Rare indeed is the organic chemical reaction that proceeds unaided at a rate sufficient to support living systems. Enzymes have extraordinary catalytic power, often far greater than that of synthetic or inorganic catalysts. They have a high degree of specificity for their substrates, they substantially accelerate chemical reactions, and they function in aqueous solutions under very mild conditions of temperature and pH. Few nonbiological catalysts have all these properties.

Enzymes are central to every cellular process. Acting in organized sequences, they catalyze the hundreds of stepwise reactions that degrade nutrient molecules, conserve and transform chemical energy, make biological macromolecules from simple precursors, and carry out the various processes of DNA and RNA metabolism.

In molecular biology, the study of enzymes has immense practical importance. In some diseases, especially hereditary genetic disorders related to DNA or RNA metabolism, there may be a deficiency or even a total absence of one or more enzymes. Other disease conditions may be caused by excessive activity of an enzyme. Many medicines act through interactions with enzymes. Furthermore, researchers can isolate and harness enzyme functions to suit their purposes in the laboratory. The set of methods collectively described as “biotechnology” (many of which are described in Chapter 7 and elsewhere throughout this book) is made possible by our understanding of the enzymes of DNA and RNA metabolism.

With the exception of a small group of catalytic RNA molecules (described in Chapter 16), all enzymes are proteins. Their catalytic activity depends on the integrity of their native protein conformation. Enzymes, like other proteins, have molecular weights ranging from about 12,000 to more than 1 million. Some enzymes require for their activity no additional chemical groups other than their amino acid residues. Others require an additional chemical component called a cofactor, either one or more inorganic metal ions (Table 5-3) or a complex organic or metallo-organic molecule called a coenzyme, which acts as a transient carrier of specific functional groups (Table 5-4). Most coenzymes are derived from vitamins, organic nutrients required in small amounts in the human diet. Some enzymes require both a coenzyme and one or more metal ion cofactors for activity. A coenzyme or inorganic cofactor that is very tightly or even covalently bound to the enzyme protein is known as a prosthetic group. A complete, catalytically active enzyme, with its bound coenzyme and/or inorganic cofactor, is referred to as a holoenzyme. The protein part of such an enzyme is called the apoenzyme or apoprotein.

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An enzyme catalyzes a reaction by providing a specific environment in which the reaction can occur more rapidly. Here are some key principles of enzyme-catalyzed reactions:

The enormous and highly selective rate enhancements achieved by enzymes can be explained by the many types of covalent and noncovalent interactions between enzyme and substrate. Chemical reactions of many types may take place between substrates and the functional groups (specific amino acid side chains, metal ions, and coenzymes) on enzymes. The particular reactions that occur depend on the requirements of the overall reaction to be catalyzed. An enzyme’s catalytic functional groups may form a transient covalent bond with the substrate and activate it for reaction. Or a group may be transiently transferred from the substrate to the enzyme. The most common type of group transfer involves the transfer of protons between ionizable amino acid side chains in the active site and groups on the substrate molecule, a process called general acid and base catalysis (Figure 5-9). In the enzymes important to molecular biology, phosphoryl group transfers are also common. In many cases, these group transfer reactions occur only in the enzyme active site. The capacity to facilitate multiple interactions and transfers of this type, sometimes all at once, is one of the factors contributing to the rate enhancements provided by enzymes.

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Covalent interactions are only part of the story, however. Much of the energy required to increase the reaction rate is derived from weak, noncovalent interactions between substrate and enzyme, including hydrogen bonds and hydrophobic and ionic interactions. The formation of each weak interaction is accompanied by the release of a small amount of free energy that stabilizes the interaction. The energy derived from enzyme-substrate interaction is called binding energy, ΔG_B. Its significance extends beyond a simple stabilization of the enzyme-substrate interaction. Binding energy is a major source of the free energy used by enzymes to increase the rates of reactions.

In the context of nucleic acid metabolism, one type of weak interaction merits special mention. Ionic interactions can include interactions between bound metals (such as Mg²⁺, Mn²⁺, and Fe²⁺ or Fe³⁺ ions) and substrates. About one-third of all enzymes utilize metals in their catalytic mechanisms, and that proportion is much higher for the enzymes that act on DNA and RNA. For example, the active sites of DNA and RNA polymerases universally feature two metal ions, usually two Mg²⁺ions, that help orient substrates and facilitate the overall reaction in multiple ways (see Figure 5-9). Mg²⁺ ions play key roles at the active sites of a wide range of enzymes discussed in later chapters.

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A simple enzyme reaction might be written like this:

where E, S, and P represent the enzyme, substrate, and product, and ES and EP are transient complexes of the enzyme with the substrate and with the product.

