Chapter 16

Although the catalytic properties of enzymes may seem almost magical, it is simply chemistry—the breaking and making of bonds—that gives enzymes their prowess. This chapter will explore the unique features of this chemistry. The mechanisms of hundreds of enzymes have been studied in at least some detail. In this chapter, it will be possible to examine only a few of these. Nonetheless, the chemical principles that influence the mechanisms of these few enzymes are universal, and many other cases are understandable in light of the knowledge gained from these examples.

16.1 • The Basic Principle — Stabilization of the Transition State

In all chemical reactions, the reacting atoms or molecules pass through a state that is intermediate in structure between the reactant(s) and the product(s). Consider the transfer of a proton from a water molecule to a chloride anion:

In the middle structure, the proton undergoing transfer is shared equally by the hydroxyl and chloride anions. This structure represents, as nearly as possible, the transition between the reactants and products, and it is known as the transition state.¹

¹It is important here to distinguish transition states from intermediates. A transition state is envisioned as an extreme distortion of a bond, and thus the lifetime of a typical transition state is viewed as being on the order of the lifetime of a bond vibration, typically 10^{-13
sec}. Intermediates, on the other hand, are longer-lived, with lifetimes in the range of 10^-13 sec to 10^-3 sec.

Chemical reactions in which a substrate (S) is converted to a product (P) can be pictured as involving a transition state (which we henceforth denote as X^‡), a species intermediate in structure between S and P (Figure 16.1). As seen in Chapter 14, the catalytic role of an enzyme is to reduce the energy barrier between substrate and transition state. This is accomplished through the formation of an enzyme - substrate complex (ES). This complex is converted to product by passing through a transition state, EX^‡ (Figure 16.1). As shown, the energy of EX^‡ is clearly lower than X^‡. One might be tempted to conclude that this decrease in energy explains the rate acceleration achieved by the enzyme, but there is more to the story.

Figure 16.1 • Enzymes catalyze reactions by lowering the activation energy. Here the free energy of activation for (a) the uncatalyzed reaction, DG_u ^‡, is larger than that for (b) the enzyme-catalyzed reaction, DG_e^‡.

The energy barrier for the uncatalyzed reaction (Figure 16.1) is of course the difference in energies of the S and X^‡ states. Similarly, the energy barrier to be surmounted in the enzyme-catalyzed reaction, assuming that E is saturated with S, is the energy difference between ES and EX^‡. Reaction rate acceleration by an enzyme means simply that the energy barrier between ES and EX^‡ is less than the energy barrier between S and X^‡. In terms of the free energies of activation, DGe^‡ , DGu^‡.
There are important consequences for this statement. The enzyme must stabilize the transition-state complex, EX^‡, more than it stabilizes the substrate complex, ES. Put another way, enzymes are “designed” by nature to bind the transition-state structure more tightly than the substrate (or the product). The dissociation constant for the enzyme-substrate complex is

Enzyme catalysis requires that K_T<K_S. According to transition-state theory (see references at end of chapter), the rate constants for the enzyme-catalyzed (k_e) and uncatalyzed (ku) reactions can be related to K_S and K_T by:

Thus, the enzymatic rate acceleration is approximately equal to the ratio of the dissociation constants of the enzyme-substrate and enzyme-transition-state complexes, at least when E is saturated with S.

Enzymes are powerful catalysts. Enzyme-catalyzed reactions are typically 10⁷ to 10¹⁴ times faster than their uncatalyzed counterparts (Table 16.1). (There is even a report of a rate acceleration of >10¹⁶ for the alkaline phosphatase -catalyzed hydrolysis of methylphosphate !)

These large rate accelerations correspond to substantial changes in the free energy of activation for the reaction in question. The urease reaction, for example,

shows an energy of activation some 84 kJ/mol smaller than the corresponding uncatalyzed reaction. To fully understand any enzyme reaction, it is important to account for the rate acceleration in terms of the structure of the enzyme and its mechanism of action. There are a limited number of catalytic mechanisms or factors that contribute to the remarkable performance of enzymes. These include the following:

Any or all of these mechanisms may contribute to the net rate acceleration of an enzyme-catalyzed reaction relative to the uncatalyzed reaction. A thorough understanding of any enzyme would require that the net acceleration be accounted for in terms of contributions from one or (usually) more of these mechanisms. Each of these will be discussed in detail in this chapter, but first it is important to appreciate how the formation of the enzyme-substrate (ES) complex makes all these mechanisms possible.

How is it that X^‡ is stabilized more than S at the enzyme active site? To understand this, we must dissect and analyze the formation of the enzyme-substrate complex, ES. There are a number of important contributions to the free energy difference between the uncomplexed enzyme and substrate (E + S) and the ES complex (Figure 16.2). The favorable interactions between the substrate and amino acid residues on the enzyme account for the intrinsic binding energy, DG_b. The intrinsic binding energy ensures the favorable formation of the ES complex, but, if uncompensated, it makes the activation energy for the enzyme-catalyzed reaction unnecessarily large and wastes some of the catalytic power of the enzyme.

Figure 16.2 • The intrinsic binding energy of the enzyme-substrate (ES) complex ( DG_b ) is compensated to some extent by entropy loss due to the binding of E and S (T D S) and by destabilization of ES ( DG_d) by strain, distortion, desolvation , and similar effects. If DG_b were not compensated by T D S and DG_d, the formation of ES would follow the dashed line.

Compare the two cases in Figure 16.3. Because the enzymatic reaction rate is determined by the difference in energies between ES and EX^‡, the smaller this difference, the faster the enzyme-catalyzed reaction. Tight binding of the substrate deepens the energy well of the ES complex and actually lowers the rate of the reaction.

Figure 16.3 • (a) Catalysis does not occur if the ES complex and the transition state for the reaction are stabilized to equal extents. (b) Catalysis will occur if the transition state is stabilized to a greater extent than the ES complex (right). Entropy loss and destabilization of the ES complex DG_d ensure that this will be the case.

