS‡, the entropy of activation, is calculated for a fixed temperature according to DG‡ = DH‡ – T DS‡ (31) 7.5.4.3 Chemical Nature of
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 Prototropic Group(s) in Active Site of Enzyme The effect of temperature on ionization of prototropic groups in the active site of the free enzyme or E.S complex can be determined from the change in pKa values for the groups. This is important in determining the chemical nature of ionizable groups. Shown in Figure 19 are the effects of temperature and pH on relative Vmax/Km for hydrolysis of a-N-benzoyl-Largininamide by papain [97]. There is no effect of temperature on ionization of the prototropic group of pKa1 = 4 in the active site of the free enzyme, consistent with the group being a carboxyl group, active in the carboxylate form. The second prototropic group has pKa2 values of 9.0, 8.2, and 7.4 at 5, 38, and 66°C, respectively. The DHion calculated from a plot of -log Kion versus 1/T(K) is 33,000 J/mol (Fig. 20 gives an example of this type of plot). Both pKa2 values and DHion (Table 9) indicate that the group is probably an -SH group of Cys25, known to be involved in activity of papain. FIGURE 19 Effect of temperature (and pH) on relative k1 for papain-catalyzed hydrolysis of a-N-benzoyl-L-arg ininamide. The k1 values were calculated from where the maximum value at each temperature (5, 38, and 66°C) was set to 100. (From Ref. 97, p. 20, by courtesy of the American Society of Biolog ical Chemists.) Pag e 468 FIGURE 20 Effect of temperature on ionization constant, Kion, of Tris. pK (-log Kion) values are plotted as a function of 1/T(K) to permit calculation of DH ion. (From Ref. 111, p. 310.) 7.5.4.4 Enzyme Activity at Low Temperatures (See also Chap. 2.) Intuitively, we might suppose that enzyme activity ceases at temperatures below 0°C, especially after the solution appears to be frozen. If so, this would be an important way of preserving our food indefinitely. Also, perhaps enzymes are denatured by freezing. Figure 21 shows the effect of temperature on invertase-catalyzed hydrolysis of sucrose from 49.6 to – 19.4°C, and b-galactosidase-catalyzed hydrolysis of o- and p-nitrophenyl-b-galactosides from 25 to -60.2°C. Note that these two enzymes have activity over the temperature ranges studied, Pag e 469 FIGURE 21 Effect of temperature (49.6 to -60.2°C) on velocities of enzyme-catalyzed reactions. (A) Invertase-catalyzed hydrolysis of sucrose in Aqueous buffer system. (Adapted from Refs. 61 and 94.) The intersection of the two dashed lines indicate the point of chang e in rates at the freezing point of the solution. (B) b-Galactosidasecatalyzed hydrolysis of o-nitrophenyl- b-g alactoside, pH 6.1. (C) , b-Galactosidase-catalyzed hydrolysis of p-nitrophenyl-b-g alactoside, pH 7.6. Solvent in (B) and (C) was 50% dimethyl sulfoxide-water. (Adapted from Ref. 36.) even though the activities decreased by 105 for invertase and 104.3 for b-galactosidase. There is a change in slope of plot A at – 3°C, where the solution began to freeze. One interpretation for the change in slope is the phase change. Others [29,30] have suggested that the change in slope is due to a change in the rate-determining step, formation or stabilization of intracellular hydrogen bonds in the enzyme, association of the enzyme into polymers, or increased hydrogen bonding between the substrate and water. The b-galactosidase-catalyzed hydrolysis of o- and p-nitrophenyl-b-galactosides was performed in 50% dimethyl sulfoxide to prevent freezing. There were no changes in the slopes, and kcat and Km changed in a linear fashion. Therefore b-galactosidase acted in a predictable fashion over the range of 25 to –60.2°C, provided ice was absent. Storage of foods at or just below the freezing point of water should be avoided. As water freezes the enzyme and substrate become more concentrated (solute is rejected from the ice phase), which may lead to enhanced activity. In addition, freezing and thawing disrupt the tissues, permitting greater access of enzyme to substrate. As shown in Figure 22, phospholipase activity in cod muscle is about five times greater at -4°C, below the freezing point, than at -2.5°C. 7.5.5 Water Activity/Concentration Enzyme activities usually occur in aqueous media in vitro although in vivo enzyme reactions can occur not only in the cytoplasm but in cell membranes, in lipid depots, and in the electron Pag e 470 FIGURE 22 Rate constants (k) of phospholipasecatalyzed hydrolysis of phospholipids in cod muscle at subfreezing temperatures. (From Ref. 77, with permission of Institute of Food Technolog ists.) transport system, where transfer of electrons is known to occur in a lipid matrix. There are three major ways of studying the effect of water activity on enzyme activity. The first method is to carefully dry an unheated biological sample (or model system) containing active enzymes, then to equilibrate it to various water activities and measure the velocity of enzyme activity. An example of this approach is shown in Figure 23. Below 0.35aw (< 1% total water) there is no phospholipase activity on lecithin. Above 0.35aw there is a nonlinear increase in activity. Maximum activity was still not reached at 0.9aw (about 12% total water content). b-Amylase had no activity on starch until about 0.8aw (~2% total water); activity then increased 15 times FIGURE 23 Enzyme activity as a function of a w (A) , Phospholipase-catalyzed hydrolysis of lecithin (adapted from data of Ref. 1). (B) , Percent total water content of soluble solids (50:50 (w/w) protein and carbohydrate). (Adapted from data of Ref. 32.) (C) , b-Amylase-catalyzed hydrolysis (as maltose equivalents) of starch. (Adapted from data of Ref. 31.) Pag e 471 FIGURE 24 Effect of g lycerol concentration in water on peroxidase and lipoxyg enase reaction velocities. (Adapted from Ref. 11.) by 0.95 aw (~ 12% total water). From these examples, it can be concluded that the total water content must be < 1–2% to prevent enzyme activity. The second method of determining the concentration of water needed for enzymatic activity is to replace some of the water with organic solvents. Replacement of water with water-miscible glycerol reduces the activities of peroxidase and lipoxygenase when water content is reduced below 75% (Fig. 24). At 20% and 10% water, lipoxygenase and peroxidase have zero activity. Viscosity and specific effects of glycerol may have a bearing on these results. In the third method, most of the water can be replaced by immiscible organic solvents in lipase-catalyzed transesterification of tributyrin with various alcohols [118]. The “dry” lipase particles (0.48% water), suspended in dry n-butanol at 0.3, 0.6, 0.9, and 1.1% water (w/w) over- all, gave initial velocities of 0.8, 3.5, 5, and 4 mmol transesterification/h.100 mg lipase. Therefore, porcine