To understand catalysis, we must recall the important distinction between reaction equilibria and reaction rates. The equilibrium between S and P reflects the difference in the free energies of their ground states. Any reaction, such as S ⇌ P, can be described by a reaction coordinate diagram, a picture of the energy changes during the reaction (Figure 5-10a). Energy in biological systems is described in terms of free energy, G (see Chapter 3). In the diagram, the free energy of the system is plotted against the progress of the reaction (the reaction coordinate). In this example, the free energy of the ground state of P is lower than that of S, so the biochemical standard free-energy change, or ΔG′°, for the reaction is negative and the equilibrium favors P.

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A favorable equilibrium does not mean that the S → P conversion will occur at a fast or even detectable rate. An unfavorable equilibrium does not mean that the reaction will be slow. Instead, the rate of a reaction depends on the height of the energy hill that separates the product from the substrate. At the top of this hill lies the transition state (denoted by ‡ in Figure 5-10a). The transition state is not a stable species but a transient moment when the alteration in the substrate has reached a point corresponding to the highest energy in the reaction coordinate diagram. The difference between the energy levels of the ground state and the transition state is the activation energy, ΔG^‡. A higher activation energy corresponds to a slower reaction.

The function of a catalyst is to increase the rate of a reaction. Catalysts do not affect reaction equilibria, and enzymes are no exception. The bidirectional arrows in Equation 5-6 make this point: any enzyme that catalyzes the reaction S → P also catalyzes the reaction P → S. The role of enzymes is to accelerate the interconversion of S and P. The enzyme is not used up in the process, and the equilibrium point is unaffected. However, the reaction reaches equilibrium much faster when the appropriate enzyme is present, because the rate of the reaction is increased. Enzymes increase reaction rates by lowering the activation energy of the reaction. To achieve this, enzymes utilize noncovalent and covalent interactions in somewhat different ways.

Two fundamental and interrelated principles provide a general explanation for how enzymes use noncovalent binding energy to accelerate a reaction:

When the enzyme active site is complementary to the reaction transition state, some of the noncovalent interactions between enzyme and substrate occur only in the transition state. The free energy (binding energy) released by the formation of these interactions partially offsets the energy required to reach the top of the energy hill. The summation of the unfavorable (positive) activation energy ΔG^‡ and the favorable (negative) binding energy ΔG_B results in a lower net activation energy (see Figure 5-10a). Even on the enzyme, the transition state is not a stable species but a brief point in time that the substrate spends atop an energy hill. The enzyme-catalyzed reaction is much faster than the uncatalyzed process, however, because the hill is much smaller. The groups on the substrate that are involved in the weak interactions between the enzyme and transition state can be at some distance from the substrate bonds that are broken or changed. The weak interactions formed only in the transition state are those that make the primary contribution to catalysis (see Figure 5-10b).

Covalent interactions can accelerate some enzyme-catalyzed reactions by creating a different, lower-energy reaction pathway. When the reaction occurs in solution in the absence of the enzyme, the reaction takes a particular (and usually very slow) path. In the enzyme-catalyzed reaction, if a group on the enzyme is transferred to or from the substrate during the reaction, the reaction path is altered. The new pathway results in acceleration of the reaction only if its overall activation energy is lower than that of the uncatalyzed reaction.

The oldest approach to understanding enzyme mechanisms, and the one that remains most important, is to determine the rate of a reaction and how it changes in response to changes in experimental parameters, a discipline known as enzyme kinetics. We provide here a brief review of key concepts related to the kinetics of enzyme-catalyzed reactions (for more advanced treatments, see the Additional Reading list at the end of the chapter.)

Substrate concentration affects the rate of enzyme-catalyzed reactions. Studying the effects of substrate concentration [S] in vitro is complicated by the fact that it changes during the course of the reaction, as substrate is converted to product. One simplifying approach in kinetics experiments is to measure the initial velocity, designated V₀ (Figure 5-11a). In a typical reaction, the enzyme may be present in nanomolar quantities, whereas [S] may be five or six orders of magnitude higher. If just the beginning of the reaction is monitored (often no more than the first few seconds), changes in [S] can be limited to a small percentage, and [S] can be regarded as constant. V₀ can then be explored as a function of [S], which is adjusted by the investigator. The effect on V₀ of varying [S] when the enzyme concentration is held constant is shown in Figure 5-11b. At relatively low concentrations of substrate, V₀ increases almost linearly with an increase in [S]. At higher substrate concentrations, V₀ increases by smaller and smaller amounts in response to increases in [S]. Finally, a point is reached beyond which increases in V₀ are vanishingly small as [S] increases. In this plateau-like V₀ region, the reaction approaches its maximum velocity, V_max.