The message of Figure 16.3 is that raising the energy of ES will increase the enzyme-catalyzed reaction rate. This is accomplished in two ways: (a) loss of entropy due to the binding of S to E, and (b) destabilization of ES by strain, distortion, desolvation , or other similar effects. The entropy loss arises from the fact that the ES complex (Figure 16.4) is a highly organized (low-entropy) entity compared to E + S in solution (a disordered, high-entropy situation). The entry of the substrate into the active site brings all the reacting groups and coordinating residues of the enzyme together with the substrate in just the proper position for reaction, with a net loss of entropy. The substrate and enzyme both possess translational entropy, the freedom to move in three dimensions, as well as rotational entropy, the freedom to rotate or tumble about any axis through the molecule. Both types of entropy are lost to some extent when two molecules (E and S) interact to form one molecule (the ES complex). Because DS is negative for this process, the term -T D S is a positive quantity, and the intrinsic binding energy of ES is compensated to some extent by the entropy loss that attends the formation of the complex.

Figure 16.4 • Formation of the ES complex results in a loss of entrophy. Prior to binding, E and S are free to undergo translational and rotational motion. By comparison, the ES complex is a more highly ordered, low-entrophy complex.

Destabilization of the ES complex can involve structural strain, desolvation , or electrostatic effects. Destabilization by strain or distortion is usually just a consequence of the fact (noted previously) that the enzyme is designed to bind the transition state more strongly than the substrate. When the substrate binds, the imperfect nature of the “fit” results in distortion or strain in the substrate, the enzyme, or both. This means that the amino acid residues that make up the active site are oriented to coordinate the transition-state structure precisely, but will interact with the substrate or product less effectively.
Destabilization may also involve desolvation of charged groups on the substrate upon binding in the active site. Charged groups are highly stabilized in water. For example, the transfer of Na⁺ and Cl^- from the gas phase to aqueous solution is characterized by an enthalpy of solvation, DH_solv, of -775 kJ/mol. (Energy is given off and the ions become more stable.) When charged groups on a substrate move from water into an enzyme active site (Figure 16.5), they are often desolvated to some extent, becoming less stable and therefore more reactive.

Figure 16.5 • Substrates typically lose waters of hydration in the formation of the ES complex. Desolvation raises the energy of the ES complex, making it more reactive.

When a substrate enters the active site, charged groups may be forced to interact (unfavorably) with charges of like sign, resulting in electrostatic destabilization (Figure 16.6). The reaction pathway acts in part to remove this stress. If the charge on the substrate is diminished or lost in the course of reaction, electrostatic destabilization can result in rate acceleration.

Figure 16.6 • Electrostatic destabilization of a substrate may arise from juxtaposition of like charges in the active site. If such charge repulsion is relieved in the course of the reaction, electrostatic destabilization can result in a rate increase.

Whether by strain, desolvation , or electrostatic effects, destabilization raises the energy of the ES complex, and this increase is summed in the termDG_d, the free energy of destabilization. As noted in Figure 16.2, the net energy difference between E + S and the ES complex is the sum of the intrinsic binding energy, DG_b; the entropy loss on binding, - T D S; and the distortion energy, DG_d. ES is destabilized (raised in energy) by the amount DG_d - T D S. The transition state is subject to no such destabilization, and the difference between the energies of X^‡ and EX^‡ is essentially DG_b , the full intrinsic binding energy.

Although not apparent at first, there are other important implications of Equation 16.3. It is important to consider the magnitudes of K_S and K_T. The ratio k_e/ k_u may even exceed 10¹⁶, as noted previously. Given a typical ratio of 10¹² and a typical K_S of 10^-3 M, the value of K_T should be 10^-15 M! This is the dissociation constant for the transition-state complex from the enzyme, and this very low value corresponds to very tight binding of the transition state by the enzyme.
It is unlikely that such tight binding in an enzyme transition state will ever be measured experimentally, however, because the transition state itself is a “moving target.” It exists only for about 10^-14 to 10^-13 sec, less than the time required for a bond vibration. The nature of the elusive transition state can be explored, on the other hand, using transition-state analogs, stable molecules that are chemically and structurally similar to the transition state. Such molecules should bind more strongly than a substrate and more strongly than competitive inhibitors that bear no significant similarity to the transition state. Hundreds of examples of such behavior have been reported. For example, Robert Abeles studied a series of inhibitors of proline racemase (Figure 16.7) and found that pyrrole-2-carboxylate bound to the enzyme 160 times more tightly than L-proline, the normal substrate.

Figure 16.7 • The proline racemase reaction. Pyrrole-2-carboxylate and D-1-pyrroline-2-carboxylate mimic the planar transition state of the reaction.

This analog binds so tightly because it is planar and is similar in structure to the planar transition state for the race-mization of proline. Two other examples of transition-state analogs are shown in Figure 16.8. Phosphoglycolohydroxamate binds 40,000 times more tightly to yeast aldolase than the substrate dihydroxyacetone phosphate. Even more remarkable, the 1, 6-hydrate of purine ribonucleoside has been estimated to bind to adenosine deaminase with a K_i of 3 x 10^-13 M!

Figure 16.8 • (a) Phosphoglycolohydroxamate is an analog of the enediolate transition state of the yeast aldolase reaction. (b) Purine riboside, a potent inhibitor of the calf intestinal adenosine deaminase reaction, binds to adenosine deaminase as the 1, 6-hydrate. The hydrated form of purine riboside is an analog of the proposed transition state for the reaction.

It should be noted that transition-state analogs are only approximations of the transition state itself and will never bind as tightly as would be expected for the true transition state. These analogs are, after all, stable molecules and cannot be expected to resemble a true transition state too closely.

formation of covalent bonds between enzyme and substrate. Consider the reaction:

and an enzymatic version of this reaction involving formation of a covalent intermediate:

If the enzyme-catalyzed reaction is to be faster than the uncatalyzed case, the acceptor group on the enzyme must be a better attacking group than Y and a better leaving group than X. Note that most enzymes that carry out covalent catalysis have ping-pong kinetic mechanisms.
The side chains of amino acids in proteins offer a variety of nucleophilic centers for catalysis, including amines, carboxylates, aryl and alkyl hydroxyls, imidazoles, and thiol groups. These groups readily attack electrophilic centers of substrates, forming covalently bonded enzyme-substrate intermediates. Typical electrophilic centers in substrates include phosphoryl groups, acyl groups, and glycosyl groups (Figure 16.9).