pancreatic lipase had a maximum vo for transesterification at 0.9% water concentration. Organic solvents can have two major effects on enzyme-catalyzed reactions: an effect on stability, and an effect on direction of reversible reactions. These effects are different in water-immiscible and water-miscible solvents. In immiscible organic solvents there is a shift in specificity form hydrolysis to synthesis. Rates of lipase-catalyzed lipid transesterification reactions are increased more than sixfold while there is up to 16 times decrease in rate of hydrolysis when “dry” (~1% water) enzyme particles are suspended in the immiscible solvent [4,91, 118]. Alkylation of lipases and trypsin to make them more hydrophobic has similar effects in shifting from hydrolysis to synthesis as does the use of immiscible solvents [4, 91]. The rate of trypsincatalyzed esterification of sucrose by oleic acid was increased six times by alkylation of some of the amino groups of trypsin [4]. There is also a change in the stereospecificity of the products formed in organic solvents [40]. The ratio of v R /v S chiral isomers was 75 and 6 when lipasecatalyzed transesterification of vinyl butyrate with sec-phenethyl alcohol was performed in nitromethane versus decane, respectively [40]. Enzymes can be more stable in organic solvents than in aqueous buffers. Ribonuclease and lysozyme become much more stable as the water content is reduced, whether this is done by drying (ribonclease) or by adding water-immiscible organic solvents (Table 10). At 6% water content, ribonuclease has a thermal transition temperature (Tm) of 124°C and a half-life of 2.0 h at 145°C. The stability decreases as the water content increases; a dilute solution of ribonuclease has a Tm of 61°C and a half-life too short to measure [103]. Lysozyme is nearly as stable in Pag e 472 TABLE 10 Stability of Lysozyme and Ribonuclease as a Function of W ater Content Enzyme W ater content (%) Thermal transition temperature, Tm Half-life of activity (h) Ribonuclease a 6 124 2.0 11 111 0.83 13 106 0.5 16 99 0.17 20 92 0.07 Dilute solution 61 Too fast to measure Lysozyme b Dry powder 200 Cyclohexane 140 Hexadecane 120 1-heptanol 100 Buffer,c pH 4 1.42 Buffer, pH 6 0.17 Buffer, pH 8 0.01 aAdapted from Ref. 103. Half-life determined at 145°C. bAdapted from Ref. 64. The dry powder contained 0.5% moisture. The dry powder was dispersed in anhydrous org anic solvent. cThe lysozyme powder was dissolved in aqueous buffer to g ive 1% solution. water-immiscible organic solvents as in the dry powder, but the half-life of dilute solutions is very short [64]. Subtilisin crystals placed in acetonitrile have the same crystal structure as does subtilisin crystallized from aqueous solutions [41]. Enzymes do not turn inside-out when placed in immiscible organic solvents. The stability and catalytic activity of enzymes in water/water-miscible organic solvent systems are different from those in waterimmiscible organic solvent systems. Kang et al. [55–57] showed that protease-catalyzed hydrolysis of casein, in either 5% ethanol/95% aqueous buffer or 5% acetonitrile/95% aqueous buffer systems, caused an increase in Km, a decrease in Vmax, and a decrease in stability (determined by circular dichroism and differential scanning calorimetry methods) compared to controls in aqueous buffer only. It is well known that protic solvents, such as alcohols and amines, compete with water in hydrolytic enzyme reactions [5, 9]. 7.5.6 Why Enzymes are Effective Catalysts Enzymes are very effective catalysts as indicated by their ability to lower the activation energy, Ea, of reactions (Table 2). This results in larger values of no. But these facts provide no indication of how the enzymes achieve such “magic.” Be assured that enzymes act by the same principles of all chemical reactions. They just do it better [67]. Listed in Table 11 are the factors that account for the catalytic efficiency of enzymes. Not all enzymes use all possibilities listed, but all enzymes do bind substrates stereospecifically into the active site where essential groups (side chain of specific amino acids or required cofactors) perform from the reaction chemistry either by general acid-general base or nucleophilic-electrophilic processes. It is apparent that the most impressive enhancement of the rate is due to formation of the enzyme-substrate complex, where the rate enhancement is on the order of 104 for a two-substrate reaction, 109 for a three-substrate reaction, and 1015 (or more) for a four-substrate reaction. The seven factors listed in Table 11 can account for 1018 to 1036 rate enhancements, if all Pag e 473 TABLE 11 Factors Accounting for Catalytic Effectiveness of Enzymes Factor Rate enhancementa 1. Formation of stereospecific enzyme-substrate complex (conversion form inter- to intramolecular reaction) 104 ; 109 ; 1015b 2. Decreased entropy of reaction 103 3. Concentration of reactive catalytic g roups 103 –104 4. Distortion of substrate 102 –104 5. General acid/g eneral base catalysis 102 –103 6. Nucleophilic/electrophilic catalysis 102 –103 7. Using several steps 102 –104 Overall rate enhancement 1018–1036 a In some cases, these values can be approximated by model experiments. In others, they are the best estimates available. Source: Adapted from Ref. 111, pp. 129–142. factors are operative. An important factor not included is the role of the increased hydrophobicity of the active site when the substrates are bound. The role of hydrophobicity in the active site during catalysis is one of the least known and appreciated factors in enzyme catalysis. A good example is synthesis of glutathione from g-glutamylcysteine and glycine, using ATP as the energy source (a three-substrate reaction). This reaction is catalyzed by glutathione synthetase. After the three substrates bind stereospecifically into the active site, the active site is closed by a “lid” consisting of a 17-amino-acid loop in the enzyme. This protects the transition state intermediates from competition with water [60]. The rate constant ko is 151 sec-1 for the synthesis. Using recombinant DNA technology, Kato et al. [60] replaced the 17-amino-acid loop