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When the enzyme is first mixed with a large excess of substrate, there is an initial period called the pre-steady state, when the concentration of ES (enzyme-substrate) builds up. This period is usually too short to be observed easily, lasting just microseconds, and is not evident in Figure 5-11a. The reaction quickly achieves a steady state in which [ES] (and the concentration of any other intermediates) remains approximately constant over time. The concept of a steady state was introduced by G. E. Briggs and J. B. S. Haldane in 1925. The measured V₀ generally reflects the steady state, even though V₀ is limited to the early part of the reaction, and analysis of these initial rates is referred to as steady-state kinetics.

The curve expressing the relationship between [S] and V₀ (see Figure 5-11b) has the same general shape for most enzymes (approaching a rectangular hyperbola). It can be expressed algebraically by an equation developed by Leonor Michaelis and Maud Menten, called the Michaelis-Menten equation:

The important terms are [S], V₀, V_max, and a constant called the Michaelis constant, K_m. All these terms are readily measured experimentally.

The Michaelis-Menten equation is the rate equation for a one-substrate enzyme-catalyzed reaction. It states the quantitative relationship between the initial velocity V₀, the maximum velocity V_max, and the initial substrate concentration [S], all related through the Michaelis constant K_m. Note that K_m has units of concentration. Does the equation fit experimental observations? Yes; we can confirm this by considering the limiting situations where [S] is very high or very low, as shown in Figure 5-11b. The K_m is functionally equivalent to the [S] at which V₀ is one-half V_max. At low [S], K_m ≫ [S], and the [S] term in the denominator of the Michaelis-Menten equation (Equation 5-7) becomes insignificant. The equation simplifies to V₀ = V_max[S]/K_m, and V₀ exhibits a linear dependence on [S]. At high [S], where [S] ≫ K_m, the K_m term in the denominator of the Michaelis-Menten equation becomes insignificant and the equation simplifies to V₀ = V_max; this is consistent with the plateau observed at high [S]. The Michaelis-Menten equation is therefore consistent with the observed dependence of V₀ on [S], and the shape of the curve is defined by the terms V_max/K_m at low [S] and V_max at high [S].

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Kinetic parameters are used to compare enzyme activities. Many enzymes that follow Michaelis-Menten kinetics have different reaction mechanisms, and enzymes that catalyze reactions with six or eight identifiable intermediate steps within the enzyme active site often exhibit the same steady-state kinetic behavior. Even though Equation 5-7 holds true for many enzymes, both the magnitude and the real meaning of V_max and K_m can differ from one enzyme to the next. This is an important limitation of the steady-state approach to enzyme kinetics. The parameters V_max and K_m can be obtained experimentally for any given enzyme, but by themselves they provide little information about the number, rates, or chemical nature of discrete steps in the reaction. Nevertheless, steady-state kinetics is the standard language with which biochemists compare and characterize the catalytic efficiencies of enzymes.

Figure 5-11b shows a simple graphical method for obtaining an approximate value for K_m. The K_m, as noted, can vary greatly from enzyme to enzyme, and even for different substrates of the same enzyme. The magnitude of K_m is sometimes interpreted (often inappropriately) as an indicator of the affinity of an enzyme for its substrate. However, K_m is not equivalent to the substrate K_d (the simple binding equilibrium for the enzyme substrate) for many enzymes.

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The term V_max depends on both the concentration of enzyme and the rate of the rate-limiting step in the reaction pathway. Because the number of steps in a reaction and the identity of the rate-limiting step can vary, it is useful to define a general rate constant, k_cat, to describe the limiting rate of any enzyme-catalyzed reaction at saturation. If the reaction has several steps and one is clearly rate-limiting, k_cat is equivalent to the rate constant for that limiting step. When several steps are partially rate-limiting, k_cat can become a complex function of several of the rate constants that define each individual reaction step. In the Michaelis-Menten equation, V_max = k_cat[E_t], where [E_t] is the total concentration of enzyme, and Equation 5-7 becomes:

The constant k_cat is a first-order rate constant and hence has units of reciprocal time. It is equivalent to the number of substrate molecules converted to product in a given unit of time on a single enzyme molecule when the enzyme is saturated with substrate. Hence, this rate constant is also called the turnover number. A k_cat may be 0.01 s⁻¹ for an enzyme with an intrinsically slow function (some enzymes with regulatory functions act very slowly) or as high as 10,000 s⁻¹ for an enzyme catalyzing some aspect of intermediary metabolism.

In cells, there are many situations in which enzyme activity is inhibited by specific molecules, including other proteins. From a practical standpoint, the development of pharmaceutical and agricultural agents almost always involves the development of inhibitors for particular enzymes. Some aspects of enzyme inhibition are reviewed in Highlight 5-1.

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An understanding of the complete mechanism of action of a purified enzyme requires the identification of all substrates, cofactors, products, and regulators. Moreover, it requires a knowledge of (1) the temporal sequence in which enzyme-bound reaction intermediates form, (2) the structure of each intermediate and each transition state, (3) the rates of interconversion between intermediates, (4) the structural relationship of the enzyme to each intermediate, and (5) the energy contributed by all reacting and interacting groups to intermediate complexes and transition states. As yet, there is probably no enzyme for which we have an understanding that meets all these requirements.