Figure 16.9 • Examples of covalent bond formation between enzyme and substrate. In each case, a nucleophilic center (X:) on an enzyme attacks an electrophilic center on a substrate.

The covalent intermediates thus formed can be attacked in a subsequent step by a water molecule or a second substrate, giving the desired product. Covalent electrophilic catalysis is also observed, but usually involves coenzyme adducts that generate electrophilic centers. Well over 100 enzymes are now known to form covalent intermediates during catalysis. Table 16.2 lists some typical examples, including that of glyceraldehyde-3-phosphate dehydrogenase, which catalyzes the reaction:

Glyceraldehyde-3-P + NAD⁺ + P_i ® 1 , 3-Bisphosphoglycerate + NADH + H⁺

Table 16.2
Enzymes That Form Covalent Intermediates
Enzymes	Reacting Group	Covalent Intermediate
1. Chymotrypsin Elastase Esterases Subtilisin Thrombin Trypsin
2. Glyceraldehyde-3-phosphate dehydrogenase Papain
3. Alkaline phosphatase Phosphoglucomutase
4. Phosphoglycerate mutase Succinyl-CoA synthetase
5. Aldolase Decarboxylases Pyridoxal phosphate-dependent enzymes

As shown in Figure 16.10, this reaction mechanism involves nucleophilic attack by -SH on the substrate glyceraldehyde-3-P to form a covalent acylcysteine (or hemithioacetal) intermediate. Hydride transfer to NAD⁺ generates a thioester intermediate. Nucleophilic attack by phosphate yields the desired mixed carboxylic-phosphoric anhydride product, 1,3-bisphosphoglycerate. Several examples of covalent catalysis will be discussed in detail in later chapters.

Figure 16.10 • Formation of a covalent intermediate in the glyceraldehyde-3-phosphate dehydrogenase reaction. Nucleophilic attack by a cysteine —SH group forms a covalent acylcysteine intermediate. Following hydride transfer to NAD⁺, nucleophilic attack by phosphate yields the product, 1 ,3-bisphosphoglycerate.

Nearly all enzyme reactions involve some degree of acid or base catalysis. There are two types of acid-base catalysis: (1) specific acid-base catalysis, in which H⁺ or OH^- accelerates the reaction, and (2) general acid-base catalysis, in which an acid or base other than H⁺ or OH^- accelerates the reaction. For ordinary solution reactions, these two cases can be distinguished on the basis of simple experiments. As shown in Figure 16.11, in specific acid or base catalysis, the buffer concentration has no effect. In general acid or base catalysis, however, the buffer may donate or accept a proton in the transition state and thus affect the rate.

Figure 16.11 • Specific and general acid - base catalysis of simple reactions in solution may be distinguished by determining the dependence of observed reaction rate constants (k_obs) on pH and buffer concentration. (a) In specific acid-base catalysis, H⁺ or OH^- concentration affects the reaction rate, k_obs is pH-dependent, but buffers (which accept or donate H⁺/OH^-) have no effect. (b) In general acid - base catalysis, in which an ionizable buffer may donate or accept a proton in the transition state, k_obs is dependent on buffer concentration.

By definition, general acid-base catalysis is catalysis in which a proton is transferred in the transition state. Consider the hydrolysis of p-nitrophenylacetate with imidazole acting as a general base (Figure 16.12). Proton transfer apparently stabilizes the transition state here. The water has been made more nucleophilic without generation of a high concentration of OH^- or without the formation of unstable, high-energy species. General acid or general base catalysis may increase reaction rates 10- to 100-fold. In an enzyme, ionizable groups on the protein provide the H⁺ transferred in the transition state. Clearly, an ionizable group will be most effective as a H⁺ transferring agent at or near its pK_a. Because the pK_a of the histidine side chain is near 7, histidine is often the most effective general acid or base. Descriptions of several cases of general acid - base catalysis in typical enzymes follow.

Figure 16.12 • Catalysis of p-nitrophenylacetate hydrolysis by imidazole—an example of general base catalysis. Proton transfer to imidazole in the transition state facilitates hydroxyl attack on the substrate carbonyl carbon.

Many enzymes require metal ions for maximal activity. If the enzyme binds the metal very tightly or requires the metal ion to maintain its stable, native state, it is referred to as a metalloenzyme . Enzymes that bind metal ions more weakly, perhaps only during the catalytic cycle, are referred to as metal activated. One role for metals in metal-activated enzymes and metalloenzymes is to act as electrophilic catalysts, stabilizing the increased electron density or negative charge that can develop during reactions. Among the enzymes that function in this manner (Figure 16.13) is liver alcohol dehydrogenase. Another potential function of metal ions is to provide a powerful nucleophile at neutral pH. Coordination to a metal ion can increase the acidity of a nucleophile with an ionizable proton:

The reactivity of the coordinated, deprotonated nucleophile is typically intermediate between that of the un-ionized and ionized forms of the nucleophile. Carboxypeptidase (Chapter 5) contains an active site Zn²⁺, which facilitates deprotonation of a water molecule in this manner.

Figure 16.13 • Liver alcohol dehydrogenase catalyzes the transfer of a hydride ion ( H: ^-) from NADH to acetaldehyde (CH₃CHO), forming ethanol (CH₃CH₂OH). An active-site zinc ion stabilizes negative charge development on the oxygen atom of acetaldehyde, leading to an induced partial positive charge on the carbonyl C atom. Transfer of the negatively charged hydride ion to this carbon forms ethanol.