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 with a sequence of three glycine residues, so that the active site could not close. The ko for the mutant enzyme is 0.163 sec-1, which is 1 × 10-3 that of the wildtype enzyme. Loop replacement did not appear to cause any other change in the physical structure of the enzyme. 7.6 Enzyme Cofactors Some enzymes, especially the hydrolases, are composed only of amino acids. The catalytic activity of the active site is the result of specific binding and catalysis due to specific side chains of amino acid residues (Table 9). These prototropic groups alone, such as the imidazole group of His57, carboxyl group of Asp102, and the hydroxyl group of Ser195 of chymotrypsin, are sufficient for the catalytic performance of some enzymes (Fig. 25). The pathway for chymotrypsin-catalyzed reactions has been described [10]. A stereospecific adsorptive complex is formed between the substrate and enzyme (Fig. 25, structure A). The imidazole group of His57 acts as a general base to stretch the H-O bond of the hydroxyl group of Ser195, thus facilitating the nucleophilic attack of the O of Ser-OH on the carbonyl carbon of the peptide bond of the substrate. This leads initially to formation of a tetrahedral intermediate (structure B) followed by formation of the acylenzyme intermediate by expulsion of the R’ group of the substrate (structure C). In the deacylation process, the imidazole group acts as a general base to extract a proton from water to facilitate the attack of the active site-generated hydroxide ion at the carbonyl group of the acylenzyme (structure D). In formation of the second transition-state intermediate (structure E), the imidazole group acts as a general acid in deacylation of the acylenzyme (structure F). The action of His57 as a general base (acylation step) and Pag e 474 FIGURE 25 Proposed mechanism for a-chymotrypsin-catalyzed reactions. The top reactions show acylation of the Ser-OH of the enzyme by the substrate; the bottom reactions show removal of the acyl g roup by hydrolysis. (Modified from Ref. 12.) general acid (deacylation step) is facilitated by hydrogen bonding with the Asp102 carboxyl group. The acyl part of the substrate diffuses away and the enzyme is ready to accept another substrate molecule (structure A). Each cycle requires about 10 msec to complete, with either the acylation (for amide bonds) or deacylation (for ester bonds) being the rate-determining step. Many enzymes require cofactors (nonprotein organic compounds or inorganic ions) for activity. Cofactors include coenzymes, prosthetic groups, and the inorganic ions. Many of the coenzymes and prosthetic groups require a vitamin and often phosphate, ribose, and a nucleotide as part of the cofactor (Table 12). The nucleotide binds into the active site, specifically placing the cofactor so it can participate in the binding and/or catalytic step. Enzymes associated with these cofactors are also listed in Table 12. The essential vitamins, cations, and anions must come from our foods, since we cannot synthesize them. Zn2+, one of the essential cations, is part of the active site of at least 154 different enzymes in our bodies. 7.6.1 Distinguishing Features of Organic Cofactors The coenzymes and prosthetic groups can be distinguished in two important ways. The coenzymes are loosely bound to the active site and dissociate from the enzyme at the end of each catalytic cycle, as shown for the reaction catalyzed by alcohol dehydrogenase (Eq. 32). They also are lost during purification of coenzyme-requiring enzymes and must be added back to the enzymes in the in vitro systems. (32) Pag e 475 TABLE 12 Importance of Phosphate, Ribose, and Purine and Pyrimidine Bases in Cofactors Enzyme Cofactor Vitamin Phosphate Ribose Base Oxidoreductases NAD+ Niacin + + Adenine Oxidoreductases NADP+ Niacin + + Adenine Oxidoreductases FMN Riboflavin + + — Oxidoreductases FAD Riboflavin + + Adenine Lig ases ATP —a + + Adenine Lig ases UTP — + + Uridine Lig ases CTP — + + Cytidine Transferases CoA Pantothenic acid + + Adenine Transferases Acetyl phosphate — + — — Transferases Carbamyl phosphate — + — — Transferases S-Adenosyl methionine — — + Adenine Adenosine-3′-phosphate-5′-phosphosulfate — + + Adenine Transferases and lig ases Pyridoxal phosphate Thiamine pyrophosphate Pyridoxine Thiamine + + — — — — a—, Not present. Source: Adapted from Ref. 111, p. 331. Pag e 476 where Kinetically, the NAD+ behaves as a second substrate in the reaction (Eq. 32). It forms the product NADH, which can be recycled back to NAD+ only by a second enzyme, not alcohol dehydrogenase. The velocity of the reaction is best followed spectrophotometrically based on absorbance of NADH at 340 nm (eM = 6.2 × 103 M-1 cm-1). In contrast to coenzymes, prosthetic groups are tightly bound (often covalently) to the enzyme, and they end up at the end of the cycle in the same oxidation state as they started. A typical example is flavin adenine dinucleotide (FAD), the prosthetic group of glucose oxidase (Eq. 33). (33) 7.6.2 Coenzymes We shall continue with the role of NAD+ as coenzyme for alcohol dehydrogenase. Figure 26 is a schematic diagram of NAD+ bound in the active site of yeast alcohol dehydrogenase, with the adenine ribose phosphate (ADPR) moiety bound at the ADPR binding site. The nicotinamide moiety, the functional part of NAD+ in accepting H from the ethanol , is bound into the lipophobic binding site. Zn2+, a cation cofactor, is required also; it is shown liganded to three amino acid side chains on the protein and coordinately bound to the O of the OH group of ethanol. A tetrahedral intermediate is formed in step 2 followed by transfer of the H from ethanol to from NADH. The turnover number is about 103 moles of substrate converted to product permole enzyme per second. Therefore, the recycle time is about 1 msec. The primary role of the alcohol dehydrogenase is as a specific template, binding ethanol, NAD+ , and Zn2+ stereospecifically at the active site. As discussed in Section 7.5.7, that is a very significant role, accounting for ~1015 rate enhancement. It must also be remembered that the enzyme provides stereospsecific treatment of the substrate. Even though ethanol does not have an asymmetric carbon atom, the enzyme always recognizes ethanol as being asymmetric because of the three-point attachment in the transition state (Fig. 26). The protein also provides the environment, often hydrophobic, in which the reaction takes place. FIGURE 26 Schematic representation of binding NAD + and ethanol in the active site of yeast alcohol dehydrog enase followed by oxidation of ethanol to acetaldehyde. (From Ref. 99, p. 552, by courtesy of Spring er-Verlag .) Pyridoxal phosphate is selected as the prosthetic group to discuss because of its involvement in five types of reactions, one of them being the development of off aroma in broccoli and cauliflower and another the development of the desired aroma of onions and garlic. The reactions catalyzed by pyridoxal phosphate (PALP) are shown in Figure 27. The first step, common to all five reactions, is reaction of the aldehyde group of PALP with the a-amino group of serine (used as an example) to give a Schiff base intermediate. This is stabilized by an acidic group B+ in the enzyme active site. Then, depending on the specific nature of the protein to which PALP is bound, decarboxylation (decarboxylase), deamination (transaminase), a,b- elimination (lyase) or racemization (isomerase) of serine occurs. The intermediate