It is impractical, of course, to cover all possible classes of enzyme chemistry, and we focus here on an enzyme reaction important to molecular biology: the reaction catalyzed by DNA ligases. The discussion concentrates on selected principles, along with some key discoveries that have helped bring these principles into focus. We also use the DNA ligase example to review some of the conventions used to depict enzyme mechanisms. Many mechanistic details and pieces of experimental evidence are necessarily omitted; an entire book would be needed to document the rich experimental history of enzyme research.

DNA ligases were discovered in 1967, and reports were published from four different research groups in that year. These enzymes catalyze the joining of DNA ends at strand breaks (also called nicks) that have a phosphorylated 5′ terminus and a 3′ terminus with a free hydroxyl group. In cells, these enzymes provide the critical links between discontinuous segments of replicated DNA (called Okazaki fragments; see Chapter 11) and carry out the final step in most DNA repair reactions (see Chapter 12). Since their discovery, many details of the reaction mechanism of these enzymes have been elucidated. DNA ligases have become essential tools of biotechnology, used by laboratories around the world to covalently join DNA segments to create recombinant DNA (see Chapter 7). RNA ligases have also been characterized; they use a similar reaction mechanism.

DNA ligases make use of two cofactors: Mg²⁺ ions and either ATP or nicotinamide adenine dinucleotide (NAD⁺). ATP-dependent DNA ligases are found in eukaryotes, viruses, and some bacteria and archaea. NAD⁺-dependent ligases are found in most bacteria, as well as in some viruses and archaea, but are not found in eukaryotes. Commonly, NAD⁺ is a cofactor participating in oxidation-reduction reactions. Its role in DNA ligase reactions is quite different, however, and it parallels the role of ATP in the ATP-dependent ligases.

All DNA ligases promote a reaction that involves three chemical steps (Figure 5-12). In step 1, an adenylate group, adenosine 5′-monophosphate (AMP), is transferred from either ATP or NAD⁺ to a Lys residue in the enzyme active site. This process occurs readily in the absence of DNA. In step 2, the enzyme binds to DNA at the site of a strand break and transfers the AMP to the 5′ phosphate of the DNA strand. This activates the 5′ phosphate for nucleophilic attack by the 3′-hydroxyl group of the DNA, in step 3, leading to displacement of the AMP and formation of a new phosphodiester bond in the DNA that seals the nick. Each of the three steps has a highly favorable reaction equilibrium that renders it effectively irreversible. In the absence of DNA, the adenylated enzyme formed in step 1 is quite stable, and it is likely that most DNA ligases in a cell are already adenylated and ready to react with DNA. In addition to illustrating the reaction pathway, Figure 5-12 introduces the conventions commonly used to describe enzyme-catalyzed reactions.

The overall picture of the DNA ligase reaction mechanism is a composite derived from kinetic and structural studies of many closely related enzymes, including those isolated from bacteriophages T4 and T7, bacteria, eukaryotic viruses (that is, viruses with eukaryotic host cells), and mammals. The ligases typically consist of a DNA-binding domain, a nucleotidyltransferase (NTase) domain, and an OB-fold domain. In DNA ligases, the OB fold, normally associated with binding to single-stranded DNA (see Section 5.1), interacts with the minor groove of double-stranded DNA. These domains provide a flexible structure that closes to completely encircle a nicked DNA molecule, with all three domains in contact with the DNA (see Figure 5-12). During step 1, some residues in the OB fold become part of the active site for transfer of AMP to the active-site Lys residue. As the covalent link between the enzyme and AMP is formed, the adenine base of AMP is fixed in a binding site in the NTase domain, where it stays throughout the remaining steps. In steps 2 and 3, a conformational change rearranges this part of the OB fold so that the same residues now face the solvent, while other parts of the OB fold bind to the DNA and interact primarily with the strand adjacent to the 5′-phosphate end of the strand break. At the same time, the conformational change closes the enzyme around the nicked DNA, and the N-glycosyl bond between the adenine base and the ribose moiety of AMP is rotated, thereby realigning the phosphate of AMP for reaction, in step 2, with the 5′ phosphate of DNA. Step 3 follows closely behind step 2 within the same complex.

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This one example cannot provide a complete overview of the broad range of strategies that enzymes use, but it does serve as a good introduction to the complexity of enzyme reaction mechanisms. In addition, it illustrates the transfer of phosphoryl groups, a reaction catalyzed by protein enzymes and RNA enzymes (ribozymes) involved in almost every aspect of molecular biology—from DNA polymerases and RNA polymerases to nucleases, topoisomerases, spliceosomes, and ligases.