Chemical reactions go faster when the reactants are in proximity, that is, near each other. In solution or in the gas phase, this means that increasing the concentrations of reacting molecules, which raises the number of collisions, causes higher rates of reaction. Enzymes, which have specific binding sites for particular reacting molecules, essentially take the reactants out of dilute solution and hold them close to each other. This proximity of reactants is said to raise the “effective” concentration over that of the substrates in solution, and leads to an increased reaction rate. In order to measure proximity effects in enzyme reactions, enzymologists have turned to model studies comparing intermolecular reaction rates with corresponding or similar intramolecular reaction rates. A typical case is the imidazole-catalyzed hydrolysis of p-nitrophenylacetate (Figure 16.14a). Under certain conditions the rate constant for this bimolecular reaction is 35 M^-1 min^-1. By comparison, the first-order rate constant for the analogous but intramolecular reaction shown in Figure 16.14b is 839 min^-1. The ratio of these two rate constants

has the units of concentration and can be thought of as an effective concentration of imidazole in the intramolecular reaction. Put another way, a concentration of imidazole of 23.9 M would be required in the intermolecular reaction to make it proceed as fast as the intramolecular reaction.

Figure 16.14 • An example of proximity effects in catalysis. (a) The imidazole-catalyzed hydrolysis of p-nitrophenylacetate is slow, but the corresponding intramolecular reaction is 24-fold faster (assuming [ imidazole ] = 1 M in [a]).

There is more to this story, however. Enzymes not only bring substrates and catalytic groups close together, they orient them in a manner suitable for catalysis as well. Comparison of the rates of reaction of the molecules shown in Figure 16.15 makes it clear that the bulky methyl groups force an orientation on the alkyl carboxylate and the aromatic hydroxyl groups that makes them approximately 250 billion times more likely to react. Enzymes function similarly by placing catalytically functional groups (from the protein side chains or from another substrate) in the proper position for reaction.

Figure 16.15 • Orientation effects in intramolecular reactions can be dramatic. Steric crowding by methyl groups provides a rate acceleration of 2.5 x 10¹¹ for the lower reaction compared to the upper reaction. (Adapted from Milstien , S., and Cohen, L. A., 1972. Stereopopulation control I. Rate enhancements in the lactonization of o- hydroxyhydrocinnamic acid. Journal of the American Chemical Society 94:9158 - 9165.)

Clearly, proximity and orientation play a role in enzyme catalysis, but there is a problem with each of the above comparisons. In both cases, it is impossible to separate true proximity and orientation effects from the effects of entropy loss when molecules are brought together (described the Section 16.4). The actual rate accelerations afforded by proximity and orientation effects in Figures 16.14 and 16.15, respectively, are much smaller than the values given in these figures. Simple theories based on probability and nearest-neighbor models, for example, predict that proximity effects may actually provide rate increases of only 5- to 10-fold. For any real case of enzymatic catalysis, it is nonetheless important to remember that proximity and orientation effects are significant.

The balance of this chapter will be devoted to several classic and representative enzyme mechanisms. These particular cases are well understood, because the three-dimensional structures of the enzymes and the bound substrates are known at atomic resolution, and because great efforts have been devoted to kinetic and mechanistic studies. They are important because they represent reaction types that appear again and again in living systems, and because they demonstrate many of the catalytic principles cited above. Enzymes are the catalytic machines that sustain life, and what follows is an intimate look at the inner workings of the machinery.

Serine proteases are a class of proteolytic enzymes whose catalytic mechanism is based on an active-site serine residue. Serine proteases are one of the best-characterized families of enzymes. This family includes trypsin, chymotrypsin, elastase, thrombin, subtilisin, plasmin, tissue plasminogen activator, and other related enzymes. The first three of these are digestive enzymes and are synthesized in the pancreas and secreted into the digestive tract as inactive proenzymes, or zymogens. Within the digestive tract, the zymogen is converted into the active enzyme form by cleaving off a portion of the peptide chain. Thrombin is a crucial enzyme in the blood-clotting cascade, subtilisin is a bacterial protease, and plasmin breaks down the fibrin polymers of blood clots. Tissue plasminogen activator (TPA) specifically cleaves the proenzyme plasminogen, yielding plasmin. Owing to its ability to stimulate breakdown of blood clots, TPA can minimize the harmful consequences of a heart attack, if administered to a patient within 30 minutes of onset. Finally, although not itself a protease, acetylcholinesterase is a serine esterase and is related mechanistically to the serine proteases. It degrades the neurotransmitter acetylcholine in the synaptic cleft between neurons.

Trypsin, chymotrypsin, and elastase all carry out the same reaction — the cleavage of a peptide chain — and although their structures and mechanisms are quite similar, they display very different specificities. Trypsin cleaves peptides on the carbonyl side of the basic amino acids, arginine or lysine (see Table 5.6). Chymotrypsin prefers to cleave on the carbonyl side of aromatic residues, such as phenylalanine and tyrosine. Elastase is not as specific as the other two; it mainly cleaves peptides on the carbonyl side of small, neutral residues. These three enzymes all possess molecular weights in the range of 25,000, and all have similar sequences (Figure 16.16) and three-dimensional structures.

Figure 16.16 • Comparison of the amino acid sequences of chymotrypsinogen, trypsinogen, and elastase. Each circle represents one amino acid. Numbering is based on the sequence of chymotrypsinogen. Filled circles indicate residues that are identical in all three proteins. Disulfide bonds are indicated in yellow. The positions of the three catalytically important active-site residues (His⁵⁷, Asp¹⁰², and Ser¹⁹⁵) are indicated.

The molecule is ellipsoidal in shape and contains an a-helix at the C-terminal end (residues 230 to 245) and several b-sheet domains. Most of the aromatic and hydrophobic residues are buried in the interior of the protein, and most of the charged or hydrophilic residues are on the surface. Three polar residues — His⁵⁷, Asp¹⁰², and Ser¹⁹⁵ — form what is known as a catalytic triad at the active site (Figure 16.18).