of serine FIGURE 27 Schematic representation of reactions catalyzed by pyridoxal phosphate. Acommon Schiff base intermediate between the amino g roup of serine and the carbonyl g roup of pyridoxal phosphate is shown in the upper center. This results in conversion of substrate to one of five types of products, depending on the enzyme involved. The lower scheme shows b,g-elimination involving the-SH g roup of cysteine. The intermediate is also a Schiff base. B+ is an acidic g roup on the enzyme. PALP and PAP are pyridoxal phosphate and pyridoxamine phosphate, respectively. (From Ref. 111, p. 351.) Pag e 478 formed in the a,b-elimination reaction can react with indole to give tryptophan, or pyruvate and ammonia (one off-aroma compound in broccoli and cauliflower). If serine is replaced with cystine the products are H2S, NH3, pyruvate and thiolcysteine. H2S and NH3 are the two major off-aroma compounds of broccoli. Alliinase is the key enzyme in aroma and flavor development in onions and garlic when they are cut or mashed. Alliinase is a PALP-requiring enzyme. The substrates S-1-propenyl-L- cysteine sulfoxide and S-allyl-L-cysteine sulfoxide undergo a,belimination by alliinase to produce 1-propenylsulfenic acid (a lachrymator) or allysulfenic acid which then nonenzymatically forms the aroma and flavor components of onions and garlic, respectively [113]. 7.6.4 Inorganic lons Both cations and anions can serve as cofactors. The cations include Ca2+, Mg2+, Zn2+, Fe2+, Cu2+, Co2+, Ni2+, Na+ , K+ , and others. The anions include Cl- , Br- , F- , and I- , among others. The cations may be involved directly in catalysis, as shown earlier for yeast alcohol dehydrogenase (Fig. 26), in binding of substrate to the active site, in maintaining the conformation and stability of the substrate, such as Ca2+ in a-amylase, or as part of the substrate, such as MgATP2- as required by kinases. Zn2+ in Escherichia coli milk alkaline phosphatase (APase) will be used as an example because Zn+ is a major essential cation in enzyme systems. Alkaline phosphatase requires four Zn2+ ions per molecule of enzyme (80,000 MW). Two of the Zn2+ serve a structural function, helping to hold the two subunits of the enzyme together (maintaining quaternary structure). The other two Zn2+ are located in the two active sites of the enzyme, where the role of the Zn2+ is as a “super acid” group to activate the SerOH group involved in the hydrolysis of the p-nitrophenylphosphate substrate: (34) The in vivo role of alkaline phosphatase is to help recycle phosphate, an essential compound. Alkaline phosphatase is ideally designed for this, as it is specific for the orthophosphate group only and it does not really matter what the other group (pnitrophenyl here) is. Chloride ion is an important anionic factor. Cl- is essential for the activity of salivary and pancreatic a-amylases of humans and other animals and for some microbial a-amylases. Its role in a-amylase appears to be twofold. The binding constant for Ca2+
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 (another cation cofactor) in pancreatic a-amylase is increased ~100 times in the presence of Cl- , thereby increasing the effectiveness of Ca2+ as a stabilizer of the tertiary structure of the protein [71]. Cl- also causes a shift in the pH optimum of human salivary a-amylase [83] from pH 6 in the absence of Cl- (much lower levels of activity) to pH 6.8 in the presence of Cl- (0.005–0.04 M Cl- gives maximum activity). It is postulated that the pH shift results from Cl- masking an unwanted positively charged group in or near the active site of the enzyme [83]. Perhaps this represents a primitive control mechanism in some aamylases. 7.7 Enzyme Inactivation and Control Enzymes are responsible for the myriad reactions associated with reproduction, growth, and maturation of all organisms. In most cases, these are desired activities. In some cases, too much enzyme activity, such as polyphenol oxidase-caused browning, can lead to major losses in fresh Pag e 479 fruits and vegetables. In many humans, absence of or too little of an enzyme is responsible for many genetically related diseases [104]. Microbially caused diseases present another problem. The best way to treat these types of diseases is through inhibition of one or more key enzymes of the microorganisms, resulting in their death. The inhibitors might compete reversibly with substrates or cofactors for binding to the active site, or the inhibitor might form a covalent bond with active site groups (affinity labeling inhibitor), or the compound might be treated as a substrate and be catalyzed to a product that, while still in the active site, forms a convalent bond with a group in the active site (kcat inhibitors) [8]. The last type is the most specific and desirable in medicine and in food because the inhibitor can be targeted specifically for the enzyme. Enzymes continue to catalyze reactions in raw food materials after they reach maturity. These reactions can lead to loss of color, texture, flavor and aroma, and nutritional quality. Therefore, there is need for control of these enzymes to stabilize the product as food. Enzyme inhibitors are also important in the control of insects and microorganisms that attack raw food. They also are used as herbicides in the control of unwanted weeds, grasses, and shrubs. Enzyme inhibitors are an important means of controlling enzyme activity. An enzyme inhibitors is any compound that decreases vo when added to the enzyme-substrate reaction. There are many enzyme inhibitors, both naturally occurring and synthetic [106]. Some inhibitors bind reversibly to enzymes and others form irreversible, covalent bonds with the enzymes. Some inhibitors are large proteins or carbohydrates, and others are as small as HCN. Products of enzyme-catalyzed reactions can be inhibitory. Change in pH can alter activity by making conditions less optimum for enzyme activity. Elevated temperatures can decrease enzyme activity by denaturing some of the enzyme, but at the same time increasing the velocity of conversion of substrate to product by the active enzyme. Most enzymologists do not consider either of these variables to be enzyme inhibitors. Denaturation of the enzyme eliminates its activity, and this can be accomplished by shear forces, very high pressures, irradiation, or miscible organic solvents. Enzyme activity can also be decreased by chemical modification of essential active site groups of the enzyme. Enzymes are also inactive when their substrate(s) are removed. All of these inhibitory approaches are valid ways of controlling enzyme activities in foods. 