These three residues are conserved in trypsin and elastase as well. The active site in this case is actually a depression on the surface of the enzyme, with a small pocket that the enzyme uses to identify the residue for which it is specific (Figure 16.19). Chymotrypsin, for example, has a pocket surrounded by hydrophobic residues and large enough to accommodate an aromatic side chain. The pocket in trypsin has a negative charge (Asp¹⁸⁹) at its bottom, facilitating the binding of positively charged arginine and lysine residues. Elastase, on the other hand, has a shallow pocket with bulky threonine and valine residues at the opening. Only small, nonbulky residues can be accommodated in its pocket. The backbone of the peptide substrate is hydrogen-bonded in antiparallel fashion to residues 215 to 219 and bent so that the peptide bond to be cleaved is bound close to His⁵⁷ and Ser¹⁹⁵.

Figure 16.19 • The substrate-binding pockets of trypsin, chymotrypsin, and elastase. (Irving Geis )

Much of what is known about the chymotrypsin mechanism is based on studies of the hydrolysis of artificial substrates — simple organic esters, such as p-nitrophenylacetate, and methyl esters of amino acid analogs, such as formylphenylalanine methyl ester and acetylphenylalanine methyl ester (Figure 16.20).

Figure 16.20 • Artificial substrates used in studies of the mechanism of chymotrypsin .

p-Nitrophenylacetate is an especially useful model substrate, because the nitrophenolate product is easily observed, owing to its strong absorbance at 400 nm. When large amounts of chymotrypsin are used in kinetic studies with this substrate, a rapid initial burst of p-nitrophenolate is observed (in an amount approximately equal to the enzyme concentration), followed by a much slower, linear rate of nitrophenolate release (Figure 16.21). Observation of a burst, followed by slower, steady-state product release, is strong evidence for a multistep mechanism, with a fast first step and a slower second step.

Figure 16.21 • Burst kinetics observed in the chymotrypsin reaction. A burst of nitrophenolate production is followed by a slower, steady-state release. After an initial lag period, acetate release is also observed. This kinetic pattern is consistent with rapid formation of an acyl-enzyme intermediate (and the burst of nitrophenolate ). The slower, steady-state release of products corresponds to rate-limiting breakdown of the acyl-enzyme intermediate.

In the chymotrypsin mechanism, the nitrophenylacetate combines with the enzyme to form an ES complex. This is followed by a rapid second step in which an acyl-enzyme intermediate is formed, with the acetyl group covalently bound to the very reactive Ser¹⁹⁵. The nitrophenyl moiety is released as nitrophenolate (Figure 16.22), accounting for the burst of nitrophenolate product. Attack of a water molecule on the acyl-enzyme intermediate yields acetate as the second product in a subsequent, slower step. The enzyme is now free to bind another molecule of p-nitrophenylacetate, and the p-nitrophenolate product produced at this point corresponds to the slower, steady-state formation of product in the upper right portion of Figure 16.21. In this mechanism, the release of acetate is the rate-limiting step, and accounts for the observation of burst kinetics — the pattern shown in Figure 16.21.

Figure 16.22 • Rapid formation of the acyl-enzyme intermediate is followed by slower product release.

Serine proteases like chymotrypsin are susceptible to inhibition by organic fluorophosphates, such as diisopropylfluorophosphate (DIFP, Figure 16.23). DIFP reacts rapidly with active-site serine residues, such as Ser¹⁹⁵ of chymotrypsin and the other serine proteases (but not with any of the other serines in these proteins), to form a DIP-enzyme. This covalent enzyme-inhibitor complex is extremely stable, and chymotrypsin is thus permanently inactivated by DIFP.

Figure 16.23 • Diisopropylfluorophosphate (DIFP) reacts with active-site serine residues of serine proteases (and esterases), causing permanent inactivation.

A likely mechanism for peptide hydrolysis is shown in Figure 16.24. As the backbone of the substrate peptide binds adjacent to the catalytic triad, the specific side chain fits into its pocket. Asp¹⁰² of the catalytic triad positions His⁵⁷ and immobilizes it through a hydrogen bond as shown. In the first step of the reaction, His⁵⁷ acts as a general base to withdraw a proton from Ser¹⁹⁵, facilitating nucleophilic attack by Ser¹⁹⁵ on the carbonyl carbon of the peptide bond to be cleaved. This is probably a concerted step, because proton transfer prior to Ser¹⁹⁵ attack on the acyl carbon would leave a relatively unstable negative charge on the serine oxygen. In the next step, donation of a proton from His⁵⁷ to the peptide’s amide nitrogen creates a protonated amine on the covalent, tetrahedral intermediate, facilitating the subsequent bond breaking and dissociation of the amine product. The negative charge on the peptide oxygen is unstable; the tetrahedral intermediate is short-lived and rapidly breaks down to expel the amine product. The acyl-enzyme intermediate that results is reasonably stable; it can even be isolated using substrate analogs for which further reaction cannot occur. With normal peptide substrates, however, subsequent nucleophilic attack at the carbonyl carbon by water generates another transient tetrahedral intermediate (Figure 16.24). His⁵⁷ acts as a general base in this step, accepting a proton from the attacking water molecule. The subsequent collapse of the tetrahedral intermediate is assisted by proton donation from His⁵⁷ to the serine oxygen in a concerted manner. Deprotonation of the carboxyl group and its departure from the active site complete the reaction as shown.

Until recently, the catalytic role of Asp¹⁰² in trypsin and the other serine proteases had been surmised on the basis of its proximity to His⁵⁷ in structures obtained from X-ray diffraction studies, but it had never been demonstrated with certainty in physical or chemical studies. As can be seen in Figure 16.17, Asp¹⁰² is buried at the active site and is normally inaccessible to chemical modifying reagents. In 1987, however, Charles Craik , William Rutter , and their colleagues used site-directed mutagenesis (see Chapter 13) to prepare a mutant trypsin with an asparagine in place of Asp¹⁰². This mutant trypsin possessed a hydrolytic activity with ester substrates only 1/10,000 that of native trypsin , demonstrating that Asp¹⁰² is indeed essential for catalysis and that its ability to immobilize and orient His⁵⁷ is crucial to the function of the catalytic triad.