7.7.1 Reversible Inhibitors Reversible inhibitors are distinguished from irreversible inhibitors by the following criteria: (a) reversible inhibitors rapidly (within milliseconds) form noncovalent diffusion-controlled equilibrium complexes with enzymes; the complex can be dissociated and enzyme activity restored by displacing the equilibrium by dialysis or by gel filtration; (b) irreversible inhibitors slowly form covalent derivatives of the enzyme that cannot be dissociated by dialysis or by gel filtration. The reversible inhibitors can be treated kinetically by the methods described in Section 7.4 provided the equilibrium dissociation constant (Ki) is not less then 10-8 M. Four types of reversible inhibitors can be identified based on their effects on the slopes and intercepts of Lineweaver-Burk plots, Vmax and Km, or allosteric effects: (a) competitive inhibitors, (b) noncompetitive inhibitors, (c) uncompetitive inhibitors, and (d) allosteric inhibitors. Allosteric #inhibitors not only decrease the velocity of the enzymecatalyzed reactions, they cause allosteric (sigmoidal) plots of vo versus [I], similar to the effect of [S]o (Fig. 11). The type of reversible inhibitor must be determined by kinetic methods. Fortunately, all the mathematical equations differ in slope and/or y-intercept only by 1 + [I]o/Ki from the Michaelis-Menten and Lineweaver-Burk equations in the absence of inhibitor. Pag e 480 7.7.1.1 Competitive Inhibition In addition to the assumptions made in derivation of the Michaelis-Menten equation (Sec. 7.4.2.2), it is assumed for competitive inhibition that [I]o >> [E]o, such that [I]@ [I]o. In competitive inhibition the substrate and the competitive inhibitor compete for the binding to the enzyme, as shown in Figure 28 and Equations 35 and 36: (35) (36) where [I] is the free inhibitor concentration, and E.I is the enzyme-inhibitor complex, and Ki = k3/k3. E.I does not bind S, and it does not form a product. The conservation equation with respect to enzyme concentration is FIGURE 28 Schematic representation of inhibition of enzymes by various types of inhibitors. The model of E shows only the active site with the binding locus (inner void) and the transforming locus, with catalytic g roups A and B. The symbols for the species involved (in parentheses) are: E, free enzyme; E.A, enzyme-substrate complex; E-A’, acylenzyme intermediate; C, competitive inhibitor; N, noncompetitive inhibitor; U, uncompetitive inhibitor;P 1 and P2 are products formed from the substrate, A, and E.N, E.A’.N, E.C, and E.A’.U are complexes with respective enzyme species and inhibitors. All complexes formed involve noncovalent bonds except E-A’, where a covalent bond is formed with catalytic g roup A. (From Ref. 111, p. 235.) Pag e 481 [E]o = [E] + [E.S] + [E.I] (37) Inclusion of Equations 36 and 37 in derivation of the Michaelis-Menten equation gives (38) where vo’ is the initial velocity in the presence of a fixed concentration of inhibitor. The Lineweaver-Burk equation derived from Equation 38 is (39) A plot of experimental data in the presence and absence of a competitive inhibitor is shown in Figure 29. Reactions with variable [S]o must be run in the absence and presence of a fixed concentration of [I]o, and vo and vo’ must be determined. Linear competitive inhibition is indicated experimentally when the y-intercept is the same in the presence and absence of inhibitor, but the slope in the presence of inhibitor is greater than that in the absence of inhibitor by 1 + [I]o/Ki. 7.7.1.2 Noncompetitive Inhibition In this type of inhibition, the inhibitor does not compete with the substrate for binding with enzyme, so the inhibitor and substrate can bind to the enzyme simultaneously (Fig. 28; Eqs. 40–43). FIGURE 29 Linear competitive inhibition. The 1/ v o and 1/v o’ values are plotted versus 1/[A]o. Data are plotted in reciprocal form according to Equation 39. [I] o1 = Ki. [A]o is initial substrate concentration. (From Ref. 111, p. 228.) Pag e 482 (40) (41) (42) (43) The assumptions are that k-1/k1 is the dissociation constant, KS, for both E.S and E.S.I, and k-3/k3 is the dissociation constant, Ki, for both E.I. and E.S.I. Further, E.S.I. does not form products from the substrate. These assumptions lead to simple linear noncompetitive inhibition where (44) and the corresponding Lineweaver-Burk equation is (45) The enzyme conservation equation is [E]o = [E] + [E.S] + [E.I.] + [E.S.I] (46) In the case of simple linear noncompetitive inhibition, both the y-intercept and the slope are increased by 1 + [Io]/Ki (Fig. 30). 7.7.1.3 Uncompetitive Inhibition Unlike the cases of competitive and noncompetitive inhibition, in uncompetitive inhibition, the inhibitor cannot bind to the free enzyme but only with one or more of the intermediate complexes as shown in Fig. 28. Equations 47 and 48 apply: (47) (48) where E.S.I does not form product from the substrate. The appropriate form of the Michaelis- Menten equation is Pag e 483 FIGURE 30 Simple linear noncompetitive inhibition. The 1/ v o and 1/v o’ values are plotted versus 1/[A] o according to Equation 45. [Io]1 =Ki, where k intercept’ = k slope’ [A]o is initial substrate concentration. (From Ref. 111, p. 229.) (49) and the corresponding Lineweaver-Burk and conservation equations are Equations 50 and 51: (50) [E]o = [E] + [E.S] + [E.S.I] (51) The associated plot is shown in Figure 31. The y-intercept changes by 1 + [I]o/Ki from the reaction with no inhibitor, while the slopes in the presence and absence of an uncompetitive inhibitor are the same. 7.7.1.4 Allosteric Inhibition Allosteric inhibition usually results from the binding of inhibitor to multi-subunit enzymes, in the same way as described for allosteric behavior on substrate binding (Eq. 28). Whenever Ki2 and subsequent Ki values are smaller (tighter binding) than Ki1, positive allosteric inhibition results. Negative allosteric inhibition results when Kil, is smaller than Ki2 and subsequent Ki values. Allosteric inhibition can be quantified by the methods of Monod et al. [80] and Koshland et al. [66] or by the modified Hill equation [47]. The Hill equation, developed for O2 or CO2 binding to hemoglobin, is applicable to substrate or inhibitor binding to enzymes, Pag e 484 FIGURE 31 Linear uncompetitive inhibition. Data plotted in reciprocal form according to Equation 49. [I o]1 = 2Ki. [A]o is initial substrate concentration. (From Ref. 111, p. 232.) to give allosteric kinetics. The Hill equation for effect of [S]o on an enzyme-catalyzed reaction is (52) where n is the apparent number of interacting binding sites. Equation 52 can be written in a linear transform as FIGURE 32 Effect of inhibitor concentration on activity of an allosteric-behaving enzyme system plotted according to Equation 54. (From Ref. 111, p. 233.) Pag e 485 (53) with the analogous equation for an inhibitor being (54) A plot of experimentally obtained v ‘o values as a function of [S]o concentration is shown in Figure 32. The [I]0.5 for 50% inhibition of the enzyme is shown at the common cross point of two sets of data obtained at two different, not saturating, inhibitor concentrations. The slope values, m indicate the number of apparent interacting binding sites with inhibitor, modified by the strength of interaction, n. 7.7.2 Irreversible Inhibitors Many inhibitors form covalent derivatives with enzymes; therefore, they should be treated by rate equations, not by Ki. Kinetically, irreversible inhibition also occurs when Ki<1 1.2="" 10-9="" 55="" 7.7.1.