Mammals, fungi, and higher plants produce a family of proteolytic enzymes known as aspartic proteases. These enzymes are active at acidic (or sometimes neutral) pH, and each possesses two aspartic acid residues at the active site. Aspartic proteases carry out a variety of functions (Table 16.3), including digestion (pepsin and chymosin), lysosomal protein degradation (cathepsin D and E), and regulation of blood pressure (renin is an aspartic protease involved in the production of angiotensin, a hormone that stimulates smooth muscle contraction and reduces excretion of salts and fluid). The aspartic proteases display a variety of substrate specificities, but normally they are most active in the cleavage of peptide bonds between two hydrophobic amino acid residues. The preferred substrates of pepsin, for example, contain aromatic residues on both sides of the peptide bond to be cleaved.

Most aspartic proteases are composed of 323 to 340 amino acid residues, with molecular weights near 35,000. Aspartic protease polypeptides consist of two homologous domains that fold to produce a tertiary structure composed of two similar lobes, with approximate twofold symmetry (Figure 16.25). Each of these lobes or domains consists of two b-sheets and two short a-helices. The two domains are bridged and connected by a six-stranded, antiparallel b-sheet. The active site is a deep and extended cleft, formed by the two juxtaposed domains and large enough to accommodate about seven amino acid residues. The two catalytic aspartate residues, residues 32 and 215 in porcine pepsin, for example, are located deep in the center of the active site cleft. The N-terminal domain forms a “flap” that extends over the active site, which may help to immobilize the substrate in the active site.

Figure 16.25 • Structures of (a) HIV-1 protease, a dimer , and (b) pepsin (a monomer). Pepsin’s N-terminal half is shown in red; C-terminal half is shown in blue.

On the basis, in part, of comparisons with chymotrypsin, trypsin, and the other serine proteases, it was hypothesized that aspartic proteases might function by formation of covalent enzyme-substrate intermediates involving the active-site aspartate residues. Two possibilities were proposed: an acyl-enzyme intermediate involving an acid anhydride bond and an amino-enzyme intermediate involving an amide (peptide) bond (Figure 16.26). All attempts to trap or isolate a covalent intermediate failed, and a mechanism (see following paragraph) favoring noncovalent enzyme-substrate intermediates and general acid-general base catalysis is now favored for aspartic proteases.

Figure 16.26 • Acyl-enzyme and amino-enzyme intermediates originally proposed for aspartic proteases were modeled after the acyl-enzyme intermediate of the serine proteases.

A crucial datum supporting the general acid-general base model is the pH dependence of protease activity (see Critical Developments in Biochemistry: The pH Dependence of Aspartic Proteases and HIV-1 Protease, page 525). Enzymologists hypothesize that the aspartate carboxyl groups function alternately as general acid and general base. This model requires that one of the aspartate carboxyls be protonated and one be deprotonated when substrate binds. X-ray diffraction data on aspartic proteases show that the active-site structure in the vicinity of the two aspartates is highly symmetric. The two aspartates appear to act as a “catalytic dyad” (analogous to the catalytic triad of the serine proteases). The dyad proton may thus be covalently bound to either of the aspartate groups in the free enzyme or in the enzyme-substrate complex. Thus, in pepsin, for example, Asp³² may be deprotonated while Asp²¹⁵ is protonated, or vice versa.
In the most widely accepted mechanism (Figure 16.27), substrate binding is followed by a step in which two concerted proton transfers facilitate nucleophilic attack on the carbonyl carbon of the substrate by water. In the mechanism shown, Asp³² acts as a general base, accepting a proton from an active-site water molecule, whereas Asp²¹⁵ acts a general acid, donating a proton to the oxygen of the peptide carbonyl group. By virtue of these two proton transfers, nucleophilic attack occurs without explicit formation of hydroxide ion at the active site. The resulting intermediate is termed an amide dihydrate. Note that the protonation states of the two aspartate residues are now opposite to those in the free enzyme (Figure 16.27).
Breakdown of the amide dihydrate occurs by a mechanism similar to its formation. The ionized aspartate carboxyl (Asp³² in Figure 16.27) acts as a general base to accept a proton from one of the hydroxyl groups of the amide dihydrate , while the protonated carboxyl of the other asparate (Asp²¹⁵ in this case) simultaneously acts as a general acid to donate a proton to the nitrogen atom of one of the departing peptide products.

Figure 16.27 • A mechanism for the aspartic proteases. In the first step, two concerted proton transfers facilitate nucleophilic attack of water on the substrate carbonyl carbon. In the third step, one aspartate residue (Asp³² in pepsin) accepts a proton from one of the hydroxyl groups of the amine dihydrate , and the other aspartate (Asp²¹⁵) donates a proton to the nitrogen of the departing amine.

Recent research on acquired immune deficiency syndrome (AIDS) and its causative viral agent, the human immunodeficiency virus (HIV-1), has brought a new aspartic protease to light. HIV-1 protease cleaves the polyprotein products of the HIV-1 genome, producing several proteins necessary for viral growth and cellular infection. HIV-1 protease cleaves several different peptide linkages in the HIV-1 polyproteins, including those shown in Figure 16.28. For example, the protease cleaves between the Tyr and Pro residues of the sequence Ser- Gln-Asn-Tyr-Pro-Ile-Val, which joins the p17 and p24 HIV-1 proteins.

Figure 16.28 • HIV mRNA provides the genetic information for synthesis of a polyprotein . Proteolytic cleavage of this polyprotein by HIV protease produces the individual proteins required for viral growth and cellular infection.

The HIV-1 protease is a remarkable viral imitation of mammalian aspartic proteases: It is a dimer of identical subunits that mimics the two-lobed monomeric structure of pepsin and other aspartic proteases. The HIV-1 protease subunits are 99-residue polypeptides that are homologous with the individual domains of the monomeric proteases. Structures determined by X-ray diffraction studies reveal that the active site of HIV-1 protease is formed at the interface of the homodimer and consists of two aspartate residues, designated Asp²⁵ and Asp^25', one contributed by each subunit (Figure 16.29). In the homodimer, the active site is covered by two identical “flaps,” one from each subunit, in contrast to the monomeric aspartic proteases, which possess only a single active-site flap.

Figure 16.29 • (left) HIV-1 protease complexed with the inhibitor Crixivan ^Ò(red) made by Merck. The flaps (residues 46 - 55 from each subunit) covering the active site are shown in green and the active site aspartate residues involved in catalysis are shown in white. (right) The close-up of the active site shows the interaction of Crixivan ^Ò with the carboxyl groups of the essential aspartate residues.

Enzyme kinetic measurements by Thomas Meek and his collaborators at SmithKline Beecham Pharmaceuticals have shown that the mechanism of

HIV-1 protease is very similar to those of other aspartic proteases (Figure 16.30). Two concerted proton transfers by the aspartate carboxyls facilitate nucleophilic attack by water on the carbonyl carbon of the peptide substrate. If the protease-substrate complex is incubated with H₂¹⁸O, incorporation of ¹⁸O into the peptide carbonyl group can be measured. Thus, not only is the formation of the amide dihydrate reversible, and the two hydroxyl groups of the dihydrate equivalent, but the protonation states of the active-site carboxyl groups must interchange (Figure 16.30). The simplest model would involve direct proton exchange across the Asp-Asp dyad as shown. The symmetrical nature of the active-site aspartyl groups is consistent with the idea of facile exchange of the proton between the two Asp residues.

Figure 16.30 • A mechanism for the incorporation of ¹⁸O from H₂¹⁸O into peptide substrates in the HIV protease reaction. (Adapted from Hyland, L., et al., 1991. Biochemistry 30:8441 - 8453.)

The observation that ¹⁸O can accumulate in the substrate from H₂¹⁸O also implies that formation and reversal of the amide dihydrate must be faster than its reakdown to form product. Kinetic studies by Meek and his coworkers are consistent with a model in which breakdown of the dihydrate intermediate is the rate-determining step for the protease reaction. These studies also show that the transition state of this step involves two proton transfers — with one aspartate acting as a general acid to donate a proton to the departing proline and the other aspartate acting as a general base to abstract a proton from one of the hydroxyl groups, facilitating collapse of the dihydrate to a carboxyl group. Note that the events of the breakdown to form product are simply the reversal of the steps that formed the intermediate.

Lysozyme is an enzyme that hydrolyzes polysaccharide chains. It ruptures certain bacterial cells by cleaving the polysaccharide chains that make up their cell wall. Lysozyme is found in many body fluids, but the most thoroughly studied form is from hen egg whites. The Russian scientist P. Laschtchenko first described the bacteriolytic properties of hen egg white lysozyme in 1909. In 1922, Alexander Fleming, the London bacteriologist who later discovered penicillin, gave the name lysozyme to the agent in mucus and tears that destroyed certain bacteria, because it was an enzyme that caused bacterial lysis .
As seen in Chapter 9, bacterial cells are surrounded by a rigid, strong wall of peptidoglycan, a copolymer of two sugar units, N-acetylmuramic acid (NAM) and N-acetylglucosamine (NAG). Both of these sugars are N-acetylated analogs of glucosamine, and in bacterial cell wall polysaccharides, they are joined in b (1 ® 4) glycosidic linkages (Figure 16.31). Lysozyme hydrolyzes the glycosidic bond between C-1 of NAM and C-4 of NAG, as shown in Figure 16.31, but does not act on the b(1 → 4) linkages between NAG and NAM.

Lysozyme is a small globular protein composed of 129 amino acids (14 kD ) in a single polypeptide chain. It has eight cysteine residues linked in four disulfide bonds. The structure of this very stable protein was determined by X-ray crystallographic methods in 1965 by David Phillips (Figure 16.32). Although X-ray structures had previously been reported for proteins (hemoglobin and myoglobin ), lysozyme was the first enzyme structure to be solved by crystallographic (or any other) methods. Although the location of the active site was not obvious from the X-ray structure of the protein alone, X-ray studies of lysozyme-inhibitor complexes soon revealed the location and nature of the active site. Since it is an enzyme, lysozyme cannot form stable ES complexes for structural studies, because the substrate is rapidly transformed into products.

Figure 16.32 • The structure of lysozyme. Glu³⁵ and Asp⁵² are shown in white.

On the other hand, several substrate analogs have proven to be good competitive inhibitors of lysozyme that can form complexes with the enzyme stable enough to be characterized by X-ray crystallography and other physical techniques. One of the best is a trimer of N-acetylglucosamine, (NAG)₃ (Figure 16.33), which is hydrolyzed by lysozyme at a rate only 1/60,000 that of the native substrate (Table 16.4).

Figure 16.33 • (NAG)₃, a substrate analog, forms stable complexes with lysozyme .

(NAG)₃ binds at the enzyme active site by forming five hydrogen bonds with residues located in one-half of a depression or crevice that spans the surface of the enzyme (Figure 16.34).

Figure 16.34 • The lysozyme -enzyme-substrate complex. (Photo courtesy of John Rupley , University of Arizona )

The few hydrophobic residues that exist on the surface of lysozyme are located in this depression, and they may participate in hydrophobic and van der Waals interactions with (NAG)₃, as well as the normal substrate. The absence of charged groups on (NAG)₃ precludes the involvement of electrostatic interactions with the enzyme. Comparisons of the X-ray structures of the native lysozyme and the lysozyme -(NAG)₃ complex reveal that several amino acid residues at the active site move slightly upon inhibitor binding, including Trp⁶², which moves about 0.75 Å to form a hydrogen bond with a hydroxymethyl group (Figure 16.35).