="" about="" are="" at="" be="" because="" bind="" but="" by="" called="" cannot="" complexes="" confusion="" covalent="" derivatives="" described="" dissociation="" e.i.="" enzymes="" eq.="" form="" from="" in="" inhibitors.="" inhibitors="" is="" k-1="" m="" methods="" min.="" noncovalent="" of="" or="" q.="" rapidly="" rate="" rates="" section="" since="" slow="" slower="" so="" some="" sometimes="" strike="" t0.5="" the="" there="" these="" this="" tight-binding="" tight="" to="" treated="" very="">One needs to know which step(s) leads directly to loss of enzyme activity (rate = k1[E]o[I]o or k2 [E.I]) in order to interpret the data and identify the mechanism of the reaction. The reaction shown by Equation 55 is typical of affinity labeling inhibitors. Sometimes when compounds bind into the active site of an enzyme they are catalyzed to products, which then react covalently with the enzyme to irreversibly inactivate it: (56) When k3 > k4 the enzyme is rapidly inactivated. Should the substrate be called an inhibitor or a substrate? These types of inhibitors are called kcat inhibitors and are among the most specific inhibitors known. They are highly valued in medicine, and could be very useful in control of enzymes of foods. Other irreversible enzyme inhibitors are specific for certain amino acid side-chain groups of the enzyme, such as the sulfhydryl group, the amino group, the carboxyl group, and the imidazole group. Generally, these types of reactions are called protein modification. A lot of different compounds have been studied in this connection, in an attempt to determine the nature of one or more groups in the active site of enzyme or to label a specific group so that its location in the primary amino acid sequence can be determined. Some examples are given in Table 13. They are often not specific for a single type of group and may react also with the same type of specific group outside the active site, unless the one in the active site is more reactive than the Pag e 486 TABLE 13 Some Reag ents Used in Chemical Modification of Enzymes Group modified Ser Reag ent Amino Carboxyl Sulfhydryl Thr His Trp Tyr Met Acetic anhydride + + + + Acetyl imidazole ± + + Aryl sulfonylchlorides + + ± + + NN-Bromosuccinimide + + + + Carbodiimides + Diazomethane + + + + 2,4-Dinitrofluorobenzene + + + + N-Ethyl maleimide ± + ± Haloacetate, haloamide + + + + Hydrog en peroxide ± ± + Hydroxylamine + Hydroxynitrobenzylbromide + Mercuribenzoate ± + ± Photooxidation + + + + ± Tetranitromethane + + ± Trinitrobenzene sulfonate + Source: Adapted from Ref. 48. same groups outside the active site. The reader is referred to Whitaker [111] for more detailed discussion of these types of inhibitors. 7.7.3 Naturally Occurring Inhibitors Some organisms have evolved enzyme inhibitors as metabolic and physiological regulatory systems. They also produce inhibitors to prevent premature activation of proenzymes and to protect the organism against insect and microbial attacks. Ryan [89] has shown that insect larvae feeding on leaves or stems of tomatoes and potatoes stimulates rapid protease inhibitor production, which causes the larvae to quit feeding. The elicitors appear to be pectin fragments produced by action of pectic enzymes, in response to tissue damage. Most of the elicited protease inhibitors are proteins. 7.7.3.1 Protease Inhibitors Trypsin inhibitors appear to be ubiquitous in all tissues, and they have been most intensively studied in the legumes and cereals. Soybean seeds contain two types of trypsin inhibitors, the Kunitz inhibitor of 21,000 MW that is specific for trypsin only (1:1 complex) and the Bowman-Birk inhibitor of 8300 MW that binds independently and simultaneously to trypsin and chymotrypsin (1:1:1 complex). Most legumes contain the Bowman-Birk type inhibitors, with considerable homology among them, while most legumes do not produce the Kunitz-type Pag e 487 inhibitor. The Kunitz inhibitor, with two disulfide bonds, is much more heat labile than the Bowman-Birk inhibitors, with seven disulfide bonds. About 1 h of cooking is required to completely inactivate the Bowman-Birk inhibitor, unless a reducing compound, such as cysteine, is added [39]. Several isoinhibitors of trypsin are often found in higher plants [116]. The legume inhibitors are known to be significant, nutritionally, at least in some animals. As shown in Fig. 33, the PER (protein efficiency ratio) increases as the amount of inhibitor decreases. As stated earlier, the inhibitor can be inactivated by prolonged cooking. The four red kidney bean protease inhibitors have Ki values for bovine trypsin ranging from 3.4 × 10-10 to 8.4 × 10-10 M, while Ki for bovine chymotrypsin is 5.5 × 10-10 to 4.0 × 10-9 M [116]. These are slow, very tight-binding inhibitors. The Ki values can differ significantly with trypsin from different sources. Chicken egg white contains three types of protease inhibitors, the ovomucoids specific for trypsin, the ovoinhibitor with specificity for trypsin, chymotrypsin, subtilisin, and Aspergillus oryzae protease, and the papain inhibitor (cystatin) with specificity for papain and ficin [107]. Protease inhibitors are found in the pancreas and in the blood, where they protect against premature activation of the pro-forms of the digestive proteases and the blood clotting proteases, respectively. Microorganisms contain a variety of low-molecular-weight peptide inhibitors of proteases. These inhibitors include the leupeptins active against trypsin, plasmin, papin, and cathepsin B, the chymostatins active against chymotrypsin and papain, elastatinal active against elastase, pepstatins active against pepsin and several other carboxyl proteases and phosphoramidon active against thermolysin. The reader is referred to Whitaker [107] for more detailed information. These small peptide inhibitors are very stable and can be produced economically by fermentation. Therefore, they could be used to control some enzyme activities in foods. 7.7.3.2 a-Amylase Inhibitors There are three types of a-amylase inhibitors: (a) proteins produced by higher
———————————————————————————————————————- plants, (b) small polypeptides produced by several species of Streptomyces, and (c) small N-containing carbohydrates produced by Streptomyces [50]. Several higher plants contain inhibitors of aamylases of mammals and insects, with a few FIGURE 33 Effect of heat treatment on the trypsin inhibitory activity and nutritive value, as measured by the protein efficiency ratio (PER), of soybean meal. (From Ref. 87, p. 164A, with permission of American Oil Chemists Society.) Pag e 488 of the inhibitors active on microbial a-amylases and host-specific a-amylases. Sources include wheat, corn, barley, millet, sorghum, peanut, and beans. The proteins range from 9000 to 63,000 MW. Most slowly form 1:1 stoichiometric complexes with the a-amylases. The Ki values for the been a-amylase inhibitors are 10-10 to 10-11 M; therefore, these inhibitors are slow, tight-binding inhibitors. They are quite heat stable, so that some of the inhibitor can survive breadmaking (for example) and up to 1 h cooking of the seeds. Whether these a-amylase inhibitors are nutritionally important is controversial. They certainly slow down the rate of digestion of starch in saliva and small intestine of humans and rats, and some undigested starch is present in the feces of rats fed the inhibitors. Glucose is not released as rapidly from starch digestion into the blood of humans and rats in the presence of the inhibitors. However, there is little effect of the inhibitor on growth of rats and chickens. Relatively small polypeptide inhibitors of aamylase ranging from 3936 to 8500 MW are produced by several species of Streptomyces. There is much homology among the five polypeptide inhibitors that have received the most attention. The microbially derived polypeptide a-amylase inhibitors do not appear to have been tested with plant, insect, and most animal aamylases. Their reaction with human a-amylases has been studied extensively at a clinical level. The Streptomyces also produce three types of N-containing carbohydrates that are effective a-amylase and a-glucosidase inhibitors. These are the oligostatins, the amylostatins, and the trestatins. All have in common a pseudodisaccharide unit, oligobioamine or dehydro-oligobioamine, a variable number of a-D-glucose units linked a-1,4, and in one type a-1,1. Substantial clinical research has been done on their effects in modulating the effects of diabetes and hyperglycemia. 7.7.3.3 Invertase Inhibitors Irish potatoes and sweet potatoes (yams) contain invertase inhibitors [107]. Invertase inhibitor in the Irish potato is a 17,000- MW protein. It inhibits potato invertase and several other plant invertases [86]. The level of invertase inhibitor in stored potatoes is temperature dependent, increasing at higher temperatures and decreasing at lower temperatures. The in vivo function of the inhibitor in potatoes is thought to be regulation of invertase activity. Potatoes stored at lower, nonfreezing temperatures are sweeter (more sucrose) than those stored at higher temperatures (where the potato is more starchy). These enzymatically caused changes are important both for quality and cost. 7.7.3.4 Other Enzyme Inhibitors Inhibitors against several other enzymes have been reported [107]. Probably the best understood, physiologically, are the protein phosphatase inhibitors and the protein kinase inhibitors. Phosphorylase and glycogen synthase are the key enzymes involved in the metabolic breakdown of glycogen to glucose, and biosynthesis of glucose glycogen in animal tissues, respectively. The role of the protein phosphatase inhibitors and the protein kinase inhibitors appears to be regulation of the amount of phosphorylase a present by the mechanism shown in Equation 57. (57) Pag e 489 7.7.3.5 Chemically Tailored Inhibitors Chemical tailoring of inhibitors is a major activity in pharmaceutical research laboratories. In the 1950–1970 era the affinity labeling inhibitors (Eq. 55) were the big thing because of the increased specificity in covalent binding to the enzyme that needed to be inhibited. Then came the kcat-type inhibitors (Eq. 56) with their double specificity of binding selectively to the active site, and catalytic conversion to a product that irreversibly inactivates the enzyme while still in the active site. Now two new thrusts are to target malignant cells, as in cancer, specifically with appropriate antibodies, and to destroy the cells either by a radio label or by tailoring the target molecule to enter the cell and inhibit a key enzyme(s) in the cell division process. The other major area is the use of antisense mRNA designed to “turn off” a specific gene [25], as is done in the Flavr Savr tomato in suppressing the translation of the polygalacturonase gene. This is a general method that, in theory, can be used to suppress the production of any detrimental enzyme in our raw food products or those responsible for diseases of plants and animals. This method is much more specific than decreasing the level of an enzyme by breeding, as has been done, for example, with polyphenol oxidase in peaches. 7.7.4 Inactivation and/or Control by Physical Methods Several handling methods used in food processing inactivate enzymes, intentionally or not. These include exposure to interfaces formed along the walls of transport tubes or the effects of stirring, whipping, shearing, high temperatures, pressure, and heat. However, not all of these lead to “instantaneous” or complete
———————————————————————————————————————-
 inactivation, and there can be an initial period when the activity of the enzyme, in contact with its substrate, can be enhanced. Chemical changes, enzymatically catalyzed or not, intended or not, are of major importance to the quality and safety of foods. 7.7.4.1 Interfacial Inactivation of Enzymes Air/water and lipid/water interfaces are regions of high energy in which proteins tend to align themselves, with hydrophilic segments oriented into the water phase and hydrophobic segments oriented into the air or lipid phase. Unfolding of the protein at the interface is thermodynamically favored. Proteins are ideal surface-active molecules and very important in foaming and emulsifying in foods (see Chaps. 3 and 6). Mild shaking of a solution of enzyme can cause rapid denaturation. This effect can be a disadvantage in orange and tomato juice products and in the cold break process in tomato paste production. Sometimes, foaming is used in enzyme or protein purification, where one protein is more stable to foaming than the other. For example, the purification of invertase inhibitor from Irish potatoes is a problem because the inhibitor complexes with the invertase present. By controlled foaming the invertase can be denatured, enhancing recovery of undenatured inhibitor [15]. 