Figure 16.35 • Enzyme-substrate interactions at the six sugar residue-binding subsites of the lysozyme active site. (© Irving Geis )

Model Studies Reveal a Strain-Induced Destabilization of a Bound Substrate on Lysozyme

One of the premises of lysozyme models is that the native substrate would occupy the rest of the crevice or depression running across the surface of the enzyme, because there is room to fit three more sugar residues into the crevice and because the hexamer (NAG)₆ is in fact a good substrate for lysozyme (Table 16.4). The model building studies refer to the six sugar residue-binding subsites in the crevice with the letters A through F, with A, B, and C representing the part of the crevice occupied by the (NAG)_{3inhibitor (Figure
16.35)}. Modeling studies clearly show that NAG residues fit nicely into subsites A, B, C, E, and F of the crevice, but that fitting a residue of the (NAG)₆ hexamer into site D requires a substantial distortion of the sugar (out of its preferred chair conformation) to prevent steric crowding and overlap between atoms C-6 and O-6 of the sugar at the D site and Ile⁹⁸ of the enzyme. This distorted sugar residue is adjacent to the glycosidic bond to be cleaved (between sites D and E), and the inference is made that this distortion or strain brings the substrate closer to the transition state for hydrolysis. This is a good example of strain-induced destabilization of an otherwise favorably binding substrate (Section 16.4). Thus, the overall binding interaction of the rest of the sugar substrate would be favorable (DG<0), but distortion of the ring at the D site uses some of this binding energy to raise the substrate closer to the transition state for hydrolysis, an example of stabilization of a transition state (relative to the simple enzyme-substrate complex). As noted in Section 16.4, distortion is one of the molecular mechanisms that can lead to such transition-state stabilization.

Figure 16.36 • The C₁-O bond, not the O-C₄ bond, is cleaved in the lysozyme reaction. ¹⁸O from H₂¹⁸O is thus incorporated at the C₁ position.

Studies using ¹⁸O-enriched water showed that the C₁—O bond is cleaved on the substrate between the D and E sites. Hydrolysis under these conditions incorporates ¹⁸O into the C₁ position of the sugar at the D site, not into the oxygen at C₄ at the E site (Figure 16.36).

Model building studies place the cleaved bond approximately between protein residues Glu³⁵ and Asp⁵². Glu³⁵ is in a nonpolar or hydrophobic region of the protein, whereas Asp⁵² is located in a much more polar environment. Glu³⁵ is protonated , but Asp⁵² is ionized (Figure 16.37). Glu³⁵ may thus act as a general acid, donating a proton to the oxygen atom of the glycosidic bond and accelerating the reaction. Asp⁵², on the other hand, probably stabilizes the carbonium ion generated at the D site upon bond cleavage. Formation of the carbonium ion may also be enhanced by the strain on the ring at the D site. Following bond cleavage, the product formed at the E site diffuses away, and the carbonium ion intermediate can then react with H₂O from the solution. Glu³⁵ can now act as a general base, accepting a proton from the attacking water. The tetramer of NAG thus formed at sites A through D can now be dissociated from the enzyme.
On the basis of the above, the rate acceleration afforded by lysozyme appears to be due to (a) general acid catalysis by Glu³⁵; (b) distortion of the sugar ring at the D site, which may stabilize the carbonium ion (and the transition state); and (c) electrostatic stabilization of the carbonium ion by nearby Asp⁵². The overall k_catfor lysozyme is about 0.5/sec, which is quite slow (Table 14.4) compared with that for other enzymes. On the other hand, the destruction of a bacterial cell wall may only require hydrolysis of a few polysaccharide chains. The high osmotic pressure of the cell ensures that cell rupture will follow rapidly. Thus, lysozyme can accomplish cell lysis without a particularly high k_cat.

Table 16.1
A Comparison of Enzyme-Catalyzed Reactions and Their Uncatalyzed Counterparts
Reaction	Enzyme	Uncatalyzed Rate, v_u (sec^-1)	Catalyzed Rate, v_e(sec^-1)	v_e/v_u
	Alkaline phosphatase	1 x 10^-15	14	1.4 x 10¹⁶
	Urease	3 x 10^-10	3 x 10⁴	1 x 10¹⁴
	Chymotrypsin	1 x 10^-10	1 x 10²	1 x 10¹²
Glycogen + P_i → Glycogen + Glucose-1-P (n) (n - 1)	Glycogen phosphorylase	<5 x 10^-15	1.6 x 10^-3	>3.2 x 10¹¹
Glucose + ATP → Glucose-6-P + ADP	Hexokinase	<1 x 10^-13	1.3 x 10^-3	<1.3 x 10¹⁰
	Alcohol dehydrogenase	<6 x 10^-12	2.7 x 10^-5	>4.5 x 10⁶
CO₂ + H₂O → HCO₃^- + H⁺	Carbonic anhydrase	10^-2	10⁵	1 x 10⁷
Creatine + ATP → Cr-P + ADP	Creatine kinase	<3 x 10^-9	4 x 10^-5	>1.33 x 10⁴
Adapted from Koshland , D., 1956. Journal of Cellular Comparative Physiology, Supp. 1, 47:217.

Table 16.3
Some Representative Aspartic Proteases
Name	Source	Function
Pepsin^*	Animal stomach	Digestion of dietary protein
Chymosin ^†	Animal stomach	Digestion of dietary protein
Cathepsin D	Spleen, liver, and many other animal tissues	Lysosomal digestion of proteins
Renin ^‡	Kidney	Conversion of angiotensinogen to angiotensin I; regulation of blood pressure
HIV-protease^§	AIDS virus	Processing of AIDS virus proteins
^*The second enzyme to be crystallized (by John Northrup in 1930). Even more than urease before it, pepsin study by Northrup established that enzyme activity comes from proteins. ^†Also known as rennin, it is the major pepsinlike enzyme in gastric juice of fetal and newborn animals. ^‡A drop in blood pressure causes release of renin from the kidneys, which converts more angiotensinogen to angiotensin. ^§A dimer of identical monomers, homologous to pepsin.

Table 16.4
Hydrolysis Rate Constants for Model Oligosaccharides with Lysozyme
Oligosaccharide	Rate Constant, k_cat(s^-1)
(NAG-NAM)₃	0.5
(NAG)₆	0.25
(NAG)₅	0.033
(NAG)₄	7 x 10^-5
(NAG)₃	8 x 10^-6
(NAG)₂	2.5 x 10^-8