7.7.4.2 Pressure Effect on Enzymes Normal food processing practices do not create high enough pressures alone to inactivate enzymes. In combination processes, such as pressure combined with high-temperature treatment and/or high shear rates, the latter two aspects are primarily responsible for any enzyme inactivation. Extruders operate at pressures greater than those normally encountered during food processing, and this results in greater fluidity, texturization of the food through shear forces, chemical reactions, and high temperatures. It is doubtful that most enzymes would survive extruder conditions. Research has been done on the use of high pressures to inactivate microorganisms. Pag e 490 Hydrostatic pressures that are compatible with intact food tissues almost certainly will not completely inactivate enzymes of interest. It is anticipated that pressure treatment of food will have greater effect on multi-subunit enzymes than on single polypeptide enzymes, since pressure, if high enough, will favor the monomer form of a protein, that is, will cause more dissociation to the monomer. Gutfreund and his colleagues [44] studied the effect of a pressure jump on the dissociation of hemoglobin. They reported that 107 Pa pressure caused 4% dissociation into two a,b-dimers, a relatively easy dissociation (i.e., 0.1 M NaCl will do this). 7.7.4.3 Inactivation of Enzymes by Shearing Considerable shearing occurs in food processing during mixing, transfer through tubes, and extrusion. Some attention has been given to the shear conditions required for enzyme inactivation. Charm and Matteo [20] determined that when the shear value [shear rate (sec-1) times the exposure time (sec)] is greater than 104 some inactivation of rennet is detectable (Fig. 34), and the rate of inactivation is very much faster at 7 × 105 . About 50% inactivation of catalase, rennet and carboxypeptidase was found at shear values of about 107 [20]. Rennet regained some activity after shear inactivation, similar to regaining of activity following heat inactivation. 7.7.4.4 lonizing Radiation and Enzyme Inactivation Much attention has been given to the effect of ionizing radiation of food on the inactivation of enzymes [34]. It soon became clear that about 10 times greater dose of ionizing radiation is required to inactivate enzymes than is needed to destroy microbial spores. For example, irradiated meat in which the microbial population was reduced to quite low numbers still developed textural defects due to protease activity during storage. Some of the factors that influence the rate of trypsin inactivation by irradiation are shown in Figure 35. Trypsin is more stable in the dry state than in the wet state, because the indirect effect of free radicals (HO. and H.) produced from water are very important in enzyme FIGURE 34 Inactivation of rennet by shearing at 4°C. (Reprinted from Ref. 20, p. 505, couurtesy of Academic Press.) Pag e 491 FIGURE 35 Inactivation of trypsin by irradiation under various conditions: (1) dry trypsin, (2) 10 mg /ml of trypsin solution at pH 8 and – 78°C, (3) 80 mg /ml of trypsin solution at pH 8 and ambient temperature, (4) 10 mg /ml of trypsin at pH 2.6 and -78°C, (5) 80 mg /ml of trypsin at pH 2.7 and ambient temperature, (6) 10 mg /ml of trypsin at pH 2.6 and ambient temperature, (7) 1 mg /ml of trypsin at pH 2.6 and ambient temperature. (Redrawn from Ref. 69.) inactivation. In general, the more dilute the trypsin solution, the more effective the radiation treatment. A given dose of ionizing radiation is also more effective at ambient temperature than at -78°C. At least part of this difference in effect results from immobilization of water free radicals at -78°C (frozen state). Differences in pH do not seem to have a pronounced effect on irradiation inactivation of trypsin, but these differences have been shown to be important in activating enzymes by irradiation in meat. Other factors that enhance inactivation by ionizing radiation are the presence of O2, metal ions such as Cu2+, unsaturated lipids, fatty acids, and purity of the enzyme, since other proteins protect against inactivation. Individual enzymes differ substantially in their resistance to inactivation by ionizing radiation, with catalase being about 60 times more resistant than carboxypeptidase (both metal-containing enzymes; Fe3+ is the cofactor in catalase, Zn2+ is the cofactor in carboxypeptidase). Ionizing radiation causes most damage to tyrosyl, histidyl, tryptophanyl, and cysteinyl residues in proteins. Sulfhydryl compounds in food tend to protect enzymes against inactivation by ionizing radiation. In order to avoid using higher doses of ionizing radiation than required to destroy microorganisms, heat treatment (to inactivate enzymes) is usually combined with ionizing radiation in those foods where this preservation method is permitted. 7.7.4.5 Solvent Inactivation of Enzymes The effect of solvents on enzyme activity was discussed for model systems in Section 7.5.6. In general, water immiscible solvents, by displacing water, stabilize enzymes, just as does removal of water by drying. However, water-miscible solvents, when present at concentrations exceeding about 5–10%, generally inactivate enzymes. This effect is, of course, temperature dependent (more stable at lower temperatures). Solvent treatment, for example with ethanol, can be quite effective in inactivating Pag e 492 microorganisms on the surface of grains and legume seeds. However, the efficacy of this treatment probably does not depend on inactivation of enzymes. 7.7.5 Removal of Substrate and/or Cofactor It is obvious that removal of one or more of the required substrates or a required cofactor will eliminate enzyme activity. It is well known that polyphenol oxidase browning can be prevented by excluding O2. This is effectively done by natural selective permeability of the surface of most fruits and vegetables. Browning catalyzed by polyphenol oxidase can also be prevented by removing or altering the phenols (substrates). Binding of phenols to polyethylene glycol, polyvinylpyrrolidone, or Sephadex is effective. Polyphenol oxidase activity can be prevented by enzymatic methylation of one or more of the hydroxyl groups of the phenolic substrate [37]. This might be effective in juice, for example. Another approach to preventing browning caused by polyphenol oxidase is to reduce the initial product, o-benzoquinone, back to the substrate before the o-benzoquinone can undergo further (non-enzyme-catalyzed) oxidative polymerization to melanin. Compounds that reduce o-benzoquinone include ascorbic acid, sodium bisulfite, and thiol compounds. It has been shown that these compounds also directly inactivate polyphenol oxidase by free radical degradation of histidine residues at the active site (ascorbic acid and Cu2+), and by reducing Cu2+ to Cu+ in the active site of the enzyme, thereby causing Cu+ to dissociate more readily from the enzyme [84]. Enzymes are generally more stable in the presence of substrates, cofactors and competitive inhibitors. The general conclusion is that binding of these compounds into the active site of the enzyme has a stabilizing effect.

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